**Methods in Molecular Biology 2533**

# Ribosome Biogenesis

Methods and Protocols

# M ETHODS IN M OLECULAR B IOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651 For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

# Ribosome Biogenesis

# Methods and Protocols

Edited by

# Karl-Dieter Entian

Institute for Molecular Biosciences, Goethe University Frankfurt, Frankfurt am Main, Hessen, Germany

Editor Karl-Dieter Entian Institute for Molecular Biosciences Goethe University Frankfurt Frankfurt am Main, Hessen, Germany

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2500-2 ISBN 978-1-0716-2501-9 (eBook) https://doi.org/10.1007/978-1-0716-2501-9

© The Editor(s) (if applicable) and The Author(s) 2022

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# Dedication

This book is dedicated to my wife Heike Gru¨ter and my son Christoph Gru¨ter for all their great support and understanding.

# Preface

Ribosomes are the organelles that synthesize proteins according to the genetically encoded information. Whereas prokaryotic ribosomes were even reconstituted in vitro, eukaryotic ribosomes are more complex. In yeast—as a eukaryotic model organism—about 5–10% of the genomic capacity is needed for the biogenesis of a ribosome. Worldwide a remarkable number of laboratories is investigating all aspects of ribosome biogenesis. As shown by the increasing number of participants in the respective meetings, approximately 3,000 scientists are working on structural, functional, and biomedical aspects of ribosome biogenesis. This book covers most of the significant steps during eukaryotic ribosome biogenesis. The research areas are introduced by reviews followed by chapters covering the respective methods of investigation.

A comparative survey about the unity and diversity of ribosome biogenesis in pro- and eukaryotic cells is provided in Part I (Chapter 1). The genomic organization of eukaryotic rDNA and the role of RNA polymerase I in ribosome biogenesis are summarized in Part II (Chapters 2 and 3). In vitro methods to study RNA polymerase I structure and its function are outlined in Part III (Chapters 4–6). Ribosome assembly in the nucleolus and a method to analyze the nucleo-cytoplasmic transport of assembled ribosomes and RNP complexes are dealt with in Part IV (Chapters 7 and 8). Various modifications increasing the complexity of rRNAs and the methods to analyze these modifications are given in Part V (Chapters 9–12). Finally, Part VI provides a review of eukaryotic translation and several methods to analyze translation in vivo (Chapters 13–16). Remarkably, for the first time a fully reconstituted eukaryotic yeast translation system is described together with the methods to purify the respective proteins.

This book is a valuable resource for scientists and all those interested in key questions in ribosome biogenesis. It provides an overview of the abundant literature and is supposed to stimulate collaborations.

Frankfurt am Main, Hessen, Germany Karl-Dieter Entian

# Acknowledgments

The majority of articles was supported by Deutsche Forschungsgemeinschaft SFB 960 (RNP Biogenesis: Assembly of Ribosomes and Non-ribosomal RNPs and Control of their Function) and SPP 1784 (Chemical Biology of Natural Nucleic Acid Modifications). The editor is extremely grateful to his scientific teachers and friends Friedrich K. Zimmermann, Darmstadt, Dieter Mecke, Tu¨bingen, Michael Ciriacy, Du¨sseldorf, James A. Barnett, Norwich, Piotr Slonimski, Gif-sur-Yvette, Jim Matoon, Colorado Springs, and Bernhard Prior, Stellenbosch for all their support and stimulating scientific discussions. I also thank John Walker, Anna Rakowski, Patrick Marton and Tessy Pria for their excellent support during editing this book.

The majority of articles in this book were supported by Deutsche Forschungsgemeinschaft. Collaborative network SFB 960 "Biogenesis of Ribosomes" and priority programme SPP1784 "Chemical Biology of Native Nucleic Acid Modifications". I also acknowledge the great computing support of Christoph Gru¨ter.

# Contents


#### PART IV RIBOSOME ASSEMBLY, TRANSPORT AND RNP COMPLEXES


# Contributors


UTE FISCHER • Institute of Biochemistry, ETH Zurich, Zurich, Switzerland


CLAUDIA HO¨ BARTNER • Institute of Organic Chemistry, Julius-Maximilians-University Wu¨rzburg, Wu¨rzburg, Germany

MONA HO¨ CHERL • Regensburg Center for Biochemistry (RCB), University of Regensburg, Regensburg, Germany


JORGE PEREZ-FERNANDEZ • Biochemistry III—Institute for Biochemistry, Genetics and Microbiology, University of Regensburg, Regensburg, Germany; Department of Experimental Biology, University of Jaen, Jae´n, Spain

MICHAEL PILSL • Universitat Regensburg, Regensburg Center for Biochemistry (RCB), € Lehrstuhl Biochemie III, Regensburg, Germany

SOPHIA PINZ • Institute for Biochemistry, Genetics and Microbiology, University of Regensburg, Regensburg, Germany


WOLFGANG SEUFERT • Institute for Biochemistry, Genetics and Microbiology, University of Regensburg, Regensburg, Germany


HERBERT TSCHOCHNER • Universitat Regensburg, Regensburg Center for Biochemistry € (RCB), Lehrstuhl Biochemie III, Regensburg, Germany


# Part I

Ribosome Biogenesis

# A Comparative Perspective on Ribosome Biogenesis: Unity and Diversity Across the Tree of Life

## Michael Ju¨ttner and Se´bastien Ferreira-Cerca

#### Abstract

Ribosomes are universally conserved ribonucleoprotein complexes involved in the decoding of the genetic information contained in messenger RNAs into proteins. Accordingly, ribosome biogenesis is a fundamental cellular process required for functional ribosome homeostasis and to preserve satisfactory gene expression capability.

Although the ribosome is universally conserved, its biogenesis shows an intriguing degree of variability across the tree of life. These differences also raise yet unresolved questions. Among them are (a) what are, if existing, the remaining ancestral common principles of ribosome biogenesis; (b) what are the molecular impacts of the evolution history and how did they contribute to (re)shape the ribosome biogenesis pathway across the tree of life; (c) what is the extent of functional divergence and/or convergence (functional mimicry), and in the latter case (if existing) what is the molecular basis; (d) considering the universal ribosome conservation, what is the capability of functional plasticity and cellular adaptation of the ribosome biogenesis pathway?

In this review, we provide a brief overview of ribosome biogenesis across the tree of life and try to illustrate some potential and/or emerging answers to these unresolved questions.

Key words Ribosome biogenesis, Ribosome assembly, rRNA, Maturation, RNA modifications, Eukaryotes, Archaea, Bacteria, Tree of life, Comparative biology, Evolution, Adaptation

#### 1 In Search of Unity?

From an historical perspective, the search for unifying concepts in Science in general and in Biology in particular has been a key step to our fundamental and general understanding of molecular processes across the tree of life [1–7]. This idea can be easily grasped by famous aphorism variations around this theme: "From the elephant to butyric acid bacterium—it is all the same!" ([8], cited in [2]) or "Anything found to be true of E. coli must also be true of elephants" (attributed to Jacques Monod, 1954 [2]). However, there are also valid arguments to think that elephants and bacteria are characterized, to some extent, by distinguishable biological properties. Accordingly, molecular processes, including ribosome

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_1, © The Author(s) 2022

biogenesis, have been dissected from two albeit different and in part contra intuitively but cross-fertilizing viewpoints: a unifying and a dividing functional perspective [9–11]. As such comparative—ribosome biogenesis—biology may be torn apart between defining the real weight of functional similarities and differences which biological systems may adopt. In any case, these similarities and differences can only be appreciated in the light of detailed knowledge about the scrutinized biological system across a larger number of entities.

In this chapter, we attempt to provide a short comparative overview on the molecular principles required for ribosome biogenesis. In addition, we like to highlight few challenges and surprises that may alter our unifying/differential view on ribosome biogenesis across the tree of life.

#### 2 Ribosome Biogenesis

#### 2.1 Once Upon a Time ... Ribosome Basic Facts

Ribosomes are universally conserved ribonucleoprotein particles allowing the decoding of the genetic information carried within messenger RNAs into amino-acid chains, the proteins [12]. Cytosolic ribosomes are composed of two ribosomal subunits, the small and the large ribosomal subunit (SSU and LSU, respectively) [12]. Strikingly, ribosomes are formed around a universally conserved structural core composed of three ribosomal RNA (rRNAs) molecules and 33 universally conserved ribosomal proteins (r-proteins) [13, 14]. Cytosolic ribosomes isolated from prokaryotic and eukaryotic organisms differ by the numbers and composition of their structural components, the r-proteins and rRNAs. Typically, cytosolic bacterial and archaeal 70S ribosomes are formed by the 30S (SSU) and 50S (LSU) ribosomal subunits [15–17]. Those are themselves composed of varying amounts of r-proteins (Figs. 1 and 2) which interact with the SSU 16S rRNA and LSU 23S and 5S rRNAs. These rRNAs also present various degree of organism's specific sequence size variations [54–56].

In eukaryotic cells, cytosolic 80S ribosomes are formed by the 40S (SSU) and 60S (LSU) ribosomal subunits [57, 58]. Concerning the amounts of ribosomal proteins, eukaryotic ribosomal subunits show also some, however less pronounced intra domain variations, compared to those observed across the bacterial and archaeal kingdoms (Figs. 1 and 2) [13, 14]. A striking feature of eukaryotic ribosomes is the presence of longer and additional rRNAs, the SSU 18S rRNA and the LSU 25/28S, 5.8S and 5S rRNAs [44, 59, 60].

In eukaryotes, rRNAs size expansion occurs by virtue of incorporation of additional rRNA sequences, the expansion segments, within the universally conserved prokaryotic-like rRNA core [23, 24, 61]. These expansion segments are varying in size and composition across eukaryotes [23, 24, 61, 62] and may have

Fig. 1 | Ribosome and ribosome biogenesis key features overview across the tree of life. (a) Summary of ribosome and ribosome biogenesis key features. Modified from [18] according to <sup>1</sup> [19–22]; <sup>2</sup> [23–27]; <sup>3</sup> [13, 14, 28]; <sup>4</sup> [29–35]; <sup>5</sup> [36–43]; <sup>6</sup> [10, 44–48]. Sso—Saccharolobus solfataricus; Hv—Haloferax volcanii; Tko—Thermococcus kodakarensis; Hs—Homo sapiens; Sc—Saccharomyces cerevisiae. (b, <sup>c</sup>) Summary of shared ribosomal proteins (b) and ribosome biogenesis factors (c) across the three domains of life. Numbers of r-proteins and putative ribosome biogenesis factors sequence homologues shared between bacteria, archaea, and eukarya (BAE); bacteria, archaea (BA), archaea and eukarya (AE), bacteria and eukarya (BE), or unique to bacteria (B), or archaea (A), or eukarya (E), are indicated [based on [10, 13, 14, 28, 41, 44–51] and our unpublished results]. (Modified from Londei and Ferreira-Cerca [52])

originated early on during rRNA evolution, since some progenitors of these expansion segments have been traced within modern archaeal but also in some case in bacterial rRNAs [23–25, 63–66].

The diverse composition of r-proteins which is, up to now, apparently more predominant in bacteria and archaea [13, 14, 55], could indicate that in these cellular contexts, ribosome assembly, that is, the assembly of r-proteins with the rRNAs, and ribosome function may tolerate a higher degree of flexibility than in most eukaryotes. It is also interesting that reductive evolution (loss) of r-proteins seems to prevail in archaea [13, 14, 25].

Fig. 2 | Exemplary conservation of selected ribosomal proteins and putative ribosome biogenesis factors involved in small ribosomal subunit biogenesis in archaea. (a) Exemplary repartition of selected archaeal ribosomal proteins shared between archaea and eukaryotes across two major archaeal Phyla. Black circle denotes the presence, and open circle denotes the absence of sequence homologue for the indicated ribosomal protein of the small (S) or large (L) ribosomal subunits, respectively (adapted from [13, 14] using the nomenclature proposed in [49]). (b) Phylogenetic conservation profile of the indicated known or putative small ribosomal subunit ribosome biogenesis factors across 1500 archaeal genomes were generated using AnnoTree (http:/annotree.uwaterloo.ca) [53]. Archaeal classes are annotated in a phylogenetic tree (upper left) as provided by AnnoTree. Note the absence of significant homology for Nep1 (e.g., Thermoplasmata, Halobacteria and more) or Tsr3 (e.g., Thermococcales) in a large group of organisms, in contrast to the more widespread distribution of KsgA/Dim1, Rio1, and Nob1 archaeal homologs. Modified from Londei and Ferreira-Cerca [52]

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From a compositional point of view archaea and eukaryotes do share common r-proteins which are absent in bacteria, whereas bacteria do possess domain specific r-proteins [49, 61]. This correlates with the increased structural similarities between archaeal and eukaryotic ribosomal subunits in comparison to their bacterial counterparts [25, 61, 67].

This structural similarity has been observed early on by the group of James Lake, using electron microscopy, thereby, suggesting a closer evolutionary relationship of archaea and eukaryotes [15, 67, 68]. Recent phylogenetic analysis [69–71] and higher resolution structure analysis of ribosomal subunits [25, 61, 67] essentially confirm this idea but also provide additional insights into structural differences between the different domains of life, like for example differences in the peptide exit tunnel geometry [72], or species-specific structural alteration which may be related to organism-specific environmental adaptations [62, 73].

#### 3 The Ribosome Assembly Process

The ribosome assembly process, that is, the assembly of r-proteins with rRNAs, has been analyzed very early in the history of ribosome research. Early work from the Nomura laboratory in the 1960–1970s aiming to understand the individual contribution of the r-proteins/rRNA to the protein synthesis process, has led to the first in vitro reconstitution of bacterial ribosomal subunits from its isolated structural components [29, 30, 74–77]. These studies were then followed by r-proteins omission experiments which culminated in the establishment of the first ribosomal proteins assembly maps describing r-proteins assembly dependencies [29– 31]. Beyond being biochemical masterpieces, these studies have revealed key features of the ribosomal assembly process in bacteria. Notably, the self-assembling nature of ribosomal subunit formation, and the fact that ribosome assembly proceeds via a combination of cooperative and hierarchical mechanisms [29–31, 78, 79]. However, these in vitro assembly experiments have been mitigated by the fact that they occur under nonphysiological conditions, thereby suggesting the existence of in vivo facilitating mechanisms which were discovered later [29–31, 78, 79]. Remarkably, in vitro reconstitution of ribosomal subunits has not only been achieved using structural components isolated from different bacterial sources, but also from two evolutionary divergent representative archaea [32–34]. In contrast, similar in vitro reconstitution of eukaryotic ribosomal subunits solely using purified structural components has not been accomplished to date.

Despite this fundamental biochemical difference, some aspects of ribosome assembly in bacteria and eukaryotes follow rather similar molecular principles, for example the hierarchical and cooperative assembly, or stepwise stabilization of r-proteins [18, 78–87]. Together these similarities suggest that ribosome assembly has likely evolved around a self-assembling (presumably self-replicating) ancestor ribosome [26, 88], which has retained some of its original assembly properties and constraints. Not surprisingly, some of these ancestral properties/constraints are most probably universally shared. In addition, existing molecular mechanisms have been modified (adapted or optimized), new ones implemented, or some maybe lost, due to organisms or common ancestor specific requirements. All these evolutionary contributions are not trivial to disentangle, but functional and structural analysis of the ribosome assembly pathway, in model and probably most importantly in nonmodel organisms, will help us to further clarify the inherited molecular constraints and properties underlying the assembly of ribosomal subunits.

#### 4 Facilitating Ribosome Assembly

As mentioned above, efficient ribosome assembly in vivo depends on ribosome biogenesis factors which are collectively believed to facilitate various aspects of the ribosome biogenesis process [44, 59, 60, 78, 79]. These ribosome biogenesis factors can be subdivided into different protein classes according to their respective structural organization and/or enzymatic activity. For example, ribosome biogenesis progression depends on the presence of energy consuming enzymes, like GTPases, ATPases (AAA ATPase, RNA helicase, etc.). However, it is important to note that the ensemble of ribosome biogenesis factors differs in numbers and nature from bacteria to eukaryotes [10, 18, 44, 59, 60, 78, 79] (Fig. 1). Accordingly, the relative domain-specific repartition of structural features and/or enzyme activities implicated in ribosome biogenesis progression may vary considerably between different groups and may reflect functional adaptations within the different domains of life. For example, GTPases seem to be enriched in the bacterial ribosome biogenesis pathway, whereas ATP-dependent processes, or β-propeller containing proteins are enriched in the eukaryotic ribosome biogenesis context [10, 18, 44, 59, 60, 78, 79].

In fact, and with the exception of the (almost) universally conserved dimethyl-transferase KsgA/Dim1 [89, 90], ribosome biogenesis factors are not well conserved between bacteria and archaea or between bacteria and eukaryotes [18, 45]. In contrast, a substantial portion of eukaryotic ribosome biogenesis factors are found in archaeal genomes even though our understanding of their respective functions in archaea remains still limited [45]. Nevertheless, we and others could demonstrate some functional analogy with their eukaryotic counterparts in vivo and/or in vitro [18, 91–93]. These observations suggest that probably more (if not most) of these eukaryotic-like ribosome biogenesis sequence signatures present in archaea might be authentic ribosome biogenesis factors shared between archaea and eukaryotes. In comparison, eukaryotic ribosome biogenesis is characterized by a large increase of eukaryotes-specific ribosome biogenesis factors (>200) [46, 59, 60]. The functional requirements for this "sudden" increased complexity of eukaryotic ribosome biogenesis remains to be fully understood (Fig. 1).

In addition, to composition and number variations observed in bacteria, archaea, and eukaryotes, organisms specific variations can be observed [18, 36, 45, 90, 94, 95]. For example, the set of ribosome biogenesis factors vary across the archaeal phylum and seems to follow the general trend of reductive evolution previously observed for archaeal r-proteins (Fig. 2) [13, 14, 52]. In eukaryotes, ribosome biogenesis factors diversity further increases from unicellular to multicellular eukaryotes with the addition of factors implicated in ribosome biogenesis [46, 59, 60]. Moreover, recent studies have provided new insights into ribosome biogenesis plasticity thereby suggesting that the order of functional requirement of some assembly factors/r-proteins can vary or be functionally bypassed in some conditions [90, 96–100].

These observations have various implications for our understanding of ribosome biogenesis evolution and plasticity. For one, the presence/absence of certain molecular components can be tolerated owing that the proper rescue mechanisms are implemented (coevolving). Furthermore, these imply a higher functional plasticity of the order of events within the ribosome biogenesis pathway, whereby an alternative assembly landscape might be used or kinetically favored depending on the cellular context [81, 87, 98, 101]. However, it should be noted that this apparent diversity/plasticity may still converge to the formation of essential assembly intermediates that are functionally and/or structurally equivalent, thereby fulfilling critical inherited molecular events required for ribosome biogenesis.

Accordingly, and despite differences in the nature and amounts of the ribosome biogenesis factor ensemble, it is conceivable that the core function supported by some or all ribosome biogenesis factors are functionally equivalent across the tree of life, thereby suggesting evolutionary constraints which would have favored the establishment of dedicated functional mimicry rather than functional divergence around the universal ribosome core [18]. It is for example striking, that some divergent ribosome biogenesis factors implicated in the formation of the SSU in model bacteria, archaea and eukaryotes, are binding at very similar locations within the nascent pre-ribosomal subunits and may fulfill similar molecular tasks (see further discussion in [18]). For instance, the SSU rRNA 3<sup>0</sup> end processing follows a very similar pattern which involves a KH-domain containing ribosome biogenesis factor which interact and presumably stabilized the 30 end of the 16S/18S rRNA, thereby enabling efficient and presumably controlled endonucleolytic cleavage. In E. coli, the Era GTPase, which contains a KH-domain [102–104] interact with the endonuclease YbeY and the r-protein uS11 [105], thereby facilitating 3<sup>0</sup> end maturation. In eukaryotes, Pno1/Dim2, a KH-domain containing protein, interacts with the endonuclease Nob1 and both are located in proximity of uS11. Moreover, mutational analysis revealed functional implication of uS11, Pno1/Dim2 and Nob1 for 18S rRNA maturation [106–110]. Furthermore, archaeal homologues for Pno1/Dim2, Nob1 and uS11 are present in most archaeal genomes [111]. Considering that both the endonucleases and KH-domains (type I vs. type II) are evolutionary distinct and presumably unrelated, these observations suggest a functional convergence/mimicry at the basis of the maturation of the 16S/18S rRNA 3<sup>0</sup> end. Whereas, the origin of this divergence at the molecular level is poorly understood, evolutionary constraints have remarkably selected a very similar mode of action in its principle (see further discussion in [18]).

Further supporting the existence of functional convergence enabling ribosomal subunit synthesis, we and others have proposed that pseudocircularization events might represent an early common feature of ribosomal subunits biogenesis [82, 112, 113]. However, the implicated molecular machineries are to some extent very different.

In prokaryotic organisms, stabilization of the 5<sup>0</sup> -3<sup>0</sup> mature ends of the nascent rRNA precursors in a topologically limited environment is enabled by the formation of double-stranded RNA structures, the processing stems [114–121]. In all archaeal organisms analyzed so far, this environment is further stabilized by the formation of a true covalent circularization of the pre-rRNA, in form of precircular rRNA intermediates [112, 122, 123]. Finally, in eukaryotes, recent cryo-EM studies have revealed stabilization of a pseudocircular intermediate of the pre-LSU [124]. The formation of this intermediate requires the participation of a distinct eukaryotesspecific ribosome biogenesis subcomplex which may stabilize the LSU root helix bundle prior to the assembly of the universally conserved r-protein uL3 [124, 125]. Noteworthy, early maturation of the pre-23S rRNA by mini-RNase III, which liberates the nascent 23S rRNA from its processing stem in B. subtilis, is stimulated by the presence of uL3 [126]. In addition, uL3 is critical to initiate in vitro assembly of bacterial 50S [127, 128].

In the case of the SSU, the snoRNA U3 and its associated proteins provide a scaffold that brings distant rRNA elements in close proximity within an encapsulated environment described as for the 90S/SSU Processome [129–131]. However, the relative orientation of the future mature 50 -30 ends in these structures is not resolved.

Finally, the formation of the SSU central pseudoknot is a universal feature required for SSU biogenesis and function [132, 133]. In eukaryotes, its formation is facilitated by the snoRNA U3, which is not present in bacteria and archaea [44, 59, 60, 134, 135]. However alternative (U3-independent) mechanisms, enabling the formation of the SSU central pseudoknot in these cellular contexts have been proposed [135–138]. For example, sequences present in the 5<sup>0</sup> end of the pre-16S rRNA show potential complementarity, similar to U3, which could hybridize with the region required for central pseudoknot formation [136, 139–141].

#### 5 Processing and Modifications of Ribosomal RNA

Concomitantly to the assembly process, rRNAs are matured by ribonuclease activities and modified at various positions [18, 37, 44, 59, 60, 78, 79, 114, 142].

Despite billion years of independent evolution most rRNAs are predominantly transcribed as a polycistronic operon [19]. It is believed that this organization is required for the efficient coordinated assembly of ribosomal subunits. However, this idea has been challenged on the one hand by the presence of naturally occurring independent rDNA production units, and on the other hand, by early genetic engineering experiments which have successfully separated the polycistronic eukaryotic SSU and LSU rRNAs [143, 144]. The immature precursor-rRNA contains flanking regions that need to be matured by the action of various ribonucleases. These maturations events are timely ordered during the ribosomal subunit biogenesis process. This relative ordering presumably depends on specific ribosomal subunit assembly statuses which in turn control substrate accessibility or its relative positioning [18, 37, 44, 59, 60, 78, 79, 114, 143, 145]. The inherent irreversible property of these processing steps may also impose various degrees of "quality control" constraints to the ribosome biogenesis process in order to avoid the irreparable formation of improperly assembled pre-ribosomal subunits.

Similar to the ribosome biogenesis factors, the set of ribonucleases used in bacteria, eukaryotes and presumably in archaea are not well conserved between these domains [18, 37, 44, 59, 60, 78, 79, 94, 114, 142]. In bacteria, whereas promiscuous ribonucleases are used, eukaryotic cells have developed a set of specific enzymes to mature their rRNAs, some of which are also present in archaea [18, 37, 44, 52, 59, 60, 78, 79, 94, 114, 142]. Based on our current knowledge, it is difficult to properly extract functional similarities between the different biological systems. However, we have previously noticed some peculiar common molecular principles required for the maturation of the SSU rRNA 30 -end (see above and [18]). However, additional structural and functional information capturing these events "in action" will be necessary to provide invaluable insights into the structural properties of their respective substrates [145].

Since the 1950s, ribosomal RNA modifications, which are mostly concentrated within or closed to the ribosomal subunit functional centers, have been known [37, 146, 147]. These modified rRNAs residues are found, to various extents, in all domains of life [37–40]. However, the mechanisms by which these modifications are added diverge across the tree of life. On the one hand, bacterial rRNAs are modified by stand-alone enzymes that are dependent on a specific assembly status to recognize and modify their respective substrates. On the other hand, and in addition to stand-alone enzymes, archaea and eukaryotic organisms utilize an RNA-guided modification machinery, whereby RNA-protein complexes carrying methyltransferase or pseudouridylation activity are formed (C/D and H/ACA snoRNPs, respectively). In this context, the RNA part, which contains a sequence complementary to the targeted rRNA region, guides the enzymatic activity to its substrate [37, 39, 40, 148–150] (Fig. 1).

These different modes of action have important consequences regarding substrate recognition, timing of modifications and structural constraints that may be imposed by the formation of snoRNA::rRNA duplexes during ribosome assembly, and have thereby probably (re)shaped several aspects of the ribosome biogenesis pathway [18, 37]. Whereas the relative positions of the conserved modified residues are usually similar, the nature of the modification itself may vary across the tree of life [18, 151]. Finally, rRNA modifications appear to be dynamic across the tree of life, whereby significant variation in nature and number of modifications is observed, and may also vary during the organisms life time [36, 38, 39, 41, 90, 152, 153]. These variations may not only influence ribosome function but also the ribosome biogenesis pathway itself [18, 38, 39, 90, 154].

#### 6 Learning from Organelle Ribosome Biogenesis?

Eukaryotic organelles also contain ribosomes, which are very distinct from cytoplasmic ribosomes. Organelle's ribosomes and their ribosome biogenesis pathways, which have not been discussed so far, may represent important resources to better extricate key ribosomal subunits biogenesis features. Organelle ribosomal subunits are interesting from several perspectives. First, the ribosomal subunits composition is rather diverse across different organisms. Second and in contrast to cytosolic ribosomal subunits, organelle ribosomal subunits contain a reduced amount of rRNA over r-proteins. Third, organelle ribosomal subunits contain organellespecific ribosomal proteins, expending the possible diversity of the ribosome assembly landscape. These r-proteins may replace lost rRNA elements or stabilize the reduced rRNA core. Fourth, organelle ribosomal subunits have followed complicated independent evolutionary trajectories enabling the formation of ribosomal subunits optimized for the translation of a limited set of mRNAs [49, 155–157]. In addition to ribosome biogenesis factors shared between bacteria and organelles, recent studies have revealed the existence of specific dedicated multiprotein machineries required for the progression of organelle ribosome biogenesis [155, 158– 166]. Despite this apparent sequence/structural specificity, intriguing functional similarity between cytosolic and organelle ribosome biogenesis pathway has been proposed. Altogether, these results suggest that despite very different evolutionary path ribosomal subunits biogenesis may proceed via functionally equivalent assembly intermediates and requires similar but diverse functional innovations facilitating ribosomal subunits assembly [155, 158–166]. Lessons from these and future studies will certainly reveal new insights on the evolution and adaptation of ribosomal subunit biogenesis.

#### 7 Concluding Remarks

Despite undeniable differences between the ribosome biogenesis pathways as known from various model organisms, it is striking that some (key) ribosome biogenesis features have been maintained across billions of years of evolution. However, the evolutionary events which have led to components diversification while conserving functional similarities instead of consolidating a core of conserved ribosome biogenesis components remain rather enigmatic. Moreover, the extent of true functional divergence or functional convergence, along the ribosome biogenesis pathway, needs to be properly identified, promising exciting perspectives and challenges for comparative ribosome biology analysis in the future. Correspondingly, in the recent years metagenomics have revealed an unexpected microbial biodiversity [167], which awaits its biochemical and functional examination, and will certainly provide new insights into conserved principles of ribosome biogenesis.

In addition, our increased understanding of supposedly simplified ribosome biogenesis pathway present in symbionts, organelles, and organisms harboring reduced genomes, which for simplicity is not discussed in depth, will provide supplementary functional insights into common and specific principles of ribosome biogenesis [47, 55, 62, 160–162, 168].

Finally, thanks to the massive development in the field of genetic engineering we are probably at the very beginning of a massive biological revolution, which will ease the characterization of nonmodel organisms, to develop synthetic biology approaches, and to uncover some of the most fundamental secrets of life. How we will deal with this information will however be crucial to leverage the significance of these discoveries for our understanding of ribosome biogenesis. When reaching this point, emphasis toward functional similarity and diversity of the ribosome biogenesis process, or its plasticity, will have to be carefully appreciated and will require to understand the genuine functional implication of these molecular features across different organisms' lifestyle and organisms' specific evolutionary history [169].

#### Acknowledgments

We are indebted to our many colleagues for their writing and/or invaluable discussions without which this essay would not have been possible. We also sincerely apologize to our colleagues, whose work we failed to discuss or highlight. We are grateful to Prof. Dr. Karl-Dieter Entian (University of Frankfurt, Germany), Dr. Brigitte Pertschy (University of Graz, Austria) and Dr. Robert Knu¨ppel (University of Regensburg, Germany) for comments and suggestions.

Work in the Ferreira-Cerca laboratory is supported by the Chair of Biochemistry III "House of the Ribosome"—University of Regensburg, by the DFG-funded collaborative research center CRC/SFB960 "RNP biogenesis: assembly of ribosomes and non-ribosomal RNPs and control of their function" (SFB960- B13) and by an individual DFG grant to S.F.-C. (FE1622/2-1; Project Nr. 409198929).

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Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part II

Genomic Organization

# Establishment and Maintenance of Open Ribosomal RNA Gene Chromatin States in Eukaryotes

## Christopher Sch€achner, Philipp E. Merkl, Michael Pilsl, Katrin Schwank, Kristin Hergert, Sebastian Kruse, Philipp Milkereit, Herbert Tschochner, and Joachim Griesenbeck

#### Abstract

In growing eukaryotic cells, nuclear ribosomal (r)RNA synthesis by RNA polymerase (RNAP) I accounts for the vast majority of cellular transcription. This high output is achieved by the presence of multiple copies of rRNA genes in eukaryotic genomes transcribed at a high rate. In contrast to most of the other transcribed genomic loci, actively transcribed rRNA genes are largely devoid of nucleosomes adapting a characteristic "open" chromatin state, whereas a significant fraction of rRNA genes resides in a transcriptionally inactive nucleosomal "closed" chromatin state. Here, we review our current knowledge about the nature of open rRNA gene chromatin and discuss how this state may be established.

Key words Nucleolus, Nucleolar organizer region (NOR), Ribosomal DNA, Ribosomal RNA genes, RNA polymerase I, Transcription, Chromatin, Nucleosome, Preinitiation complex (PIC), High mobility group (HMG) box proteins, Upstream binding factor (UBF), Hmo1, Psoralen cross-linking, Chromatin immunoprecipitation (ChIP), Chromatin endogenous cleavage (ChEC), Electron microscopy (EM)

#### 1 Introduction

This short review aims at summarizing research which has contributed to our current understanding of characteristic chromatin states at eukaryotic rRNA gene loci sharing many conserved features from yeast to human. This subject has also (partly) be covered by other reviews in the past, which are recommended for further reading [1–5]. The regulation of RNAP I transcription by epigenetic mechanisms (including posttranslational covalent modifications of histones and other components of the RNAP I transcription machinery, as well as DNA-methylation) has been reviewed in great detail in the past [6–9] and will not be subject of this article.

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_2, © The Author(s) 2022

In the nucleus of eukaryotic cells, the genetic information encoded in the DNA is assembled in the structure of chromatin. The basic unit of chromatin is called the nucleosome and consists of approximately 146 bp of DNA wrapped around a histone octamer (reviewed in [10–12]). The tight wrapping of the nucleic acid around the protein core renders the DNA in part inaccessible which plays an important role for the regulation of essential nuclear processes like replication, DNA-repair, and transcription. To deal with nucleosomal DNA, eukaryotic cells developed mechanisms altering chromatin structure at distinct loci in specific situations (reviewed in [13, 14]). Accordingly, characteristic changes in chromatin structure and posttranslational covalent modification state of chromatin components correlate with transitions in the transcriptional status of individual genes. Understanding the relationship between chromatin structure and transcription will be essential to fully comprehend the complex process of eukaryotic gene expression.

#### 2 Visualization of rRNA Transcription

2.1 Actively Transcribed rRNA Genes Are Prominent Structures in Nuclear Chromatin Spreads

In eukaryotes, there are at least three different nuclear RNAPs, numbered I–III, each of which has a distinct set of target genes (reviewed in [15], see also short reviews by Merkl et al. and Pilsl et al., this issue). Whereas RNAP II transcribes all protein coding genes, RNAP III is dedicated to the synthesis of small noncoding RNAs including the 5S rRNA and tRNAs. In all organisms—with only one known exception [16]—RNAP I has only one acknowledged genomic target locus from which it synthesizes a large precursor transcript encompassing the sequences of three out of four rRNAs (hereafter called rRNA gene). Remarkably, rRNAs produced by RNAP I account for more than 60% of cellular RNA synthesis in exponentially growing S. cerevisiae cells (hereafter called yeast, reviewed in [17]). This is achieved by a strong promoter with a stably bound preinitiation complex (PIC) (reviewed in [18]), as well as the multimerization of the genomic templates (reviewed in [19]). Thus, rRNA genes exist as clusters of repeated transcription units, the so-called nucleolar organizer regions (NORs), at one or more genomic locations depending on the organism. Actively transcribed NORs are part of the nucleolus, the dominant nuclear substructure of early ribosome biogenesis, whereas inactive NORs do not associate with nucleoli (reviewed in [20]).

Even slightly before the different nuclear RNAPs were biochemically defined [21, 22], it was possible to visualize the enzymes transcribing their target loci in chromatin spreads by electron microscopy (EM) [23–25]. The first transcription units that were unambiguously identified by this method were the actively transcribed extrachromosomal ribosomal RNA genes isolated from amphibian oocytes [25]. These transcription units showed a characteristic "Christmas-tree" like appearance with tightly packed elongating RNAPs forming the stem, and protein-coated nascent rRNAs extending from the polymerases forming the branches of the trees. Depending on the preparation, Christmas trees can be decorated with "terminal balls" representing preribosomal assembly intermediates [26]. Ever since, rRNA gene Christmas-trees have been observed in chromatin spreads from different cell types of many organisms (reviewed in [27, 28]). These studies yielded important insights in aspects of RNAP I transcription at the single molecule level. More recently, the spreading technique combined with cryo-EM tomography allowed to obtain more detailed structural information about yeast RNAP I transcribing its native template [29]. In fact, shortly after the first Christmas trees had been described, the conserved repeated "beads-on-the-string" nature of nontranscribed eukaryotic chromatin came into the focus of the researchers [30, 31]. The beads-on-the-string seen in EM were biochemically defined as complexes of DNA and histone octamers, the fundamental units of chromatin named nucleosomes [32–34].

