Common principles of protein biosynthesis

A current theory states that it was RNA that stood at the beginning of life on our planet (Gesteland, R.F., Cech, T.R., 1999). Yet, the later appearing proteins outperformed RNA in so many fields that those early, self-sufficient RNA constructs are now extinct. Whereas RNA is composed by a combination of basically four different types of nucleotides and is relatively limited in its ability to form tertiary structure, proteins are highly flexible chains built from twenty amino acids that can fold in a larger variety of three-dimensional shapes. The amount of building blocks (twenty compared to only four in RNA – leaving aside base modifications) equips them with a high degree of adaptability to different tasks and might be the reason for their superiority to RNA in many fields ( Figure 1 ).

To build such a peptide chain, amino acids have to be linked in the correct order via peptide bond formation. Although there exist proteins which are able to catalyze peptide bonds, like the sortase (Mazmanian et al., 1999), in all kingdoms of life the responsibility for building these peptide chains lies in a ribonucleoprotein particle: a macromolecular machine called the ribosome.

General features of the ribosome

The ribosome consists of a large and a small subunit. Both are made up of ribosomal RNA (rRNA) and ribosomal proteins ( Figure 1 ). The basic mechanism of protein synthesis is very similar in all domains of life: A messenger RNA (mRNA) contains the sequence of the protein and is bound and read by the small ribosomal subunit in collaboration with specific transfer RNAs (tRNAs). tRNAs carry the amino acids and contain characteristic anticodons, which establish base pairing with the respective codons on the mRNA presented to them by the ribosome. Matching allows for addition of the amino acid to the growing peptide chain. After all amino acids have been added, the peptide chain is released from the ribosome and, if needed, further processed by other cell components to become the final, folded protein.

Although the three domains of life, bacteria, eukaryotes, and archaea, all possess ribosomes for protein synthesis, the composition of the ribosomes varies, leading to sometimes very different overall appearance of ribosomes from different kingdoms of life (Amunts et al., 2015; Behrmann et al., 2015; Dunkle et al., 2011; Melnikov et al., 2012; Ramrath et al., 2018) ( Figure 2 ).

The eukaryotic cytoplasmic 80S ribosome is larger than the bacterial 70S ribosome, containing additional RNA segments and additional proteins. Comparison reveals that the eukaryotic ribosome possesses the same conserved structures as the bacterial ribosome in its core (Figure 2A) and an outer shell where eukaryote-specific elements are located ( Figure 1 , Figure 2 ) (Melnikov et al., 2012).

Not only the ribosomes themselves differ by certain features and are differently sensitive to antibiotics (Yusupova and Yusupov, 2017), but also the interacting factors that render translation possible are for some stages of translation remarkably different (Andersen et al., 2006) and might be an expression of the way the ribosome has been optimized for its bacterial, archaean, or eukaryotic cell environment.

Anatomy of the (mammalian) ribosome

The size of the assembled 80S mammalian ribosome is about 4.3 MDa (Wool, 1979). Both subunits are made of ribosomal RNA (rRNA) and ribosomal proteins (Wool, 1979). Upon joining of the large 60S (50S in bacteria) subunit and the small 40S (30S in bacteria) subunit to the 80S ribosome (70S in bacteria), a functionally important compartment is formed, the so-called intersubunit space (Figure 1), which is one of the main sites of action during protein synthesis. Across it span three tRNA-binding sites, named A (Aminoacyl)-, P (Peptidyl)-, and E (Exit)-sites. Additionally, the ribosome has specific factor binding sites for the interaction with protein factors, like the P-stalk ( Figure 3 C-D).

rRNA is the catalytically active component of the ribosome

The rRNA possesses the catalytic activity to perform peptide bond formation and is the main player in protein synthesis. The large (60S) subunit contains three rRNA molecules: the 28S rRNA, the 5S rRNA and the 5.8S rRNA (Supplemental Figure 1, Supplemental Figure 2). The small (40S) subunit contains only one rRNA molecule, the 18S rRNA (Supplemental Figure 3). The rRNA regions responsible for mRNA-recognition, tRNA-binding and peptidyl-transfer are highly conserved in all kingdoms of life (Gesteland, R.F., Cech, T.R., 1999). Among these conserved regions are the sarcin ricin loop (SRL) on the large subunit, interacting with GTP-hydrolyzing protein factors that catalyze certain steps of translation. In the peptidyltransferase center (PTC), also located on the large subunit, the RNA alone is responsible for catalyzing the formation of the peptide bond (Nissen et al., 2000; Spahn et al., 2000). The small subunit 18S rRNA contains the decoding center (DC), which monitors correct tRNA-anticodon matching to the mRNA codon.

