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 50 -to-30 -oriented manner [4–7]. During nucleolar maturation, the 50 domain of the 18S rRNA achieves a mature conformation, and the central domain is correctly positioned relative to the 50 domain. In contrast, the 30 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 30 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 ¼ 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 4 μM Slx9 for 1 h at 4 C (lanes 2–4 and 6). Pse1-Slx9 complex was subsequently incubated with 1.5 μM 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 α-Slx9 antibodies. L ¼ input. (b) RanGTP-independent release of eS26 from Kap123 importin. Immobilized GST-Kap123 on Glutathione Sepharose was washed and incubated with 4 μM 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 μM His6-RanQLGTP (left panel lanes 3–6) or 0.375 μM 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 α-eS26 and α-RanGTP antibodies. L ¼ 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 ¼ 5 μm. 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 bud20Δ and yrb2Δ

	- 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 ¼ 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 α-RanGTP and α-Crm1 antibodies. L ¼ 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 ¼ 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 μg 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 α-Mex67/Mtr2 antibody. L ¼ 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

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# 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] b The sizes of the 50 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 30 -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 50 -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 50 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 30 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 50 to the upstream priming sequence in the upstream oligonucleotide. For the downstream oligonucleotide, more than 40 nucleotides of sequence just 30 of the genes stop codon are added 50 of the priming sequence in reverse complement orientation. Thereby the priming sequence continues directly 30 after this reverse complement stretch of the genes 30 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 (20 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 20 -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/D0 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 D0 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 20 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 (ac4 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 m6 2A1781, m6 2A1782, and a single m7 G1575, and two base acetylation—ac4 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 ac4 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 ac4 C1280 and ac4 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 m5 C (5-methyl cytosine), and two m3 U (3-methyl uridine) (Fig. 1 and Table 4). Two m5 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 m1 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 m1 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 (m1 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 m5 C2870 Methylation

Yeast 25S rRNA contains two m5 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 m5 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 m5 C2870, loss of m5 C 2278 did not exhibit any defects in 60S biogenesis and 35S rRNA processing. As far as the biological role of m5 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 m1 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 m1 A2142 and m5 C 2278 reside in the domain IV of the 25S rRNA [66].

To understand how 25S/28S rRNA cytosine methylation (m5 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 m5 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 m3 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, m3 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 m5 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 m5 U residues [73]. This has been also confirmed for higher eukaryotes, whereas m5 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 m5 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 m5 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

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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

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 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 N6 -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 N6 -methyladenosine (m6 A) and N6 isopentenyladenosine (i6 A) [23, 24]. The m6 A-sensitive DNA enzymes have been shown to be generally applicable to analyze the presence of m6 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 50 -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 50 -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 30 -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].  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, 20 -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 (ac4 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 m3 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+ )

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. (b and c) 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.

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# 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 50 UTRs in the absence of eIFs [9], attachment of natural mRNAs requires a coordinated activity of eIF4F protein complex, which unwinds the 50 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 50 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 50 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 50 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].