#### 2.2 rRNA Genes Transcribed by RNAP I Are Nucleosome Depleted

The above analyses of chromatin spreads provided first insights into how different RNAPs deal with the chromatin template (reviewed in [27]). Thus, nonribosomal chromatin moderately transcribed by (presumably) RNAP II remained at least partially covered with nucleosomes [35–37]. This indicated that nucleosomes may either persist or are reestablished upon RNAP II transcription. In contrast, there was evidence that RNAP I and RNAP III transcription occurred exclusively on nucleosome depleted templates even in situations when transcription rate was reduced (reviewed in [27, 38–40]). Complementing EM analyses, endonucleases were used as molecular probes to investigate chromatin structure (reviewed in [41–43]). Endonucleases can only poorly access DNA assembled into a nucleosome. Therefore, endonuclease cleavage at a genomic region of interest can be used to deduce information about local nucleosome occupancy. In such assays, DNA in actively transcribed rRNA gene chromatin was more accessible than DNA in nontranscribed rRNA gene chromatin, supporting the view of nucleosome depletion at RNAP I transcribed regions [44, 45]. This notion was corroborated by in vivo and in vitro cross-linking of chromosomal DNA with psoralen (reviewed in [46] and references therein). Psoralen is a parent compound of naturally occurring substances which can intercalate in DNA (reviewed in [47] and references therein). Upon exposure to longwave ultraviolet (UVA) radiation, psoralen incorporation leads to DNA-interstrand cross-links. As observed for nucleases, nucleosome formation prevents psoralen intercalation into nucleosomal DNA tightly interacting with the histone octamer [48, 49]. Therefore, psoralen cross-links are restricted to accessible nucleosome-free DNA regions. After DNA isolation from psoralen cross-linked chromatin, nucleosomes leave a characteristic footprint of approximately 146 bp of non–cross-linked DNA surrounded by cross-linked linker DNA. This can be analyzed by EM of the cross-linked DNA under denaturing conditions, in which the non–cross-linked DNA regions are visualized as single stranded DNA bubbles. Consistent with nucleosome depletion, DNA isolated from psoralen treated actively transcribed rRNA gene chromatin was heavily cross-linked and largely devoid of single-stranded DNA bubbles [50].

Psoralen cross-linking alters the mobility of deproteinized DNA fragments in native agarose gel electrophoresis [50]. A high-degree of psoralen incorporation in nucleosome-depleted DNA leads to a strong retardation of the corresponding fragment, whereas lower psoralen incorporation in nucleosomal DNA yields faster migrating fragments. In combination with Southern blot analysis this technique can be used to monitor nucleosome occupancy at specific restriction fragments obtained from psoralen cross-linked chromosomal DNA [50–52]. DNA-fragments deriving from psoralen cross-linked transcribed regions of rRNA genes from a human cell line and yeast cells migrated as two major bands with low and high mobility in native agarose gel electrophoresis, indicating that these genes adapt at least two different chromatin states [51, 52]. Nascent RNA was exclusively cross-linked to the fragment of low mobility, suggesting that nucleosome occupancy at actively transcribed rRNA genes is strongly decreased [51, 52]. This led to the model that rRNA genes may coexist at least in a nucleosome depleted, "open" chromatin state and a transcriptionally inactive nucleosomal "closed" chromatin state. It should be noted, that psoralen cross-linking measures nucleosome occupancy at selected loci, but does not allow straightforward conclusions about the transcriptional state of a gene (reviewed in [2, 3] and references therein). Therefore, open rRNA genes are not necessarily actively transcribed, although actively transcribed rRNA genes appear to be always in an open chromatin state. To date, no other chromosomal locus with a psoralen accessibility similar to open rRNA genes has been identified. Strikingly, psoralen cross-linking of heavily transcribed RNAP II-dependent genes did not yield fragments with the low mobility expected for fully cross-linked DNA [53]. This may corroborate the observations in EM that RNAP II transcribed gene regions retain a significant number of nucleosomal particles [35, 37]. Thus, the open rRNA gene chromatin state is probably unique regarding the extent of nucleosome depletion and the size of the nucleosome depleted region.

With the advent of chromatin immunoprecipitation (ChIP) the nucleosome-depleted nature of actively transcribed rRNA genes was challenged (reviewed in [2, 3]). In ChIP experiments, histone

2.3 rRNA Genes Coexist in At Least Two Different Chromatin States

molecules coprecipitated rRNA gene fragments from extracts obtained from yeast cells carrying a reduced number of rRNA gene copies [54]. Because under these conditions, the majority of rRNA genes were transcribed [55], it was concluded that RNAP I transcribes a dynamic, nucleosomal chromatin template. A subsequent study based on chromatin endogenous cleavage (ChEC) experiments in yeast supported rather the model of robust histone/nucleosome depletion at actively transcribed rRNA genes [56]. ChEC is performed in yeast strains in which a protein of interest is expressed in fusion with Micrococcal nuclease (MNase) from Staphylococcus aureus [57]. MNase is a secreted DNA and RNA endo- and exonuclease which has long been used to study chromatin structure (reviewed in [41, 43], see chapter of Teubl et al. in this issue). MNase activity strictly depends on calcium. Thus, due to the low intracellular calcium concentrations recombinantly expressed MNase fusion proteins are not active. After isolation of crude nuclei from cells expressing MNase-fusion proteins and addition of calcium, the tethered nuclease will cut neighboring DNA. Specific cleavage events in genomic DNA can be monitored by Southern blot analysis, primer extension or high-throughput sequencing to reveal which DNA regions were in the proximity of the MNase fusion proteins [58, 59]. ChEC experiments can also be performed in combination with psoralen cross-linking to determine if a factor of interest preferentially associates with the open or the closed rRNA gene chromatin state [56, 60]. Using this method, it could be shown that RNAP I subunits fused to MNase specifically degraded the highly psoralen cross-linked fragments derived from open rRNA gene chromatin in yeast. In contrast, histone-MNase fusion proteins preferentially degraded the poorly psoralen crosslinked fragments derived from closed rRNA gene chromatin. These results supported the hypothesis that RNAP I transcribes a histone/nucleosome depleted chromatin template [56]. The apparent contradiction of these results to the conclusions derived from ChIP experiments [54] may be explained by the notion that the term histone depletion does not exclude that a few histone molecules occasionally associate with open rRNA gene chromatin. Additionally, even in yeast cells with a lower rRNA gene copy number, a small subpopulation of rRNA genes resides in the closed nontranscribed nucleosomal chromatin state [55, 61]. This nucleosomal subpopulation might then be detected in ChIP analyses, which unlike the combination of ChEC with psoralen cross-linking analyses—does not distinguish between DNA fragments derived from open or closed rRNA gene chromatin. In higher eukaryotes ChEC combined with psoralen cross-linking analyses have not been conducted so far. Nevertheless, endonuclease accessibility of human rRNA gene loci combined with high throughput sequencing (HTS) suggested that nucleosome occupancy is reduced at rRNA genes when compared to intergenic regions [62]. More recently, a deconvolution histone ChIP-HTS analysis combined with genetic manipulation of RNAP I transcription in mice strongly supported that histone depletion at active rRNA genes is conserved from yeast to mammals [63, 64].

2.4 HMG-Box Proteins Are Architectural Components of Open rRNA Gene Chromatin States

The high mobility group (HMG) box is a structural motif first described in eukaryotic proteins with a high electrophoretic mobility (reviewed in [65]). HMG box proteins are involved in the regulation of many important DNA-dependent processes in the nucleus presumably due to their acknowledged function as architectural chromatin components. Along these lines, HMG-box proteins are conserved constituents of open rRNA gene chromatin (reviewed in [2, 63, 66, 67]). In vertebrates, the upstream binding factor (UBF) was initially identified to be required for proper recruitment of the basal RNAP I PIC at the rRNA gene promoter [68–70]. UBF has a characteristic arrangement of up to six tandemly repeated HMG-boxes (reviewed in [71]). Upon dimerization UBF may wrap DNA in one single loop in a structure called the "enhancesome" in vitro [72]. It was suggested that this structure establishes a characteristic architecture at the rRNA gene promoter (reviewed in [71]). In vivo, UBF molecules spread along the entire rRNA gene region transcribed by RNAP I [62, 63, 66, 73, 74]. Additionally, UBF is preferentially recruited to repetitive enhancer DNA elements preceding the rRNA gene promoter in higher eukaryotes [62, 63, 69, 73, 74]. UBF was shown to localize to NORs harboring active rRNA genes [75]. During the cell division cycle, UBF stays associated with NORs which have been transcriptionally active even upon RNAP I transcription shutdown at mitosis [76–78]. In this condition, UBF likely maintains NORs bearing active rRNA gene clusters in a hypocondensed, open chromatin state leading to "secondary constrictions" visualized by light microscopy in plant mitotic chromosomes early in the twentieth century [79–81]. Yeast contains a bona fide UBF homologue Hmo1 [82, 83]. As UBF, Hmo1 is a chromatin component of actively transcribed rRNA gene regions but has only one HMG-box and—in contrast to UBF—no reported role in RNAP I PIC formation [56, 84–86]. Both murine UBF and yeast Hmo1 are required to maintain the nucleosome depleted open rRNA gene chromatin state in the absence of RNAP I transcription [61, 63]. In agreement, both UBF and Hmo1 can destabilize nucleosomal templates in vitro [87, 88]. Furthermore, Hmo1 mediated DNA compaction, bridging and looping observed in vitro was suggested to be implicated in the maintenance of the nucleosome depleted rRNA gene chromatin state in vivo [89]. UBF and Hmo1 bind to various DNA sequences in vitro [90–92] and genome wide ChIP analyses suggested that both proteins additionally associate with nucleosome depleted promoter regions of highly transcribed RNAP II dependent genes [84–86, 93]. This indicates that UBF and Hmo1 may generally recognize structures associated with highly transcribed DNA templates. This hypothesis was substantiated in in vitro studies in which Hmo1 bound preferentially to DNA templates thought to mimic transcription intermediates [91]. Along these lines it has also been suggested that Hmo1 protects negatively supercoiled DNA at gene boundaries in vivo [94], indicating that (r)DNA topology may be important for Hmo1 recruitment. Whereas there is evidence that recruitment of Hmo1 to rRNA gene sequences is dependent on prior RNAP I transcription [61, 86], RNAP I transcription may not be required to recruit UBF to the rRNA gene promoter or enhancer elements [95]. Thus, integration of rRNA gene enhancer repeats from Xenopus laevis at different loci in human chromosomes led to the formation of "pseudo-NORs" resulting in secondary constrictions in mitotic chromosomes (reviewed in [67, 96]). These structures were bound by human UBF, other components of the basal RNAP I transcription machinery and ribosome biogenesis factors [95, 97] but appear to be transcriptional inactive. Although, psoralen accessibility at pseudo-NOR sequences has not been tested to date, it is assumed that they reside in an open chromatin structure. Taken together, both Hmo1 and UBF are architectural proteins of open rRNA gene chromatin and involved in the maintenance of the nucleosome depleted state [56, 61, 63, 73]. Additionally, UBF binding at the rRNA gene promoter (and likely at enhancer regions) is likely related to its role in RNAP I PIC formation (reviewed in [18, 66]). As an essential PIC component, UBF but not Hmo1—is required for the establishment of the open rRNA gene chromatin state in vivo [56, 63, 73].

#### 2.5 Molecular Requirements to Establish the Open rRNA Gene Chromatin State

As indicated above, there is a tight correlation between RNAP I transcription and the establishment of the open chromatin state. In fact, studies in yeast indicated that the equilibrium of open and closed chromatin states observed in asynchronously dividing cells may be explained as the result of RNAP I transcription dependent opening and replication dependent closing of rRNA genes [61]. In each synthesis phase of the cell division cycle, replication leads to nucleosome deposition and chromatin closing at rRNA genes on both sister chromatids [98]. After rRNA genes have been replicated, opening of rRNA genes correlates with the onset of RNAP I transcription. In higher eukaryotes, cell division cycle dependent rRNA gene chromatin transitions were likely dismissed in early psoralen cross-linking studies in which interphase cells were compared with metaphase cells [51]. However at least in one study, time course experiments during the cell division cycle in a human cell line revealed very similar rRNA gene chromatin state transitions to those observed in yeast [99]. In agreement with a requirement for RNAP I transcription for establishment of the open rRNA gene chromatin state in replicating yeast cells, all rRNA genes adapt the closed chromatin state when RNAP I transcription is impaired [61]. In addition, there was good correlation between downregulation of RNAP I transcription and closing of rRNA gene chromatin in various physiological relevant situations. Thus, rRNA gene chromatin closing occurs when cells grow to stationary phase [51, 52, 100, 101], upon UV-damage [102, 103], or when cells differentiate [104]. On the other hand, upregulation of RNAP I transcription upon transfer of stationary yeast cells in fresh growth media correlates with rapid opening of rRNA genes [100, 101]. Furthermore, the observed opening of rRNA genes when replicationmediated chromatin closing was prevented strictly depends on ongoing RNAP I transcription [61].

It remains unclear if RNAP I transcription alone suffices to establish open, nucleosome-depleted rRNA gene chromatin. Thus, additional factors may assist RNAP I to convert genes from the closed to the open chromatin state. The "Facilitates Chromatin Transcription" (FACT) complex, known to support RNAP II transcription, was shown to associate with rRNA genes in yeast and human, copurified with human RNAP I and enhanced RNAP I nonspecific transcription from chromatin templates in vitro [39, 105]. In yeast, the chromatin remodeling factors Ino80, Isw1, and Isw2, as well as the Swi/Snf complex associate with rRNA genes in vivo and ex vivo and may modulate rRNA gene chromatin structure [106–108]. Furthermore, bona fide RNAP II transcription (elongation) factors TFIIH, Paf1, Spt4/5, Spt6, and THO were reported to support RNAP I transcription [109– 114]. In addition, early ribosome biogenesis factors might be components of rRNA gene chromatin in yeast and higher eukaryotes perhaps supporting efficient RNAP I transcription [97, 115]. Finally, a constantly growing number of factors which are involved in (epigenetic) posttranslational covalent modifications of chromatin components (including proteins and nucleic acids), as well as noncoding RNAs may influence RNAP I transcription and the establishment of rRNA gene chromatin states (reviewed in [6– 9]). Since most of the above factors have acknowledged roles in various nuclear processes, it is, however, often difficult to distinguish between direct and indirect effects on RNAP I transcription and rRNA gene chromatin structure.

#### 3 Conclusions

Our current knowledge of molecular mechanisms required for establishment of the open rRNA gene chromatin state is mostly based on in vivo observations or ex vivo analyses of relatively crude cellular fractions. In the future, analyses with defined in vitro reconstituted chromatin will probably allow to stringently test current hypotheses and to describe molecular mechanisms which are responsible for chromatin opening (see [116] and chapter by Merkl et al., this issue). One promising approach may involve the investigation of isolated native rRNA gene chromatin templates. Chromosomal rRNA gene domains in their native chromatin context can be purified from yeast, and initial analyses indicate that important features of the in vivo chromatin structure are maintained upon isolation [107]. The purified chromatin can be subjected to functional biochemical assays, or single molecule structural analyses [107, 117]. The reconstitution of transcription using highly purified components of the RNAP I transcription machinery and the native chromatin template in vitro might closely reflect the in vivo situation. Together, these studies may help to derive a detailed understanding about the interplay between chromatin structure and transcription on a molecular level. Given the tight connection between RNAP I transcription and early steps in ribosome biogenesis [115], it should be attempted to include selected ribosome biogenesis factors in the purified system. Perhaps, this may lead us soon to the first Christmas trees "grown" in vitro.

#### Acknowledgments

We are grateful to all scientists who contributed to this exciting field of research and apologize to those colleagues whose work has not been cited. We thank the members of the department of Biochemistry III for constant support and discussion. This work was funded by the Deutsche Forschungsgemeinschaft (DFG) in the context of the SFB960. P. E. M. was partly supported by a fellowship of the German National Academic Foundation.

#### References


the RNA polymerase I transcription factor hUBF during the cell cycle. J Cell Sci 104 (Pt 2):327–337


elongation factors Spt4p and Spt5p play roles in transcription elongation by RNA polymerase I and rRNA processing. Proc Natl Acad Sci U S A 103:12707–12712


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 3

# Analysis of Yeast RNAP I Transcription of Nucleosomal Templates In Vitro

## Philipp E. Merkl, Christopher Sch€achner, Michael Pilsl, Katrin Schwank, Kristin Hergert, Gernot L€angst, Philipp Milkereit, Joachim Griesenbeck, and Herbert Tschochner

#### Abstract

Nuclear eukaryotic RNA polymerases (RNAPs) transcribe a chromatin template in vivo. Since the basic unit of chromatin, the nucleosome, renders the DNA largely inaccessible, RNAPs have to overcome the nucleosomal barrier for efficient RNA synthesis. Gaining mechanistical insights in the transcription of chromatin templates will be essential to understand the complex process of eukaryotic gene expression. In this article we describe the use of defined in vitro transcription systems for comparative analysis of highly purified RNAPs I–III from S. cerevisiae (hereafter called yeast) transcribing in vitro reconstituted nucleosomal templates. We also provide a protocol to study promoter-dependent RNAP I transcription of purified native 35S ribosomal RNA (rRNA) gene chromatin.

Key words Transcription, Chromatin, Nucleosomes, Histones, RNA polymerase I, RNA polymerase II, RNA polymerase III, Ribosomal RNA gene chromatin, Rrn3, Core factor, Net1, In vitro assays, Saccharomyces cerevisiae, Yeast

#### 1 Introduction

In all eukaryotes there are at least three different nuclear multisubunit RNAPs (RNA polymerases) termed I–III (reviewed in [1], see also short reviews from Merkl et al., and Pilsl et al. in this issue). The enzymes share common subunits and a similar core structure but clearly differ in their composition and function (reviewed in [1], see also short reviews from Merkl et al., and Pilsl et al. in this issue). The template transcribed by RNAPs I–III in vivo is chromatin. In chromatin, approximately 146 bp of DNA are wrapped around an octameric core of histone proteins forming a basic repeating unit called the nucleosome (reviewed in [2–4]). This

Philipp E. Merkl and Christopher Sch€achner contributed equally with all other contributors.

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_3, © The Author(s) 2022

leads to compaction and impairs access to the genetic information. Therefore, chromatin structure must be transiently altered allowing transcription at defined genomic loci. Accordingly, characteristic chromatin transitions have been observed in vivo, when genes switch from an transcriptionally inactive to an actively transcribed state (reviewed in [5, 6]). There is evidence that RNAPs I–III deal differently with the chromatin template ([7], see also short reviews from Merkl et al. and Sch€achner et al. in this issue).

Defined in vitro systems including purified components contributed substantially to our current understanding of mechanisms of eukaryotic transcription (reviewed in [8]). The possibility to use in vitro reconstituted nucleosomal templates provided a minimal model system for transcription in the context of chromatin (reviewed in [9]). Here, we describe a protocol for comparative analyses of yeast RNAPs I–III in promoter-independent transcription of in vitro reconstituted chromatin templates. To this end, we developed a purification procedure for affinity purification of the three enzymes under identical conditions [10, 11]. The purified enzymes were devoid of cross-contamination with other RNAPs or transcription factors. For promoter-independent transcription, tailed DNA templates containing a 3<sup>0</sup> overhang can be employed (Fig. 1a) [12]. From 3<sup>0</sup> overhangs all RNAPs can initiate transcription without additional transcription factors. These templates in combination with the strong 601 nucleosome positioning sequence (601 templates) [13] were used for nucleosome assembly at defined positions (Fig. 1b). To determine the transcription efficiency through an assembled nucleosome, radioactively labeled RNA synthesized by the different RNAPs from the nucleosomal 601 templates are detected and quantified (Fig. 1c, d). Each transcription reaction contains another nucleosome-free reference template of different size, which serves as an internal control for the transcriptional activity of the respective RNAP. Additionally, transcription reactions are performed containing the naked 601 templates and the reference template (Fig. 1c). The transcription efficiency through a nucleosome is then calculated by dividing the quotient of transcripts from nucleosomal 601 template to reference template by the quotient of transcripts from naked 601 template to reference template. These analyses revealed marked differences between transcription of nucleosomal templates by RNAPs I–III ((Fig. 1d), and [7]). Whereas RNAPs I and III could readily transcribe through an in vitro assembled nucleosome RNAP II transcription was strongly impaired.

Nucleosomal templates obtained after salt dialysis have the advantage that they are fully defined. However, it is unclear in how far they reflect endogenous chromatin, the native template of nuclear transcription. Thus, we established a technique allowing the excision of a genomic region of interest in its native chromatin context by site-specific recombination in specifically engineered

Fig. 1 Promoter-independent transcription of in vitro assembled nucleosomal templates by RNAPs I–III. (a) Schematic representation of the template for nucleosome assembly containing a 601 nucleosome positioning sequence (601 template). The reference template lacks the nucleosome positioning sequence but is otherwise identical. Restriction sites used to release the templates from plasmids K1253 and K1573 are indicated. (b) Electrophoretic mobility shift assay after nucleosome assembly. Purified core histones H2A, H2B, H3, and H4 from chicken erythrocytes were assembled on the 601 template shown in (a) by salt gradient dialysis. Assembly reactions with different histone to DNA ratios (Table 7) were analyzed in a native polyacrylamide gel as described in Subheading 3.2. The position of the nucleosome-free template and the K1253 vector backbone and the chromatin template after assembly are indicated on the left. Sizes of selected DNA fragments in DNA markers (M) are indicated on the right. (c, <sup>d</sup>) Purified RNAPs I–III transcribe through a nucleosome on tailed templates with different efficiencies. (c) Transcription reactions were performed in the presence of a reference template and the 601 template before (-) or after (+) nucleosome assembly. The cartoon on the left indicates the length of the transcripts derived from the different templates. In vitro transcription assays were performed using 7.5 nM of purified RNAPs, identical buffer conditions and 10 nM of each of the different templates. Radiolabeled transcripts were separated on a denaturing polyacrylamide gel and visualized as described in Subheading 3.5.2. Sizes of selected RNA fragments in an RNA marker (M) are indicated on the left. (d) Quantification of the experiments shown in (c) was performed as described in Subheading 3.5.3

yeast strains (Fig. 2a) [17–19]. To assist affinity purification of the released chromatin domain, the genomic region of interest contains a cluster of DNA binding sites for the bacterial LexA protein. In the yeast strains used for recombination, recombinant LexA C-terminally fused to a tandem affinity purification (TAP) tag is constitutively expressed. The protein binds to its binding sites

Fig. 2 Promoter-dependent transcription of purified native rRNA gene chromatin by RNAP I. (a) Schematic representation of purification and restriction enzyme digest of yeast native 35S rRNA gene chromatin. (b) DNA analysis of samples withdrawn during the purification procedure (shown in <sup>a</sup>, and described in Subheadings 3.3.1 and 3.3.2). DNA of the indicated fraction of individual samples depicted on the top was isolated as described in Subheading 3.3.4, linearized with SacII and separated by electrophoresis in a 1% agarose gel. An image of the agarose gel stained with SYBR safe is shown on the top. The same gel was subjected to the Southern blot procedure with a radioactively labeled probe described in Subheading 2.3. The autoradiogram of the blot is shown at the bottom. The positions and respective length of DNA marker fragments (NEB 1 kb ladder, not shown) are depicted on the left of the gel picture. The position of the rDNA fragment derived from the 35S rRNA gene chromatin domain in the gel and on the blot is depicted on the right. (c) Quantitative Southern blot analysis of the "beads post EcoRV digest" sample (BpD) together with a titration from 0.1 to 1 fmol of plasmid K375. The isolated DNA was digested with PflmI prior to electrophoresis in a 1.5% agarose gel which was subjected to the Southern blot procedure. The positions and respective length of DNA marker fragments (NEB 1 kb ladder, not shown) are depicted on the left of the autoradiogram. The positions of PflmI fragments derived from the EcoRV digested or undigested 35S rRNA gene chromatin domain, as well as from the PflmI/EcoRV digested plasmid K375 are depicted on the right of the autoradiogram. (d) Promoterdependent in vitro transcription of native rRNA gene chromatin. Promoter-dependent in in vitro transcription was performed as described in [14] using either 10 nM of EcoRV (E) or PvuII (P) linearized plasmid K375 or within the chromatin domain and serves as bait for subsequent affinity purification of the genomic target region (Fig. 2b). The isolated native chromatin domains are suitable substrates for compositional analysis by mass spectrometry, structural analysis by electron microscopy or biophysical molecular tweezer analysis [20– 23]. Furthermore, they can be used in functional assays to study transcription factor binding, chromatin remodeling, or transcription on native templates in vitro [18, 24, 25]. Here, we show that purified native 35S rRNA gene chromatin can be used as template for promoter-dependent RNAP I transcription. To this end, the chromatin domain bound to the affinity matrix is linearized by restriction enzyme digest to produce transcripts of a defined length (Fig. 2c). Promoter-dependent transcription of the purified chromatin domain requires at least purified RNAP I, as well as two components of the RNAP I transcription machinery, the threesubunit core factor (CF) and the initiation factor Rrn3 (Fig. 2d). We find that the Net1 protein, which has previously been characterized as an activator of RNAP I transcription in vitro and in vivo [15, 26], leads to a robust stimulation of RNAP I dependent native chromatin transcription (Fig. 2d).

In summary, combining analyses involving strictly defined in vitro assembled nucleosomal templates and ex vivo purified native chromatin will likely help to gain further insights in mechanisms of nuclear transcription by eukaryotic RNAPs.

#### 2 Materials

#### 2.1 Preparation of Tailed Templates

Details about construction of plasmids K1253 (pUC19 tail g— 601) and K1573 (pUC19 tail g—w/o BS) can be found elsewhere [10, 11]. Plasmids and related sequence information are available upon request.


Fig. 2 (continued) 10 nM of EcoRV linearized rRNA gene chromatin as template. Note that only 20% of the chromatin template were properly digested by EcoRV and produce transcripts with a defined length. Individual transcription reactions were performed in the presence or absence of RNAP I (5 nM), CF (20 nM), Rrn3 (70 nM), and the RNAP I transcription activator Net1 (20 nM) [15, 16], as indicated on top. Radiolabeled transcripts were separated on a denaturing polyacrylamide gel and visualized as described in Subheading 3.5.2. The positions and respective lengths of RNAs synthesized from PvuII or EcoRV digested plasmid K375 or chromatin templates are indicated on the left and the right of the autoradiogram


#### Table 1 Buffers for chromatin assembly

(5<sup>0</sup> - CGAGTAAGTATAGGGTAAGGTGAT -3<sup>0</sup> ) as the 24 nt overhang generated by Nb.BsmI/KasI digest of plasmid K1253 or K1573.

	- 2. Bovine serum albumin (10 mg/mL), RNase free.
	- 3. DNA template (see Subheading 3.1).
	- 4. Siliconized micro tubes (Eppendorf).
	- 5. Buffers see Table 1.
	- 6. Polyacrylamide gel for electrophoretic mobility shift assay (EMSA) (Table 2).
	- 7. 6 EMSA buffer (60% glycerol, Orange G).
	- 8. FLA3000 (Fuji) Imager or equivalent imaging system for Ethidium bromide staining.
	- 9. Gel dryer (Drystar).
	- 10. Whatman filter paper (Macherey-Nagel, MN 827 B).

#### 2.3 Purification and Restriction Enzyme Digest of Native Yeast Chromatin

Materials for strain construction, cell growth and harvesting (preparation of yeast cell "spaghetti"), as well as for coupling of rabbit immunoglobulin G (IgG) to epoxy-activated magnetic beads, and protein analysis have been described elsewhere [19, 20]. Strain y2381 used for the purification of a 35S rRNA gene chromatin domain and plasmid K375 which contains an entire rDNA repeat were used in the experiments shown in Fig. 2 and have been


#### Table 2 Composition of native polyacrylamide gel for EMSA


2.4 Purification of RNAPs I, II, and III from

S. cerevisiae



#### 2. IgG coupled to magnetic beads, and BcMag Separator, rotating wheel as described in Subheading 3.3.2.


#### Table 3 Yeast strains used for RNA polymerase purification



#### Table 4 Buffers for RNA polymerase purification

#### 2.5 In Vitro Transcription of Chromatin Templates


#### 3 Methods

Tailed Templates

	- 2. DNA is precipitated by the addition of 0.1 volume of 3 M sodium acetate and 2.5 volumes of 100% EtOH and incubation at -20 C for at least 30 min. The template DNA is pelleted by centrifugation at 16,000 g for 20 min at 4 C, washed with 70% EtOH and resuspended in RNase-free water to result in a final concentration of 2 mg/mL.


#### Table 5 Buffers for in vitro transcription experiments

#### Table 6

#### Composition of denaturing polyacrylamide gel for RNA electrophoresis




#### Table 7 Assembly reaction setup


#### 3.2 In Vitro Nucleosome Assembly

Nucleosomes are reconstituted on tailed templates which contain one or multiple 601 positioning sequences [13]. The 601 sequence directs precise nucleosome positioning upon assembly, which is a prerequisite to obtain homogeneous chromatin templates.


Details on strain construction, cell growth and harvesting (preparation of yeast cell "spaghetti,") as well as instructions for coupling of rabbit IgGs to epoxy-activated magnetic beads analyses, and protein analysis can be found elsewhere [19, 20]. Subheadings 3.3.1, 3.3.2, and 3.3.4 have been adapted from [19] with modifications.

Unless noted otherwise all manipulations are carried out at 4 C.


#### 3.3 Purification and Restriction Enzyme Digest of Native Yeast Chromatin

3.3.1 Preparation of Cellular Lysates


Unless noted otherwise all manipulations are carried out at 4 C.


3.3.2 Affinity Chromatography


#### 3.3.4 DNA Analysis 1. Samples are adjusted to a total volume of 100 μL using TE buffer, followed by the addition of 100 μL of IRN buffer.

2. Samples are supplemented with 10 μL of 10% SDS and 2 μL of Proteinase K (20 mg/mL), mixed and incubated for 1 h at 56 C.

3.3.3 Restriction Enzyme Digest of Purified Native Chromatin Domains


3.4 Purification of RNAPs I, II, and III from S. cerevisiae Wild-type RNAPs I, II, and III are purified from yeast strains y2423 (RNAP I), y2424 (RNAP II), and y2425 (RNAP III) (Table 3) via IgG-protein A affinity purification [10, 11]. In each strain, the second largest subunit of the respective RNAP is expressed as a C-terminal fusion protein with protein A tag. A recognition site for TEV protease located between the C-terminus of the RNAP subunit and the protein A tag enables efficient elution of the different RNAPs (see below).

> 1. A 20 mL YPD culture is grown to stationary phase at 30 C. From this culture, 2 L of YPD are inoculated to an OD600 such that it results in an OD600 of 1.5 after overnight cultivation at 30 C.



#### Table 8 Composition of a standard in vitro transcription reaction


3.5.2 Denaturing Gel Electrophoresis and Visualization of Radiolabeled Transcripts Transcripts are separated in a denaturing polyacrylamide gel (20 cm 0.3 cm 17 cm).

	- 2. The relative radioactive signal intensity of a single band is background corrected and divided by the signal area.
	- 3. Relative signal intensities of transcripts derived from 601 templates with and without assembled nucleosome are divided by the signal intensities of transcripts derived from the reference template. This yields the normalized transcript levels derived from nucleosomal and nucleosome-free 601 templates. The quotient of the normalized transcript levels derived from the nucleosomal 601 template and derived from the nucleosomefree 601 template is taken as measure for the efficiency of transcription through a nucleosome.

#### 4 Notes

3.5.3 Transcript Quantification

> 1. Strong shaking of the coffee mill prevents sticking of the dry ice powder to the inside wall of the grinder. (Cryo-)gloves should be used to protect the hands during grinding.


#### Acknowledgments

We thank the members of the Department of Biochemistry III for constant support and discussion. This work was funded by the Deutsche Forschungsgemeinschaft (DFG) in the context of the SFB960. P. E. M. was partly supported by a fellowship of the German National Academic Foundation.

#### References


analysis of selected chromosomal domains from Saccharomyces cerevisiae. Nucleic Acids Res 42:e2


Le´ger I, Gadal O, Milkereit P, Griesenbeck J, Tschochner H (2012) The Reb1-homologue Ydr026c/Nsi1 is required for efficient RNA polymerase I termination in yeast. EMBO J 31:3480–3493


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part III

RNA Polymerases

# Specialization of RNA Polymerase I in Comparison to Other Nuclear RNA Polymerases of Saccharomyces cerevisiae

## Philipp E. Merkl, Christopher Sch€achner, Michael Pilsl, Katrin Schwank, Catharina Schmid, Gernot L€angst, Philipp Milkereit, Joachim Griesenbeck, and Herbert Tschochner

#### Abstract

In archaea and bacteria the major classes of RNAs are synthesized by one DNA-dependent RNA polymerase (RNAP). In contrast, most eukaryotes have three highly specialized RNAPs to transcribe the nuclear genome. RNAP I synthesizes almost exclusively ribosomal (r)RNA, RNAP II synthesizes mRNA as well as many noncoding RNAs involved in RNA processing or RNA silencing pathways and RNAP III synthesizes mainly tRNA and 5S rRNA. This review discusses functional differences of the three nuclear core RNAPs in the yeast S. cerevisiae with a particular focus on RNAP I transcription of nucleolar ribosomal (r)DNA chromatin.

Key words RNA polymerase I, RNA polymerase II, RNA polymerase III, Transcription, Chromatin, Nucleosomes, Ribosomal RNA genes, Transcription factors, Gene expression, Yeast, Saccharomyces cerevisiae

#### 1 RNA Polymerase I Has Only One Essential Genomic Target

Nuclear yeast RNAPs are protein complexes consisting of 12 (RNAP II), 14 (RNAP I) and 17 (RNAP III) subunits. They all share a conserved architecture of the RNAP core, which catalyzes the highly accurate polymerization of RNA from single NTP molecules [1–3] (see review by Pilsl and Engel in this issue). Based on structural analyses and on functional in vitro assays, on the one hand the molecular mechanisms driving the RNA polymerization appear to be similar in all RNAPs. On the other hand, all three enzymes have special features, supporting their specific in vivo tasks [1, 2]. Thus, RNAP II has to transcribe thousands of different genes and produces transcripts from a few hundred to several thousand of bases in length. To account for dynamic changes in cellular gene expression RNAP II has (a) to recognize many

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_4, © The Author(s) 2022

promoters, (b) to access differently modified chromatin templates, which probably requires to adjust its elongation and termination properties. To fulfill these multiple tasks, RNAP II interacts with many different factors at each step of the transcription process [4, 5]. In contrast, RNAP III recognizes only three distinct classes of promoters and has probably a distinct transcription termination mechanism [6]. RNAP III synthesizes mainly short noncoding RNAs (typically <200 bp) with high efficiency in rapidly growing cells. Accordingly, the RNAP III transcription machinery is rather well defined [7–10]. Finally, the yeast RNAP I transcription machinery has only one known genomic target, the multicopy 9.1 kb 35S rRNA genes [11]. The rRNA genes are transcribed at very high rates accounting for up to 60% of RNA synthesis upon cellular growth [12]. Accordingly, electron micrographs of chromatin spreads show an extremely high density of RNAP I molecules at rRNA genes, whereas most RNAP II-dependent genes are only sparsely covered with polymerases [13–15]. Furthermore, RNAP I and RNAP III-dependent genes are constitutively transcribed in actively dividing cells and—as opposed to the majority of RNAP II transcribed genes—apparently devoid of nucleosomes (see as review [16–18] and Sch€achner et al., within this issue).