Different from bacterial rRNA, eukaryotic 18S rRNA and 28S rRNA contain several expansion segments, long elements of additional rRNA that are to a great deal responsible for the big difference in size between bacterial and mammalian ribosomes. The function of these expansion segments is not yet clear, and their structural investigation is hindered by their high flexibility and peripheral location, making it very difficult to obtain high-resolution structural information (Ramesh and Woolford, 2016; Yusupova and Yusupov, 2017). There is some evidence, however, that expansion segments may play a role in ribosome biogenesis (Ramesh and Woolford, 2016).

Despite the rRNA’s prominent role in translation, the ribosome would not function without ribosomal proteins. They are important for rRNA folding and assembly and stabilize the tertiary structure of rRNA and the ribosome’s functional centers. There are 33 ribosomal proteins on the 40S and 47 ribosomal proteins on the 60S subunit of the mammalian ribosome. Following a recent convention (Ban et al., 2014) universally conserved proteins are prefixed ‘u’, unique bacterial ones ‘b’, unique eukaryotic ones ‘e’, and unique archaeal ones ‘a’ (Supplemental Figure 4, Supplemental Figure 5).

The three-dimensional shape of the ribosome is optimized for its function

The 40S subunit can be morphologically divided into several regions, named after the 40S subunit’s resemblance in shape to a bird: Looking at it from the solvent site, on top is the 40S ‘head’ with its prominent ‘beak’, below follows the ‘neck’, and the ‘body’ is supplemented by the ‘platform’, ‘shoulder’ and ‘foot’ domains (Figure 3A). In this work, a rough division into two parts will be used: the 40S head (including beak) and the 40S body/platform, comprising the remaining domains. Importantly, the link between the 40S head and 40S body/platform is flexible and allows for intrasubunit motions. Visible from the intersubunit space, there are the three 40S tRNA binding sites that are distributed among the 40S head and 40S body/ platform: A, P, and E ( Figure 3 B). The rRNA residues that constitute the tRNA binding sites are well-conserved (Supplemental Figure 1, Supplemental Figure 2, Supplemental Figure 3).

The large (60S) subunit is characterized by several landmarks; The central protuberance, the P-stalk/stalk base and the L1 stalk ( Figure 3 C-D). From the solvent side, one can see the ribosomal exit tunnel, from which the newly synthesized protein emerges. The solvent side is to a large degree covered by expansion segment ES7, the largest expansion segment of the 28S rRNA ( Figure 3 C). Looking on the 60S from the intersubunit space reveals the A-, P-, and E-tRNA binding sites and the sarcin-ricin loop (SRL) ( Figure 3 D).

The main interaction partners of the ribosome

To carry out translation of an RNA template into a protein, the ribosome is dependent on a large set of molecules – mRNAs, tRNAs, protein factors and energy carriers.

Messenger RNA (mRNA) contains the protein sequence

The information on the sequence of amino acids is encoded in the DNA of a cell and has to be transferred to RNA first (paradigm of molecular biology (Crick, 1970)). Such RNA, which serves as template for protein synthesis, is called messenger RNA (mRNA). Its production involves transcription by polymerase II and processing steps such as splicing, polyadenylation and capping (Meister, 2011). The mRNA pool accessible to the ribosome is strictly controlled by a balance between production and decay, variation of polyadenylation, storage in P-bodies, and many other mechanisms that are not well understood yet (Meister, 2011). In eukaryotes, mature mRNA is characterized by a 5’ cap, a 5’ UTR (untranslated region), the coding region, a 3’ UTR, and a poly-A-tail ( Figure 4 ). Notably, the mRNA is intensely decorated with proteins that regulate its transport, stability, decay and translation, for example poly-A-binding proteins (PABPs). Thus, mRNA really is a messenger ribonucleoprotein (mRNP) (Mitchell and Parker, 2014). Moreover, mRNA forms secondary structure that influences its interaction with the ribosome ( Figure 4 B). The entirety of the cellular mRNAs produced for translation at a given time-point is called transcriptome, and the actually translated mRNAs in a cell at a given timepoint constitute the translatome.