#### 2 RNA Polymerase I Contains Additional Subunits Resembling Transcription Factors of RNAP II

RNAPs I, II, and III contain ten conserved subunits which form the catalytic core. In addition, all three enzymes contain a 2-subunit stalk structure, which is distantly related and consists of subunits A14/A43, Rpb4/Rpb7 and C17/C25 in RNAPs I, II and III, respectively. In RNAP III an additional heterotrimer C82/C34/ C3 connects the stalk with the RNAP III clamp, which probably helps to open the DNA duplex [19]. The overall architecture of the three RNAPs differs mainly in vicinity of the lobe structure, which is formed by the second largest subunits Rpa135, Rpb2 and Rpc128, respectively. Only one subunit—Rpb9—is bound to the RNAP II lobe, whereas the heterodimer A34.5/49 and subunit A12.2 bind to the lobe of RNAP I, and the homologous C17/C25 and C11 subunits to the lobe of RNAP III. RNAP I subunits A34.5 and A49 consist of three subdomains: a dimerization module formed by A34.5 and the N-terminal part of A49 (full length A34.5 and aa 1–110 of A49); the A49 linker (aa 105–187 of A49); and the C-terminal part of A49 (aa 187–415). The dimerization module binds to the "lobe" and "external" domains of the second largest Pol I subunit A135 on the core module side [20– 22]. In contrast, the C-terminal part of A49 which contains a tandem winged helix can be detected at the upstream face of the clamp core in several states of transcriptional active RNAP I molecules and seems to be flexible attached [20, 21, 23–31]. Biochemical and cell biological experiments showed that A34.5/A49 support transcription initiation, enhance RNAP I elongation and stimulate the intrinsic RNAP I RNA cleavage activity [26, 28, 32– 36]. RNA cleavage activity depends on the dimerization module which is located in close proximity to A12.2 whose C-terminal part is also important for efficient RNA cleavage [26]. Deletion of A12.2 results in growth inhibition at elevated temperature, sensitivity to nucleotide-reducing drugs, and inefficient transcription termination; hampers the assembly of the RNAP I enzyme; and leads to incorporation of wrong NTPs [37–40]. The lack of A12.2 may also lead to the loss of A34.5/A49, and might influence the intrinsic stability of elongation and termination complexes [40, 41].

Based on amino acid sequence similarities, position on the enzyme and function, the heterodimer formed by A34.5 and the N-terminus of A49 was suggested to be homologous to the RNAP II transcription factor TFIIF [26, 33]. TFIIF is predominantly found at promoter-proximal regions suggesting a crucial role in transcription initiation [42, 43]. On the other hand, TFIIF was suggested to leave the promoter—at least transiently—in complex with RNAP II, likely supporting early elongation [42–45]. A role of TFIIF in RNAP II elongation is further corroborated by in vitro studies, where it increases transcription rates by suppressing RNAP II pausing [46–48]. Several studies propose that TFIIF promotes transcription elongation in concert with the RNA cleavage supporting factor TFIIS, which structurally resembles the C-terminus of the RNAP I subunit A12.2 (reviewed in [3, 48–51]. In the RNAP II system, TFIIF and TFIIS are independently capable to release arrested RNAP II to resume productive elongation. However, these factors may synergistically enhance resumption of RNAP II transcription especially in conditions when the paused enzyme has additionally backtracked on the template [49]. Backtracked RNAP I requires the C-terminal, RNA cleavage activating part of A12.2 to resume elongation [52]. It is, however, unknown if the heterodimer A34.5/A49 participates in this process.

Finally, the C-terminus of A49 structurally and functionally resembles the tandem winged helix of TFIIE [33]. As TFIIE in RNAP II transcription, the C-terminal domain of A49 binds DNA and supports promoter-dependent transcription initiation in vitro [28] and RNAP I promoter recruitment in vivo [32]. In contrast to TFIIE, the A49 subunit stays associated with the enzyme after promoter clearance in vivo [32], and supports elongation of RNA from a DNA/RNA scaffold in vitro [33]. Recent studies suggest a more dynamic association of the heterodimer A34.5/A49 to the RNAP I lobe, since the heterodimer was absent from the core enzyme and A12.2 C-terminus was rearranged when RNAP I elongation was artificially blocked by addition of a nonhydrolyzable nucleotide [53].

A specific challenge for all elongating RNAPs is the transcription of chromatin templates. Since RNAP I and III transcribe nucleosome-depleted chromatin templates ( [13–15] see short review of Sch€achner et al. this issue). This indicates that nucleosomes are displaced from the chromatin template in the initial round of transcription, and it is possible that the lobe associated subunits of RNAP I and III may be involved in the process of nucleosome depletion.

#### 3 Nuclear RNAPs Transcribe Chromatin Templates

In eukaryotic cells, nuclear DNA is assembled into repeated units called nucleosomes consisting of 146 bp of DNA wrapped around an octameric complex of histone proteins [54]. Nucleosomes generally provide a strong barrier for elongating RNAP II in vitro [46, 55, 56]. DNA attached to nucleosomes recoils on the octamer, locking the enzyme in an arrested state [57] thereby providing four major superhelical pausing sites [58]. Additional factors are required for passage of RNAP II through this barrier (see below). The mechanism how purified RNAP II complexes passes nucleosomes in vitro was thoroughly studied [59, 60]. Whether assembled nucleosomes stay associated or are evicted during RNAP II transcription depends on the formation of a small intranucleosomal DNA loop and on the transcription efficiency (rate) [61]. Accordingly, various RNAP II complexes can remodel chromatin to a different extent [59, 60].

It was suggested, that RNAP II can only pass nucleosomes if uncoiling of the DNA from the surface of the octamer is facilitated and if transcription elongation factors keep the polymerase in a transcriptionally competent state [62]. TFIIF and TFIIS may prevent the release of RNAP II at nucleosomal barriers and thereby support transcription through a nucleosome in a synergistic manner [63]. Passage of purified RNAP II through in vitro assembled nucleosomes was also supported by elongation factor Spt4/Spt5 [64] and in the presence of TFIIS together with Spt4/5 and elongation factor Elf1 [65]. Insights in the molecular mechanism how Spt4/5 together with Elf1 facilitate progression of RNAP II through a nucleosome were recently obtained using high resolution structures by cryo-EM of different stalled elongation complexes [65]. Other factors that were reported to partially disassemble nucleosomes similar to Spt4/Spt5 are the histone chaperone FACT and Paf1c [66–68].

Much less is known about how RNAP I and RNAP III interact with nucleosomes in vitro. However, in vivo there is ample evidence that RNAP I and RNAP III genes are largely devoid of nucleosomes (reviewed in [16–18], see short review of Sch€achner et al. in this issue). It is an open question how RNAP I and RNAP III deal with nucleosomal genes in the initial round of transcription. In contrast to RNAP II which has only subunit Rpb9 associated to the lobe structure, yeast RNAP I and RNAP III have the TFIIFand TFIIS-homologous subunits A34.5/A49, A12.2 (RNAP I), and C37/C53, C11 (RNAP III) tightly associated to the lobe. Similar to TFIIF and TFIIS in RNAP II transcription, the homologous Pol I subunits A34.5/A49 and A12.2 facilitate RNAP I passage through nucleosomes [35]. Depletion of either the heterodimeric subunits A34.5/A49 or the cleavage supporting activity of A12.2 resulted in both reduced Pol I processivity and impaired passage though nucleosomes [35]. It is tempting to speculate that the homologous RNAP III subunits could play a similar role in RNAP III chromatin transcription. Furthermore, additional factors like FACT, Spt4/Spt5, or Paf1c have all been suggested to support RNAP I elongation [69–72]. Appropriate in vitro transcription system using highly purified factors and defined nucleosome templates will be the key to elucidate details of the molecular mechanisms of RNAP I and RNAP III transcription in the context of chromatin (see chapter by Merkl et al. in this issue).

#### Acknowledgments

We thank all the members of the department of Biochemistry III for constant support and discussion. This work was funded by the Deutsche Forschungsgemeinschaft (DFG) in the context of the SFB960. P. E. M. was partly supported by a fellowship of the German National Academic Foundation.

#### References


basis of rRNA cleavage. Nucleic Acids Res 40: 5591–5601


enhancement of sequence-specific pausing. Genes Dev 5:683–696


polymerase II with elongation factors. Science 363:744–747


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 5

# Structural Studies of Eukaryotic RNA Polymerase I Using Cryo-Electron Microscopy

## Michael Pilsl and Christoph Engel

#### Abstract

Technical advances have pushed the resolution limit of single-particle cryo-electron microscopy (cryo-EM) throughout the past decade and made the technique accessible to a wide range of samples. Among them, multisubunit DNA-dependent RNA polymerases (Pols) are a prominent example. This review aims at briefly summarizing the architecture and structural adaptations of Pol I, highlighting the importance of cryo-electron microscopy in determining the structures of transcription complexes.

Key words RNA polymerase I, Cryo-electron microscopy, Pre-rRNA transcription

#### 1 Cryo-Electron Microscopy: The New Standard in Transcription Research

The visualization of macromolecular complexes is essential to our understanding of their function. This is especially true for eukaryotic RNA polymerases (Pol) I, II, and III. These enzymes play a pivotal role within the central dogma of molecular biology by synthesizing the 35S ribosomal RNA precursor (Pol I), messenger and many noncoding RNAs (Pol II), and tRNAs, 5S rRNA, U6 snRNA as well as other small, structured RNAs (Pol III).

Almost 20 years ago, advances in cryo-crystallography allowed solving the structure of RNA polymerase II, first in its 10-subunit form [1], later comprising all 12 subunits [2], providing insights into the function of this molecular machine at an unprecedented level of detail. This gave rise to a number of follow-up studies, resulting in structures of an actively elongating Pol II form [3, 4] and elongation [5, 6] or initiation [7, 8] factor–bound Pol II. Thereby, the molecular mechanisms of transcription, as well as regulatory and catalytic functions of transcription factors, could be deciphered.

However, X-ray diffraction analysis relies on the availability and quality of the analyzed crystals. It is therefore not surprising that a

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_5, © The Author(s) 2022

number of transcription factors could never be successfully studied in complex with their respective polymerase by crystallography. This includes the general Pol II initiation factors TFIIF and TFIIE, which are involved in initiation complex formation and promoter DNA melting [9]. A crystal structure of Pol I was solved 10 years after Pol II in an inactive conformation [10, 11], a Pol III crystal structure is still lacking to date. Crystallization depends on high amounts of purified material. Whereas conformational heterogeneity usually is problematic for crystal formation, cryo-EM allows for visualization of different functional states of proteins in vitrified ice, therefore capturing close-to-native states. Technical improvements such as the development of direct electron detectors [10], highly stable microscopes and improved processing software [11, 12] pushed previous limitations of the technique and led to the often quoted "resolution revolution" in cryo-EM [13]. Within this book, we describe protocols for a sample preparation and single-particle cryo-EM screening workflow, adapted to RNA polymerase complexes (Chapter 6 ).

Here, we aim to briefly outline the benefits and limitations of single-particle cryo-EM for the analysis of transcription complexes at the example of RNA polymerase I in the context of functional characterization reviewed by Merkl et al. in the same issue.

#### 2 Pol I Specific Subunits Resemble Built-in Transcription Factors

Yeast Pol I has a molecular weight of 590 kDa and consists of 14 protein subunits. A core of ten subunits includes the large subunits A190 and A135, the subcomplex AC40/AC19 (shared with Pol III), the common subunits Rpb5, Rpb6, Rpb8, Rpb10, and Rpb12 (also included in Pol II and Pol III) and the subunit A12.2. Pol I also contains the specific subunit complex A14/A43 forming the "stalk" and the specific heterodimer subcomplex A49/A34.5. In Fig. 1, we present a "hybrid model" of Pol I, constructed using the software package COOT [14]. The model combines structural information obtained from cryo-EM reconstructions of elongation and initiation complexes and the dimer crystal structure (see below). The Pol I subunit complex A49/A34.5 structurally and functionally resembles a built-in version of subunits Tfg1/2 constituting the Pol II initiation factor TFIIF [15] and is involved in open complex stabilization as well as promoter escape. Nevertheless, subcomplex A49/A34.5 may also contribute to transcript cleavage activity [16] and may play a role in elongation [17], as reviewed by Merkl et al. in this book. The Pol I subunit A12.2 displays features of the Pol II subunit Rpb9, as well as the Pol II elongation/cleavage factor TFIIS, explaining the intrinsic ability of Pol I for transcript cleavage [18] and efficient recovery from deep backtracks [19].

Fig. 1 Ribbon model of RNA polymerase I highlighting specific subunits and –domains. Hybrid model constructed using COOT [14] for demonstration purposes. The structure of the Pol I elongation complex (PDB 5M3F) was extended by adding (a) a model of the C-terminal domain of subunit A12.2, (b) the "connector" domain of subunit A43 from the crystal structure (both from PDB 4C2M) and (c) the linker and tandem-winged helix domains [15] from an ITC reconstruction (PDB 5 W66). The "front view" looks along the incoming ("downstream") DNA. Subunits not visible in the front view but present in the model are AC19, Rpb10, and Rpb12. Cyan spheres depict coordinated zinc atoms

The constitutive association of subunits A12.2 and A49/A34.5 may be a symptom of the enzyme's adaptation to transcribing one specific, extraordinarily long gene. A differential regulation of Pol I transcription similar to the conditional, gene-specific regulation of Pols II and III is most likely not required. Instead, a binary activity switch can be achieved by posttranslational modification of the polymerase or its initiation factors [20–23].

#### 3 The Pol I Transcription Cycle Visualized In Vitro

Throughout transcription of a DNA template, Pols pass through three main phases, which are more or less conserved throughout various organisms. First, the Pol is recruited to the template sequence, melts the DNA duplex with or without the help of additional factors and begins the synthesis of an RNA strand (initiation). Thereafter, the initial RNA is extended according to the DNA template strand (elongation). Finally, Pol dissociates from its DNA template and releases its RNA product (termination) [24].

The Pol I transcription cycle requires few basic transcription factors [25, 26]. Basal initiation in yeast can be achieved with only the factor Rrn3 and the Core Factor (CF) complex, consisting of proteins Rrn6, Rrn7, and Rrn11 [27, 28]. Elongation may involve regulatory factors in vivo, but can commence in vitro without them [25, 26]. Termination finally requires a Myb-domain containing protein, Nsi1 in yeast [29, 30]. Pol I can adopt a dimeric state specific to this enzyme, [31], in which both molecules are inactivated [32] and prevented from binding initiation factors Rrn3 and CF [33–35]. Dimerization is reversible [33] and may play a role in storage of Pol I molecules during starvation [36, 37]. Using cryo-EM in many variations, our understanding of the Pol I transcription cycle has been significantly improved in recent years. A structural description of the Pol I transcription cycle allowed a detailed structure–function analysis and the comparison to other transcription systems [38–41].

Specifically, single-particle cryo-EM analyses showed how Pol I monomers are bound by the initiation factor Rrn3 [42], which allows for recruitment of CF subunit Rrn7 [37, 43, 44] that is related to the general Pol II initiation factor TFIIB [45, 46]. The interaction with Rrn3 prevents the formation of transcriptionally inactive Pol I dimers. In baker's yeast (Saccharomyces cerevisiae) Pol I dimerization depends on the specialized, nonconserved "connector" domain of subunit A43, suggesting that this mode of regulation is specific to S. cerevisiae [37]. However, recent cryo-EM studies of Pol I from fission yeast (Schizosaccharomyces pombe) demonstrated that inactive dimers can form independent of the connector domain utilizing divergent structure elements [47]. It is therefore possible that different organisms evolved different dimer interfaces, but use the same strategy of Pol I hibernation by dimerization during periods of starvation.

Monomeric Pol I bound to Rrn3 can be recruited by promoterengaged CF. Whereas the structure of CF was determined by X-ray crystallography [48], its interaction with the Pol I–Rrn3 complex was apparently too flexible for this method to succeed. Instead, three independent studies used cryo-EM to visualize a reconstituted initially transcribing complex (ITC) that relies on an artificially stabilized, mismatched transcription bubble with a short initial product RNA [48–50]. Comparison of these studies shows that seemingly similar experimental cryo-EM approaches may yield different, though highly complementary results. This demonstrates the influence of sample preparation and experimental conditions on the outcome of a cryo-EM experiment and the strength of singleparticle cryo-EM to elucidate dynamic processes. Specifically, some complexes exist in conformations that fail to bind Rrn3 [49], which is required essential to the initiation process and dissociates after promoter escape [42, 51, 52]. These complexes may represent later initiation intermediates. In another structure, the RNA primer is lost and the C-terminal domain of subunit A12.2 is inserted into the funnel domain of Pol I, as would be expected during RNA cleavage events [50]. A third reconstruction showed local resolution differences suggesting high flexibility in distal CF regions [48]. More recently, consolidating structures of close-to-native preinitiation complexes were reported [53, 54]. A closed and an open complex could be reconstructed from a small fraction of recorded particles but yielded important information about the roles of a flexible loop in Rrn3 in CF engagement, and the role of the linker/tWH domain of subunit A49 in template DNA melting, open bubble stabilization, and Rrn3 dissociation [53].

Following initiation, Pol I adopts an actively elongating conformation. This conformation was never successfully crystallized but was reconstructed by single-particle cryo-EM [55, 56]. It is still under debate, whether dissociation/transient association of the A49/A34.5 heterodimer and formation of a 12-subunit polymerase has a physiological function under some circumstances. Chromatin immunoprecipitation (ChIP) data does not suggest subunit depletion along the 35S gene body in vivo [57], although a lack of density in cryo-EM reconstructions of S. cerevisiae and S. pombe Pol I ECs suggests that subdomain relocation can take place [47, 58]. Upon initiation, the central DNA-binding cleft contracts and tightly binds downstream DNA and the DNA–RNA hybrid region, thus facilitating processive elongation. Contraction upon activation is apparently common to Pol I regulation in different organisms [47] but differs from Pol II [3] and III [59]. The various Pol I EC reconstructions demonstrated the versatile nature of the specific subunits A12.2, A49, and A34.5 [47, 55, 56, 58], and the importance of a Pol I-specific arginine residue in the "bridge-helix" in stalling at DNA lesions caused by UV radiation [60]. Furthermore, the physiological relevance of a contracted Pol I state was confirmed by the 3D-reconstruction of the enzyme from ex vivo purified, actively transcribing Pol I using electron cryo-tomography [55]. Interestingly enough, the underlying preparation technique has been used to describe Pol I functionality since the 1960s [61].

Structural information on transient and highly dynamic states of the Pol I transcription cycle, such as termination, are still lacking to date.

#### 4 Outlook

Continuing investigation of the Pol I transcription system currently aims at defining the structural basis of promoter recruitment in the context of TATA-binding protein [62] and yeast upstream activating factor (UAF) which are important to achieve full initiation levels under physiological conditions [27, 63]. The structural investigations of these complexes and their interplay with the Pol I enzyme will be the key to understand this initiation systems in comparison with Pol II [64, 65] and Pol III [66, 67]. Ultimately, such comparative structure–function analyses will enable us to understand the particular adaptation of the Pol I transcription system to pre-rRNA transcription.

Cryo-EM has now firmly established itself as the method of choice to analyze the structure of such mega-Dalton sized, dynamic, macromolecular complexes. However, the method has also proven to be prone to certain errors. Especially the lack of tools for intrinsic validation of density interpretation may be an issue for initial sequence assignment at resolutions worse than 4 A˚ . Continuing method development at the experimental and the computational level continues and may soon overcome issues of flexibility and density assignment. For the time being, however, the interpretation of cryo-EM reconstructions should be carefully evaluated and supported by functional data or mutational analysis. Results can also be strengthened by crystallization or small angle X-ray scattering analysis of flexible domains [68], native mass spectrometry [15], hydrogen–deuterium exchange mass-spectrometry evaluation [69], or single-molecule FRET analysis [70]. Especially distance restrains obtained from protein crosslinking coupled to mass spectrometry provides architectural information and may assist low-resolution density assignment [71, 72].

#### Acknowledgements

The authors thank all members of the Structural Biochemistry group and Biochemistry I/III departments of the University of Regensburg for help and discussions. This work was supported by the Emmy-Noether-Programme (DFG grant no. EN 1204/1-1 to CE) and SFB 960 TP-A8.

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recognition and promoter melting. Nat Commun 10:5543. https://doi.org/10.1038/ s41467-019-13510-w


polymerase I using recombinant core factor. Gene 492:94–99. https://doi.org/10.1016/ j.gene.2011.10.049


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 6

# Preparation of RNA Polymerase Complexes for Their Analysis by Single-Particle Cryo-Electron Microscopy

## Michael Pilsl, Florian B. Heiss, Gisela Po¨ ll, Mona Ho¨ cherl, Philipp Milkereit, and Christoph Engel

#### Abstract

Recent technological progress revealed new prospects of high-resolution structure determination of macromolecular complexes using cryo-electron microscopy (cryo-EM). In the field of RNA polymerase (Pol) I research, a number of cryo-EM studies contributed to understanding the highly specialized mechanisms underlying the transcription of ribosomal RNA genes. Despite a broad applicability of the cryo-EM method itself, preparation of samples for high-resolution data collection can be challenging. Here, we describe strategies for the purification and stabilization of Pol I complexes, exemplarily considering advantages and disadvantages of the methodology. We further provide an easy-to-implement protocol for the coating of EM-grids with self-made carbon support films. In sum, we present an efficient workflow for cryo-grid preparation and optimization, including early stage cryo-EM screening that can be adapted to a wide range of soluble samples for high-resolution structure determination.

Key words RNA polymerase I, Preparation of transcription complexes, Single-particle cryo-electron microscopy, Grid preparation, Plunge freezing, Negative staining

#### 1 Introduction

During the past years, development of more sophisticated microscopes, new direct electron detectors, and advances in computational image processing caused a "resolution revolution" [1] in the field of cryo-electron microscopy (cryo-EM). This culminated in the award of the Nobel Prize in Chemistry to three scientists driving these developments: Joachim Frank, Richard Henderson, and Jacques Dubochet in 2017. Recent developments have been summarized in detail [2–8]. With its increasing popularity, the technique is now more easily accessible and widely used in many areas of structural biology research, such as the analysis of transcription complexes [9]. Among many others, cryo-EM studies on RNA polymerase (Pol) I complexes became possible and advanced our understanding of the molecular mechanisms employed by this

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_6, © The Author(s) 2022

highly specialized enzyme [10–19]. Here, we describe the workflow of biochemical purification and sample preparation of Pol I complexes and the optimization of cryo-grid preparation containing frozen, hydrated single particle specimens with the goal of acquiring high resolution images for structure determination. We focus on grid preparation, quality control, and EM screening in an iterative manner. We also emphasize the steps necessary for establishment of a customized workflow that can be tailored to the needs of any sample suitable for single-particle analysis.

The success of structure determination by single-particle cryo-EM depends on high-quality biochemical preparation and characterization of the molecule(s) of interest. Buffer optimization and stabilization of complexes should be done in the initial phase of a project, strategies are described in detail [20]. We previously presented protocols for the purification of 14-subunit, 590 kDa Pol I complexes and their characterization in vitro [21, 22]. Starting from this, we detail the specimen features which are important for successful structure determination using single-particle cryo-EM and suggest approaches for their optimization. For this purpose, we compare complex preparation- and assembly strategies using endogenously purified Pol I and recombinant transcription factors on nucleic acid templates. Transcription factor complexes can be (a) assembled on a biotinylated DNA, enriched using the interaction with bead-coupled streptavidin and eluted with restriction enzymes [23]. Alternatively (b), size exclusion chromatography (SEC) may yield homogenous and stable complexes that are wellsuited for cryo-grid preparation. Large macromolecules can also be enriched using density gradient centrifugation protocols (c). As such, the gradient-fixation (GraFix)-method relies on a sedimentation step coupled with an intra- and intermolecular cross-linking step combining purification and complex stabilization [24, 25]. Following GraFix, a buffer exchange is required to remove sucrose or glycerol that would otherwise interfere with the subsequent freezing process and may increase background noise in cryo-EM images. Generally, cross-linking of protein-protein or protein-nucleic acid complexes can improve their stability during the grid preparation process. This cross-linking can be coupled to a purification step (as in GraFix), or performed directly before grid-plunging (d). These preparation techniques can also be combined, for example, carrying out an SEC run after batch cross-linking. Such a strategy combines the advantages of sample stabilization and purification while directly including the transition to a suitable buffer system but requires larger quantities of sample.

In addition to sample preparation and stabilization, we discuss the use of different grid types and support materials. General cryo-EM sample preparation techniques applicable to a wide range of samples were described in detail [26, 27]. Adsorption of particles to a thin support film can aid sample concentration, alter bias in orientation distribution, or protect fragile particles from denaturation on air–water interfaces [28]. To this end, grids covered with thin carbon layer substrates are commonly used. Additionally, graphene oxide or hydrogenated graphene were reported to be wellsuited support materials as they are almost transparent to electrons [29–31]. While holey carbon grids covered with ultrathin support layers are commercially available (see Subheading 2), many support types can be manually prepared more economically at similar or higher quality. We also describe a technical gadget to float an ultrathin carbon film on holey carbon grids. This is similar to other floatation techniques [32], but allows for sample adsorption on precoated grids directly prior to plunging and may thus be helpful to some users. This device is based on [33] and is commercially available in a modified form (SKU 10840; LADD Research Industries, Williston, VT, USA).

In addition to sample optimization and the choice of grid type, various physical parameters influence the preparation of cryo-EM grids. Glow discharge settings, grid type variation, buffer choice, and the mechanics of the blotting device should be carefully considered for each individual sample.

In general, we recommend initial optimization using negative staining EM, followed by two stages of cryo-EM screening (Fig. 1). The first cryo-screening stage aims at an evaluation of grid types, support films, or blotting conditions and gives information on sample behavior in ice. In a second phase of cryo-screening, intermediate-resolution single-particle maps may be reconstructed. Phase I cryo-screening results yield insights into sample behavior, whereas results of the second cryo-screening phase indicate whether the sample is suitable for high-resolution data collection by identifying flexibilities within the macromolecular complex and bias in orientation distribution.

For screening, we recommend the acquisition of low-magnification grid maps for navigation and evaluation of overall ice distribution. Low dose acquisition strategies to avoid beaminduced particle damage are used. This workflow includes focusing and exposure-dose measurement at higher magnifications on grid areas adjacent to the foil-hole of interest using image/beam shift functions, as for the acquisition of high-resolution images. Overviews of sample preparation, the use of cryo-holders and imaging strategies have been described [34–37]. For automated data collection, the SerialEM software includes many more features that have been recently summarized [38]. The open-source software is freely available and can be adapted to a wide range of microscopes and camera systems. Recent developments include the implementation of "on-the-fly" data processing and evaluation strategies into the software packages RELION, Warp, cryoSPARC, and SPHIRE, that may be useful in sample evaluation [39–42]. Here, we focus on the sample and grid preparation, as well as phase I cryo-EM screening approaches for multisubunit RNA polymerases.

Fig. 1 Overview of a workflow for (cryo-)EM sample screening and grid optimization. We suggest five phases, starting with sample preparation and iterative evaluation of screening results. Detailed protocols for individual steps are referenced and listed in Subheadings 3.1–3.6

#### 2 Materials


Fig. 2 Schematic representation of carbon film transfer onto EM grids as detailed in Subheading 3.4 (QF: Quantifoil, EM: Electron Microscopy)





#### 3 Methods


	- 2. Vent and open the carbon coater and place glass slide with mica on stage.
	- 3. Check carbon rods: left rod should have a smooth tip, right rod should have a plain face, both rods have to be in physical contact.
	- 4. Close the recipient chamber. Turn on Cressington carbon coater device and wait for the vacuum to be under <sup>4</sup> <sup>10</sup><sup>5</sup> mbar.
	- 5. Degas carbon rods: Slowly turn up voltage to 2 V, hold until carbon rod glows red. Wait for vacuum to recover and repeat two more times.
	- 6. Switch on the thickness monitor and evaporate carbon at 3.6 V for 10 s. Do not remove shield to ensure indirect evaporation. Wait at least 30 s before reading the carbon film thickness from the monitor, the film should be ~2 nm. Repeat evaporation if necessary (see Note 9).
	- 7. Vent the recipient chamber, wait at least 5 min before opening it (evaporation source will become hot during evaporation). Store carbon films on mica carrier for >3 days (ideally >10 days) in a petri dish before use (see Note 10).

10840) of a similar apparatus is available from LADD Research Industries (Williston, VT, USA). We also use this flotation chamber to transfer surface assembled graphene oxide [44] onto EM grids.

	- 2. Check Vitrobot settings and perform a test run without grid or forceps. Vitrobot settings: 4 C, 100%, Blotting settings: wait 0 s, blot 5 s, drain 0 s, Blot force 12 (see Note 14).

#### 3.5 Cryo-Grid Preparation

3.6 Cryo-Electron Microscopy


#### 1. Place Gatan 626 cryo holder in transfer station and fill the transfer station with liquid nitrogen. Fill the holder Dewar (which will keep the specimen cold during the transfer and microscopy process) with liquid nitrogen directly afterward (see Note 21).


Optional: Prepare electron microscope by tilting the stage 30 to avoid spilling liquid nitrogen during holder insertion.


#### 4 Notes


#### Acknowledgments

We thank Herbert Tschochner and Joachim Griesenbeck for discussions, Lena Heuschneider for technical assistance, and Reinhard Rachel for practical advice. We acknowledge funding by SFB960 of the DFG (TP-A8 to CE and TP-B1 to PM) and by the "Emmy-Noether-Programme" (DFG grant no. EN 1204/1-1 to CE).

#### References


using single-particle electron microscopy. Methods Mol Biol 718:3–22


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part IV

# Ribosome Assembly, Transport and RNP Complexes

# Chapter 7

# Eukaryotic Ribosome assembly and Nucleocytoplasmic Transport

## Michaela Oborska´-Oplova´ , Ute Fischer, Martin Altvater, and Vikram Govind Panse

This chapter is dedicated to the memory of our former graduate student and friend Dr. Cohue Pen˜a.

#### Abstract

The process of eukaryotic ribosome assembly stretches across the nucleolus, the nucleoplasm and the cytoplasm, and therefore relies on efficient nucleocytoplasmic transport. In yeast, the import machinery delivers ~140,000 ribosomal proteins every minute to the nucleus for ribosome assembly. At the same time, the export machinery facilitates translocation of ~2000 pre-ribosomal particles every minute through ~200 nuclear pore complexes (NPC) into the cytoplasm. Eukaryotic ribosome assembly also requires >200 conserved assembly factors, which transiently associate with pre-ribosomal particles. Their site(s) of action on maturing pre-ribosomes are beginning to be elucidated. In this chapter, we outline protocols that enable rapid biochemical isolation of pre-ribosomal particles for single particle cryo-electron microscopy (cryo-EM) and in vitro reconstitution of nuclear transport processes. We discuss cell-biological and genetic approaches to investigate how the ribosome assembly and the nucleocytoplasmic transport machineries collaborate to produce functional ribosomes.

Key words Budding Yeast, Ribosome Assembly, Nuclear Import, Nuclear Export, preribosome structure

#### 1 Introduction

Eukaryotic ribosome assembly takes place across multiple cellular compartments: the nucleolus, the nucleoplasm and the cytoplasm (Fig. 1). This dynamic and energy consuming process requires the coordination of three RNA polymerases (I, II, and III), the RNA splicing, and nucleocytoplasmic transport machineries [1]. Despite this complexity, ribosome biogenesis is an incredibly efficient process, with yeast producing up to 60 ribosomes every second [2].

The small subunit (SSU) processome is the first ribosome precursor assembled cotranscriptionally in the nucleolus through stepwise association with UTP-A, UTP-B, and UTP-C complexes

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_7, © The Author(s) 2022

Fig. 1 Current model for eukaryotic ribosome assembly. Transcription of primary 35S rRNA transcript, the common precursor of 18S, 5.8S, and 25S rRNAs, by Pol I from rDNA repeats together with cotranscriptional joining of U3 snoRNP, r-proteins and 40S assembly factors form the 90S ribosomal precursor. 5S rRNA is transcribed separately by Pol III for assembly of 5S RNP before joining the 60S pre-ribosome. Cotranscriptional cleavage of 35S rRNA at A2 site separates the 60S and 40S r-subunit maturation pathways. Pre-ribosomal particles undergo a cascade of maturation steps in the nucleoplasm by transient association with assembly factors until reaching nuclear export competency. Once exported to the cytoplasm, last maturation events and quality control steps can occur resulting into functional r-subunits production. (Adapted from Pen˜ a et al. 2017)

and U3 snoRNP [1, 3]. Cryo-electron microscopy (cryo-EM) studies are revealing how UTP-A, UTP-B complexes, U3 snoRNP and additional biogenesis factors encapsulate and guide pre-rRNA folding in a hierarchical 5<sup>0</sup> -to-3<sup>0</sup> -oriented manner [4–7]. During nucleolar maturation, the 5<sup>0</sup> domain of the 18S rRNA achieves a mature conformation, and the central domain is correctly positioned relative to the 5<sup>0</sup> domain. In contrast, the 3<sup>0</sup> major domain is buried within the SSU processome core, and its conformation is distinct as in the mature 40S subunit. Release of the 40S pre-ribosome requires endonucleolytic cleavages within the pre-rRNA [8–11], the release of U3 snoRNA and associated proteins catalyzed by the RNA helicase Dhr1 and its cofactor Utp14 [12, 13]. The RNA exosome mediates degradation of 50 -ETS (external transcribed spacer) rRNA within the 50 -ETS rRNA– UTP complex to disassemble and recycle the UTPs for a new round of 40S assembly [14].

Following 40S preribosome release the growing 27S pre-rRNA associates with 60S-specific r-proteins and maturation factors to initiate 60S pre-ribosome assembly. Cryo-EM structures of early states of the 60S pre-ribosome suggest a sequence of events during nucleolar 60S assembly. Early states the nucleolar 60S pre-ribosome show a characteristic arch-like morphology, whereas the later states adopt a compact shape closer to a mature 60S subunit. The 50 domains (I and II) within 27S pre-rRNA adopt a near mature conformation, and connect with the 3<sup>0</sup> terminal domain VI, thus forming the solvent-exposed backside of the 60S subunit. In all states, the pre-rRNA spacer ITS2, located in the 27S pre-rRNA between 5.8S and 25S rRNA, and the associated ITS2 factors form the "foot" structure. The central domains III, IV and V, which form the subunit interface, are not visible in the early states. Although cotranscriptional assembly of the 60S pre-ribosome occurs in a sequential manner, it undergoes nonlinear compaction of domains I, II, and VI, which then allows domains III, IV, and V to fall onto the arch-like structure, thus preparing the pre-ribosomal cargo for nuclear export [15–17].

In yeast, before every cell division, ~200,000 pre-ribosomes are transported to the cytoplasm through ~200 NPCs (nuclear pore complexes) by the exportin Crm1 that recognizes nuclear export sequences (NESs) on cargos and interacts with the FG-rich (phenylalanine/glycine-rich) meshwork of the NPC transport channel [18–22]. Nmd3 is the only identified essential adaptor for Crm1 mediated 60S pre-ribosome export [23, 24]. In contrast, no essential NES-containing export adaptor has been identified for 40S pre-ribosome export. 40S pre-ribosome bound shuttling assembly factors Ltv1 and Rio2 can recruit Crm1 in the presence of RanGTP through a leucine-rich NES. Pre-ribosomes also employ multiple export factors (Mex67-Mtr2, Arx1, Ecm1, Bud20) that directly interact with the FG-meshwork of the transport channel [1, 21, 22]. In addition, a non-FG pathway involving the mRNA export factor Gle2 also facilitates 60S pre-ribosome export [25].

Exported pre-ribosomes undergo final maturation and proofreading before initiating translation [26]. Cytoplasmic proofreading of 60S pre-ribosomes involves release of assembly factors that block binding of r-proteins, interactions with translation factors, or pairing with the 40S. Tif6 prevents binding of immature 60S pre-ribosomes to mature 40S subunits, ensuring that only properly assembled subunits engage in translation. Quality control of the 40S pre-ribosome relies on assembly factors that prevent the premature binding of initiation factors, mRNA, tRNA, and the 60S subunit. Ltv1 and Enp1 directly bind uS3 on its solvent side, thereby blocking the mRNA channel opening. Rio2, Tsr1, and Dim1 bind the subunit interface, thus preventing joining of the mature 60S subunit and translation initiation factor eIF1A. Nob1 and Pno1 block the binding of eIF3, thereby interfering with translation initiation [27]. After release of Rio2, Tsr1, and Dim1 initiated by the reorganization of the beak structure, the 40S pre-ribosome becomes competent to interact with a mature 60S subunit. This translation-like interaction is thought to test the ability of a 40S pre-ribosome to engage with a mature 60S subunit, and only then triggers Nob1 to cleave 20S pre-RNA to mature 18S rRNA in vitro [28, 29]. Cryo-EM studies are revealing how these late factors interact with the 40S pre-ribosome and have provided a structural framework for the ordering of cytoplasmic maturation events [30–32].

In addition to >200 assembly factors, ribosome assembly relies on efficient nucleocytoplasmic transport [1]. All r-proteins and assembly factors need to be imported into the nucleus, and correctly assembled pre-ribosomal particles need to be transported through nuclear pore complexes into the cytoplasm. How the ribosome assembly machinery collaborates with the cellular trafficking pathways is currently under intense investigation. Here, we outline biochemical approaches that permit rapid screening and analyses of pre-ribosomal particles for single particle cryo-EM studies, and reconstitution of nucleocytoplasmic transport processes. We discuss genetic and cell-biological approaches that will enable the unveiling of the functional interface between the ribosome assembly and nucleocytoplasmic machineries.

#### 2 Materials, Reagents, and Yeast Media



#### 2.2 Reagents (Listed in Alphabetical Order)

	- 2. SC plates (SC, 2% agar).
	- 3. Yeast-extract peptone dextrose (YPD) media: 1% yeast extract, (ForMedium, YEM03), 2% peptone (ForMedium, PEP03), 2% glucose (Sigma-Aldrich, G8270).
	- 4. YPD plates (YPD, 2% agar).

#### 3 Pre-ribosome Isolation for Single Particle Cryo-Electron Microscopy

During the last decade, affinity-purification protocols combined with sensitive mass spectrometry have dramatically altered our understanding of eukaryotic ribosome assembly [33]. By employing the powerful "tandem affinity-purification" (TAP), several groups have unravelled the compositions of the 90S, 60S, and 40S pre-ribosomes and thus expanded the inventory of the assembly machinery and ordered the 40S and 60S maturation pathways [34–40]. Recently, improvement in these biochemical approaches together with advances of cryo-EM have driven structural studies of pre-ribosomal particles, thus providing high-resolution snapshots of the process of eukaryotic ribosome assembly [4–6, 15– 17, 30–32, 41, 42] (Fig. 2a, b).

3.1 Preparing Yeast Cells for Cryogenic Lysis (Modified from Rout Lab Protocol: Harvesting Cells and Making Yeast Noodles [43])

3.1.1 Buffers and Solutions

3.1.2 Yeast Cell Preparation

Resuspension Buffer: 1.2% PVP-40 (polyvinylpyrrolidone), 20 mM HEPES pH 7.4, Supplemented with 1:100 complete protease inhibitor cocktail tablet (Roche), 2 mM PMSF, 1 mM DTT (see Note 1).


Fig. 2 Purification and structure of a late 40S pre-ribosome. (a) Protein composition of 80S ribosomes Nob1- D15N particles, purified via ProteinA (pA)-tag. Proteins were separated by SDS-PAGE and visualized by silver staining. Labelled protein bands were characterized by mass spectrometry. (b) Front and back view of cryo-EM structure of a cytoplasmic 40S preribosome (PDB: 6FAI). The 20S rRNA is shown in light gray, r-proteins in dark gray—except of uS3 in yellow. Assembly factors are shown in color: Tsr1 in orange, Rio2 in blue, Pno1 in green, Ltv1 in purple, and Enp1 in red. (Adapted from Scaiola et al. 2018)


3.1.3 Cryogenic Lysis of Yeast Cells (See Also Note 2) (Modified from Rout Lab Protocol: Cryogenic Lyses of yeast Cells [43])


M-IgG Buffer: 20 mM HEPES pH 7.4, KOAc, 40 mM NaCl, 0.5% Triton-X, 0,1% Tween 20.

Supplemented with (see Note 1) 2 mM PMSF, 1 mM DTT, 2–10 mM MgCl2 (optional).


3.1.4 Isolating Preribosomes Using Magnetic Beads (Modified from Oeffinger et al. 2007 [43])

Buffers and Solutions

Method (See Also Notes 5–8)


3.2 Nuclear Import Assays for Ribosomal Proteins

It is assumed that like a typical import cargo, RanGTP dissociates the r-protein from the importin after arriving in the nuclear compartment. However, r-proteins contain disordered regions, which make them prone to non-specific interactions with other nucleic acids, aggregation and degradation in their non-assembled state. During a single 90 min generation time ~14 million r-proteins synthesized in the cytoplasm, need to be targeted to the yeast nucleus and handed over to the ribosome assembly machinery [1, 44]. Thus, safe and efficient transport of r-proteins to their rRNA binding site is a logistical challenge.

A number of groups have uncovered dedicated chaperones that interact with newly synthesized r-proteins in the cytoplasm [44– 55]. The dedicated chaperone–r-protein complexes recruit the import machinery and are transported to the nucleus where they are released by the action of RanGTP.

A different mechanism is employed by the r-protein eS26 to reach the 90S pre-ribosome. Like a typical import cargo protein, eS26 uses importins for targeting to the nucleus. However, after reaching the nuclear compartment, eS26 is removed from the importin by an unloading factor, the escortin Tsr2, without the aid of RanGTP. Tsr2 shields eS26 from proteolysis and enables its safe transfer to the 90S pre-ribosome [56]. Below, we outline a step-by step protocol to investigate the canonical RanGTP dependent disassembly of a Pse1–Slx9 complex (Fig. 3a), and non-canonical Tsr2 dependent disassembly of a Kap123–eS26 complex (Fig. 3b). Slx9 is a predominantly nuclear localized protein, yet it shuttles between the nucleus and the cytoplasm [57, 58]. The import receptor transports Slx9 to the nuclear compartment, where Slx9 facilitates assembly of a Crm1-export complex and efficient export of 40S pre-ribosomes from the nucleus [57–60].

3.2.1 A. RanGTP Mediated Disassembly of a PBS-KMT buffer (pH 7.3): 150 mM NaCl, 25 mM Sodium phosphate, 3 mM KCl, 1 mM MgCl2, 0,1% Tween 20.

Pse1–Slx9 Complex

Buffers and Solutions

	- 2. Meanwhile thaw recombinant GST-Pse1 and GST-alone sample on ice.
	- 3. Centrifuge the GST-Pse1 and GST-alone or lysates for 10 min at 4 C at 16,000 x g to pellet aggregates and store the tubes on ice until use.
	- 4. Add 12 μg GST-Pse1 or GST-alone (per reaction) to washed GSH-Sepharose beads suspended in 500 μL of PBS-KMT buffer.
	- 5. Incubate on a rotating platform for 1 h at 4 C to immobilize GST-Pse1 or GST-alone on GSH-Sepharose beads.

Fig. 3 In vitro binding assays for nuclear import. (a) RanGTP-dependent release of Slx9 from Pse1 importin. Left panel: Workflow overview of the importin binding assay. POI <sup>¼</sup> protein of interest. Right panel: GST–Pse1 complex formation with Slx9 dissociated in the RanGFP-dependent manner. GST–Pse1 (lanes 1–6) or GST alone (lanes 7–8) was immobilized on Glutathione Sepharose, washed with PBS-KMT and incubated with <sup>4</sup> <sup>μ</sup>M Slx9 for 1 h at 4 C (lanes 2–4 and 6). Pse1-Slx9 complex was subsequently incubated with 1.5 <sup>μ</sup>M RanGTP under same conditions (lane 5). After washing with PBS-KMT, bound proteins were eluted in SDS sample buffer, separated by SDS-PAGE and visualized by Coomassie Blue staining and Western analyses using <sup>α</sup>-Slx9 antibodies. L <sup>¼</sup> input. (b) RanGTP-independent release of eS26 from Kap123 importin. Immobilized GST-Kap123 on Glutathione Sepharose was washed and incubated with 4 <sup>μ</sup>M eS26FLAG for 1 h at 4 C. GST-Kap123:eS26FLAG complex was washed and incubated with either buffer alone (lane 2 and 8), 0.375 <sup>μ</sup>M His6-RanQLGTP (left panel lanes 3–6) or 0.375 <sup>μ</sup>M His6-Tsr2 (right panel lanes 9–12). Samples were withdrawn at the indicated time points and washed with PBS-KMT. Bound proteins were eluted in SDS sample buffer, separated by SDS-PAGE and visualized by Coomassie Blue staining and Western analyses using <sup>α</sup>-eS26 and <sup>α</sup>-RanGTP antibodies. L <sup>¼</sup> input. GST-Kap123 is indicated by asterisk. (Adapted from Schu¨tz et al. 2014)


10 min to elute the bound proteins. Separate the eluted denatured proteins on an 12–15% SDS-PAGE and analyze by Coomassie Blue staining.


3.2.2 Tsr2 Mediated Disassembly of a Kap123– eS26FLAG Complex (See Also Notes 9–14)


Nuclear Import It is complicated to monitor the nuclear targeting of r-proteins in vivo since at steady state they reside in the cytoplasm. Therefore, one strategy is to uncouple r-protein nuclear import from their export by selectively impairing recruitment to the pre-ribosome. Fusing GFP to either the N- or C- terminus can prevent r-protein incorporation into the pre-ribosome. These non-functional proxy constructs can be employed to determine nuclear localization signals within r-proteins [56]. Further, by monitoring their mislocalization to the cytoplasm in different importin-deficient (Table 1) and/or mutant strains, these non-functional r-protein fusions (listed in Table 3) can pinpoint their nuclear pathway(s).

3.2.3 Cell Biological Assays for r-protein



#### 3.3 Nuclear Export Assays for Preribosomes

3.3.1 Monitoring Nuclear Export of Pre-ribosomes (See Also Notes 15 and 16)

Transport of 60S and 40S pre-ribosomes from the nucleus into the cytoplasm can be investigated by monitoring the localization of r-protein fusions with GFP (uL23-GFP, uL5-GFP, uL18-GFP for the 60S subunit and uS5-GFP for the 40S subunit) in distinct mutant strains [72–75]. These r-protein fusions are stably incorporated into pre-ribosomes, and their steady-state localization in wild type (WT) cells is cytoplasmic. Nuclear accumulation of these reporters indicates either an early pre-ribosome assembly and/or export defect. Next to the r-protein GFP reporters, nuclear accumulation of GFP fused export adaptor factors serve as reporters for impairment in late assembly steps and/or of export of 60S (Nmd3-GFP) and 40S (Rrp12-GFP and Rio2-GFP) pre-ribosomes, respectively (Fig. 4).

Fig. 4 Monitoring ribosome export defect by fluorescence microscopy. GFP-reporters for large r-subunit (uL18-GFP or Nmd3-GFP) and small r-subunit (uS5-GFP or Rrp12-GFP) were transformed into WT or indicated mutant strains and they localization was analyzed by fluorescence microscopy. Scale bar <sup>¼</sup> 5 <sup>μ</sup>m. Output of the experiment is indicated on the left scheme: accumulation of the reporter in the nucleus in case of export defect as seen for bud20<sup>Δ</sup> and yrb2<sup>Δ</sup>

	- 2. The transformants expressing reporters are grown in appropriate liquid media (for mutants at permissive temperature) until saturation (overnight).
	- 3. The next day, dilute cells into 10 mL fresh media and grown until mid-log phase for at least 2–3 divisions. (In the case of cold or temperature sensitive mutants, shift the cultures to the appropriate restrictive temperature after 2–3 divisions of growth at permissive temperature).
	- 4. Pellet cells by centrifugation in a 15 mL Falcon tube ( 450 g, 3 min) and wash once with 10 mL dH2O.
	- 5. Pour off the supernatant and resuspend the cell pellet in remaining liquid.
	- 6. Place 2 μL of cell suspension on a slide, press a coverslip over before inspection under a fluorescence microscope.


Fig. 5 In vitro binding assays for nuclear export. (A) Ltv1 NES-dependent export complex formation. Left panel: Workflow overview of the binding assay. POI <sup>¼</sup> protein of interest. Right panel: GST-Ltv1 or GST-Ltv1 lacking NES sequence (450-463aa) was immobilized on the Glutathione Sepharose, washed with PBS-KMT and incubated with buffer alone (lanes 1 and 5), buffer with 2μM His6-RanQLGTP (lanes 2 and 6), 50nM Crm1-His6 (lanes 3 and 7) or 50nM Crm1-His6 and 2μM His6-RanQLGTP simultaneously (lane 4 and 8) for 1h at 4C. Bound proteins were washed with PBS-KMT, eluted in 2XLDS sample buffer, separated by SDS-PAGE and visualized by Coomassie Blue staining and Western analyses using <sup>α</sup>-RanGTP and <sup>α</sup>-Crm1 antibodies. L <sup>¼</sup> input. Adapted from Fischer et al., 2015. (B) Heterodimeric transport receptor Mex67-Mtr2 interacts with FG repeat sequences of different nucleoporins. Left panel: Workflow overview of the binding assay. POI <sup>¼</sup> protein of interest. Right panel: Indicated GST-Nucleoporin fusion proteins were immobilized on Glutathione Sepharose and washed with PBS-KMT before 1 h incubation with 3.6 <sup>μ</sup>g His6-Mex67-Mtr2 at 4C. Bound proteins were washed, eluted with 2X LDS buffer and analyzed by SDS-PAGE followed by Coomassie staining or Western blotting using <sup>α</sup>-Mex67/Mtr2 antibody. L <sup>¼</sup> Load; Asterisks indicate Mex67 bands. Adapted from Altvater et al., 2012

Method (See Also Notes 22–28)


	- 1. Wash 80 μL of GSH-Sepharose slurry (per reaction) into a 1.5 mL reaction tube and wash 3 times with 1 mL universal buffer; spin down in between for 30 s at 3000 rpm at 4 C. Wash GSH-Sepharose beads for all reactions together.
		- 2. Meanwhile thaw lysates containing GST-nucleoporins (Nup1, Nup100, Nup116, Nup42) or GST alone on ice.
		- 3. Centrifuge the lysates for 10 min at 4 C at 16,000 g to pellet aggregated proteins and store the tubes on ice until use.
		- 4. Add 1 mL of each lysate to washed GSH-Sepharose beads (combine GSH-Sepharose beads for reactions with same GST-fusion protein) (see Note 22).
		- 5. Incubate tubes on a rotating platform for 1 h at 4 C to immobilize GST and GST fusion protein on GSH-Sepharose beads.
		- 6. Wash samples 3- (850 g, 30 s, 4 C) with 1.5 mL universal buffer and resuspended in 1 mL buffer each.
		- 7. Split beads, by pipetting 500 μL, into two new 1.5 mL eppendorf tubes.

3.4 Genetic Analyses of Nuclear Export Pre-ribosomal particles are among the largest and most abundant cargoes that need to cross the permeability barrier of the NPC. Rapid transit through the NPC is especially important, as any delay within the channel may hinder transport of other cargoes. Therefore, pre-ribosomal particles deploy a set of multiple redundant export receptors for their transport to the cytoplasm [1, 21]. High-copy suppressor screens and synthetic lethal interactions and in budding yeast have uncovered ancillary factors that also mediate efficient pre-ribosome nuclear export. These approaches have proved to be powerful tools to either discover new components of the ribosome export machinery or to directly test functional overlap of a factor of interest with the process of nuclear export.

The essential export adaptor Nmd3 was identified as a high copy suppressor of the bud20Δ mutant slow growth and 60S pre-ribosome export defect [72]. Further, the bud20Δ strain exhibits synthetic lethal or synthetic enhanced growth defects when combined with mutants of established 60S pre-ribosome export factors (nmd3ΔNES1, xpo1-1, arx1Δ, gle2-1, and ecm1Δ), included mex67 and mtr2 mutant alleles (mex67kraa and mtr2-33) that are specifically impaired in 60S subunit [72]. These analyses culminated in the discovery of new FG-repeat binding proteins that functions directly to the process of nuclear export of 60S pre-ribosomes.

Similarly, a high-copy suppressor screen aimed at identifying genes that suppress the impaired growth of the slx9Δ mutant, led to the discovery of the mRNA and 60S export receptor Mex67-Mtr2 as an auxiliary export receptor for in 40S pre-ribosome export [57]. Mex67-Mtr2 mutants that fail to bind to the 40S pre-ribosome were synthetically lethal when combined with the slx9Δ mutant [57]. Further, the slx9Δ mutant displayed a synthetic growth defect when combined with a strain expressing Rrp12-GFP.

#### Table 2 Yeast strains used for ribosomal nuclear export studies


Rrp12 is a 40S pre-ribosome export factor that directly interacts with FG-rich nucleoporins [79]. Finally, Yrb2 that stimulate the assembly of Crm1-export complexes on certain NES-containing cargos by cooperatively binding Crm1 and RanGTP also showed a strong genetic interaction with Slx9 [57]. This genetic link culminated in the identification of a new type of RanGTP binding protein that binds to a specific NES-containing cargo and RanGTP and promotes Crm1 recruitment.

Tables 2 and 3 provide a comprehensive list of mutants and their sources to investigate 60S and 40S preribosome export using standard yeast genetic approaches.

#### 4 Conclusions

Diverse approaches in budding yeast have greatly advanced our understanding of ribosome assembly and transport. Genetic perturbation of energy-consuming enzymes can now be combined with biochemical affinity purifications to obtain high-resolution snapshots of the ribosome assembly pathway. We also anticipate that traditional genetic, biochemical, and cell-biological approaches will provide eagerly awaited integration of the ribosome pathway with other biological pathways including nucleocytoplasmic transport, cellular quality control and signaling.


Table 3 Plasmids for genetic studies of nucleo-cytoplasmic transport

#### 5 Notes


uL18-GFP reporter plasmids (LEU2, URA3, TRP1 and HIS3) can be requested from the Panse laboratory [72]. All the above reporters are expressed under the endogenous promoters. The ribosomal protein uL18 can be C-terminally tagged with GFP at the genomic locus for monitoring the nuclear export of the large pre-ribosomal subunit.


#### Acknowledgments

V.G.P. is supported by grants from the Swiss National Science Foundation, the NCCR RNA & Disease, the Novartis Foundation and the Olga Mayenfisch Stiftung. M.O-O. is a recipient of a doctoral fellowship from the Boehringer Ingelheim Fonds.

#### References


(2016) Insertion of the biogenesis factor Rei1 probes the ribosomal tunnel during 60S maturation. Cell 164:91–102


pre-ribosome is regulated by its eukaryotespecific extension. Mol Cell 58:854–862


60S subunit export machinery. Mol Cell Biol 32:4898–4912


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Tethered MNase Structure Probing as Versatile Technique for Analyzing RNPs Using Tagging Cassettes for Homologous Recombination in Saccharomyces cerevisiae

## Fabian Teubl, Katrin Schwank, Uli Ohmayer, Joachim Griesenbeck, Herbert Tschochner, and Philipp Milkereit

#### Abstract

Micrococcal nuclease (MNase) originating from Staphylococcus aureus is a calcium dependent ribo- and desoxyribonuclease which has endo- and exonucleolytic activity of low sequence preference. MNase is widely used to analyze nucleosome positions in chromatin by probing the enzyme's DNA accessibility in limited digestion reactions. Probing reactions can be performed in a global way by addition of exogenous MNase, or locally by "chromatin endogenous cleavage" (ChEC) reactions using MNase fusion proteins. The latter approach has recently been adopted for the analysis of local RNA environments of MNase fusion proteins which are incorporated in vivo at specific sites of ribonucleoprotein (RNP) complexes. In this case, ex vivo activation of MNase by addition of calcium leads to RNA cleavages in proximity to the tethered anchor protein thus providing information about the folding state of its RNA environment.

Here, we describe a set of plasmids that can be used as template for PCR-based MNase tagging of genes by homologous recombination in S. cerevisiae. The templates enable both N- and C-terminal tagging with MNase in combination with linker regions of different lengths and properties. In addition, an affinity tag is included in the recombination cassettes which can be used for purification of the particle of interest before or after induction of MNase cleavages in the surrounding RNA or DNA. A step-by-step protocol is provided for tagging of a gene of interest, followed by affinity purification of the resulting fusion protein together with associated RNA and subsequent induction of local MNase cleavages.

Key words RNA, RNP, Ribosome, Structure probing, Enzymatic probing, Micrococcal nuclease, Fusion protein, Chromatin endogenous cleavage, ChEC, Saccharomyces cerevisiae

Fabian Teubl and Katrin Schwank contributed equally to this work.

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_8, © The Author(s) 2022

#### 1 Introduction

Numerous methods have been developed over the years to chemically or enzymatically probe the structure of chromatin and of ribonucleoprotein (RNP) complexes, two major manifestations of nucleic acid–protein complexes in eukaryotic cells [1–4]. Depending on the respective agent or enzyme applied, the accessibility, flexibility, secondary structure, or tertiary fold of the respective nucleic acid components can be analyzed either in vitro or in vivo. Combined with a high throughput sequencing readout these analyses can be performed genome- or transcriptome wide, respectively.

One of the enzymes which are routinely used to characterize nucleosome positions and chromatin states is the micrococcal nuclease (MNase) which is secreted by the bacterium Staphylococcus aureus. MNase has endo- and exonucleolytic activity which is strictly dependent on the presence of calcium. It cleaves both DNA and RNA with some preference for single stranded and for A/T- and A/U-rich regions (reviewed in [5–7]. In a typical nucleosome mapping experiment exogenous MNase is added to chromatin to perform a limited digest. Subsequently, positions of DNA cleavages and regions which were less accessible to the enzyme are determined. Information on the exact positioning of nucleosomes on DNA can be deduced if the size of a protected fragment is close to 146 base pairs which are typically protected by a nucleosome core particle [8–10].

Laemmli and colleagues have introduced a variation of this approach which is termed "chromatin endogenous cleavage" (ChEC) [11]. Here, a DNA-binding protein of interest is expressed in S. cerevisiae in fusion with MNase which remains inactive in the cell due to low calcium concentrations. Increasing the calcium concentration after cell breakage activates the enzyme which cleaves then close by accessible DNA. The resulting cuts at specific genomic loci can be analyzed by Southern blotting, or by high throughput sequencing for obtaining a genome wide data set [11– 14]. ChEC and related MNase tethering approaches [15, 16] provide a valuable alternative to techniques as CHIP-Seq or DAM-ID [17, 18] for mapping of DNA binding sites of specific proteins. In addition, occupancies of fusion proteins at selected genomic loci can be analyzed in a quantitative way [13].

Protein components of RNPs have been as well expressed in fusion with MNase in yeast. They are potentially assembled in vivo at their endogenous location in the respective RNPs. The activation of MNase after cell breakage by the addition of calcium leads then to specific RNA cleavages in proximity to the fusion proteins which can be mapped with nucleotide resolution either by local [19] or by transcriptome wide primer extension analyses [20]. Experiments using a ribosomal model substrate showed that the radius of the enzymatic probe can be customized through introduction of differently sized linkers between the RNP-protein and MNase [19]. Cleavages up to around 6 nm surface distance from the anchor protein could be observed with longer linkers while more proximal cuts were strongly favored when using a short linker. Thus, in analogy to the ChEC methodology, probing of the RNA in proximity of a MNase fusion protein can reveal if, and in which region this protein binds to cellular RNPs. Importantly, additional information about the RNP's folding state in the surrounding of the MNase fusion proteins can be obtained. In this regard, the general accessibility of the substrate for the enzyme's active center and the preference of MNase for single stranded and A/U-rich regions seem to be major determinants of spatially restricted cleavage positions [19, 20]. Consequently, a low number of defined cleavages can be expected in RNP's with a compact fold and thereby low accessibility. A/U-rich loops of spatially flexible hairpins which are often hardly detectable in structural analyses by X-ray crystallography or single molecule cryo-electron microscopy, were observed to be preferred substrates of MNase tethered to RNPs [20].

To further facilitate the application of MNase fusion proteins for both chromatin and RNP research we established a set of vectors that can be used as templates for PCR based chromosomal N- or C-terminal MNase tagging of genes by homologous recombination in S. cerevisiae. The tagging cassettes include regions coding for a variety of protein linkers, the MNase itself, a 3x-FLAG affinity tag as well as a heterologous selection marker (see Tables 1, 2 and 3). The cassettes were designed in a modular way allowing for straightforward exchange of each of the individual functional elements. This can be used to replace for example the rather strong RPS28B promoter present in the N-terminal tagging cassettes with other promoters better reflecting the endogenous expression level of the protein to be tagged. Too high expression levels might cause an excess of free MNase fusion proteins not assembled in RNPs or chromatin. This in turn could favor the appearance of nontethered cleavages. The plasmids further code for a set of different linker regions separating the anchor protein from the MNase in the fusion protein. The resulting protein linker regions vary in length from 6 to 63 amino acids and contain elements of different predicted conformational flexibility [22]. Finally, the cassettes contain sequences coding for the strong 3x-FLAG affinity tag which can be used for detection of the fusion proteins by Western blotting. In addition, the tag enables efficient affinity purification of the fusion proteins before MNase activation. This can be useful to facilitate the downstream analyses of the resulting cleavages in RNPs.

#### Table 1 Plasmids containing tagging cassettes


a The total length of the region between the protein of interest and the tethered MNase is indicated. This includes amino acids encoded by the priming sequences needed for PCR of the tagging cassettes. K2373 does not contain the MNasecoding sequence. Hence, no linker length is indicated for K2373

b Hygromycin resistance gene with TEV promoter and CYC1 terminator

#### Table 2 Amino acid sequences of linker modules referred to in Table 1


In the following, we provide instruction for PCR based MNase tagging of a gene of interest in S. cerevisiae using the described plasmid set. We then outline a step-by-step protocol for the analysis of MNase tagged RNPs, including their purification using the built-in 3x-FLAG tag and the subsequent activation of the tethered MNase to induce local RNA cleavages. For the use of generated fusion protein expressing strains in chromatin analyses we refer to recently published protocols [13, 14, 16, 23].


#### Table 3 Primer design for PCR based homologous recombination

a The priming sequences for all vectors except K2515 and K2628 are the same as the ones described in [21] <sup>b</sup> The sizes of the 5<sup>0</sup> regions of the oligonucleotides used for PCR which are homologous to the respective genomic sequence have to be added to these rounded values (see Subheading 3.1)

#### 2 Materials

2.1 Template Plasmids for PCR Based Amplification of Tagging Cassettes



Table 4

Plasmid

construction



Table 4


#### Table 5 Oligonucleotides used for plasmid construction




#### 3 Methods

3.1 Primer Design and Choice of the Template Plasmid for PCR Reactions

Table 1 provides an overview on different plasmids which were created as templates for PCR based amplification of different tagging cassettes. Some of the plasmids are designed for N-terminal and others for C-terminal MNase tagging of a gene of interest in S. cerevisiae. Otherwise, the MNase cassettes encoded by the different plasmids differ by the linkers between MNase and the target protein and by the yeast selection markers. Two oligonucleotides have to be designed after choosing one of the plasmids as template for PCR based amplification of a recombination cassette (see for an overview [25, 27, 28]). These two oligonucleotides, in the following referred to as upstream and downstream oligonucleotide, are designed to contain at their 3<sup>0</sup> -end vector specific priming sequences indicated in the respective columns in Table 3. The priming sequences hybridize with corresponding regions on the plasmid which flank the tagging cassette and thus serve to amplify these cassettes in the PCR reaction. Sequence stretches of the gene of interest are added at the 5<sup>0</sup> -end of the primers. They should target cellular homologous recombination reactions of the amplified cassette to the gene of interest. For N-terminal tagging more than 40 nucleotides upstream of the start codon of the gene of interest are added 5<sup>0</sup> to the upstream priming sequence to generate the upstream oligonucleotide. The start codon of the gene and more than 40 following nucleotides are added in reverse complement orientation to the 50 -end of the downstream priming sequence. Thus, the priming sequence in this downstream oligonucleotide directly follows 3<sup>0</sup> after the reverse complement sequence of the start codon (5´-CAT-30 ). For C-terminal tagging a stretch of more than 40 nucleotides ending with the last codon before the stop codon of the gene of interest is added 5<sup>0</sup> to the upstream priming sequence in the upstream oligonucleotide. For the downstream oligonucleotide, more than 40 nucleotides of sequence just 3<sup>0</sup> of the genes stop codon are added 5<sup>0</sup> of the priming sequence in reverse complement orientation. Thereby the priming sequence continues directly 3<sup>0</sup> after this reverse complement stretch of the genes 3<sup>0</sup> untranslated region.

3.2 Generation of PCR-Based Tagging Cassettes for Homologous Recombination

	- 2. Yeast cells are made chemically competent for transformation as outlined in detail in [25].
	- 3. For homologous recombination, 50 μL of competent yeast cell suspension are transformed with 10 μg of the PCR amplified tagging cassette (see Subheading 3.2). Yeast transformation is performed as described in [25].
	- 4. Cells transformed with recombination cassettes based on plasmids K2515 and K2628 are plated on YPD-Hygromycin plates. For all other recombination cassettes, cells are plated on SC-URA plates. Before spreading on YPD-Hygromycin

3.3 Transformation of Competent Yeast Cells and Screening for Positive Clones

plates, the transformed cells should regenerate for at least 6 h in 10 mL of liquid YPD medium at 30 C with shaking. Plates are incubated upside down at 30 C until appearance of colonies (2–5 days depending on the strain background and plates used).


3.4 Yeast Cell Culture and Preparation of Cellular Extracts

	- 1. The cellular extract prepared in 3.4 is supplemented with 0.1% (w/v) Tween and 0.5% (w/v) Triton X-100 (see Note 7).
	- 2. Prior to affinity purification, the Anti-Flag M2 affinity matrix is equilibrated with buffer AG100++. For preribosomal particles we usually use 200 μL of matrix suspension containing about 100 μL matrix for cellular extracts prepared from 1 L of yeast cell culture. The suspension is transferred to Poly-Prep chromatography columns, washed once with 10 mL water and four times with 3 mL of buffer AG100++.
	- 3. The equilibrated affinity matrix is transferred to the tube containing the cellular extract (step 1) and incubated for 1 h at 4 C on a turning wheel.
	- 4. The suspension is transferred into a Poly-Prep chromatography column and washed twice with 2 mL of buffer AG100++ and once with 10 mL AG100++.

3.5 Purification of RNPs from Cellular Extracts and Activation of MNase


#### 4 Notes


cleaved as determined by RNA extraction and northern blotting. Excessive cleavage conditions should be avoided since they may favor disintegration of the analyzed RNP and accumulation of nontethered cleavages.

#### Acknowledgments

We thank all members of the chair of Biochemistry III for their support of this work. This work was financially supported by the grant SFB 960 from the "Deutsche Forschungsgemeinschaft" (DFG).

#### References


Epigenetics Chromatin 7:33. https://doi. org/10.1186/1756-8935-7-33


Genet 10:669–680. https://doi.org/10. 1038/nrg2641


transcribed rRNA genes in S. cerevisiae are organized in a specialized chromatin associated with the high-mobility group protein Hmo1 and are largely devoid of histone molecules. Genes Dev 22:1190–1204. https://doi.org/ 10.1101/gad.466908


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part V

RNA Modification

# Chapter 9

# Chemical Modifications of Ribosomal RNA

#### Sunny Sharma and Karl-Dieter Entian

#### Abstract

Cellular RNAs in all three kingdoms of life are modified with diverse chemical modifications. These chemical modifications expand the topological repertoire of RNAs, and fine-tune their functions. Ribosomal RNA in yeast contains more than 100 chemically modified residues in the functionally crucial and evolutionary conserved regions. The chemical modifications in the rRNA are of three types—methylation of the ribose sugars at the C2-positionAbstract (Nm), isomerization of uridines to pseudouridines (Ψ), and base modifications such as (methylation (mN), acetylation (acN), and aminocarboxypropylation (acpN)). The modifications profile of the yeast rRNA has been recently completed, providing an excellent platform to analyze the function of these modifications in RNA metabolism and in cellular physiology. Remarkably, majority of the rRNA modifications and the enzymatic machineries discovered in yeast are highly conserved in eukaryotes including humans. Mutations in factors involved in rRNA modification are linked to several rare severe human diseases (e.g., X-linked Dyskeratosis congenita, the Bowen–Conradi syndrome and the William–Beuren disease). In this chapter, we summarize all rRNA modifications and the corresponding enzymatic machineries of the budding yeast.