Transfer RNAs (tRNAs) allow for exact selection and incorporation of amino acids

tRNAs are specialized RNA molecules that usually consist of around 76 nucleotides. Their role in translation is to deliver the amino acids to the ribosome. Structurally, a tRNA consists of two parts, top and bottom, which organize into an L-shaped tertiary structure (Gesteland, R.F., Cech, T.R., 1999).

The top half comprises the TΨC-arm and the acceptor arm with the CCA-end, which is loaded with the amino acid by specialized enzymes, aminoacyl-transferases. There are elongator tRNAs and initiator tRNAs. Initiator tRNAs (tRNAi) are always loaded with a methionine. The bottom half of a tRNA consists of the D-arm and the anticodon loop, which can decode the mRNA ( Figure 5 ). The tRNA nucleotides are often and extensively modified and these modifications are thought to be important for its structure and function (Meister, 2011). The life of tRNA outside of the ribosome is object of intensive research and much less is known about it than about its role on the ribosome. The modification of a tRNA, for example, is complex and includes big protein complexes (Dauden et al., 2017). Further, like the mRNA, the tRNA does not exist in the cytosol as free and naked RNA molecule. The eukaryotic cell operates an elaborate system that channels the tRNAs to and from the ribosome, involving aminoacyl-transferases and probably more factors (Andersen et al., 2006; Mirande, 2010; Stapulionis and Deutscher, 1995).

Translational GTPases tune the energy landscape of translation

Many protein factors are involved in translation by direct interaction with the ribosome, the mRNA or the tRNA. A special subgroup of protein factors involved in translation are translational GTPases. They are GTP-hydrolyzing proteins that control key steps of translation; for example the decoding-specific eukaryotic elongation factor 1A (eEF1A), or eukaryotic elongation facter 2 (eEF2), which is necessary for efficient translocation, eukaryotic release factor 3 (eRF3), which is responsible for stop-codon recognition and termination, and eukaryotic initiation factor 5B (eIF5B), which mediates subunit joining (see section ‘translation cycle’).

Structurally, they all have in common the GTP-hydrolyzing domain (G-domain), which forms an alpha-beta-propeller and contains the nucleotide binding pocket (Wittinghofer and Vetter, 2011). Five characteristic G-motifs (G1-G5) and three loops (P-loop, switch I, switch II) organize around the bound nucleotide and characterize the G-domain ( Figure 6 ; Supplemental Figure 6 shows alignment of the G-domain of EF-G/eEF2 from different species). The G1 motif is also known as Walker A motif and is located within the P-loop. It is responsible for phosphate binding. The Mg2+ that is required for nucleotide binding is positioned by the G2 motif and the G3 motif. The G2 motif is part of switch I, whereas the G3 motif (also known as Walker B motif), is located within switch II (Bourne et al., 1991; Wittinghofer and Vetter, 2011) (Figure 6). GTP-hydrolysis is achieved by a nucleophilic attack of an activated water molecule and is assisted by the SRL of the ribosome, which serves as GTPase-activating factor for the translational GTPases. Depending on how one interprets the role of a histidine of switch II (His 84 in eF-Tu, His 108 in eEF2) ( Figure 7 A), two mechanism are being proposed that lead there: 1) A general base mechanism, in which the histidine abstracts a proton from the water molecule ( Figure 7 B), and 2) a substrate-assisted mechanism, in which GTP itself abstracts a proton from the water molecule while the histidine serves as allosteric enhancer of this process (Figure 7C) (Liljas et al., 2011; Maracci and Rodnina, 2016; Schweins et al., 1995; Voorhees et al., 2010).