Key words rRNA modification, Ribose methylation, Pseudouridylation, Base methylation, Aminocarboxypropylation, Acetylation of cytidines, Methyltransferase

#### 1 Introduction

RNA modifications are present in all three kingdoms of life and detected in all classes of cellular RNAs. RNA modifications are diverse, with more than 100 types of chemical modifications identified to date [1]. Ribosomes are molecular assemblies of RNA and proteins and are responsible for the synthesis of all proteins in the cells [2]. Structural and functional analyses of ribosomes have revealed that it is the ribosomal RNA (rRNA) that makes the structural framework of ribosomes and catalyzes the joining of amino acids (peptidyl transfer) during translation, hence making the ribosome a ribozyme [3]. Though the chemical composition of RNA seems to be rather insufficient to provide the structural complexity to RNA, the composition analysis of rRNA has shown that rRNA contains different chemical modifications that are added

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_9, © The Author(s) 2022

both co- and posttranscriptionally [4, 5]. Ribosomal RNA (rRNA) contains three types of chemical modifications, methylation of the ribose sugars at the C2-position (Nm), isomerization of uridines to pseudouridines (Ψ), and base modifications (methylation (mN) such as acetylation (acN) and aminocarboxypropylation (acpN)) [6]. Mutations in factors involved in rRNA modifications are associated with several rare severe human diseases (e.g., X-linked dyskeratosis congenita, the Bowen–Conradi syndrome, Hutchison Gilford syndrome, and the William–Beuren disease) [7– 13]. Emerging evidences indicate that some bases are not always completely modified providing heterogeneity with respect to RNA modification [14–16]. Heterogeneity in rRNA modification has been correlated with disease etiology (cancer) and shown to play a role in cell differentiation (hematopoiesis) [17]. Remarkably, alterations in rRNA modification patterns profoundly affect the preference of ribosomes for cap- versus IRES-dependent translation having major consequences on cell physiology [18, 19]. Here, we summarize all known rRNA modifications of the budding yeast with an emphasis on base modifications (see Tables 1, 2, 3 and 4).

In Saccharomyces cerevisiae, 18S rRNA of the small subunit contains 14 Ψs, 18 Nms (2<sup>0</sup> O-methylated Ns), 4 mNs (methylated Ns), and 2 acNs (acylated Ns) (Tables 1 and 2), whereas 25S rRNA of the large subunit contains 30 Ψs, 35 Nms, and 6 base methylations (Tables 3 and 4). Mapping of these modifications has revealed that these modifications cluster in the functionally conserved regions of the ribosomes like the intersubunit and the peptidyl transferase center [6, 20, 21]. Due to technical limitations, the chemical modification profile of rRNA, especially for the base modifications, remained poorly characterized for a long time. Using state-of-the-art RP-HPLC and mass spectrometry together with "reverse genetics," we and others have recently completed the characterization and mapping of the complete set of yeast rRNA modifications, and have identified the corresponding enzymatic machinery involved in adding these modifications to the RNA [16, 22, 23].

#### 2 Ribose Methylation

Methylation of 20 -OH of ribose sugar to a 20 -O-methyl-ribose is a characteristic modification in mRNA and many noncoding RNA (ncRNA) including tRNA, rRNA and siRNAs (Fig. 1). Ribose methylation favors a 30 -endo conformation of the ribose and since 30 -endo conformations are known to stabilize A-form helices, methylation of ribose increases the rigidity of the RNA by promoting base stacking. Furthermore, ribose methylation provides RNA stability against base and nuclease hydrolysis by insulating the otherwise active 20 -OH group. Our recent analyses together with other


#### Table 1 18S rRNA modifications catalyzed by snoRNPs

#### Table 2

#### 18S rRNA modifications catalyzed by specific enzymes


#### Table 3 25S and 5.8S rRNA modifications catalyzed by snoRNPs



Table 4 25S and 5S rRNA modifications catalyzed by enzymes

groups have identified several partial rRNA modifications in eukaryotes including humans. These observations suggest the existence of a heterogeneous population of ribosomes supporting the "specialized" translation model, which could possibly play an important role during embryonic development [14–16, 22, 24, 25].

2.1 C/D Box snoRNPs All 2<sup>0</sup> -OH ribose methylation occur at distinct positions in eukaryotic rRNAs (Tables 1 and 3). The respective positions are targeted via specific C/D box snoRNAs having complementary guide sequences to the respective rRNAs together with distinguishing sequence elements called boxes C and D.

The methylation guide sequence is located upstream of the box D/D<sup>0</sup> element and consists of 10–21 nucleotides. The guide sequences direct ribose methylation to the nucleotide base paired to the fifth nucleotide (nt) upstream of the box D or D<sup>0</sup> sequence (box D + 5 rule) [26]. The C/D box snoRNPs consist of four common core proteins: Fibrillarin (human)/Nop1 (yeast), Nop58, Nop56, and Snu13 [27]. Interestingly, C/D box snoRNPs involved in functions apart from catalysis like U3, U14, U8, U22, snR4, and snR45 also contain additional proteins [27, 28]. Nop1 is a S-adenosyl methionine (SAM) dependent methyltransferase and catalyzes the 20 -O methylation reaction. Snu13 binds to the kinkturn (loop-stem structure that includes the canonical C/D elements in the loop portion) in the C/D box of snoRNAs. Nop56 and Nop58 are characterized by extensive coiled-coil domains, likely responsible for heterodimerization and providing stability to the snoRNA.

#### 3 Pseudouridylation

Pseudouridylation (ψ) is a C-glycoside rotation isomer of uridine (Fig. 1). Due to rotation, the nitrogen atom at position 1 (N1) forms no longer a glycosidic bond to the ribose and is protonated at physiological pH. Compared to uracil, in pseudouracil both N1 and N3 participate in hydrogen bonding. The N1 proton makes hydrogen bonding with a phosphate group from the same or neighboring nucleotide and provides stability of the structure.

Furthermore, like ribose methylation, pseudouridine also favors a C3-endo sugar pucker (the conformation preferred by an A-form RNA helix) and has been shown to increase the thermal stability of RNA by up to 2 -C [29]. Therefore, the presence of Ψ also plays an important role in RNA stability that may not be essential, but seemingly provides a significant advantage [5].

Fig. 1 Chemical structure of modified bases and 2<sup>0</sup> O-methylation in yeast rRNAs

3.1 H/ACA snoRNPs The majority of pseudoridines in rRNAs are added via specific H/ACA box snoRNPs having complementary guide sequences to the respective rRNAs and contain small sequence elements referred to as boxes H and ACA [30]. Similar to C/D box snoRNAs, substrate targeting involves base pairing through two short guide sequences in a loop portion of the duplex structures (Tables 1 and 2). The guide sequences are 14–15 nucleotides from the H or ACA box. The binding of the substrate places the target uridine in a pseudouridylation pocket between the flanking paired regions [30].

> H/ACA snoRNPs contain four core proteins in yeast: Cbf5p (Dyskerin in humans), Gar1, Nhp2, and Nop10. Cbf5 is the catalytic pseudouridine synthase. Although the crystal structure from the eukaryotic H/ACA snoRNP is still missing, the archaeal H/ACA structure has provided noteworthy details about the organization of various core proteins [31]. Cbf5 has been shown to contact the ACA motif and both P1 and P2 stem. Nhp2, the yeast homolog of archaeal L7Ae binds to the K loop structure in the upper half of the RNA. Gar1 does not bind to the RNA directly but rather joins the complex through its interaction with Cbf5. Further structural analyses of Cbf5 have revealed that its interaction with Gar1 is essential for substrate binding and release [27].

#### 4 Base Modifications

The rRNAs contain three different types of base modifications methylation (m), acetylation (ac), and aminocarboxypropylation (acp). Methylation is the most common base modification in rRNA. All four nitrogenous bases of RNA undergo methylation either at nitrogen (N) or carbon (C) atoms. On the other hand, only cytosine bases are acetylated, and aminocarboxypropyl is added either to a uridine or pseudouridine residues in yeast. The base modifications of RNA are primarily catalyzed by snoRNAindependent enzymes with only exception being 18S rRNA acetylation that requires particular C/D box snoRNAs (Tables 2 and 4).

4.1 Base Methylation Methylation of nitrogenous base strongly affect their physical and chemical properties. Methylation promotes base stacking by increasing the hydrophobicity and the polarizability. Furthermore, methylation also influences the structure by increasing steric hindrance, blocking canonical (Watson–Crick) hydrogen bonding and fostering noncanonical Hoogsteen base paring. This presumably helps ncRNA like rRNA to attain and maintain specific conformations, essential for their corresponding function—both with respect to their structure and their enzymatic activity.

#### 4.2 Methyl transferases

Methyltransferases are the enzymes that catalyze specific transfer of methyl group form a methyl donor to various substrates. S-adenosyl-methionine (SAM or AdoMet) is the most common methyl donor by virtue of the presence of a charged methylsulfonium center [32]. Methyltransferases that utilize the methyl group of SAM for the methylation reaction are called SAM dependent methyltransferases. Based on their structural analysis, all currently known SAM dependent methyltransferases have been divided into five distinct classes (Class I to Class V) [33].

Class I methyltransferases are characterized by a Rossmann-fold like domain and methylate a wide variety of substrates (DNA, RNA and proteins along with other small molecules). Rossmann folds are nucleotide (especially NAD(P)) binding domains that also contain an alternating α/β strands topology in which two Rossmann fold domains are linked into 6 parallel β stands sandwiched by a pair of α-helices [32]. The Rossmann-like fold comprises of alternating β-stranded and α-helical regions, with all strands forming a central relatively planar β-sheet, and helices stuffing two layers, one on each side of the plane.

Class II methyltransferases are characterized by a methionine synthase activation domain. The Met synthase activation contains an unusual fold with long, central, antiparallel β-sheet flanked by groups of helices at either end, which makes it structurally distinct from Class I methyltransferases [32].

Class III methyltransferases act on ring carbons of the large, planar precorrin substrates during cobalamin biosynthesis (e.g., CbiF) [32].

Class IV methyltransferases belong to the SPOUT family and methylate either RNA or proteins [32, 34]. These enzymes contain a six- stranded parallel β sheet flanked by seven α-helices. Interestingly the first three strands of these methyltransferases form half of a Rossmann fold. The active site of SPOUT methyltransferases is located near the subunit interface of a homodimer.

Class V methyltransferases contain typical SET domains, discovered originally as conserved domain shared by chromatin remodeling proteins Su (var) 3–9, E (Z) (short for Enhancer of Zeste) and Trithorax [32]. Most of the currently known SET methyltransferases methylate lysine residues of various nuclear proteins involved in chromatin remodeling and transcriptional regulation. The SAM binding site and the catalytic center of SET domains contains all-β (eight curved β strands forming three small sheets) and knot-like structures like Class IV methyltransferases but based on a different topology [32].

4.3 N4- Acetylation of Cytidine (ac<sup>4</sup> C) Acetylation of cytidine residues is a highly conserved base modification present in 18S rRNA, as well as leucine and serine tRNAs of yeast [35, 36]. Molecular dynamics simulation and in vitro studies using the noninitiator methionine-accepting tRNA of E. coli have reported that acetylation of cytidine residues stabilize the C30 -endo puckering conformation of ribose, and stabilizes the G–C base pairing in the RNA, which has been highlighted as an important function of this modification in counteracting mistakes that may occur during translation due to misreading of the isoleucine AUA codon by tRNAMet [37–39]. Interestingly, in the 18S rRNA both ac4 C residues [ac4 C 1280 (helix 34) and ac4 C1773 (helix 45)] are also involved in C-G base pairings that are fundamental for ribosome functionality [6, 40]. The disruption of these base pairing was found to be lethal for yeast cells (unpublished data).

Structural and functional analysis of bacterial RNA acetyltransferase TmcA has revealed that RNA acetyltransferases utilize acetyl-CoA as an acetyl group donor, which is transferred in an ATP-dependent manner to the cytosine residues [41]. RNA acetyltransferases contain an N- terminal RNA helicase domain similar to that of DEAD-box RNA helicases paired with a C- terminal Gcn5-related N-acetyltransferase (GNAT) fold. TmcA is a standalone enzyme and does not necessitate auxiliary factors for substrate specificity, which is likely due to similar substrates that it acetylates [42].

In contrast to TmcA, the yeast and higher eukaryotes TmcA homologs Kre33/NAT10 utilize protein factors Tan1 and THUMPD1, respectively, for tRNA acetylation and snoRNAs snR4 and snR45 for 18S rRNA acetylation [28, 43, 44].

4.4 3-Amino carboxy propylation acp The addition of 3-aminocarboxypropyl (acp) to the RNA is another highly conserved modification in eukaryotic cellular RNAs derived from S-adenosyl-L-methionine (SAM) [45]. This modification is mostly added to the uridine and pseudouridine residues and impacts the ability of these bases to establish hydrogen bonding with otherwise complementary adenosine residue [46, 47].

RNA aminocarboxypropyl transferase is a novel class of SAM dependent acp transferase with a significant homology to the SPOUT-class RNA-methyltransferases [46, 47]. Structure analysis of its archaeal Tsr3 homologs has revealed that these enzymes contains the same fold as in SPOUT- class-RNA methyltransferase [34], but due to a discrete SAM binding arrangement acp transferases transfer the acp instead of the methyl group of SAM to its substrate [47].

#### 5 Base Modifications of 18S rRNA

The 18S rRNA of ribosome small subunit (SSU) contains four base methylation: one m1 acp3Ψ1191, two m<sup>6</sup> 2A1781, m6 2A1782, and a single m7 G1575, and two base acetylation—ac<sup>4</sup> C 1280 and 1773 (Fig. 1 and Table 3).


already observed in case of Bud23 and Dim1, the methylation appeared not to be the essential function of the Nep1. Instead, the BSC-mutated yeast Nep1 protein showed enhanced dimerization propensity and increased affinity for its RNA-target in vitro. Furthermore, the BCS mutation prevented nucleolar accumulation of Nep1, which could be the reason for reduced growth in yeast and the Bowen-Conradi syndrome [8]. Furthermore, loss of methylation leads to defects in 40S subunits and affects antibiotic sensitivity [8, 56, 57]. Genetic analysis of Nep1 has provided understanding of its possible essential function [56, 58]. Overexpression of Rps19 protein has been demonstrated to suppress the deletion of otherwise essential Nep1 [58]. This has led to the hypothesis that perhaps the essential function of Nep1 is to assist the incorporation of Rps19 protein during 40S biogenesis. Nevertheless, this model awaits the biochemical experiments in its support [59].

Whereas Ψ (snR35) and N1-CH3 (Nep1) are introduced in the nucleus, it was shown that the addition of 3-amino-3-carboxypropyl to the Ψ1191 takes place in the cytoplasm [53]. The enzyme responsible for the transfer of 3-amino-3-carboxypropyl (acp) to the N3 atom of Ψ1191 was recently identified as Tsr3 [47]. A homologous enzyme was also identified in E. coli that catalyzes acp3 U modification at position 47 in the variable loop of eight E. coli tRNAs [46].

Eukaryotes contain only one cytidine acetyltransferase Kre33 (yeast) and NAT10 (human) that catalyzes both rRNA and tRNA cytidine acetylation [28, 43, 44]. Kre33 contains a N terminal helicase domain fused to the C terminal acetyltransferase domain related to GCN5 [42, 44]. In S. cerevisiae Tan1 was initially identified as a protein that is required for acetylation of ac4 C formation at position 12 of tRNA-Ser (CGA) [36]. Nevertheless, Tan1 does not contain any catalytic (acetyltransferase) domain for ac<sup>4</sup> C formation but rather contains the THUMP domain, responsible for tRNA binding.

Interestingly, the acetylation of two cytosine residues in 18S rRNA catalyzed by Kre33 are guided by the two box C/D snoR-NAs snR4 and snR45 which are not known to be involved in methylation [28]. This is in contrast to tRNA acetylation where protein Tan1 likely guides Kre33 to the acetylated cytidine [28, 43, 44]. These results highlighted yet another example of an incredible cellular modularity, where a single enzyme is targeted to diverse substrates by means of either protein or RNA adaptors. Both snR4 and snR45 establish extended bipartite complementarity around the cytosines targeted for acetylation [28].

The viability of catalytically deficient Kre33 mutants demonstrated that acetylation of 18S rRNA is dispensable for cell viability. Although both catalytic deficient mutants, kre33-H545A and kre33-R637A exhibited severe growth defects at 37 -C, similar to

5.4 Kre33/NAT10 Catalyzes ac<sup>4</sup> C1280 and ac<sup>4</sup> C1773 Acetylation of the 18S rRNA in a snoRNA Dependent Manner

what was observed previously for the Tan1 deletion mutant [36, 44]. Since snoRNA deletions did not exhibit this phenotype at 37 -C, we concluded that this is probably due to the loss of tRNA acetylation.

#### 6 Base Modifications of 25S rRNA

The 25S rRNA of the ribosome large subunit (LSU) in yeast contains 6 base methylation: two m1 A (1-methyl adenosine), two m<sup>5</sup> C (5-methyl cytosine), and two m<sup>3</sup> U (3-methyl uridine) (Fig. 1 and Table 4). Two m<sup>5</sup> U (5-methyl uridine) methylations were also reported previously but the presence of these in the 25S rRNA have been disproven.

6.1 Rrp8 (Bmt1) Catalyzes m<sup>1</sup> A 645 Methylation Rrp8, a protein previously shown to be involved in the processing of 35S rRNA at site A2 [60]. Rrp8 catalyzes m1 A645 base methylation of 25S rRNA and is a Class I SAM dependent rRNA methyltransferase [61]. The mapping of m1 A645 on the ribosomal RNA revealed that this residue is present in helix 25.1 of domain II of the 25S rRNA. Helix 25.1 is highly conserved in eukaryotes including humans [62]. The in vivo structural probing of 25S rRNA, using both DMS and SHAPE, revealed that the loss of the Rrp8-catalyzed m<sup>1</sup> A modification alters the conformation of domain I of yeast 25S rRNA causing translation initiation defects detectable as halfmer formation, likely because of incompetent loading of 60S on the 43S-preinitiation complex [62]. Surprisingly, quantitative proteomic analysis of the yeast Δrrp8 mutant strain using 2D-DIGE have exhibited that loss of m1 A645 impacts production of a specific set of proteins involved in carbohydrate metabolism, translation, and ribosome synthesis [62]. These findings in yeast point to a role of Rrp8 in primary metabolism. In conclusion, the m1 A modification is crucial for maintaining an optimal 60S conformation, which in turn is important for regulating the synthesis of key metabolic enzymes. 6.2 Bmt2 Catalyzes Bmt2 catalyzes the m1 A2142 methylation located in helix

#### A2142 Methylation (m<sup>1</sup> A2142)

65 (H65) of the 25S rRNA [63]. H65 belongs to domain IV of 25S rRNA, which makes most of the intersubunit surface of the large subunit. Interestingly, L2, a highly conserved protein makes physical contact with H65, especially with its SH3-β barrel globular domain. Like Rrp8, Bmt2 also belongs to Class I Rossmann-like fold methyltransferases family [63].

A deletion mutant of BMT2 was previously identified to extend hibernating life span and to exhibit peroxide sensitivity [64]. A methyltransferase dead mutant (bmt2-G180R) and rDNA point mutants (A2142G, A2142C, and A2142T) also exhibited hydrogen peroxide sensitivity. Taken together this indicates that the biosynthetic pathways of this modifications interact with the cellular response toward oxidative stress stimulated by hydrogen peroxide.

#### 6.3 Rcm1 (Bmt3) and Nop2 (Bmt4) Catalyze m5 C2278 and m<sup>5</sup> C2870 Methylation

Yeast 25S rRNA contains two m<sup>5</sup> C residues at positions 2278 and 2870 [65]. Rcm1 and Nop2 were shown to be responsible for two distinct m5 C methylations of 25S rRNA at position C2278 and C2870, respectively [66]. Both Rcm1 and Nop2 belong to Class I methyltransferases characterized by Rossmann-like folds. Interestingly, biochemical and structural analyses of other RNA cytosine methyltransferases have provided an important insight into their catalytic mechanism [67]. All known RNA cytosine methyltransferases utilize two highly conserved cysteine residues in the motif IV and motif VI for the addition of methyl group at C5 of cytosine [65, 67]. The cysteine in motif VI makes a nucleophilic attack on to the C-6 of cytosine and form a covalent adduct with the RNA, whereas the cysteine of motif IV is important for the release of the substrate [67]. For both Rcm1 and Nop2 exchange of cysteine in motif VI with the alanine led to methyltransferase-dead mutants, whereas replacement of cysteine in motif IV with alanine turned out to be lethal [66].

The lethality of both rcm1 and nop2 motif IV cysteine mutant proteins suggested that due to substitution of cysteines in motif IV, the mutant proteins fail to separate themselves from their corresponding targets and forms a stable protein-rRNA complex. This fixing of the mutant protein then blocks the 25S rRNA processing and probably causes cell death. This model was further supported by suppression of this lethality upon exchange of both motif IV and motif VI cysteine residues together in case of Rcm1 and also Nop2, as this precluded the formation of any covalent complex [66].

Nop2 is an essential protein in S. cerevisiae [68]. Previous biochemical analyses using the hypomorphic expression system have shown that Nop2 depletion causes severe defects in the rRNA processing and 60S biogenesis. The viability of the nop2- C478A methyltransferase-deadmutant demonstrated, that methylation at C2870 is not the essential function of Nop2. In other words m5 C2870 is not essential for viability. Nevertheless, the polysome profiles and Northern-blotting analysis of the methyltransferase-dead mutant of Nop2 demonstrated that loss of m<sup>5</sup> C2870 strongly impairs the 35S rRNA processing and 60S biogenesis; the phenotypes observed previously upon depletion of Nop2 [66]. Therefore, this suggested that the Nop2 depletion phenotypes observed previously are apparently the result of reduced m5 C modification.

Absence of m5 C2278 causes anisomycin hypersensitivity [66]. In contrast to m<sup>5</sup> C2870, loss of m<sup>5</sup> C 2278 did not exhibit any defects in 60S biogenesis and 35S rRNA processing. As far as the biological role of m<sup>5</sup> C2278 is concerned, loss of m5 C2278 in helix 70 (H70) causes anisomycin hypersensitivity, indicating that the loss of this methylation disrupts its optimal conformation. Together with the data from the m<sup>1</sup> A2142 base modification in helix 65, the anisomycin hypersensitivity after the loss of m5 C 2278 also suggests that the conformation of domain IV of 25S rRNA is crucial for anisomycin sensitivity as both m<sup>1</sup> A2142 and m<sup>5</sup> C 2278 reside in the domain IV of the 25S rRNA [66].

To understand how 25S/28S rRNA cytosine methylation (m<sup>5</sup> C) and its corresponding enzyme (Rcm1/NSUN5) impact biological functions, in a collaboration with Johannes Grillari's lab, we found, that reduced levels of the enzyme increase the life span and stress resistance in yeast, worms, and flies [69]. Loss of m<sup>5</sup> C alters the structural conformation of the ribosome and promotes translational reprogramming by favoring translation of a distinct subset of oxidative stress-responsive mRNAs [69]. Thus, rather than merely being a static molecular machine executing translation, the ribosome exhibits functional diversity by modification of just a single rRNA nucleotide, resulting in an alteration of organismal physiological behavior, and linking rRNA-mediated translational regulation to the modulation of life span, and differential stress response.

#### 6.4 Bmt5 and Bmt6 Catalyze m3 U2634 and m3 U2843 Methylation

Bmt5 and Bmt6 catalyze the m<sup>3</sup> U methylations at positions 2634 and 2843 of the 25S rRNA [70]. Both enzymes belong to the Rossmann-like fold protein family. The substitution of highly conserved glycine residues in the SAM binding motif of both Bmt5 and Bmt6 with arginine abolished their catalytic activity. Surprisingly, m<sup>3</sup> U methylation in the 16S rRNA of E. coli is catalyzed by proteins of the SPOUT methyltransferase family which had led to the assumption that m3 U methylation could only be performed by SPOUT methyltransferases [71]. With the identification of Bmt5 and Bmt6 this convention is no more valid.

Both Bmt5 and Bmt6 are highly conserved in lower eukaryotes including yeasts like Schizosaccharomyces pombe, Neurospora crassa, and Kluyveromyces lactis. The Bmt5 and Bmt6 homologs are likely performing the same function in these organisms.

6.5 25S rRNA of Yeast Does Not Contain any m<sup>5</sup> U Residues Yeast 25S rRNA was predicted to contain two m5 U residues at position 956 and 2924 [20, 72]. Using both RP-HPLC and mass spectrometry this was disproved [70]. Similarly, the 25S rRNA of C. albicans and S. pombe also did not contain any m<sup>5</sup> U residues [73]. This has been also confirmed for higher eukaryotes, whereas m<sup>5</sup> U residues have been only observed in tRNAs, which is in consent with the presence of only one m5 U methyltransferase, Trm2 (in yeast and its homologs), responsible for m<sup>5</sup> U modification of tRNAs at position 54. Up till now, Trm2 has not been observed to display any interaction with the rRNA in yeast. Taken together, this would apparently suggest that during the course of evolution m<sup>5</sup> U methylation have disappeared in rRNA of eukaryotes.

#### Acknowledgments

We thank Jun Yang, Peter Ko¨tter, and Britta Meyer for fruitful discussions and suggestions. We would also like to thank Deutsche Forschungsgemeinschaft (DFG), Goethe-University, Frankfurt/ M., and the state of Hesse for financial support.

#### References


specific methyltransferase. Nucleic Acids Res 38:2387–2398


snoRNA snR9. Nucleic Acids Res 44: 8951–8961


(Topt, 95 degrees C). J Bacteriol 185: 5483–5490


helicase module in an acetyltransferase that modifies a specific tRNA anticodon. EMBO J 28:1362–1373


subunit rRNA impacts locally its structure and the translation of key metabolic enzymes. Sci Rep 8:11904


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# Chapter 10

# In Vitro Selection of Deoxyribozymes for the Detection of RNA Modifications

## Anam Liaqat, Maksim V. Sednev, and Claudia Ho¨ bartner

#### Abstract

Deoxyribozymes are artificially evolved DNA molecules with catalytic abilities. RNA-cleaving deoxyribozymes have been recognized as an efficient tool for detection of modifications in target RNAs and provide an alternative to traditional and modern methods for detection of ribose or nucleobase methylation. However, there are only few examples of DNA enzymes that specifically reveal the presence of a certain type of modification, including N<sup>6</sup> -methyladenosine, and the knowledge about how DNA enzymes recognize modified RNAs is still extremely limited. Therefore, DNA enzymes cannot be easily engineered for the analysis of desired RNA modifications, but are instead identified by in vitro selection from random DNA libraries using synthetic modified RNA substrates. This protocol describes a general in vitro selection stagtegy to evolve new RNA-cleaving DNA enzymes that can efficiently differentiate modified RNA substrates from their unmodified counterpart.

Key words RNA, Deoxyribozymes, Modified RNA nucleotides, Catalytic DNA, Epitranscriptomics, In vitro selection, RNA cleavage

#### 1 Introduction

Cellular RNAs can be modified posttranscriptionally through chemical modifications on nucleobases or the ribose-phosphate backbone. The flourishing field of "epitranscriptomics" explores the modifications that are functionally relevant to RNA structure, stability, base pairing, and binding potential to proteins and other ligands [1, 2]. Besides the reversible chemical modifications of DNA and proteins, posttranscriptional RNA modifications provide another layer of regulation for gene expression [3]. Although modified nucleotides are found in many different types of coding and noncoding transcripts and the precise roles are far from being completely understood, several tRNA and rRNA modifications are expected to play specific structural and functional roles [4]. For example, since ribosomal rRNA modifications are installed early in rRNA processing, it is possible that they assist RNA folding

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_10, © The Author(s) 2022

or are involved in the recruitment of chaperone proteins [5]. Interestingly, many ribosomal RNA modifications are evolutionary conserved but individual modifications are often not essential, while a global lack of rRNA modifications diminishes viability. Recently it was found that some rRNA modifications are installed substoichiometrically, resulting in different subpopulations of ribosomes with altered fitness or fine-tuned activity for translation of specific mRNA [6, 7].

Although more than 120 different modifications in various types of RNAs are known for decades, the field is still lagging on technologies for transcriptome-wide mapping for most of these modifications. Several techniques such as two-dimensional thin layer chromatography (2D-TLC), high-resolution liquid chromatography coupled to mass spectrometry (HPLC-MS), methylated RNA immunoprecipitation, and reverse transcriptase–based signatures followed by deep sequencing have been used to map modifications in RNA [8–12]. These techniques, however, suffer from various limitations such as the loss of sequence information upon digestion into mononucleotides as required for TLC and MS approaches, and from low specificity and selectivity of antibodies used for immunoprecipitation. Therefore a combination of different methods is necessary to obtain reliable insights into presence, distribution and abundance of certain modified nucleotides, and simple analytical tools are needed for validation [13].

Deoxyribozymes (alternatively called DNA enzymes) are attractive tools to expand the repertoire of methods for detecting RNA modifications in a sequence-specific manner. Deoxyribozymes are in vitro selected DNA molecules that have the potential to catalyze various chemical reactions, including protein modifications, DNA/RNA ligation, and DNA/RNA cleavage [14– 16]. Among these, DNA enzymes that catalyze a site-specific RNA cleavage reaction are the most prominent group [17, 18].

Deoxyribozymes have a catalytic core that originates from a random region of 20–40 nucleotides and that is flanked by two binding arms complementary to the RNA substrate. DNA enzymes require metal ions (Mg2+, Mn2+, Zn2+) for their catalytic activity. RNA-cleaving deoxyribozymes catalyze the cleavage reaction by mediating the attack of the 20 -hydroxyl group onto the adjacent phosphodiester linkage, which results in the formation of 20 ,3- 0 -cyclic phosphate and 50 -hydroxyl termini [18, 19]. This reaction gives the possibility to detect modifications that directly block the functional group in the cleavage reaction, such as 20 -O-methylation. In fact, this resulted in the first application of DNA enzymes to analyze ribosomal RNA modifications [20]. Alternatively, deoxyribozymes have been used to detect RNA modifications on the 50 -terminus of the cleavage site. DNA enzymes 8–17 and 10–23 [21] oriented to cleave various dinucleotide junctions served as tool for cleavage of phosphodiester linkage followed by radioactive labeling and 2D TLC for site-specific analysis of pseudouridine [22], one of the most important ribonucleotide modifications in rRNA. Recently, deoxyribozymes have been explored to sense other modifications such as N<sup>6</sup> -methyladenosine (m<sup>6</sup> A) and N<sup>6</sup> isopentenyladenosine (i6 A) [23, 24]. The m6 A-sensitive DNA enzymes have been shown to be generally applicable to analyze the presence of m<sup>6</sup> A in DGACH sequence motifs, as demonstrated for lncRNAs and a set of C/D box snoRNAs that function as guides for 20 -O ribose methylation of ribosomal RNA. These recent examples demonstrated that catalytic DNA can likely be developed for various modifications as a tool to differentiate modified from unmodified RNA in a sequence-specific manner.

Deoxyribozymes are identified through in vitro selection based on the systematic evolution of ligands by exponential enrichment (SELEX) technique from a random pool of DNA through repetitive cycles of selection and amplification as shown in Fig. 1.

This chapter gives a detailed protocol for the gel-based SELEX technique to evolve catalytically active DNA species from a random pool that is capable of specifically detecting RNA modification, thus leading to an enhanced/reduced cleavage of the target RNA depending on the modification state. The in vitro selection cycle begins with the ligation of the DNA pool to the RNA substrate containing the desired RNA modification by T4 DNA ligase using a

Fig. 1 Overview of the in vitro selection scheme for the generation of modification-sensitive RNA cleaving deoxyribozymes. The red dot represents the modified nucleotide in the RNA substrate (green), in between Watson–Crick-base-paired regions. The constant regions of the DNA are shown in light blue, and the random region of the DNA library in dark blue. The yellow star represents a 5<sup>0</sup> -label on the DNA facilitating detection on PAGE

DNA splint. The ligated product is then incubated with Mg2+ at 37 -C to initiate the cleavage reaction. The active fraction is isolated by PAGE due to the change in size. Active DNA enzymes are then amplified through PCR using a 50 -labeled forward primer (fluorescein or 32P-labeled for detection of bands on PAGE) and a tailed reverse primer containing a nonextendable ethylene glycol spacer to allow separation of sense and antisense strands. After isolation of the single-stranded PCR product through denaturing PAGE, the enriched active species are ligated to the RNA substrate to initiate the next round of selection. To increase the specificity of the resulting deoxyribozymes, negative selection rounds are introduced, in which the DNA library is challenged with the unmodified RNA. Other factors that can be adapted to enhance specificity are metal ion concentration, selection time and temperature. After several rounds of in vitro selection, the enriched pool is tested for its ability to discriminate modified from unmodified RNA. Finally, individual deoxyribozymes are identified by traditional cloning and Sanger sequencing and/or from Illumina NGS datasets that allow a deeper analysis of the enrichment of certain sequence motifs. New candidate DNA enzymes are characterized by analyzing cleavage kinetics for modified and unmodified RNA substrates and mutational analyses are performed to identify key sequence motifs responsible for recognition of the RNA modification. In exceptional cases, the modified nucleotide in the RNA may lead to switch in the cleavage site of the endonuclease deoxyribozyme, providing an additional opportunity for quantitative readout of the modification level [24].

#### 2 Materials

	- 2. Purify oligonucleotides by denaturing PAGE before use and store all oligonucleotide solutions and buffers at 20 -C.
	- 3. RNA substrates with and without modification (for counter selection), prepared by solid-phase synthesis on 0.5–1 μmol scale, using commercially available or in-house synthesized phosphoramidites of modified nucleotides. A fraction of RNA substrate is labeled at 30 -end or 50 -end for kinetic assays.
	- 4. Deoxyribozyme library: 0.5 μmol synthesis scale, 100 μM stock solution.
	- 5. Primers: 50 -fluorescein labeled forward primer and PEG3 linked tailed reverse primer. 0.5 μmol synthesis scale, 100 μM solution.
	- 6. Splint DNA for ligation of DNA library to RNA substrate. 0.5 μmol synthesis scale, 100 μM solution.




#### 3.3 In Vitro Selection

3.3.1 Phosphorylation of RNA Selection Substrates

In order to ligate RNA selection substrates with DNA library, RNA has to be phosphorylated at 5<sup>0</sup> -end.


Ligation of ssRNA substrate with ssDNA library is facilitated by complementary DNA splint in the presence of T4 DNA ligase.


3.3.3 DNA-Catalyzed Cleavage of DNA–RNA Hybrids (Key Selection Step)

3.3.2 Splint Ligation of RNA Substrate to Deoxyribozyme Selection

Pool

Selection of active DNA enzymes is based on a change in size upon cleavage of the RNA substrate in the vicinity of the modified nucleotide.




	- 2. Assemble the reaction: 5 μL of DNA template, forward primer (200 pmol), reverse primer (50 pmol), 1.25 μL of 20 mM dNTP mixture, 10 μL of 10 Dream taq buffer, 0.5 μL of Dream taq DNA polymerase (5 U/μL), and water up to 100 μL.
	- 3. PCR conditions are: 95 -C for 4 min [30 (95 -C for 30 s, 60 -C for 30 s, 72 -C for 1 min), 72 -C for 5 min].
	- 4. Purify the PCR product using denaturing PAGE.
	- 5. Isolate the fluorescein labeled forward strand by extraction and precipitation as described under Subheading 3.2.

Take the single-stranded PCR product from the previous step and ligate it to the RNA substrate using the DNA splint (see Note 3) and T4 DNA ligase, essentially as described under Subheading 3.3.2, but on a smaller scale (i.e., use only 100 pmol of RNA and 75 pmol of DNA splint in a final volume of 10 μL for the ligation reaction). The isolated ligation product is then subjected to the next round of incubation as described under Subheading 3.3.3.

After 6–8 rounds of in vitro selection, include negative selection rounds to enrich DNA enzymes having high selectivity toward modified RNA and eliminate DNA enzymes that can cleave both modified and unmodified RNA. For this purpose, ligate the DNA pool with the unmodified analog of the RNA substrate. During the selection step, perform all steps in identical manner and analyze the

3.3.5 Continuation of Selection and Identification of Deoxyribozyme Sequences

Fig. 2 Schematic presentation of polyacrylamide gel electrophoresis (PAGE) for positive and negative selection rounds. The diagram presents the area of the gel from which the corresponding cleaved (positive selection round) or uncleaved fraction (negative selection round) is isolated

catalytic activity of the DNA pool by denaturing PAGE. Note that the uncleaved fraction on gel corresponds to the specific DNA enzymes that remain inactive toward unmodified substrate (see Note 5). The band signal observed at the height of the size marker corresponds to non-specific DNA enzymes that can cleave both modified and unmodified substrates. Therefore, in the negative selection round, the uncleaved fraction is isolated from the gel (Fig. 2) and amplified by PCR as described above. Each negative round is followed by a positive round (during these positive rounds, increase the stringency of the selection by decreasing the incubation time, i.e., 6 h, 3 h, and 1 h) and continue the cycle at least for four rounds of alternating negative and positive selections.

Afterward, amplify the active DNA pool through PCR. Due to non–template-dependent terminal transferase activity of Taq polymerase, the product will contain a 3<sup>0</sup> -A overhang that allows TOPO-TA cloning (commercially available kit). Select the individual clones and subject them to Sanger sequencing. Each sequence obtained from Sanger Sequencing data corresponds to individual deoxyribozyme candidates. These sequences are then synthesized by solid-phase syntheses and their catalytic potential and discrimination power for RNA modification are assessed by kinetic assays.

3.4 Kinetic Characterization of Deoxyribozymes Kinetic characterization analyzes the trans-activity of individual deoxyribozyme to cleave modified and unmodified RNA (that are not ligated to the DNA library). Therefore, for each deoxyribozyme, the assay is performed in parallel with both, modified and unmodified RNA substrates of otherwise the same sequence. This allows to compare the cleavage rates and to determine calibration curves that are needed for quantitative estimation of modification levels.

> 1. Take 100 pmol of deoxyribozyme, 10 pmol of fluorescent labeled RNA substrate and adjust to a final volume of 7.5 μL.


#### 4 Notes


#### Acknowledgments

This work was supported by the European Research Council (ERC consolidator grant No. 682586) and by the German Academic Exchange Service (Deutscher Akademischer Austauschdienst, DAAD, PhD scholarship to A.L.). M.V.S. thanks the Graduate School of Life Sciences at the University of Wu¨rzburg for a Post-Doc Plus fellowship.

#### References


baker's yeast reveals ribosome heterogeneity on the level of eukaryotic rRNA modification. PLoS One 9:e89640


direct m6A sequencing. Angew Chem Int Ed 57:417–421


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Chapter 11

# Mapping of the Chemical Modifications of rRNAs

## Jun Yang, Peter Watzinger, and Sunny Sharma

#### Abstract

Cellular RNAs, both coding and noncoding, contain several chemical modifications. Both ribose sugars and nitrogenous bases are targeted for these chemical additions. These modifications are believed to expand the topological potential of RNA molecules by bringing chemical diversity to otherwise limited repertoire. Here, using ribosomal RNA of yeast as an example, a detailed protocol for systematically mapping various chemical modifications to a single nucleotide resolution by a combination of Mung bean nuclease protection assay and RP-HPLC is provided. Molar levels are also calculated for each modification using their UV (254 nm) molar response factors that can be used for determining the amount of modifications at different residues in other RNA molecules. The chemical nature, their precise location and quantification of modifications will facilitate understanding the precise role of these chemical modifications in cellular physiology.

Key words rRNA, Chemical modifications, Ribosomes, RP-HPLC, Mung bean nuclease assay

#### 1 Introduction

Chemical modification of biomolecules is an impressive way of nature to expand their functional diversity. Like DNA and proteins, RNA molecules also undergo a variety of chemical modifications that allow them to enrich their topological potential and perform a variety of functions that go beyond their capacity to encode proteins [1, 2].

Around 163 different chemical modifications of RNA has been cataloged [3]. Both ribose sugar and the heterocyclic ring of the nucleobases are targeted for these chemical additions. Although the presence of these modifications was first acknowledged by some elegant studies in the middle of last century, it is only very recently that we have started compiling modification profiles for the cellular RNAs of model organisms, and explore the functions of few of these modifications [4]. This time lag has been primarily due to unavailability of advanced technology in studying these modifica-

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_11, © The Author(s) 2022

tions, which has been revolutionized in last two decades by introduction of both high-performance liquid chromatography (HPLC), mass spectrometry (MS), and RNA sequencing (RNA Seq) [5–8].

Ribosomes are highly conserved ribonucleoprotein complexes that synthesize cellular proteins. Eukaryotic ribosomes comprise of 4 ribosomal RNAs (rRNAs) and around 80 ribosomal proteins (r-proteins). Both enzymatic reactions of ribosomes, namely, peptidyl transfer and peptidyl hydrolysis are performed by rRNA [9]. In eukaryotes including Saccharomyces cerevisiae, a functional ribosome contains two asymmetric subunits, a small 40S and a large 60S. A small subunit (SSU) of the ribosome (the 40S) in yeast contains a single 18S rRNA of 1.8 kb together with 33 ribosomal proteins [10]. The 60S or the large subunit (LSU) of the yeast ribosome contains three rRNAs; 25S, 5.8S and 5S, and 46 ribosomal proteins. The 40S decodes the genetic information carried by mRNA, whereas the 60S catalyzes the joining of amino acids [11, 12]. Ribosomes have always been seen as homogeneous, constitutive protein-synthesizing machine, lacking any significant contribution in regulating gene expression. Typically, the efficiency of translation is suggested to be determined either by features intrinsic to the mRNAs or assisted by protein or RNA adaptors (translational factors). However, in contrast to this view, several recent studies have underscored the undeniable roles of ribosomes in gene regulation [13–17]. Emerging data have buttressed the notion that the ribosome population in cells is heterogeneous by virtue of its different components, including ribosomal proteins, rRNAs and their chemical modifications. It becomes more and more evident that this ribosomal heterogeneity provides a further regulatory level to the translation.

Ribosomal RNA contains three types of chemical modifications, 2<sup>0</sup> -O methylation of ribose sugars (Nm), base isomerization (pseudouridylation (Ψ)), and base modifications (methylation (mN) and acetylation (acN), aminocarboxypropylation (acpN)) [1, 2]. Methylated ribose sugars and pseudouridines represent the majority of rRNA modifications [18]. A methylated sugar is generated by the addition of one methyl group at the 20 -OH position of the ribose on the nucleoside and is independent of the nature of the base. Pseudouridylation results from uridine isomerization, involving a 180rotation around the N3–C6 axis [19].

Box C/D snoRNPs (small nucleolar ribonucleoproteins) catalyzes site-directed methylation at the 20 -OH position on the sugar of the targeted nucleotide, whereas the box H/ACA snoRNPs isomerize targeted uridine to pseudouridine [20]. Remarkably, the substrate specificity for both ribose methylation and pseudouridylation is dictated by the RNA component of these snoRNP complexes [21, 22]. Here the complementary sequences in the guide RNA base pair with target RNAs to decide the nucleotide for modification [21, 22]. The catalytic activities are provided by the methyltransferase (Nop1 or Fibrillarin) or the pseudouridine synthase (Cbf5), respectively [23, 24]. In contrast, majority of base modifications are catalyzed by "protein only" enzymes with only exception being rRNA cytosine acetylation (ac<sup>4</sup> C) [25].

To analyze and explore the significance of these modifications in ribosome biogenesis and ribosome function, a comprehensive analysis of their chemical nature and precise location on the ribosome is central. In this chapter, a detailed protocol for mapping these modifications on to the ribosomal RNA by mung bean nuclease (MBN) assay and RP-HPLC (reversed phase high-performance liquid chromatography) is provided [26].

MBN is a single strand specific nuclease, and can be used to isolate specific fragments of RNA by utilizing synthetic complementary DNA. These RNA fragments can then be subjected to RP-HPLC or LC-MS/MS analysis to identify the modification or modifications they harbor (Fig. 1) [26, 27]. This method also allows to achieve a single nucleotide resolution for mapping these modifications by isolating overlapping fragments [28, 29].

Although MBN assay and LC-MS/MS are work intensive, these are so far the only methods for a direct analysis of these chemical modifications. All other RNA sequencing based methods relies either on antibodies, which most of the time have issues with cross-reactivities or on the chemical reactivities of the modified residues, which are not very specific and are similar among many different modifications [30, 31]. The major limitation of the LC-MS/MS, on the other hand, is that it cannot be applied in a high throughput manner to reveal both the modification and the sequence context as in case of RNA Seq based methods. Oxford nanopore sequencing appears to be an ideal method of choice for the direct transcriptome-wide sequencing of these modifications [32].

#### 2 Materials


3. 5% and 25% sucrose solutions (w/v) in TEN buffer.

Fig. 1 Reversed phase high-performance liquid chromatography (RP-HPLC) analysis. (a) Sucrose gradient sedimentation profile of yeast total RNA separated on 5% to 25% sucrose gradient. Fractions corresponding to tRNA (predominantly)), 18S rRNA, and 25S rRNA were collected and the rRNA was isolated using 95% ethanol precipitation at 80 -C. (b) A representative 1.5% Agarose gel showing the rRNA recovered after ethanol precipitation. (c) RP-HPLC chromatogram of different commercially available nucleosides, showing peak corresponding to major nucleosides (adapted from [26])

2.2 Mung Bean Nuclease Protection Assay

1. Hybridization buffer.

250 mM HEPES pH 7.0.

500 mM KCl in nuclease-free water.


#### 3 Methods

3.1 Isolation of Intact 18S and 25S rRNA

	- 2. Extract the total RNA with hot acidic phenol exactly as described before [33].
	- 3. To isolate intact 18S and 25S rRNA, layer 500 μg of total RNA onto a freshly prepared 5–25% sucrose gradient in TEN buffer (see Note 2). Prepare the sucrose gradients using Gradient Master 107 (Biocomp).
	- 4. Centrifuge the samples at 67,000 g for 25 h at 4 -C in an SW40 rotor in an L-70 Beckman ultracentrifuge.
	- 5. Fractionate the gradients by using a Teledyne Isco density gradient fraction collector. Collect 18S and 25S rRNA fractions in separate tubes and to each add 2.5 volume of chilled ethanol. Incubate them overnight in a 20 freezer.
	- 6. Next day, centrifuge the rRNA samples at 16,000 g for 30 min at 4 -C, air-dry the pellet, and resuspend it in nuclease-free water.

2.3 Quantitative Reversed Phase High-Performance Liquid Chromatography (qRP-HPLC)





	- 2. Centrifuge the Eppendorf tubes at 4000 g for 5 min at 4 -C.
	- 3. Precipitate the rRNA fragments overnight at 20 -C by adding 350 μL of 8 M LiCl and 1 mL ethanol in 2 mL Eppendorf tubes.

#### 3.2.2 Mung Bean Nuclease Digestion


3.2.4 Mapping of Chemical Modifications to a Single Nucleotide Resolution

3.3 Quantitative Reversed Phase High-Performance Liquid Chromatography (qRP-HPLC)

3.3.1 Preparation of Nucleosides

As illustrated in Fig. 2, MBN assay could be easily used to achieve single nucleotide resolution by designing overlapping fragment around the modifications (see Note 4).

Nucleosides for RP-HPLC are prepared as described by Gehrke and Kuo, and adapted for rRNA as described previously [7, 26].


Fig. 2 Mung bean nuclease protection assay. (a) Schematic illustration of mung bean nuclease protection assay (MBN). (b) Representative Urea PAGE gel for the MBN assay showing an intact protected fragment retrieved after the mung bean nuclease digestion. The synthetic antisense oligonucleotide used for the protection is used as a marker (see Note 5) (adapted from [26])

Table 1 RP-HPLC elution protocol


	- 2. Use the following elution protocol (Table 1) described by Gehrke and Kuo for resolving majority of canonical and modified nucleosides in the rRNA (Fig. 1) [7]. For some

#### Table 2

UV 254 nm Molar Response factors for canonical and modified nucleosides (see Note 8) [26]


modifications like m3 U protocol has to be adjusted for better separation. In contrast to gradient elution for rest of the modified nucleosides, the elution conditions for m<sup>3</sup> U are changed to an isocratic mode using 50% buffer A and 50% buffer B.

3.3.3 Quantification of the Modified Nucleosides Approximate modification levels for each ribose and base modifications can be calculated from HPLC peak areas (UV254nm absorption). For the quantification, the peak areas are divided by the standard molar response factors. Table 2 contains the UV254nm molar response factors calculated in our previous study. Similarly, mole % (Seq) for yeast (S. cerevisiae) are calculated assuming that 18S rRNA (1800 nts) contains 473 unmodified adenosines (A), 494 unmodified uridines (U), 453 unmodified guanosines (G), and 343 unmodified cytidines (C), and similarly for 25S rRNA (3396 nts) assuming that it contains 885 unmodified adenosines (A), 828 unmodified uridines (U), 956 unmodified guanosines (G), and 653 unmodified cytidines (C) Table 3. Likewise, for MBN protected fragments residues per moles or extent of modifications for each residue can be established using the sequence of the isolated fragment and its comparison to the residues/moles calculated from the 18S rRNA or 25S rRNA (see Note 7).


#### Table 3 RP-HPLC quantification of Nucleosides in S.cerevisiae 18S and 25S rRNA [26]

\*m7 G has been shown to undergo partial-degradation during hydrolysis

#### 4 Notes


#### Table 4

List of oligonucleotides used for mapping base modifications (methylation and acetylation) and ribose methylation in S. cerevisiae rRNA using the MBN assay






set may have to be empirically determined by altering the Tm, either by increasing the length or sequence.


#### References


nucleosides: analytical methods for major and modified nucleosides, HPLC, GC, MS, NMR, UV, and FT-IR, Journal of chromatography library, vol 45. Elsevier, pp A3–A71


pre-rRNA processing, pre-rRNA methylation, and ribosome assembly. Cell 72:443–457


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Non-radioactive In Vivo Labeling of RNA with 4-Thiouracil

## Christina Braun, Robert Knu¨ ppel, Jorge Perez-Fernandez , and Se´bastien Ferreira-Cerca

#### Abstract

RNA molecules and their expression dynamics play essential roles in the establishment of complex cellular phenotypes and/or in the rapid cellular adaption to environmental changes. Accordingly, analyzing RNA expression remains an important step to understand the molecular basis controlling the formation of cellular phenotypes, cellular homeostasis or disease progression. Steady-state RNA levels in the cells are controlled by the sum of highly dynamic molecular processes contributing to RNA expression and can be classified in transcription, maturation and degradation. The main goal of analyzing RNA dynamics is to disentangle the individual contribution of these molecular processes to the life cycle of a given RNA under different physiological conditions. In the recent years, the use of nonradioactive nucleotide/nucleoside analogs and improved chemistry, in combination with time-dependent and high-throughput analysis, have greatly expanded our understanding of RNA metabolism across various cell types, organisms, and growth conditions.

In this chapter, we describe a step-by-step protocol allowing pulse labeling of RNA with the nonradioactive nucleotide analog, 4-thiouracil, in the eukaryotic model organism Saccharomyces cerevisiae and the model archaeon Haloferax volcanii.

Key words 4-thiouracil, RNA, Pulse labeling, RNA tagging, Haloferax volcanii, Saccharomyces cerevisiae

#### 1 Introduction

RNA homeostasis is a fundamental process for the regulation of gene expression and therefore for the physiology of living organisms [1, 2]. Several methods enable the characterization of the cellular RNA composition in a quantitative (relative amount) and qualitative manner (specific molecule identity) [1–3]. Classically, the presence and the relative amount of an RNA of interest can be easily assessed by northern blot or quantitative RT-PCR analyses

Christina Braun and Robert Knu¨ppel contributed equally to this work.

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_12, © The Author(s) 2022

[1–3]. More recently, the emergence of high-throughput technologies, like DNA microarrays [4–6] or next generation sequencing analysis [7, 8], have leveraged our capacity to obtain a faithful qualitative and quantitative overview of the ensemble of RNA molecules present at equilibrium in varying conditions. However, all these methods are inherently lacking temporal resolution and hardly permit in depth systematic analysis of the relative contribution of the individual molecular processes underlying the life cycle of an RNA [9, 10].

This problem was recognized very early in the history of molecular biology. Pioneering studies have ingeniously taken advantage of metabolic labeling to reveal the importance of RNA dynamic for gene expression. For example, the seminal work of Brenner, Jacob, and Meselson in 1961 enabled the discovery of mRNA as a dynamic intermediate molecule required for protein synthesis [11]. Metabolic labeling of RNA using radioactively labeled molecules, like nucleotides/nucleosides, has been the method of choice to explore RNA dynamics. However, country-dependent administrative regulations of radioactive isotopes constrain the use of metabolic labeling using radioactive-labeled nucleotides. In addition, the inherent difficulty of this technique to analyze dynamics of individual RNAs in a global and systematic fashion, has often confined this methodology to assess global transcription output or production and maturation of abundant cellular RNA (like ribosomal RNA maturation [12–19]). Alternative methods using nonradioactive nucleotide analogs for the metabolic labeling of RNA were explored early on [20]. However, it is only recently, thanks to improvement of chemistry, labeled-RNA enrichment and detection methods, that the full potential of this approach is unfolding [9, 21–24].

Several nonradioactive nucleotide analogs providing different advantages/disadvantages have been described in the literature [9, 21]. Among them, uracil derivatives have been widely used [9]. Due to their versatile chemical properties, thiouracil/uridine and uracil/uridine derivatives compatible with click-chemistry have emerged as molecules of choice to perform RNA dynamic analysis [9]. These uracil/uridine derivatives allow for orthogonal chemistry with a large battery of reagents to facilitate downstream visualization, isolation and system-wide characterization of labeled RNA, thereby providing an unprecedented time and resolution depth of RNA metabolism [9, 10, 21–28].

Furthermore, the chemical properties of the uracil/uridine derivatives can be further harnessed in combination with other methodologies (e.g., UV cross-linking, structure probing, mass spectrometry, next-generation sequencing) to investigate the dynamics of structure and composition of ribonucleoprotein particle (RNP) [23, 24, 28–34]. Overall, the potential of using chemical biology strategies and their combinations is still emerging and will undoubtedly provide new molecular insights into the life cycle of RNA molecules.

In this chapter, we describe step-by-step conditions to successfully perform 4-thiouracil labeling in two model organisms, Saccharomyces cerevisiae and Haloferax volcanii, and provide general technical advice, which should facilitate application of this methodology (Fig. 1).

#### 2 Materials

There are no specific preferences of sources of chemical reagents or materials, unless stated otherwise. Use ultrapure water with 18 MΩcm resistivity at 25 C, unless stated otherwise.

#### 2.1 Microbiological Cultures


2.1.2 Haloferax Volcanii Enhanced Casamino Acids (Hv-Ca<sup>+</sup> )


Fig. 1 Exemplary 4-thiouracil (4-TU) labeling of total RNA in S. cerevisiae and H. volcanii: (a) Workflow summary of 4-TU labeling. 4-thiouracil labeling and biotinylation workflow is briefly depicted. Growing cells (see Subheading 3.1.) are pulse labeled with 4-thiouracil as described in Subheading 3.2. After RNA extraction (see Subheading 3.3), sulfhydryl groups are specifically biotinylated in the presence of methylthiosulfonate activated Biotin derivate (MTS-Biotin) (see Subheading 3.4). After biotinylation RNA are separated by gel electrophoresis and visualized as described in Subheading 3.5 and below. (<sup>b</sup> and <sup>c</sup>) Exemplary 4-TU labeling applied on the eukaryotic model organism S. cerevisiae. The indicated yeast cells, wild-type (WT), cells depleted for the small and large ribosomal subunit biogenesis factors Pwp2 (GAL::PWP2) and Pol5 (GAL:: POL5), respectively [39, 40] were labeled for 20 min with 4-thiouracil as indicated in Subheading 3.2. Labeled total RNA were separated by denaturing agarose (b) or polyacrylamide (c) gel electrophoresis, respectively. Biotinylated RNA was detected as described in Subheading 3.5. Ribosomal RNA precursors (35S, 27S, and


Fig. 1 (continued) 20S rRNA) and mature ribosomal RNA (25S, 18S, 5.8 and 5S rRNA) are indicated. Please note that depletion of the small ribosomal subunit biogenesis factor, Pwp2, for 14 h leads to the reduction of small ribosomal subunit pre- and mature rRNAs (20S and 18S rRNA, respectively) [39]. In contrast depletion of the large ribosomal subunit biogenesis factor, Pol5, for 9 h leads to the reduction of the common large ribosomal subunit precursor and corresponding mature rRNA (27S, 25S, and 5.8S, respectively) [39– 41]. Additional application examples using this protocol and S. cerevisiae is described in [41]. (d) Exemplary 4-TU labeling applied on the archaeal model organism H. volcanii. Haloferax volcanii wildtype cells were labeled for 3 h with 4-thiouracil as indicated in Subheading 3.2. Labeled total RNA was separated by denaturing agarose electrophoresis. Biotinylated RNA was detected as described in Subheading 3.5. Mature ribosomal RNA (23S, 16S, and 5S rRNA) observed in these conditions are indicated. Additional application examples using this protocol and H. volcanii has been described previously [36]

#### 2.4 Separation and Immobilization of RNA

2.4.1 Agarose Gel Electrophoresis and Capillary Transfer


1. Urea.


2.4.2 Polyacrylamide Gel Electrophoresis and Electro Transfer


#### 2.5 Detection of 4-TU Labeled RNA

	- 2. IR-dye conjugated Streptavidin (1 mg/mL).
	- 3. Wash solution I: 1 PBS pH 7.5, 1 mM EDTA pH 8; 1% SDS.
	- 4. Wash solution II: 1 PBS pH 7.5, 1 mM EDTA pH 8; 0.1% SDS.
	- 5. Infrared detection system.

## 3 Methods (See Notes 4 and 5)

#### 3.1 Microbiological Methods (See Note 6)

3.1.1 Haloferax volcanii Medium and Cultivation


#### 3.1.2 Saccharomyces cerevisiae Medium and Cultivation


3.2 In Vivo Pulse Labeling with 4-TU (See Note 9)

3.2.1 Uracil and 4- Thiouracil Stocks

3.2.2 Haloferax volcanii Labeling



	- 2. Incubate for 30 min in the dark at room temperature.
	- 3. Add 250 μL AE buffer (see item 1 Subheading 2.2).
	- 4. Proceed with RNA extraction by adding the sample to a prepared tube containing 500 μL AE buffer-saturated phenol and 50 μL 10% SDS (see Subheading 3.3).
	- 5. Dissolved RNA pellet in RNA loading buffer (see item 6 Subheading 2.2).
	- 1. Mix 1.3% agarose in 0.85 final volume of H2O.
	- 2. Dissolve the agarose by heating the mixture in the microwave.
	- 3. Let cool down under agitation (~50 C) and add 0.1 volume of 10 MOPS running buffer and 0.054 volume of 37% Formaldehyde (final concentration 2%).
	- 4. Cast the gel.
	- 5. Prepare running buffer (1 MOPS, 2% formaldehyde).
	- 6. Load the RNA samples (obtained in Subheading 3.4).
	- 7. Run the gel overnight at 40 V and room temperature.

1. Prepare urea–polyacrylamide gel solution (6% acrylamide, 7 M urea solved in 0.5 TBE). Mix by stirring at room temperature until urea is properly dissolved.

2. Add 0.001% (v/v) TEMED and 0.1% (v/v) APS.

#### 3.5 Separation and Immobilization of RNA (See Note 11)

3.5.1 Preparation of Denaturing Agarose Gel

3.5.2 Preparation of Denaturing Polyacrylamide Gel


#### 4 Notes


#### Acknowledgments

We are grateful to Prof. Dr. Karl-Dieter Entian (University of Frankfurt) for comments and suggestions, to Corinna Reglin for early contribution to this project. We would like to thank our colleagues from the chair of Biochemistry III and Biochemistry I for sharing protocols, materials, equipment, and discussion. Thanks to Prof. Dr. Thorsten Allers (University of Nottingham), Prof. Dr. Anita Marchfelder (University of Ulm) for kindly sharing strains and protocols. Work in the Perez-Fernandez and Ferreira-Cerca laboratories are supported by the chair of Biochemistry III "House of the Ribosome"—University of Regensburg, by the DFG-funded collaborative research center CRC/SFB960 "RNP biogenesis: assembly of ribosomes and nonribosomal RNPs and control of their function" (project AP2 and project AP1/B13, respectively) and by an individual DFG grant to S.F.-C. (FE1622/2-1; Project Nr. 409198929). This work is dedicated to the memory of Prof. Dr. Jonathan Warner who has among other important contributions, pioneered pulse labeling of pre-rRNAs.

#### References


populations using efficient and reversible covalent chemistry. Mol Cell 59:858–866. https:// doi.org/10.1016/j.molcel.2015.07.023


with 4-Thiouracil (Ers4tU). J Vis Exp. https:// doi.org/10.3791/59952


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Part VI

Translation

# Chapter 13

# Translation Phases in Eukaryotes

Sandra Blanchet and Namit Ranjan

#### Abstract

Protein synthesis in eukaryotes is carried out by 80S ribosomes with the help of many specific translation factors. Translation comprises four major steps: initiation, elongation, termination, and ribosome recycling. In this review, we provide a comprehensive list of translation factors required for protein synthesis in yeast and higher eukaryotes and summarize the mechanisms of each individual phase of eukaryotic translation.

Key words Translation, Ribosome, mRNA, tRNA, Yeast

#### 1 Introduction

In all domains of life, information encoded in mRNA is translated to protein by a supramolecular machine called ribosome. Eukaryotic ribosome consists of a small subunit (40S) and a large subunit (60S) that together form the 80S ribosome. The ribosome reads the information one codon (three nucleotides) at a time using aminoacyl-tRNAs as adaptor molecules that recognize each codon to insert the appropriate amino acid. Ribosomes from bacteria, archaea, and eukarya share a high degree of sequence and structure conservation, indicating a common evolutionary origin. Furthermore, they share a similar central core where mRNA decoding, peptidyl transfer, and translocation of tRNA and mRNA by one codon take place. The process of translation can be divided into four main phases: initiation, elongation, termination, and ribosome recycling. During the initiation phase, eukaryotic translation initiation factors (eIFs) promote the assembly of 80S ribosomes at the AUG start codon with an initiator methionyl-tRNA bound to the P site (Table 1). During elongation, 80S ribosomes move processively along the mRNA, synthesizing the encoded protein through the coordinated actions of aminoacyl-tRNAs and the eukaryotic translation elongation factors (eEFs) (Table 2). At the end of the open reading frame, the ribosome encounters a termination codon recognized by eukaryotic release factors (eRFs) (Table 2), which

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_13, © The Author(s) 2022

#### Table 1 Eukaryotic translation initiation factors



#### Table 2

#### Eukaryotic translation elongation and termination factors


promote the release of the nascent protein from the ribosome. Finally, during the recycling phase, the ribosome complex is recycled to the 40S and 60S subunits to begin a new round of translation. In this review, we describe the four phases of eukaryotic translation and function of translation factors (Tables 1 and 2). In the accompanying methods chapter, we focus on in vitro reconstitution of initiation and elongation phases of translation. Detailed aspects of translation termination and recycling are discussed in recent reviews [1, 2].

#### 2 Initiation

The initiation phase leads to the formation of 80S ribosomes where the initiator tRNA (Met-tRNAi Met) and the start codon are positioned in the P site. This process requires at least ten eukaryotic initiation factors, eIFs (Table 1), and comprises two major steps. In the first step, Met-tRNAi Met binds to the start codon in the P site of the 40S subunit to form the 48S initiation complex. In the second step, the 60S subunit joins the 48S complex to form the 80S initiation complex that is ready for elongation (Fig. 1) [3, 4].

Formation of 48S preinitiation complex. During this step, eIF3, eIF1, and eIF1A are recruited to 40S subunits, followed by subsequent attachment of eIF5 and eIF2–GTP–Met-tRNAi Met to form 43S complexes. eIF3 binds to the 40S subunit side that faces the solvent in the 80S ribosome [5], whereas eIF1 binds between the platform of the 40S subunit and Met-tRNAi Met to the 40S side that will form the interface to the 60S subunit [6]. The structured domain of eIF1A resides in the A site, forming a bridge over the mRNA channel, and its N- and C-terminal tails extend into the P site. Binding of eIF1 and eIF1A to the 40S subunit induces conformational changes [7, 8], which result in the opening of the mRNA entry channel and the establishment of a new connection between the head and body domains of the 40S subunit on the solvent side between helix 16 of 18S rRNA and the ribosomal protein uS3. Although 43S complexes can bind model mRNAs with completely unstructured 5<sup>0</sup> UTRs in the absence of eIFs [9], attachment of natural mRNAs requires a coordinated activity of eIF4F protein complex, which unwinds the 5<sup>0</sup> cap-proximal region of mRNA. Following mRNA attachment, the 43S complex scans the mRNA downstream of the cap in search for the initiation codon. During the scanning process, the secondary structures in the 5<sup>0</sup> UTR unwind and the 43S complex moves until the initiation codon is recognized. eIF1 and eIF1A play key roles in the 43S complex formation and scanning; omission of either or both reduces or suppresses the scanning ability, indicating that the movement of 43S complexes requires the scanning-competent conformation induced by eIF1 and eIF1A [7]. eIF3, which is indispensable for

Fig. 1 Translation initiation. eIF2–GTP–Met-tRNAi Met binds to the 40S ribosomes in presence of eIF1, eIF1A, eIF3, and eIF5 to form 43S PIC. Then mRNA is recruited with the help of eIF4F complex to form the 48S PIC. After mRNA scanning and recognition of the start codon, the 60S subunit joins with the help of eIF5B to form the 80S IC [4, 59, 60]

48S complex formation, forms an extension of the mRNA-binding channel that might contribute to scanning [10]. The fidelity of start codon recognition is ensured by the discriminatory mechanism that promotes recognition of the correct initiation codon and prevents premature Met-tRNAi Met landing at near-cognate triplets in the 5<sup>0</sup> UTR. The bona fide start site is usually the first AUG triplet in an optimum nucleotide context GCC(A/G)CCAUGG, with a purine at the -3 and a G at the +4 position relative to A of the AUG codon [11]. eIF1 has an important role in initiation codon recognition: it enables 43S complexes to select the correct AUG in a poor initiation context or an AUG located within 8 nucleotides of the mRNA 5<sup>0</sup> end, and promotes dissociation of the ribosomal complexes that aberrantly assemble at such triplets in the absence of eIF1 [9, 12, 13]. Following start codon recognition and GTP hydrolysis, Pi is released from eIF2–GTP + Pi and eIF1 dissociates from the ribosome [14–16].

Ribosomal subunit joining. eIF5B, a ribosome-associated GTPase, facilitates the joining of the 60S subunit and the dissociation of eIF1A, eIF5, and eIF2–GDP [17]. GTP hydrolysis by eIF5B is essential for its own release from assembled 80S ribosomes. eIF5B occupies the region in the intersubunit cleft [18] and promotes subunit joining by burying large solvent-accessible surfaces on both subunits [19]. After the 60S subunit joining and dissociation of initiation factors, the 80S initiation complex with the anticodon of Met-tRNAi Met in the P site base-paired with the start codon becomes elongation competent.

#### 3 Elongation

The elongation phase comprises three steps: decoding of mRNA codons by the cognate aminoacyl-tRNAs, peptide bond formation, and translocation of the tRNA–mRNA complex, resulting in movement of peptidyl-tRNA from the A site to the P site and presenting the next codon in the A site [1] (Fig. 2). Four elongation factors are required the three steps (Table 2).

The eukaryotic elongation factor eEF1A binds aminoacyltRNA in a GTP-dependent manner and delivers the aa-tRNA to the A site of the ribosome (Fig. 2). Codon recognition by the aa-tRNA triggers GTP hydrolysis by eEF1A, releasing the factor and enabling the aa-tRNA to accommodate in the A site, presumably following the same pathway as described for bacterial ribosomes [20–24]. However, decoding appears to take place more slowly and accurately than in bacteria, particularly in higher eukaryotes [25]. eEF1A is a member of the GTPase superfamily that binds and hydrolyzes GTP. The dissociation of GDP from eEF1A is accelerated by a guanine nucleotide exchange factor (GEF), eEF1B, which is composed of two subunits, eEF1Bα and eEF1Bγ in yeast,

Fig. 2 Translation elongation. During decoding, eEF1A delivers aa-tRNA to the A site. After peptide bond formation, the A-site peptidyl-tRNA and P-site tRNA are translocated to the P and E sites, respectively, with the help of eEF2. Deacylated tRNA is released from the E site and the elongation cycle repeats until a stop codon is reached. In yeast, an additional factor eEF3 is required during elongation [1]

the first one containing the catalytic domain necessary for nucleotide exchange. eIF5A, which was originally identified as initiation factor, functions globally in translation elongation, especially when ribosome encounter polyproline sequences [2].

Following the accommodation of the aminoacyl-tRNA into the A site, the peptide bond formation with the P-site peptidyl-tRNA occurs rapidly in the peptidyl transferase center (PTC). The PTC consists mainly of conserved rRNA elements of the 60S subunit that position the substrates for catalysis. Structural studies have revealed that the rRNA structure of the PTC is nearly superimposable between the eukaryotic and bacterial ribosomes [26–28], supporting the notion that the mechanism of peptide bond formation is universally conserved.

The pretranslocation complex formed as a result of peptide bond formation is dynamic and can spontaneously fluctuate between several conformations, in prokaryotes and eukaryotes alike [29, 30]. The ribosomal subunits undergo a ratchet-like motion from the so-called classical to rotated state triggering movement of the tRNAs to adopt hybrid states. In this state, the anticodon ends of the tRNAs remain positioned in the P and A sites of the 40S subunit, while the tRNA acceptor ends are positioned in the E site and P site of the 60S subunit (P/E and E/P states, respectively) [31–33]. Translocation of the mRNA–tRNA in the ribosome requires elongation factor eEF2, which facilitates the return of hybrid tRNAs to the classical state, E/E and P/P (canonical E and P sites). eEF2 in its GTP-bound state facilitates and stabilizes the hybrid rotated state. Conformational changes in eEF2 upon GTP hydrolysis and Pi release unlock the ribosome allowing tRNA and mRNA movement and then lock the subunits in the posttranslocation state. In that state of the ribosome, a deacylated tRNA occupies the E site and the peptidyl-tRNA is in the P site, leaving the A site vacant and available for binding of the next aminoacyl-tRNA in complex with eEF1A [34, 35]. In addition to eEF1A and eEF2, a third factor, eEF3, is essential for elongation in fungi [36]. The elongation cycle is repeated until a complete protein has been synthesized and a stop codon is encountered by the ribosome.