Ribosome dynamics reshapes interaction sites and defines functional states

Two main large scale motions can be observed in the assembled ribosomes of all domains of life: rotation of the small subunit relative to the large one and swivel of the small subunit head domain relative to the small subunit body/platform domain (Figure 8A-B). The combination of subunit rotation and head swivel, observed in early ribosomal structures (Frank and Agrawal, 2000; Horan and Noller, 2007; Valle et al., 2003), was termed ‘ratcheting’. Subunit rotation and head swivel result in a reshaping of the intersubunit space, leading to changes in the positions of bound tRNAs and factors: Subunit rotation is known to be associated with the movement of the tRNAs on the large subunit (Moazed and Noller, 1989; Munro et al., 2007), leading to hybrid tRNAs, meaning that the tRNA is bound to one tRNA binding site on the small ribosomal subunit and to another on the large ribosomal subunit (A/P, P/E tRNAs). Head swivel and rotation combined (‘ratcheting’) are characteristic for the so-called chimeric hybrid state, an intermediate of translocation (Ramrath et al., 2013; Ratje et al., 2010; Zhou et al., 2013, 2014), where hybrid tRNAs bind to different tRNA binding sites on the small ribosomal subunit, one on the body/platform and another on the head domain (ap/P, pe/E tRNAs). Moreover, a somewhat ratcheted conformation is also observed in inactive mammalian ribosomes bound to eukaryotic elongation factor 2 (eEF2) (Khatter et al., 2015; Voorhees et al., 2014).

A motion so far only observed in the eukaryotic 80S ribosome is rolling of the small subunit, a rotation around the small subunit’s long axis (Figure 8A) (Behrmann et al., 2015; Budkevich et al., 2014). Subunit rolling takes place after decoding and is the main “net” conformational change characterizing the transition from the pre- translocational to the post-translocational state (See section 2.5).

Besides these main motions, there are some flexible parts of the ribosome that have also been observed to adopt distinct conformations: One of them is the L1 stalk (helices H76, H77, H78, and protein uL1) on the 60S subunit (Figure 3C-D, Supplemental Figure 1, Supplemental Figure 2), which is known to adopt different positions depending on the functional state of the ribosome (Mohan and Noller, 2017; Spahn et al., 2004a). Another flexible part is the stalk base (Helices H42, H43, H44, and proteins uL10, uL11) on the 60S subunit (Figure 3C-D, Supplemental Figure 1, Supplemental Figure 2). It changes its conformation by moving towards the A site upon binding of different factors (Gao et al., 2007; Schuette et al., 2009; Spahn et al., 2004b).

In 2009, Munro at al. applied the principle of the hierarchical energy landscape (Frauenfelder et al., 1991) to the dynamics of translation (Munro et al., 2009) (Figure 8C). The ribosome’s conformational degrees of freedom are directly connected to the forward movement of tRNAs during translation (Munro et al., 2009). The interplay of thermodynamically spontaneous motions (like subunit rotation), stabilizing and destabilizing contributions of the tRNAs (e.g. before and after peptide bond formation (Valle et al., 2003)) and the binding of protein factors like translational GTPases results in an ordered, organized sequence of conformational changes of the ribosome, which are coupled to efficient transport of the tRNA2•mRNA module through the ribosome.

The key steps of mammalian translation are initiation, elongation, termination and recycling

Translation can be divided into four parts: 1) Initiation, which is the binding of the very first tRNA, positioning of the mRNA on the small ribosomal subunit, and joining of the large subunit; 2) iterative elongation cycles, wherein the amino acids coded for in the mRNA are successively connected to form the peptide chain; 3) termination, resulting in the release of the peptide chain from the ribosome, and 4) recycling, which is splitting of the associated subunits into small and large one such that they are ready to be fed in a new round of translation (Figure 9).

Initiation places the start codon in the P site by ribosome scanning

Initiation is the first step of translation. Here, the small subunit is prepared for joining with the large subunit (Figure 9, Figure 10). This preparation includes binding of an initiator tRNAi Met to the small subunit P site as well as binding of the mRNA and positioning of the mRNA start codon into the P site under assistance of initiation factors. In the next step, the large subunit joins this 40S•mRNA•tRNAi Met complex, the remaining initiation factors dissociate, and elongation can start with the incorporation of the tRNA carrying the amino acid that is coded for in the second codon.