#### 4 Termination

Termination occurs when the ribosome reaches a stop codon (UAA, UGA, or UAG) [1, 37–39] (Fig. 3). Termination is catalyzed by two protein factors, eRF1, which recognizes all three stop codons, and the GTPase eRF3, which facilitates termination at a cost of GTP hydrolysis [39–41]. eRF1 contains a structural Asn-Ile-Lys-Ser (NIKS) motif and several other conserved elements, including the Gly-Thr-Ser (GTS) and YxCxxxF motifs, which are involved in the recognition of the termination codons [42–46]. In addition, the central domain of eRF1 contains a Gly-Gly-Gln (GGQ) motif that extends into the peptidyl-transferase center to promote peptidyl-tRNA hydrolysis and the release of nascent peptide [47–50]. eRF1 requires eRF3 (a specialized GTPase) for proper function (Table 2). eRF1 and eRF3 bind to one another with very high affinity and probably enter the

Fig. 3 Translation termination and recycling. Termination occurs when a stop codon enters the A site of the ribosome and is catalyzed by eRF1 and eRF3. Peptide release is promoted by ABCE1 which also induces subunits dissociation [1]

ribosome as a complex [51]. GTP hydrolysis positions the GGQ region of eRF1 in the peptidyl transferase center and triggers peptidyl-tRNA hydrolysis. eRF3 strongly enhances peptide release by eRF1 in the presence of GTP, but not GDP or nonhydrolyzable GTP analogs [39]. Moreover, other trans-acting factors are known to affect translation termination. ABCE1 (Rli1 in yeast) interacts with eRF1 to stimulate its catalytic activity by stabilizing the active conformation of eRF1 [52–55].

#### 5 Ribosome Recycling

During recycling the ribosomal subunits dissociate and the mRNA together with the deacylated tRNA are released to regenerate the necessary components for subsequent rounds of translation. Recycling of ribosomal subunits is achieved by the ATPase ABCE1 (Rli1 in yeast), an essential protein which contains two nucleotide-binding domains and an amino-terminal iron–sulfur cluster (Fe-S) and induces subunits dissociation at the cost of ATP hydrolysis [53, 54, 56]. Upon binding and hydrolysis of ATP, the Fe-S cluster undergoes a conformational change that drives eRF1 into the ribosomal intersubunit space, leading to dissociation of posttermination ribosomes into 40S and 60S subunits. Deacylated tRNA and mRNA are then released from the 40S subunits, which may be additionally promoted by Ligatin (eIF2D) [57] (Table 2). Initiation factors eIF1, eIF1A, and eIF3j can also promote tRNA and mRNA dissociation from the 40S subunit in vitro [54]. Following termination, in some cases full dissociation of the ribosomal complex will occur, whereas in other cases partial dissolution of the complex will allow for reinitiation, a term used to describe a process wherein ribosome translates two or more ORFs in a transcript without undergoing complete recycling between these events [58]. However, the mechanism of reinitiation is not fully understood.

#### Acknowledgments

We thank Prof. Marina Rodnina for critical reading of the manuscript. This work is supported by the Deutsche Forschungsgemeinschaft (DFG) in the framework of the Schwerpunktprogram (SPP1784), and by the Max Planck Society.

#### References


a key step in start codon selection in vivo. Genes Dev 21:1217–1230


translation termination in Saccharomyces cerevisiae. EMBO J 14:4365–4373


CU, Pestova TV (2010) The role of ABCE1 in eukaryotic posttermination ribosomal recycling. Mol Cell 37:196–210


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

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# Differential Translation Activity Analysis Using Bioorthogonal Noncanonical Amino Acid Tagging (BONCAT) in Archaea

## Michael Kern and Se´bastien Ferreira-Cerca

#### Abstract

The study of protein production and degradation in a quantitative and time-dependent manner is a major challenge to better understand cellular physiological response. Among available technologies bioorthogonal noncanonical amino acid tagging (BONCAT) is an efficient approach allowing for time-dependent labeling of proteins through the incorporation of chemically reactive noncanonical amino acids like Lazidohomoalanine (L-AHA). The azide-containing amino-acid derivative enables a highly efficient and specific reaction termed click chemistry, whereby the azide group of the L-AHA reacts with a reactive alkyne derivate, like dibenzocyclooctyne (DBCO) derivatives, using strain-promoted alkyne–azide cycloaddition (SPAAC). Moreover, available DBCO containing reagents are versatile and can be coupled to fluorophore (e.g., Cy7) or affinity tag (e.g., biotin) derivatives, for easy visualization and affinity purification, respectively.

Here, we describe a step-by-step BONCAT protocol optimized for the model archaeon Haloferax volcanii, but which is also suitable to harness other biological systems. Finally, we also describe examples of downstream visualization, affinity purification of L-AHA-labeled proteins and differential expression analysis.

In conclusion, the following BONCAT protocol expands the available toolkit to explore proteostasis using time-resolved semiquantitative proteomic analysis in archaea.

Key words BONCAT, L-Azidohomoalanine, L-AHA, Archaea, Time-dependent, Translation, Translatome, Proteostasis, Haloferax volcanii, Sulfolobus acidocaldarius, Escherichia coli, Click chemistry

#### 1 Introduction

Cellular adaptation depends on efficient and timely controlled variation of the cellular protein synthesis and degradation capacity [1–3]. Accordingly, the resulting qualitative and quantitative ensemble of proteins present at a given time, in a given cell or tissue, or in a given environmental condition is instrumental for proper cell fate determination. Hence, proteome plasticity is a key feature in enabling suitable gene expression modulation.

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_14, © The Author(s) 2022

Depending on the nature of intra- and extracellular cues, changes in the protein repertoire and individual protein levels may occur at different time-scales, and need to be experimentally captured with accuracy to understand the underlying cellular and physiological mechanisms [1–3].

Over the last decade(s) various approaches have been used to explore protein synthesis and degradation dynamics [1–3]. Among these methods, ribosome profiling, which takes advantage of highthroughput sequencing of ribosome associated mRNA, has emerged as the method of choice to assess the translational capacity and properties of various cellular systems across different conditions [1–3]. Even though extremely powerful, this method remains an indirect measurement of protein synthesis, as it fails to directly measure the product of translation itself. Moreover, protein degradation is not monitored using this experimental setup. An alternative, but still emerging method, is the use of time-dependent semiquantitative proteomic approaches, which allow for analysis of both protein synthesis and degradation [1]. While less popular, and still technically challenging, various mass spectrometry–based approaches have been developed and significantly improved over the years and are now able to provide semiquantitative and direct analysis of cellular protein composition in a time-dependent manner [1, 4–8]. To achieve such in-depth analysis various experimental setups can be used depending on the time resolution and desired extensiveness of the analyses [1, 3]. Early on, time-dependent proteomics have relied on isotope labeling (e.g., SILAC), whereby differential incorporation of heavy and light isotope metabolites like amino acids are used over a specified time to sort protein populations [1, 3, 9–11]. However, isotope labeling strategies have different drawbacks: (1) some labeled metabolites can be rather expensive, (2) efficiency of labeling is highly dependent on cellular uptake and metabolism capacity, and usually requires defined synthetic growth medium, (3) multiplexing is limited due to the lack of different reagents, (4) short kinetic pulse, low-abundant or proteins with a high turnover rate are difficult to be properly analyzed due to the lack of a specific enrichment step of the isotopically labeled peptides [1, 3, 9–11].

An emerging complementary strategy allowing to circumvent some of these limitations and facilitating time-dependent proteomic analysis is bioorthogonal noncanonical amino acids tagging (BONCAT) [12, 13]. In brief, BONCAT exploits the power of in vivo incorporation of bioorthogonal molecules into polypeptides and of click chemistry which enables enrichment and/or visualization of the labeled proteins synthesized over a defined time window [4, 5, 8, 13–18]. Most importantly, BONCAT can be easily combined with isotope labeling and/or multiplexing (e.g., [4, 5, 8, 17]), thereby allowing to fully harness the potential of time-dependent proteomic analysis.

Fig. 1 Optimized BONCAT workflow in H. volcanii. (a) Chemical structures of L-methionine and L-azidohomoalanine (b) Click chemistry: Strain Promoted Alkyne–Azide Cycloaddition (SPAAC) reaction. Azidecontaining polypeptide, DBCO derivate, and the results of click chemistry reaction are schematically depicted (c). Workflow summary of BONCAT in H. volcanii. Typical workflow as described in this protocol. The numbering below the different steps refers to the corresponding protocol steps described in the Methods section (see Subheading 3)

Most commonly BONCAT experiments utilize, L-azidohomoalanine (L-AHA), a methionine analogue containing a functional azide group (Fig. 1a). L-AHA is recognized, albeit with less efficiency, by the methionyl-tRNAMet acyl transferase, and is charged onto tRNAMet, thereby allowing for incorporation of L-AHA into polypeptides instead of methionine during protein synthesis [19]. The resulting L-AHA-modified polypeptides can be further altered using strain-promoted alkyne–azide cycloaddition (SPAAC) as outlined in Fig. 1b [20]. Alkyne derivative of biotin or various fluorophores can be efficiently and quantitatively coupled to the azide-containing proteins, thereby facilitating their further visualization and/or subsequent specific enrichment for downstream analysis (Fig. 1b) and exemplary results shown in Figs. 2, 3 and 4).

Interestingly, BONCAT has been successfully applied in a wide variety of organisms and/or tissues across the tree of life, thereby highlighting its potential [4, 5, 8, 13–19]. However, despite its broad application potential, proteomics analysis in general and time-dependent analysis of protein synthesis and degradation in particular, remains in its early days in archaea.

Haloferax volcanii, an extreme halophile Euryarchaeota, has emerged as a very powerful model organism to perform genetic and functional analysis in archaea [21, 22]. Its handling simplicity has attracted several scientists who aim to push the

Fig. 2 Exemplary BONCAT of prokaryotic cells. (a) L-AHA-based labeling of proteins in E. coli and H. volcanii. The corresponding cells (E. coli K12 and H. volcanii H26) were grown in synthetic medium lacking methionine (M9 and Hv-min, respectively) and pulse labeled with increasing amounts of L-AHA (0.1 mM and 1 mM) or 1 mM Methionine for 3 h. Cells were lysed, proteins were extracted and subjected to SPAAC click chemistry with DBCO-Cy7 as described in Subheading 3. Around 150 <sup>μ</sup>g of proteins were subsequently fractionated on a 4–12% polyacrylamide gradient gel and the fluorescence signal was acquired with an Odyssey Infrared Imager (right panel). Coomassie staining (loading control) is depicted in the left panel (b) Inhibition of L-AHAbased labeling in presence of translation inhibitor in H. volcanii. Same as in (a) except that cells (H. volcanii H26) were grown in synthetic medium lacking methionine and supplemented with 0.1 mM methionine or 0.1 mM of L-AHA with or without 2.5 <sup>μ</sup>M of the translation inhibitor Thiostrepton (Th) [31] for 1 h. (c) Shorttime L-AHA labeling in H. volcanii. Same as in (a) except that cells (H. volcanii H26) were grown in synthetic medium lacking methionine and supplemented with 1 mM L-AHA for the indicated time. The depicted fluorescence signal intensities across single lanes were quantified using Fiji [32] and visualized in Microsoft Excel. Adapted from [33] under CC-BY License

experimentational limits of this organism and crack open its molecular biology. Efforts in proteomics analysis do not escape this trend. In fact, proteomics analyses in H. volcanii have been making significant progress over the last years [23–27], and are currently being improved by a concerted action of several groups working on H. volcanii (see Archaeal Proteome Project—ArcPP—https:// archaealproteomeproject.org and https://github.com/arcpp/ ArcPP—[28]). Finally, recent methodological breakthroughs using isotope labeling, SILAC reagents or 15N-labeled nitrogen

Fig. 3 Enrichment of L-AHA-containing protein by affinity purification. H26 cells were grown in synthetic medium lacking methionine and supplemented with 0.1 mM of L-AHA or methionine (Met) for 5 h. Cells were lysed and proteins were extracted and subjected to click chemistry with 1 <sup>μ</sup>M DBCO-PEG4-Biotin as described in Subheading 3.4. Affinity purification was performed as described in Subheading 3.6. Sample aliquots for input, unbound fraction (Flow), wash fraction and elution were taken at each step of the AP. Elution was performed as described in Subheading 3.6.4. 0.5% of the input, unbound (Flow) and 0.25% of the wash fraction and 20% of the eluate were loaded on two SDS-PAGE gels respectively. The fractionated proteins were visualized by silver staining (a) and transferred onto a nylon membrane (b). Membrane was incubated with IRDye-coupled streptavidin to visualize the biotinylated proteins. Images were acquired either with the Epson Scanner (silver staining) (a) or the Li-COR Odyssey Infrared Imager (IRDye signal) (b). Asterisk indicates streptavidin monomers that are coeluted

source, in H. volcanii [24, 27] offer a unique opportunity to perform cutting-edge time-dependent proteomics analysis in this model archaeon.

Here we expand the H. volcanii proteomics toolkit and describe a step-by-step BONCAT protocol and downstream visualization/enrichment procedures of labeled proteins optimized for H. volcanii (summarized in Fig. 1c). In addition, this protocol has been validated in another model archaeon (i.e., Sulfolobus acidocaldarius—Ferreira-Cerca lab unpublished), another model bacterium (i.e., Escherichia coli—see below) and is likely to be suitable for various additional biological systems [4, 5, 8, 13–19].

Together with the isotope-labeling strategy mentioned above and additional multiplexing [4, 5, 8], the following BONCAT protocol allows for global inspection of protein synthesis capacity in Haloferax volcanii and paves the way to time-dependent proteomics analysis in this ever more versatile model archaeon [21, 22]. Moreover, these additional experimental possibilities position H. volcanii as a unique experimental system to explore the molecular principles of proteostasis in archaea.

Fig. 4 Differential 2D gel analysis of L-AHA-labeled proteins. (a) Differential 2D gel experimental workflow. Exponentially growing wildtype (WT) and △yfg (your favorite gene mutant/or perturbation) cells grown in Hv-min lacking methionine were pulse-labeled with 1 mM L-AHA for 45 min (representing approx. 20% of the H. volcanii doubling time in these growth conditions). Cells were lysed, protein were extracted and subjected to click chemistry with DBCO-Cy7 and DBCO-Cy5.5 as described in Subheading 3.4. Equal protein amounts of WT cells labeled with DBCO-Cy7 and △yfg cells labeled with DBCO-Cy5.5 and vice versa (dye swap to control fluorescence bias) were mixed cells with DBCO-Cy5.5 and vice versa to control fluorescence bias. Sample mixtures were used to rehydrate immobilized pH stripe and isoelectric focusing as describe in Subheading 3.7.2. Second dimension was resolved using 4–12% polyacrylamide gradient gel and the fluorescence signals were acquired with the Li-COR Odyssey Infrared Imager. (b) Exemplary results of differentially expressed proteins. Full gel scan (upper part) for the 2D gel analysis obtained for the wild-type (Cy7-red channel) and mutant H. volcanii cells (Cy5.5-green channel) mixture are provided. Zoom in of the 2D gel (boxing) showing representative differential expression between wildtype and exemplary mutant cell used in this study is provided in the lower part. Note that the dye-swap experiment (WT-Cy5.5/Δyfg-Cy7) provided very similar results (data not shown). (Adapted from [33] under CC-BY License)

#### 2 Materials

There are no specific preferences of sources of chemical reagents or materials, unless stated otherwise. Use ultrapure water with 18 MΩcm resistivity at 25 C.

#### 2.1 Microbiological Cultures

2.1.1 Strains

2.2 Haloferax volcanii Minimal Medium (Hv-min)




#### 2.3 Noncanonical Amino Acid Pulse Reagents

	- 2. Extraction Buffer (EB) without detergent: 150 mM NaCl, 100 mM EDTA, 50 mM Tris–HCl pH 8.5, 1 mM MgCl2.
	- 3. Thermoblock.
	- 4. Benchtop centrifuge.

2.6 Acetone Precipitation of Total Proteins

2.7 Protein Alkylation

	- 2. Dark box.
	- 1. Precooled (20 C) Acetone p.a.
	- 3. Precooled benchtop centrifuge.
	- 1. Extraction Buffer (EB) + 1% SDS: 150 mM NaCl, 100 mM EDTA, 50 mM Tris–HCl pH 8.5, 1 mM MgCl2, 1% SDS.
	- 2. Extraction Buffer (EB) without detergent: 150 mM NaCl, 100 mM EDTA, 50 mM Tris–HCl pH 8.5, 1 mM MgCl2.
	- 3. 10 Alkylation buffer: 10 PBS, 2 M 2-chloroacetamide.
	- 4. Thermoblock.
	- 5. Benchtop centrifuge.
	- 6. Black 1.5 mL reagent tubes.
	- 7. Rotating wheel.



#### 3 Methods (Workflow Is Summarized in Fig. 1c)

Use ultra-pure water with 18 MΩcm resistivity at 25 C.


ature in a dark cupboard.



3.4.4 Click-Chemistry: Strain Promoted Alkyne– Azide Cycloaddition

(SPAAC)



3.6.4 Elution (See Note 9) (an Exemplary Result Is Provided in Fig. 3)

Fast-Elution for Gel Electrophoresis

Competitive Elution for Downstream Processing


3.7 Gel Electrophoresis and Detection

3.7.1 SDS-PAGE

3.7.2 2D Gel Electrophoresis (an Exemplary Result Is Provided in Fig. 4)

Isoelectric Focusing (See Note 2)

Second Dimension SDS-PAGE


#### Table 1 Parameters for isoelectric focusing


1. Incubate the gel in gel fixation solution (see item 4 Subheading 2.11) for at least 20 min under agitation.


(an exemplary result is provided in Fig. 3).


3.8 In-Gel Detection (Exemplary Results Are Provided in Figs. 2 and 4)

3.9 Detection of Affinity Purified L-AHA-Labeled Proteins

#### 4 Notes


#### Acknowledgments

We are indebted of the scientists who have pioneered click chemistry, BONCAT, and other methodological aspects which have strongly inspired the establishment of this protocol. We are grateful to Prof. Dr. Karl-Dieter Entian (University of Frankfurt) for comments and suggestions. We would like to thank Dr. Robert Knu¨ppel, Michael Ju¨ttner and our colleagues from the chair of Biochemistry III and Biochemistry I for sharing protocols, materials, equipment, and discussion. Dr. Astrid Bruckmann (University of Regensburg) for assistance with 2D SDS-PAGE. Thanks to Prof. Dr. Sonja-Albers (University of Freiburg), Prof. Dr. Thorsten Allers (University of Nottingham), and Prof. Dr. Anita Marchfelder (University of Ulm) for kindly sharing strains and protocols. Work in the Ferreira-Cerca laboratory is supported by the chair of Biochemistry III "House of the Ribosome" – University of Regensburg, by the DFG-funded collaborative research center CRC/SFB960 "RNP biogenesis: assembly of ribosomes and nonribosomal RNPs and control of their function" (project AP1/B13) and by an individual DFG grant to S.F.-C. (FE1622/2-1; Project Nr. 409198929).

#### References


c5sc03340c click here for additional data file. Chem Sci 7:1797–1806. https://doi.org/10. 1039/c5sc03340c


approach to expression proteomics. Mol Cell Proteomics 1:376. https://doi.org/10.1074/ mcp.M200025-MCP200


U S A 99:19–24. https://doi.org/10.1073/ pnas.012583299


Appl Environ Microbiol 70:943–953. https:// doi.org/10.1128/AEM.70.2.943-953.2004


https://doi.org/10.1016/0300-9084(87) 90212-4


Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made.

The images or other third party material in this chapter are included in the chapter's Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter's Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

# Thermofluor-Based Analysis of Protein Integrity and Ligand Interactions

# Sophia Pinz, Eva Doskocil, and Wolfgang Seufert

#### Abstract

Thermofluor is a fluorescence-based thermal shift assay, which measures temperature-induced protein unfolding and thereby yields valuable information about the integrity of a purified recombinant protein. Analysis of ligand binding to a protein is another popular application of this assay. Thermofluor requires neither protein labeling nor highly specialized equipment, and can be performed in a regular real-time PCR instrument. Thus, for a typical molecular biology laboratory, Thermofluor is a convenient method for the routine assessment of protein quality. Here, we provide Thermofluor protocols using the example of Cdc123. This ATP-grasp protein is an essential assembly chaperone of the eukaryotic translation initiation factor eIF2. We also report on a destabilized mutant protein version and on the ATP-mediated thermal stabilization of wild-type Cdc123 illustrating protein integrity assessment and ligand binding analysis as two major applications of the Thermofluor assay.

Key words Thermofluor, Thermal shift assay, Differential scanning fluorimetry, SYPRO Orange, Protein stability, Ligand binding, ATP, Cdc123, eIF2

#### 1 Introduction

Recombinant proteins are widely used for biochemical analyses, and frequently the question comes up whether the purified or stored protein is still stable. Methods suitable to give an answer, such as CD spectroscopy or differential scanning calorimetry, typically require large protein amounts and expensive specialized instruments, which are not available in many laboratories. In contrast, Thermofluor is a low-cost and straightforward technique well suited as a routine protein quality control in most molecular biology laboratories. Such quality control is a must for batch-to-batch comparisons and to improve experimental reproducibility.

Thermofluor, a thermal shift assay also known as differential scanning fluorimetry (DSF), has become a versatile technique for the measurement of protein stability. Thermofluor makes use of an environmentally sensitive fluorescent dye, mostly SYPRO Orange

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_15, © The Author(s) 2022

Fig. 1 A typical Thermofluor profile. Data shown were obtained with SpCdc123-his6 in the presence of 5 SYPRO Orange. (a) At low temperatures the protein is well-folded (gray sphere). The fluorescent dye SYPRO Orange is quenched in the aqueous environment. Thus, only basal SYPRO Orange fluorescence emission is measured at 555 nm upon excitation at 470 nm (1). As the protein gradually unfolds (gray ravel), SYPRO Orange binds to exposed hydrophobic regions. This leads to a strong increase in fluorescence emission (2). The protein's melting temperature (Tm, arrow) is given by the inflection point where 50% of the protein is unfolded. Following the peak of fluorescence intensity (protein is completely unfolded), a decrease of intensity is observed (3). This is probably due to protein aggregation, which removes protein from the solution and prevents SYPRO Orange from interacting with hydrophobic patches. (b) First derivative of the fluorescence as a function of temperature. The protein's melting temperature (Tm, arrow) is easily identified as the peak of the curve (see Note 13)

[1, 2], to monitor the thermal unfolding of proteins. The dye is quenched in an aqueous environment, but undergoes a pronounced increase in fluorescent quantum yield upon binding to exposed hydrophobic regions of the protein as the protein unfolds (Fig. 1). The gradual increase of temperature as well as the concomitant detection of the fluorescent signal can be performed by standard real-time PCR instruments. Data fitting using the realtime PCR instrument's accompanying software quickly provides the melting temperature (Tm) of the protein under various conditions. The T<sup>m</sup> serves as a measure of protein stability [1, 3] and the shape of the curve as an indicator for protein integrity ( [2, 4] and Fig. 2).

Upon its first description in 2001 for high throughput drug discovery [5], dyes such as 1-anilino-8-naphthalenesulfonate (ANS) or dapoxyl sulfonic acid were used, but eventually the dye with the most favorable characteristics for Thermofluor turned out to be SYPRO Orange [1, 6]. It has a high increase in quantum yield, and its excitation and emission maxima of ~500 nm and ~600 nm, respectively [1, 6], are compatible with standard filter sets of most real-time PCR instruments [2].

The simplicity and economical protein requirement make Thermofluor very attractive for the routine quality control of purified recombinant proteins. Thermofluor allows to easily evaluate any adverse effects on the protein of choice that might occur during

Fig. 2 Thermal destabilization of a mutant protein. Thermofluor was performed using 5x SYPRO Orange and 2 <sup>μ</sup>g (2 <sup>μ</sup>M) SpCdc123-his6 wild-type (WT, black line) or two SpCdc123-his6 mutants: mutant 1 (M1, dark gray line) and mutant 2 (M2, light gray line). In the thermal denaturation profiles shown, fluorescence emission is plotted versus temperature to monitor protein unfolding. (a) Proteins in imidazole-containing buffer (50 mM Tris–HCl pH 8.0, 500 mM NaCl, approximately 160 mM imidazole) before dialysis. (b) Proteins in imidazolefree buffer (50 mM Tris–HCl pH 7.5, 500 mM NaCl) after dialysis (see Note 3). Shown is one representative curve per condition. The <sup>T</sup>m (calculated from duplicate reactions) was derived as the peak of the first derivative of the fluorescence as a function of temperature, calculated by the melting curve analysis of the Rotor-Gene Q Software 2.3.4. Mutant 1 is similarly stable as wild-type SpCdc123-his6. Both proteins are essentially unaffected by imidazole. Mutant 2 is less stable than wild-type SpCdc123-his6. In imidazole-free buffer, it does no longer show a defined unfolding transition. Together the data indicate that amino acid replacements can affect protein stability and sensitize a protein toward buffer composition

affinity purification, protein concentration, buffer exchange (Fig. 2), freezing or prolonged storage. In particular for mutant proteins, Thermofluor quickly shows whether or not amino acid exchanges do influence protein stability (Fig. 2). Thermofluor analysis therefore provides valuable data for the interpretation of downstream assays.

Ligand interaction usually stabilizes the native protein [1, 7, 8], thus leading to an increase in melting temperature. This can be employed, on the one hand, for ligand screenings in drug design [5, 8–10], and on the other hand, to characterize the binding of natural ligands, such as nucleotides, to proteins [11, 12] (Fig. 3). Even an approximation of Kd values is possible [9, 13–15], which provides useful preinformation for biophysical methods like isothermal titration calorimetry (ITC) that consume larger quantities of protein. Thermodynamic parameters obtained from Thermofluor assays correlate well with those determined by other biophysical methods [4, 9, 16, 17]. Not only ligands but also solvents and additives affect the stability and thus the T<sup>m</sup> of proteins. Accordingly, Thermofluor is a popular method for the determination of optimal buffer conditions for protein purification, storage and structural studies such as crystallization or NMR [2, 3, 16, 18–20].

Fig. 3 Stabilization of hD123 through ATP. Thermofluor was performed using 2 <sup>μ</sup>g hD123(1–290)-his6 (2.3 <sup>μ</sup>M) and 5 SYPRO Orange without nucleotide (no, black line), or with 1 mM ADP (ADP, dark gray line) or 1 mM ATP (ATP, light gray line). One representative curve is shown. (a) In the thermal denaturation profiles shown, fluorescence emission is plotted versus temperature to monitor the unfolding of hD123 (1-290)-his6. (b) The first derivative of the fluorescence as a function of temperature was exported from the Rotor-Gene Q Software 2.3.4 and plotted using Microsoft Excel. The <sup>T</sup>m is represented as the peak of the curve. The <sup>T</sup>m values shown in the legend were calculated as an average of duplicates from three independent experiments. The data indicate that ATP-binding stabilizes hD123 by more than 10 degrees

Recombinant proteins are critical tools in biochemical studies such as the analysis of ribosome biogenesis and mRNA translation. Recently it has become clear that the well-studied eukaryotic translation initiation factor 2 (eIF2) requires a dedicated assembly factor [21]. This protein called Cdc123 is conserved among eukaryotic organisms and indispensable for the viability of yeast and human cells [21–24]. Protein structure analysis revealed that Cdc123 is related to ATP-grasp enzymes [25]. Here we use Cdc123 as an example to present Thermofluor protocols for the analysis of protein stability and nucleotide binding. We show data illustrating a mutant protein that became unstable after removal of imidazole by dialysis, while generating a proper unfolding curve before dialysis (Fig. 2). Furthermore, as an example of a nucleotide-binding assay, we show that Cdc123 is stabilized strongly by ATP, and to some extent also by ADP (Fig. 3). While several of the technical aspects discussed here are specific to the real-time PCR machine Rotor-Gene Q 2plex Platform (Qiagen), the method can be applied to any standard real-time PCR instrument [2, 3, 9, 10, 13, 15] typically available in molecular biology laboratories.

#### 2 Materials

1. SYPRO Orange Protein Gel Stain, 5000 concentrate in DMSO (e.g., Thermo Scientific) (see Note 1), diluted 1:5 in DMSO to yield a 1000 working dilution in DMSO (see Note 2). The 1000 SYPRO Orange stock is stored at 4 C.

Fig. 4 Fluorescence dye optimization. Melting curves are shown using different SYPRO Orange concentrations as described in Subheading 3.1. Thermofluor was performed using 2 <sup>μ</sup>g his6-SpCdc123 (2 <sup>μ</sup>M) together with 1 (light gray line), 5 (gray line), 10 (dark gray line), or 20 (black line) SYPRO Orange. The gain was set to 9.33. In the thermal denaturation profiles shown, fluorescence intensity is plotted versus temperature to monitor the unfolding of his6- SpCdc123. The melting curve with the best dynamic rage was obtained with 5 SYPRO Orange

2. Purified recombinant proteins dialyzed against 2 Thermofluor (TF) buffer (see Note 3).

We used Cdc123 from Schizosaccharomyces pombe (Sp), carrying a six histidine affinity-tag (his6) at the N- or C-terminus, as indicated: his6-SpCdc123 (39.1 kDa) (Fig. 4) and SpCdc123-his6 (37.9 kDa) (wild-type (wt), mutant 1 (M1) with 2 amino acid exchanges, and mutant 2 (M2) with an additional third exchange) (Figs. 1 and 2). In addition, a C-terminally truncated version of the human homolog hD123 (hD123(1-290)-his6; 34.8 kDa) (Fig. 3) was used. All proteins were expressed in E. coli BL21- CodonPlus and purified on an A¨KTA system using Ni-NTA columns (GE Healthcare). An imidazole gradient (20–350 mM) was used for elution of the proteins from the column.


#### 3 Methods

Unless indicated otherwise, all steps are performed on ice.

3.1 Optimization of Protein Amount and SYPRO Orange Concentration (See Note 5 and Fig. 4)


#### Table 1 Protein mix for optimization of SYPRO Orange and protein concentration


#### Table 2 Buffer mix for protein stability assay



3.3 Nucleotide Binding Assay

#### Table 3 Buffer mix for nucleotide binding assay


2. The melting curves of wild-type hD123 (hD123(1-290)-his6) in the absence of nucleotide and in the presence of ADP and ATP are shown in Fig. 3. The data indicate that ATP stabilizes hD123 by more than 10 degrees.

#### 4 Notes


relative to protein leads to reduced quantum yield (see Fig. 4 20 and 1 SYPRO Orange). Choose the lowest protein amount that uses the full dynamic range of the photomultiplier, a good signal to noise ratio and sharp unfolding transition. Another approach, starting with a fixed 10 SYPRO Orange concentration and adjusting only protein amount, will in many cases consume more protein than necessary. We found that for most proteins optimal conditions are at around 2 μg protein (0.08 mg/ml) and 5 SYPRO Orange. However, in some cases protein concentrations might have to be varied from 0.01 to 0.2 mg/ml and SYPRO Orange from 1 to 20 to obtain suitable conditions.


also results in high background fluorescence, or (3) might lack hydrophobic regions. The latter is often the case for small proteins, for example, ubiquitin (data not shown) and [2].

13. Here, the T<sup>m</sup> is determined by plotting the first derivative of the fluorescence as a function of temperature, where the T<sup>m</sup> is represented as the peak of the curve. Alternatively, the Tm, as midpoint of the unfolding transition, can be determined by nonlinear fitting to a Boltzmann equation [3, 17]. The data can be easily exported from the real-time PCR instrument's software as a text file and analyzed with a method and software of choice.

#### Acknowledgments

This work was supported by the Deutsche Forschungsgemeinschaft (DFG) within the collaborative research center SFB960. We thank Wolfgang Mages and Lena Kreuzpaintner for purification of the his6-SpCdc123 protein used in Fig. 4.

#### References


RL, Jaeger EP, Devine H, Asel ED, Springer BA, Bone R, Salemme FR, Todd MJ (2005) Decrypting the biochemical function of an essential gene from Streptococcus pneumoniae using ThermoFluor technology. J Biol Chem 280:11704–11712


stability: making a good technique more robust. ACS Comb Sci 15:387–392


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# In Vitro Assembly of a Fully Reconstituted Yeast Translation System for Studies of Initiation and Elongation Phases of Protein Synthesis

Sandra Blanchet and Namit Ranjan

#### Abstract

Protein synthesis is an essential and highly regulated cellular process. To facilitate the understanding of eukaryotic translation, we have assembled an in vitro translation system from yeast using purified components to recapitulate the initiation and elongation phases of protein synthesis. Here, we describe methods to express and purify the components of the translation system and the assays for their functional characterization.

Key words Translation, Ribosome, mRNA, tRNA, Yeast

#### 1 Introduction

Protein synthesis in yeast is carried out by the 80S ribosomes that translate genetic information from messenger RNA (mRNA) into functional proteins with the help of aminoacyl-tRNAs (aa-tRNA) and a large number of specific translation factors. Translation entails four steps, initiation, elongation, termination, and ribosome recycling [1, 2]. During the initiation phase, the ribosome selects a start codon and an open reading frame for translation. Eukaryotic initiation factor 2 (eIF2), in its active GTP-bound form, binds to initiator methionyl-tRNA (Met-tRNAi Met) to form a ternary complex that delivers the tRNA to the small ribosomal subunit (40S). The 40S subunit together with eIF2–GTP–Met-tRNAi Met, eIF1, eIF1A, and eIF3 forms the 43S preinitiation complex (43S PIC), which then interacts with the mRNA that is recruited by the eIF4/ PABP complex to form the 48S complex (48S PIC). Following scanning and start codon recognition, eIF2 hydrolyzes GTP using eIF5 as a GTPase activating protein, allowing the dissociation of eIF2–GDP and recruitment of eIF5B. After large ribosomal subunit (60S) joining, eIF5B hydrolyzes GTP, the initiation factors

Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9\_16, © The Author(s) 2022

Fig. 1 Schematic of components required to in vitro reconstitute yeast mRNA translation initiation and elongation using a short unstructured mRNA, which overcomes the requirement of eIF4F complex

dissociate and the 80S initiation complex (80S IC) is ready to enter elongation (Fig. 1). eIF2–GDP is recycled to its active GTP-bound form with the help of the nucleotide exchange factor eIF2B, whereas the nucleotide exchange in eIF5B occurs spontaneously [2–4]. In the subsequent step of elongation, the eukaryotic elongation factor eEF1A binds aa-tRNA in a GTP-dependent manner and delivers the aa-tRNA to the A site of the ribosome. Codon recognition by the aa-tRNA triggers GTP hydrolysis by eEF1A, releasing the factor, and enables the aa-tRNA to accommodate in the A site. Then, peptide bond forms between the peptidyl-tRNA in the P site and the aa-tRNA in the A site. In most cases, peptide bond formation occurs spontaneously at the peptidyl transferase center of the ribosome, but with some amino acids, for example, strings of consecutive prolines, it requires the help of eIF5A. eEF1A–GDP is recycled to its active GTP-bound form with the help of nucleotide exchange factor eEF1B. Following peptide bond formation, the ribosome moves along the mRNA by one codon in a process called translocation, which is facilitated by eEF2 in the GTP-bound state and the fungal-specific eEF3 in its ATP-bound state. Once the stop codon is reached, translation is terminated with the help of the release factors eRF1 and eRF3 (Fig. 1) [1, 5]. Ribosome recycling, which requires Rli1, Ligatin and can also be promoted by eIF3j [6–8], results in the disassembly of the ribosome posttermination complex into ribosomal subunits, tRNA, and mRNA.

In this chapter, we describe the reconstitution of yeast translation system to study the initiation and elongation phases. In detail, we provide updated purification protocols for individual translation factors, ribosomes, and aa-tRNAs. Furthermore, we describe several assays for testing the activity of the purified components, the efficiency of the initiation complex formation and of peptide bond formation. This in vitro translation system utilizes an unstructured model mRNA, which overcomes the requirement for mRNA capping and for auxiliary cap-binding proteins.

## 2 Materials 2.1 Media 1. YPD media: 10 g/L yeast extract, 20 g/L peptone, 2% glucose (1.8% agar-agar for plates). 2. Selective media: 5.1 g/L (NH4)2SO4, 1.7 g/L yeast nitrogen base, 2% glucose, 0.67 g/L minimal media powder (-LEU -URA or -LEU) with 1.8% agar-agar for plates. 3. LB-media: 10 g/L tryptone; 10 g/L NaCl; 5 g/L yeast extract. 2.2 Antibiotics, DNase, and IPTG 1. 100 mg/mL ampicillin sodium salt. 2. 34 mg/mL chloramphenicol. 3. 2 units/μL DNase. 4. 1 mM puromycin. 5. 1 M Isopropyl β-D-1-thiogalactopyranoside (IPTG). 2.3 Buffers 1. Buffer 1: <sup>10</sup> Ribo Buffer A: 200 mM HEPES–KOH pH 7.5, 1 M KOAc, 25 mM Mg(OAc)2. 2. Buffer 2: 1 Buffer 1, 1 mg/mL heparin sodium salt, 2 mM DTT. 3. Buffer 3: 1 Buffer 1, 400 mM KCl, 1 M sucrose, 2 mM DTT. 4. Buffer 4: 1 Buffer 1, 400 mM KCl, 1 mg/mL heparin sodium salt, 2 mM DTT. 5. Buffer 5: 50 mM HEPES–KOH pH 7.5, 500 mM KCl, 2 mM MgCl2, 2 mM DTT. 6. Buffer 6: Same as buffer 5 with 5 mM MgCl2, 0.1 mM ethylenediaminetetraacetic acid (EDTA) pH 8.0, 5% or 30% sucrose. 7. Buffer 7: 50 mM HEPES–KOH, 100 mM KCl, 250 mM sucrose, 2.5 mM MgCl2, 2 mM DTT. 8. Buffer 8: 20 mM HEPES–NaOH pH 7.5, 150 mM NaCl, 5% glycerol, 4 mM ß-mercaptoethanol (ß-me). 9. Buffer 9: Same as buffer 8 with 1 M NaCl.


#### 2.4 Strains and Plasmids



#### 3 Methods

3.1 Purification of Ribosomal Subunits from S. Cerevisiae


3.2 Purification of Initiation Factors Unless specified otherwise, all purifications are performed using an FPLC system.

1. eIF1

Express eIF1 from pETT22b plasmid (no tag) in BL21 E. coli cells. Inoculate a preculture of 200 mL LB with 100 μg/mL carbenicillin and grow at 37 C overnight. Next day, inoculate 6 L of LB with 100 μg/mL carbenicillin with the overnight preculture at a starting OD600 of 0.1 and grow in a shaker at 37 C with 180 rpm. Induce expression with 0.5 mM IPTG when OD600 reaches 0.8 and incubate at 37 C for additional 3–4 h. Harvest the cells at 7000 rpm for 10 min at 4 C and store at 80 C. Resuspend the cell pellet in buffer 8 (3 mL/g of cells) complemented with 100 μL of DNase and one protease inhibitor tablet. Homogenize the cell suspension on ice and use Emulsiflex to lyse the cells. Centrifuge at 180,800 g for 1 h at 4 C. Collect the supernatant and filter it through 1 μm glass fiber filters or equivalent. Equilibrate the HiTrap SP column with buffer 8. Load the sample onto the column in buffer 8. Wash the column with buffer 8. Elute with a linear gradient from 0% to 100% of buffer 9 over 60 mL. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF1. Dilute the pool to decrease the salt concentration to 150 mM NaCl with buffer 11. Equilibrate the HiTrap Heparin column with buffer 8. Load the diluted sample and wash the column with buffer 8. Elute eIF1 with a linear gradient from 0% to 100% of buffer 9 over 60 mL. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF1. Concentrate eIF1 and load on a HiLoad 26/600 Superdex75 pg size-exclusion chromatography column preequilibrated with buffer 10. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF1. Concentrate the eIF1 using a 5 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 3105 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C.

2. eIF1A

Express eIF1A from pGEX-6P1 plasmid (GST tag) in BL21 E. coli cells. Inoculate a pre-culture of 200 mL LB with 100 μg/ mL carbenicillin and grow at 37 C overnight. Next day, inoculate 6 L of LB with 100 μg/mL carbenicillin with the overnight preculture at a starting OD600 of 0.1 and let it grow in a shaker at 37 C with 180 rpm. Reduce the temperature from 37 C to 16 C when OD600 reaches 0.4, induce expression with 0.5 mM IPTG when OD600 reaches 0.8 and grow for additional 16 h. Harvest the cells at 7000 rpm for 10 min at 4 C. Resuspend the cell pellets (30–50 g) in 2 mL/g with buffer 12 complemented with 100 μL of DNase and one protease inhibitors tablet. Homogenize the cell suspension on ice and use Emulsiflex to lyse the cells. Centrifuge at 180,800 g for 1 h at 4 C. Collect the supernatant and filter using 1 μm glass fiber filters or equivalent. Equilibrate the GSTrap column with buffer 13. Load the lysate and wash the column with buffer 13. Elute eIF1A with 100% of buffer 14 and collect 2 mL fractions. Analyze the fractions on a 15% SDS-PAGE. Pool the elution fractions and add PreScission protease (1 μM final) to cleave the tag while dialyzing overnight at 4 C against 2 L of buffer 13. Load the dialyzed sample on a Resource Q column and elute with 0 to 100% buffer 15 over 60 mL. Analyze the fractions on a 15% SDS-PAGE, pool the fractions containing eIF1A. Concentrate eIF1A and load on a HiLoad 26/600 Superdex75 pg sizeexclusion chromatography column preequilibrated with buffer 16. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF1A. Concentrate proteins using a 5 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 10,095 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C.

3. eIF5

Express eIF5 from pGEX-6P1 plasmid (GST tag) in BL21 E. coli cells. Inoculate a preculture of 200 mL LB with 100 μg/ mL carbenicillin and grow at 37 C overnight. Next day, inoculate 6 L of LB with 100 μg/mL carbenicillin with the overnight preculture at a starting OD600 of 0.1 and let it grow in a shaker at 37 C with 180 rpm. Reduce the temperature from 37 C to 16 C when OD600 reaches 0.4, induce expression with 0.5 mM IPTG when OD600 reaches 0.8 and grow for additional 16 h. Harvest the cells at 7000 rpm for 10 min at 4 C. Resuspend the cell pellets (30–50 g) in 2 mL/g with buffer 17 complemented with 100 μL of DNase and one protease inhibitors tablet. Homogenize the cell suspension on ice and use Emulsiflex to lyse the cells. Centrifuge at 180,800 g for 1 h at 4 C. Collect the supernatant and filter using 1 μm glass fiber filters or equivalent. Equilibrate the GSTrap column with buffer 17. Load the lysate and wash the column with buffer 17. Elute eIF5 with 100% of buffer 18 and collect 2 mL fractions. Analyze the fractions on a 15% SDS-PAGE. Pool the elution fractions and add PreScission protease (1 μM final) to cleave the tag while dialyzing overnight at 4 C against 2 L of buffer 19. Perform a second GSTrap column and eIF5 remains now in the flowthrough as the GST tag is cleaved. Concentrate eIF5 and load on a HiLoad 26/600 Superdex75 pg size-exclusion chromatography column preequilibrated with buffer 20. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF5. Concentrate proteins using a 10 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 30,285 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C. 4. eIF5A

Express eIF5A from pGEX-6P1 plasmid (GST tag) in Rosetta 2 cells. Inoculate a preculture of 200 mL LB with 100 μg/mL carbenicillin and 34 μg/mL chloramphenicol, and grow at 37 C overnight. Next day, inoculate 6 L of LB with 100 μg/mL carbenicillin with the overnight preculture at a starting OD600 of 0.1 and let it grow in a shaker at 37 C with 180 rpm. Induce eIF5A expression with 0.5 mM IPTG when OD600 reaches 0.8 and incubate at 37 C for additional 3–4 h. Harvest the cells at 7000 rpm for 10 min at 4 C. Resuspend the cell pellets (30–50 g) in 2 mL/g with buffer 21, complemented with 100 μL of DNase and one protease inhibitors tablet. Homogenize the cell suspension on ice and use Emulsiflex to lyse the cells. Centrifuge at 180,800 g for 1 h at 4 C. Collect the supernatant and filter using 1 μm glass fiber filters or equivalent. Equilibrate the GSTrap column with buffer 21. Load the lysate and wash the column with buffer 21. Elute eIF5A with buffer 22 and collect 2 mL fractions. Analyze the fractions on a 15% SDS-PAGE. Pool the elution fractions and add PreScission protease (1 μM final) to cleave the tag while dialyzing overnight at 4 C against 2 L of buffer 23. After dialysis, load eIF5A on a HiLoad 26/600 Superdex75 pg size-exclusion chromatography column preequilibrated with buffer 24. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF5A. Concentrate proteins using a 5 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 3105 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C.

5. eIF5B-397C

Express eIF5B-397C from pGEX-6P1 plasmid (GST-tag) in BL21 E. coli cells. Inoculate a preculture of 200 mL LB with 100 μg/mL carbenicillin and grow at 37 C overnight. Next day, inoculate 6 L of LB with 100 μg/mL carbenicillin with the overnight preculture at a starting OD600 of 0.1 and let it grow in a shaker at 37 C with 180 rpm. Induce eIF5B-397C expression with 0.5 mM IPTG when OD600 reaches 0.8 and incubate at 37 C for additional 3–4 h. Harvest the cells at 7000 rpm for 10 min at 4 C. Resuspend the cell pellets (30–50 g) in 2 mL/g with buffer 25, complemented with 100 μL of DNase and one protease inhibitors tablet. Homogenize the cell suspension on ice and use Emulsiflex to lyse the cells. Centrifuge at 180,800 g for 1 h at 4 C. Collect the supernatant and filter using 1 μm glass fiber filters or equivalent. Equilibrate the GSTrap column with buffer 25. Load the lysate and wash the column with buffer 25. Elute eIF5B-397C with 100% of buffer 26 and collect 2 mL fractions. Analyze the fractions on a 15% SDS-PAGE. Pool the elution fractions and add PreScission protease (1 μM final) to cleave the tag while dialyzing overnight at 4 C against 2 L of buffer 27. After dialysis, load eIF5A on a HiLoad 26/600 Superdex200 pg size-exclusion chromatography column preequilibrated with buffer 28. Collect 2 mL fractions for each, analyze on a 15% SDS-PAGE and pool the fractions containing eIF5B-397C. Concentrate proteins using a 30 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 46,215 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C.

6. eIF2

Streak eIF2 strain on a selective media (-LEU -URA) plate and incubate for 2 days at 30 C. Inoculate a preculture of 200 mL selective media (-LEU -URA) overnight in a shaker at 30 C with 180 rpm. Inoculate 100 L selective media (-LEU -URA) with the preculture at a starting OD600 of 0.1 and grow until mid-log phase in a bioreactor. Harvest the cells, wash the pellets with MilliQ water and suspend the cell pellets in 1 mL/g of cells in buffer 29. Prepare frozen cell droplets by dropping the lysate using a syringe or serological pipette in liquid nitrogen. Grind the cell droplets using an ultracentrifugal mill prechilled with liquid nitrogen. Collect the powder and store in 80 C freezer. Thaw the grinded yeast cell powder (~200 mL lysate in buffer 29) on ice and add one protease inhibitor tablet. Centrifuge at 180,800 g for 30 min at 4 C. Collect the supernatant into a fresh beaker and note down the volume. Stir the supernatant on ice or in the cold room. While stirring, gradually add grinded solid ammonium sulfate to 75% saturation (48.3 g ammonium sulfate/100 mL of lysate) and stir for 1 h. Centrifuge at 235,418 g for 1 h at 4 C. Discard the supernatant and resuspend the pellet in at least 200 mL of buffer 30. Filter the lysate through 1 μm glass fiber filters or equivalent. Equilibrate HisTrap columns with buffer 30. Load the sample onto the columns, wash the columns with buffer 30 and elute the proteins with 100% buffer 31. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF2. Dilute the pooled fractions five times by slowly adding buffer 35 with constant mixing to avoid precipitation. Equilibrate the Heparin column with buffer 32. Load the diluted sample immediately onto the Heparin column and wash with buffer 32. Elute eIF2 with a linear gradient from 0% to 100% of buffer 33 over 60 mL. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF2. Decrease the salt concentration of the pool to 100 mM KCl. Spin the sample for 20 min at 13,200 rpm at 4 C. Equilibrate the HiTrap Q column with buffer 32. Load the diluted sample and wash the column with buffer 32. Elute eIF2

with a linear gradient from 0% to 100% of buffer 33 over 60 mL. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF2. Concentrate the eIF2 protein with a 10 kDa MWCO cutoff concentrator and exchange the buffer with buffer 34. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 59,010 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C. 7. eIF3

Streak eIF3 strain on a selective media (-LEU -URA) plate and incubate for 2 days at 30 C. Inoculate a preculture of 200 mL selective media (-LEU -URA) overnight in a shaker at 30 C with 180 rpm. Inoculate 100 L selective media (-LEU -URA) with the preculture at a starting OD600 of 0.1 and grow until mid-log phase in a bioreactor. Harvest the cells, wash the pellets with MilliQ water and suspend the cell pellets in 1 mL/g of cells in buffer 36. Prepare frozen cell droplets by dropping the lysate using a syringe or serological pipette in liquid nitrogen. Grind the cell droplets using an ultracentrifugal mill prechilled with liquid nitrogen. Collect the powder and store in 80 C freezer. Thaw the grinded yeast cell powder (~200 mL lysate in buffer 36) on ice and add one protease inhibitor tablet. Centrifuge at 88,180 g for 15 min at 4 C. Collect the supernatant and filter through 1 μm glass fiber filters or equivalent. Equilibrate two HisTrap columns with buffer 36. Load the lysate on the columns; wash the columns with buffer 36 then with buffer 37. Elute the proteins with 100% buffer 38. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF3 subunits. Load eIF3 on a Hi Load 26/600 Superdex200 pg size-exclusion chromatography column preequilibrated with buffer 39. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eIF3. Concentrate the eIF3 with a 10 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 365,210 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C.

8. eEF1A

Streak the YAS2488 strain on a YPD plate and incubate for 2 days at 30 C. Inoculate a preculture of 200 mL YPD media overnight in a shaker at 30 C with 180 rpm. Inoculate 100 L YPD culture with the preculture at a starting OD600 of 0.1 and grow until mid-log phase in a bioreactor. Harvest the cells, wash the pellets with MilliQ water and suspend the cell pellets in 1 mL/g of cells in buffer 40. Prepare frozen cell droplets by dropping the lysate using a syringe or serological pipette in liquid nitrogen. Grind the cell droplets using an ultracentrifugal mill prechilled with liquid nitrogen. Collect the powder and store in 80 C freezer. Thaw grinded yeast cell powder (~200 mL lysate in buffer 40) on ice and add one protease inhibitor tablet. Check the pH of the lysate and adjust it to pH 7.7 by adding dropwise untitrated 1 M Tris to avoid protein precipitation. Centrifuge the lysate at 180,800 g for 1 h at 4 C. Collect the supernatant and filter it through 1 μm glass fiber filters or equivalent. Connect HiTrap Q column on top of HiTrap SP column as a tandem and equilibrate with buffer 41. Load the lysate onto the columns in tandem. Disconnect HiTrap Q column and wash HiTrap SP column with buffer 41 (eEF1A will bind to HiTrap SP and most impurities will bind to HiTrap Q). Elute the protein from the HiTrap SP column with a linear gradient of buffer 42 from 0% to 100% over 30 mL. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eEF1A. Load eEF1A on a Hi Load 26/600 Superdex200 pg size-exclusion chromatography column preequilibrated with buffer 43. Collect 2 mL fractions, analyze on a 15% SDS-PAGE and pool the fractions containing eEF1A. Concentrate the eEF1A pool with a 10 kDa MWCO Vivaspin concentrator. Check the final concentration at the spectrophotometer (OD280, extinction coefficient 45,295 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C (see Note 3).

9. eEF2

Streak eEF2 strain on a selective media (-LEU) plate and incubate for 2 days at 30 C. Inoculate a preculture of 200 mL selective media (-LEU) overnight in a shaker at 30 C with 180 rpm. Inoculate 100 L selective media (-LEU) with the preculture at a starting OD600 of 0.1 and grow until mid-log phase in a bioreactor. Harvest the cells, wash the pellets with MilliQ water and suspend the cell pellets in 1 mL/g of cells in buffer 44. Prepare frozen cell droplets by dropping the lysate using a syringe or serological pipette in liquid nitrogen. Grind the cell droplets using an ultracentrifugal mill prechilled with liquid nitrogen. Collect the powder and store in 80 C freezer. Thaw grinded yeast cell powder (~200 mL lysate in buffer 44) on ice and add one protease inhibitor tablet. Check the pH of the lysate and adjust it to pH 7.7 by dropwise addition of untitrated 1 M Tris to avoid protein precipitation. Centrifuge the lysate at 18,900 g for 20 min at 4 C. Collect the supernatant carefully, it should be as clear as possible. Centrifuge a second time in ultracentrifuge at 235,418 g for 1 h at 4 C. Collect the supernatant and filter it through 1 μm glass fiber filters or equivalent. Equilibrate two HisTrap columns with buffer 44 and load the lysate onto the columns. Wash the columns with buffer 45. Elute the protein with 100% buffer 46. Collect 2 mL fractions, analyze on a 12% SDS-PAGE and pool the fractions containing eEF2. Load eEF2 on a Hi Load 26/600 Superdex200 pg size-exclusion chromatography column preequilibrated with buffer 47. Collect 2 mL fractions, analyze on a 12% SDS-PAGE and pool the fractions containing eEF2. Concentrate the pool of eEF2 using 30 kDa MWCO Vivaspin concentrator. Measure the final concentration at the spectrophotometer (OD280, extinction coefficient 74,300 M<sup>1</sup> cm<sup>1</sup> ), flash-freeze aliquots, and store at 80 C (see Note 4).

Coexpress the proteins from pQLinkH plasmid (His tag) in BL21 E. coli cells. Inoculate a preculture of 200 mL LB with 100 μg/mL carbenicillin and grow at 37 C overnight. Next day, inoculate 6 L of LB with 100 μg/mL carbenicillin with the overnight preculture at a starting OD600 of 0.1 and grow it in a shaker at 37 C with 180 rpm. Induce the expression with 0.5 mM IPTG at 0.8 OD600 and incubate at 37 C for additional 4 h. Harvest the cells at 7000 rpm for 10 min at 4 C. Resuspend the cell pellet in 3 mL/ g buffer 48, add one protease inhibitor tablet and 100 μL of DNase solution. Homogenize the cell suspension on ice and use Emulsiflex to lyse the cells. Centrifuge at 180,800 g for 30 min at 4 C. Collect the supernatant and filter it through glass fiber filters or equivalent. Equilibrate two Protino Ni-IDA 2000 columns separately with buffer 48. Load the lysate over two tandem columns. Separate the columns and wash them with buffer 48. Elute each column with buffer 49 and collect 1 mL fractions in 1.5 mL Eppendorf tubes. Analyze the fractions on a 15% SDS-PAGE and pool the fractions containing Dys1 and Lia1. Dialyze the pool against 2 L of buffer 50 overnight at 4 C with stirring. Next day, perform a second dialysis with fresh 2 L of buffer 50 for 2–3 h at 4 C. Concentrate the pool using a 10 kDa MWCO Vivaspin concentrator. Measure the final concentration at the spectrophotometer (OD280), flash-freeze aliquots, and store at 80 C (see Note 6).

3.3.2 In Vitro Hypusination [14, 15] This is performed in a two-step process. In the first step, the synthesis of deoxyhypusine is catalyzed by deoxyhypusine synthase Dys1 using the reaction described in Table 1.

Mix components as indicated in Table 1 and incubate at 37 C for 1 h. Exchange the buffer on a NAP-column equilibrated with buffer 51. Load the sample and collect the flow-through. Add 1 mL of buffer 51 and collect the eluent. Repeat the elution step once more. Analyze the eluents on a 15% SDS-PAGE and pool the fractions containing eIF5A.

In the second step, the deoxyhypusine hydroxylase Lia1 catalyzes the final step of hypusination.

Mix the components indicated in Table 2 and incubate at 37 C for 2 h. Purify modified eIF5A from the enzyme mix using Protino Ni-IDA 2000 column. Equilibrate the column with buffer 52. Load the reaction sample and wash the column two times with

#### 3.3 In Vitro Hypusination of eIF5A

3.3.1 Purification of eIF5A Hypusination Enzymes Deoxyhypusine Synthase (Dys1) and Deoxyhypusine Hydroxylase (Lia1) (See Note 5)

#### Table 1 In Vitro Hypusination Step 1


#### Table 2 In Vitro Hypusination Step 2


4 mL of buffer 52. Elute three times with 3 mL buffer 53 with a fraction size of 1 mL. Analyze the fractions on a 15% SDS-PAGE and pool the fractions containing eIF5A.

Load eIF5A on a Superdex75 10/300 GL size exclusion chromatography column preequilibrated with buffer 52. Collect fractions of 250 μL, analyze on a 15% SDS-PAGE and pool the fractions containing eIF5A. Concentrate the pool using a 5kDA MWCO Vivaspin concentrator. Measure the final concentration at the spectrophotometer (OD280), flash-freeze aliquots, and store at 80 C.


AAGCGCGCAGGGCTCATAACCCTGATGTCCTCGGA TCGAAACCGAGCGGCGCTACCA30 .


#### Table 3 In Vitro Transcription Setup

Amplify the DNA using the following forward and reverse primers for in vitro transcription.

Forward primer: 5<sup>0</sup> TAATACGACTCACTATAAGCGCCG3<sup>0</sup> .

Reverse primer: 5<sup>0</sup> TmGmGTAGCGCCGCTCGGTTTC3<sup>0</sup> (see Note 2).

Perform in vitro transcription using the reaction components as described in Table 3:

Purify the transcription product on a HiTrap Q column. Load the transcription product on a HiTrap Q column preequilibrated with buffer 57. Elute the product with a linear gradient from 0% to 100% buffer 58 over 120 mL. Collect 2.5 mL fractions, analyze on a 12% UREA-PAGE and pool the fractions containing tRNAi Met. Precipitate tRNAi Met by adding 1/10 volume of 200 mM KOAc pH 5 and 2.5 volumes of ice-cold ethanol at 20 C overnight. The next day, centrifuge at 3901 g for 60 min at 4 C to pellet tRNAi Met. Dry and dissolve the pellet in water.

Perform the aminoacylation reaction at 37 C for 30 min using the components indicated in Table 4.

Quench the reaction by adding 1/10 volume of 200 mM KOAc pH 5. Extract Met-tRNAi Met by adding 1 volume of phenol (RNA grade) and collect the upper aqueous phase after 10 min centrifugation at 3901 g at room temperature. Precipitate Met-tRNAi Met by adding 1/10 volume of 200 mM KOAc pH 5 and 2.5 volumes of ice-cold ethanol at 20 C overnight. The next day, centrifuge at 3901 g for 60 min at 4 C to pellet Met-tR-NAi Met. Dry and dissolve the pellet in water. Purify Met-tRNAi Met by HPLC using a LiChrospher WP 300 RP-18 (5 μM) reverse phase column preequilibrated with buffer 60. Load the product and elute with a linear gradient from 0% to 100% buffer 61 over 255 mL. Collect 3 mL fractions and count the radioactivity of each fraction. Pool the fractions containing radioactivity and precipitate as before. Finally, dissolve the pellet in water and measure


#### Table 4 Aminoacylation Reaction Setup

concentration by dividing obtained [<sup>3</sup> H] radioactivity by the specific activity of [<sup>3</sup> H]-Methionine to obtain the pmol of methionine incorporated. Divide obtained pmol by the volume of water used to dissolve the final pellet to obtain the final concentration of [<sup>3</sup> H] Met-tRNAi Met [16].

Perform aminoacylation of individual tRNAVal and tRNAPhe using the components indicated in Table 4 with [14C]Valine or [ 14C]Phenylalanine, respectively. Purify aa-tRNA by HPLC as described for Met-tRNAi Met.

The mRNA used was purchased from IBA; the unstructured 5<sup>0</sup> UTR is underlined, the initiation codon is indicated in bold and is followed by the valine and phenylalanine codons.

50 GGUCUCUCUCUCUCUCUCUAUGGUUUUUUCU-CUCUCUCUC3<sup>0</sup> .

3.5 Nucleotide Binding and Dissociation Assay for eIF5B-397C and eEF1A To monitor the nucleotide binding and dissociation ability of eIF5B-397C and eEF1A, we used a fluorescence change that is observed upon mant-GTP binding to and dissociation from eIF5B-397C or eEF1A [17]. The experiments were carried out in a stopped-flow apparatus. Fluorescence of mant was excited at 363 nm and measured after passing KV408 long-pass filters. We collected 5–7 individual traces for each experiment, averaged them and plotted against time.

For nucleotide binding assay, prepare 800 μL of 1 μM eIF5B-397C or 0.2 μM eEF1A and 800 μL of 1 μM mant-GTP in YT buffer (buffer 54). In a stopped-flow apparatus, rapidly mix the protein with mant-GTP solutions. Follow the time course of reaction for 10 s (eIF5B-397C) or 100 s (eEF1A) by monitoring the fluorescence change.

For nucleotide dissociation assay, incubate 0.2 μM eIF5B-397C or eEF1A with 5 μM mant-GTP in YT buffer (buffer 54) with 3 mM PEP and 1% PK, for 10 min at 26 C. Prepare 800 μL of 1 mM GTP in YT buffer (buffer 54). In a stopped-flow apparatus, collect the traces over 10 s (eIF5B-397C) or 100 s (eEF1A) by rapidly mixing equal volumes of reactants and monitoring the time courses of fluorescence change.

Incubate 4 μM eIF2 (2 over tRNA) in YT buffer conditions with 3 mM PEP, 1% PK, 1 mM DTT, 2 mM GTP for 10 min at 26 C. Start the reaction by adding 2 μM of [<sup>3</sup> H]Met-tRNAi Met and incubate at 26 C. Collect 10 μL samples before the addition of Met-tRNAi Met at (0 time) and after different incubation times points. Spot them on 0.2 μM nitrocellulose filters presoaked in YT buffer (buffer 54) and wash with 5 mL of YT buffer. Transfer the filters into a polyethylene scintillation vials, add 10 mL of Quickszint 361 (Zinsser Analytic) scintillation cocktail to each tube, mix well and count the radioactivity in a scintillation counter.

> Prepare the ternary complex by incubating 4 μM of eIF2 with 3 mM PEP, 1% PK, 1 mM DTT, 1 mM GTP in YT buffer (buffer 54) in a 80 μL reaction volume for 10 min at 26 C. Add [<sup>3</sup> H]MettRNAi Met to 2 μM and incubate for additional 5 min at 26 C. Separately, prepare 80 μL of 48S IC by mixing 1 μM 40S subunits with 5 μM mRNA (5 over 40S), 5 μM eIF1 (5 over 40S), 2 μM eIF3 (2 over 40S), 2.5 μM eIF1A (2.5 over 40S), 2.5 μM eIF5 (2.5 over 40S), 2 mM DTT, 0.25 mM spermidine, and 1 mM GTP in YT buffer. Incubate for 5 min at 26 C. Add 1.5 μM 60S subunit (1.5 over 40S) and 3 μM eIF5B (2 over 60S) and incubate for additional 5 min at 26 C. Mix the ternary complex to 48S IC and load on a Biosuite450 (WATERS) size-exclusion chromatography column preequilibrated by YT buffer (buffer 54) on a HPLC. Run the sample at 0.8 mL/min for 25 min in YT buffer (buffer 54) and follow the absorbance at 290 nm. Collect 0.5 min fractions (0.4 mL) and count the radioactivity for each fraction. Pool the fractions containing 80S IC, count the radioactivity, calculate the concentration (obtained [<sup>3</sup> H] radioactivity divided by the specific activity of [<sup>3</sup> H]-Methionine to obtain the pmol of methionine incorporated. Divide obtained pmol by the volume of 80S IC to obtain the final concentration), flash-freeze aliquots, and store at 80 C.

3.7.2 Method 2: Sucrose Cushion Centrifugation This method allows preparing larger quantities of complexes; all concentrations are doubled compare to Method 1. Prepare 500 μL of ternary complex by incubating 8 μM of eIF2 with 3 mM PEP, 1% PK, 1 mM DTT, 2 mM GTP for 10 min at 26 C in YT buffer. Add 4 μM [<sup>3</sup> H]Met-tRNAi Met and incubate for additional 5 min at 26 C. Separately, prepare 500 μL of 48S IC by mixing 2 μM 40S with 10 μM mRNA (5 over 40S subunits), 10 μM eIF-mix (mixture of initiation factors eIF1, eIF1A, eIF3, eIF5) (5 over 40S), 2 mM DTT, 0.25 mM spermidine and 1 mM GTP in YT

#### 3.6 eIF2–GTP–MettRNAiMet Ternary Complex Formation Assay

3.7 80S Initiation Complex Formation

3.7.1 Method 1: Size-Exclusion Chromatography buffer (buffer 54). Incubate for 5 min at 26 C, then add 3 μM 60S subunits (1.5 over 40S) and 6 μM eIF5B (2 over 60S) and incubate for 5 min at 26 C. Mix the ternary complex with the 48S IC and adjust the MgCl2 to a final concentration of 9 mM to stabilize the 80S IC.

On ice, add 300 μL of 1 M sucrose (prepared in buffer 55 containing 9 mM MgCl2 (YT9 buffer)) into TLS-55 centrifuge tubes and carefully layer 1 mL of 80S IC reaction on top without disturbing the sucrose. Centrifuge in a TLS-55 rotor at 259,000 g for 2 h at 4 C. Invert the tubes to remove the supernatant and carefully wipe off excess of liquid without perturbing the pellet. Place the tubes on ice and dissolve the pellets in buffer 55 containing 9 mM MgCl2. Resuspend the pellet gently, count the radioactivity, calculate the concentration (as in Method 1), flash-freeze aliquots, and store at 80 C.

3.8 Peptide Bond Formation Assay Prepare the ternary complex eEF1A–GTP–[14C]Val-tRNAVal by incubating 1 μM eEF1A, 0.1 μM eEF1Bα, 3 mM PEP, 1% PK, 1 mM DTT, 0.5 mM GTP in YT buffer (buffer 54) for 15 min at 26 C. Add 0.2 μM [14C]Val-tRNAVal (5 eEF1A:1 aa-tRNA) and incubate for 5 min at 26 C. Then add 2 μM modified eIF5A to the ternary complex. Separately, prepare 1 μM 80S IC in YT buffer containing 3 mM MgCl2 by diluting 80S IC by YT0 buffer (buffer 56). Mix equal volumes of 80S IC containing [3 H]Met-tRNAi Met with the eEF1A–GTP–[14C]Val-tRNAVal ternary complex. After the desired incubation times quench the reaction by adding KOH to a final concentration of 0.5 M. Release the peptides by alkaline hydrolysis for 45 min at 37 C, separate the reaction products by LiChrospher 100 RP-8 (5 μm) reversed-phase HPLC using buffer 62 and buffer 63, and quantified by double-label ([3 H] and [14C]) radioactivity counting [17]. To form tripeptides, additionally prepare the eEF1A–GTP–[14C]Phe-tRNAPhe ternary complex by incubating 1 μM of eEF1A, 0.1 μM of eEF1Bα, 3 mM PEP, 1% PK, 1 mM DTT, 2 mM GTP for 15 min at 26 C, in YT buffer (buffer 54). After incubation add 0.2 μM of [14C]Phe-tRNAPhe (5 eEF1A:1 aa-tRNA) and incubate for additional 5 min at 26 C. Then add 1 μM eEF2 and 4 μM eEF3 to ternary complex. Mix equal volumes of ribosomes containing dipeptidyl-tRNA with the ternary complex. After the desired incubation times, quench the reaction and analyze the peptide products as described above for the dipeptides.

#### 4 Notes

1. eIF5A contains the uncommon amino acid hypusine, which is derived from lysine in two modifying steps (deoxyhypusine synthase and deoxyhypusine hydroxylase).


#### Acknowledgments

We thank Prof. Marina Rodnina for critical reading of the manuscript, and Olaf Geintzer, Tessa Hu¨bner, Theresia Niese for expert technical assistance. We thank Profs. Ralf Ficner, Alan Hinnebusch, Jon R. Lorsch, and Terri G. Kinzy for plasmids to express and purify translation factors. This work is supported by the Deutsche Forschungsgemeinschaft (DFG) in the framework of the Schwerpunktprogram (SPP1784), and by the Max Planck Society.

#### References


translation initiation. Methods Enzymol 430: 111–145


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# INDEX

#### A


#### B



#### C


Karl-Dieter Entian (ed.), Ribosome Biogenesis: Methods and Protocols, Methods in Molecular Biology, vol. 2533, https://doi.org/10.1007/978-1-0716-2501-9, © The Editor(s) (if applicable) and The Author(s) 2022

#### 282 RIBOSOME BIOGENESIS: METHODS AND PROTOCOLS Index


#### D


#### E



#### F


#### G


#### H


#### I

#### J


RIBOSOME BIOGENESIS: METHODS AND PROTOCOLS Index 283

#### K


#### L


#### M


#### 284 RIBOSOME BIOGENESIS: METHODS AND PROTOCOLS Index


#### N


#### O


#### P

Partial rRNA modification............................................153 PCR...................................................45, 53, 57, 129–131, 134–136, 138, 139, 142, 143, 170, 172, 175, 176, 248, 250–253, 256


#### Q


#### R




#### S


#### 286 RIBOSOME BIOGENESIS: METHODS AND PROTOCOLS Index


#### T


#### U


RIBOSOME BIOGENESIS: METHODS AND PROTOCOLS Index 287

#### V


#### W


#### X


#### Y