Eukaryotic and bacterial systems profoundly differ at the stage of initiation. Bacterial 16S rRNA possesses a sequence, to which the characteristic Shine Dalgarno sequence of the mRNA aligns. This alignment facilitates positioning of the mRNA start codon in the P site of the 30S subunit (Shine and Dalgarno, 1974). Initiation factor 1 (IF1) binds at the 30S A site and induces a structural change in the 30S subunit (Allen et al., 2005; Carter et al., 2001; Milon et al., 2008). A ternary complex consisting of fMet-tRNAi fMet (the first amino acid in bacterial proteins is always formyl-methionine) and IF2•GTP binds to the P site, while initiation factor 3 (IF3) ensures that it is an initiator tRNA and not an elongator tRNA that binds (Milon et al., 2008). Positioning of the initiator tRNA in the P site is followed by joining of the large subunit, GTP-hydrolysis of IF2, and finally dissociation of the initiation factors (Simonetti et al., 2008).

In contrast, in eukaryotes, the positioning of the start codon into the 40S P site occurs after ‘ribosome scanning’ of the mRNA (Jackson et al., 2010; Kozak, 1999; Shine and Dalgarno, 1974). The protein machinery required for eukaryotic initiation is by far more complex than in bacteria, where only three initiation factors are needed (Hashem and Frank, 2018; Jackson et al., 2010; Kozak, 1999; Shirokikh and Preiss, 2018).

Eukaryotic initiation starts with the formation of the 43S preinitiation complex, which consists of the 40S subunit, the ternary complex eIF2•GTP•Met•tRNAi Met, eIF3, eIF1 and eIF1A. This 43S complex is ready to bind to the mRNA, which can be circularized via the 5’-bound eIF4F that connects to the 3’ end by binding to PABPs. eIF1 and eIF1A induce an open latch conformation of the 40S that facilitates mRNA binding, while eIF4G plays a key role in loading the mRNA on the 43S preinitiation complex via interactions with eIF3. The main role of the now assembled 48S complex is the scanning of the mRNA until finding a start codon in the correct environment (Jackson et al., 2010; Kozak, 1999). It is eIF1 that enables the 48S complex to discriminate the eligible start codon against other codons or start codons with poor nucleotide context. The establishment of codon- anticodon base pairing between the mRNA start codon and the tRNA leads to eIF1 dissociation, allowing GTP hydrolysis and dissociation of eIF2 under assistance of its GTPase activating protein eIF5. Finally, eIF5B mediates the joining of the 60S subunit and the assembly of the elongation competent 80S ribosome is completed when GTP hydrolysis of eIF5B leads to its dissociation from the 80S (Figure 10).

Additionally to this classical initiation pathway, in the eukaryotic system a group of RNA structures called IRESs (Internal ribosomal entry sites) are able to employ alternative initiation pathways partially or fully independent from the 5’ cap and/or initiation factors (Pelletier and Sonenberg, 1988; Yamamoto et al., 2017).

Elongation is at the heart of translation

Elongation is an iterative process which is aimed at polymerization of the peptide chain until all amino acids encoded in the mRNA are incorporated into the peptide chain. It consists of the steps decoding/tRNA selection, peptidyl transfer and translocation. At each codon between start and stop, the ribosome must undergo one full elongation cycle with the result of one amino acid being added to the peptide chain.

The very first elongation cycle takes place right after initiation and has as its starting point the assembled 80S ribosome with a Met- tRNAi Met in the P site. The A site is empty and must be occupied by a tRNA carrying the amino acid that is encoded next in the mRNA. The selection of the correct tRNA requires eEF1A (EF-Tu in bacteria), a translational GTPase which reaches the ribosome as ternary complex in association with a tRNA and GTP. The interaction of the ribosome with the ternary complex is referred to as ‘decoding’. Here, the tRNA anticodon is brought to the small subunit A site (‘decoding center’) and interacts with the mRNA codon. Correct base pairing in case of complementary codons triggers conformational changes of the ribosome that in turn result in GTP-hydrolysis and eEF1A dissociation (Budkevich et al., 2014; Schmeing and Ramakrishnan, 2009; Schuette et al., 2009; Voorhees et al., 2010).

The dissociation of eEF1A allows the tRNA to be fully accommodated in the 60S A site. There, the CCA-end of the tRNA with the aminoacylated amino acid is positioned in proximity to the CCA-end of the P-site tRNA that carries the first amino acid, or in later rounds of elongation the entire peptide chain built so far. During peptide bond formation, this amino acid/peptide chain from the P-site tRNA is transferred to the A-site tRNA. As consequence, the ribosome contains a peptidyl tRNA in the A site, a deacylated tRNA in the P site, and in case of later elongation rounds, also a deacylated E-site tRNA (Behrmann et al., 2015). The ribosomal conformation at this point is characterized by an unrotated (canonical) 40S subunit that is rolled relative to the 60S subunit (Budkevich et al., 2011, 2014). It is referred to as the classical PRE state, where PRE means pre- translocation.

Translocation prepares the ribosome for a new elongation cycle

While the main task of the first half of the elongation cycle is the polymerization of the peptide chain, the second half serves as the preparation for the next elongation cycle. Translocation is the movement of both tRNAs as well as the mRNA, forming the tRNA2•mRNA module, from the A- and P- to the P- and E sites, respectively. It is catalyzed by eEF2 (EF-G in bacteria) and leads from the pre-translocational (PRE) to the post-translocational (POST)-state ribosome. The POST-state ribosome can then accept a new tRNA in its A site. Alternatively, in case of the very last elongation cycle, the ribosome does not bind a new tRNA, but enters termination (see below).

eEF2/EF-G belongs to the group of translational GTPases and is a five-domain protein (Figure 11). Mammalian eEF2 is very similar to its homologs from other domains of life (Supplemental Figure 6) The G-domain (or domain I) is responsible for GTP hydrolysis and possesses the earlier described characteristic switch loops and G-motifs. Domains 2 and 3 are part of a bridge between the G-domain and the small ribosomal subunit when the factor is bound to the ribosome. Domain 4 is a mimicry of a tRNA anticodon arm and protrudes into the A-site of the ribosome. Domain 4 carries three functionally important loops: loop 1, facing the tRNA (Ramrath et al., 2013), loop 2, facing the small subunit body/platform (Ramrath et al., 2013), and loop 3 in the middle (Figure 11B). A unique feature of domain 4 of the eukaryotic and archaeal elongation factors eEF2 is a diphthamide modification on histidine 715 in mammalia (H699 in yeast) in loop 3 of domain 4 (Oppenheimer and Bodley, 1981).

EF-G/eEF2 function can be specifically targeted by antibiotic agents. Fusidic acid binds between the G-domain and domain 3 of EF-G. It occupies the place of the Pi after its release following GTP-hydrolysis (Figure 11) and prevents EF-G’s dissociation from the ribosome. The concrete impact of fusidic acid on eukaryotic eEF2 is not fully understood yet. Sordarin binds between domain 2 and 3 of eEF2 and prevents dissociation of eEF2 in yeast (Figure 11) (Spahn et al., 2004a).

The diphthamide modification on loop 3 of domain 4 of eEF2 is known for being the target of toxins like diphtheria toxin and exotoxin A. When it is ribosylated, eEF2 cannot function in translation (Davydova and Ovchinnikov, 1990; Oppenheimer and Bodley, 1981).

There exists a current model on translocation based on bacterial and eukaryotic structures, however, a detailed understanding of the mechanism has not been (fully) achieved yet. Studies of bacterial translocation intermediates reveal the following sequence of events:

First, tRNA movement on the large 50S subunit occurs spontaneously after peptide bond formation which alters tRNA affinities and drives subunit rotation (Cornish et al., 2008; Valle et al., 2003). This subunit rotation is reversible and coupled to fluctuations between classical A/A, P/P states and hybrid A/P, P/E states of the tRNAs and has as well been visualized in eukaryotic ribosomes (Agirrezabala et al., 2008; Behrmann et al., 2015; Blanchard et al., 2004; Budkevich et al., 2011; Moazed and Noller, 1989; Munro et al., 2007).

Only after the tRNAs have reached their hybrid positions, EF-G/ eEF2 comes into play. It contributes to the irreversibility and directionality of the translocation reaction. eEF2/EF-G probably binds to the rotated ribosome (Brilot, 2013). According to Rodnina et al., 1997, already at this early stage and before tRNA movement has started, GTP-hydrolysis takes place. The next state visualized is the TI (translocation intermediate)-POST state, where the tRNAs are already translocated on the 30S body/platform and adopt chimeric hybrid (ap/P, pe/E) positions. The ribosome at this state adopts a partly rotated conformation and exhibits a high-degree head swivel (Ramrath et al., 2013; Ratje et al., 2010; Zhou et al., 2014) (Figure 12). After completion of translocation, EF-G dissociates. The ribosome can now accept the next tRNA.

It is not clear how exactly EF-G/eEF2 and GTP-hydrolysis contribute to translocation. Two opposing models were suggested, 1) the ‘Brownian ratchet’ model, according to which binding of EF-G/ eEF2 to the ribosome suffices to deflect the ribosome’s natural, thermodynamically driven propensity to backrotate, resulting in translocation (reviewed in (Spirin, 2009)). 2) The power stroke model, in which the energy of GTP-hydrolysis is used by eEF2/EF-G to actively push the tRNAs in the direction of translocation (Rodnina et al., 1997; Chen et al., 2016).

Termination releases the peptide chain and is followed by recycling

Usually, the last codon of the mRNA is followed by a stop codon (UAA, UAG or UGA). When this stop codon is positioned into the A site, no canonical tRNA will match it. The stop codon is recognized by class-1 release factor eRF1 (RF1 and RF2 in bacteria) which facilitates release of the peptide chain from the P-site tRNA. The activity of eRF1 is supported by class-2 release factor eRF3 (RF3 in bacteria), which belongs to the group of translational GTPases.

After termination, the ribosomal subunits dissociate and are then reused for the next round of translation. This process is called ribosome recycling. In bacteria, ribosomal recycling factor (RRF), EF-G and IF3 act together to disassemble the 80S ribosome (Hirashima and Kaji, 1970). In eukaryotes, ABCE1 splits the 80S ribosome (Jackson et al., 2012; Pisarev et al., 2010). Ligatin (also known as eIF2D) and DENR (density regulated protein) have been found to promote dissociation of tRNA and mRNA from the small subunit in eukaryotes (Skabkin et al., 2013).

Some evidence points towards an alternative event after termination. It is referred to as ‘reinitiation’: Here, recycling and dissociation of the 60S subunit takes place as well, but the 40S subunit does not leave the mRNA. Instead, it continues with scanning and thus can translate a next open reading frame (ORF; see Figure 4B) (Jackson et al., 2012; Skabkin et al., 2013).

mRNA quality control prevents production of degenerated proteins

Corrupt mRNA will lead to defective protein products. To avoid mistakes in protein translation, the cell has developed multi-level control mechanisms to check for integrity and correctness of the mRNA. Already mRNA transcription and maturation are susceptible to errors, and control points at this level are there to detect and eliminate defective mRNA products. In the nucleus, aberrant mRNAs are degraded in the 5’ to 3’ direction by exoribonuclease Xrn2 and in the 3’ to 5’ direction by the nuclear exosome before reaching the cytoplasm (Fasken and Corbett, 2009). In the cytoplasm, the ribosome and protein factors are involved in controlling the translated mRNA in three main ways:

Nonsense-mediated mRNA decay takes place when there is a premature stop codon. Downstream of the premature stop codon, the exon junction complex contains the upstream frameshift proteins (UPF) 2 and 3. Upon recognition of the stop codon, UPF1 is recruited. The interaction of UPF1 with UPF2 activates the mRNA degradation process (Isken and Maquat, 2007).

Non-stop mRNA decay is induced by the absence of a stop codon. The ribosome is stalled at the 3’ end of the mRNA because termination factors are not recruited (Isken and Maquat, 2007). Such stalled ribosomes are recognized by a mechanism that is not yet clearly understood, but apparently involving Dom34•Hbs1 (Hilal et al., 2016; Tsuboi et al., 2012). The exosome is recruited to the ribosome and degrades the mRNA.

In some cases, the ribosome cannot continue translation due to stable secondary structure. The mechanism that rescues a ribosome in this situation is referred to as no-go mRNA decay. In yeast, a Dom34•Hbs1•GTP complex is able to mediate dissociation of such stalled ribosomes (Becker et al., 2011; Shoemaker et al., 2010).

Translation is subject to multiple layers of regulation

Translation offers many levers for regulation. The pool of ribosomes that can enter translation is determined by the production, processing and modification of rRNA and ribosomal proteins and their assembly in the nucleus. The actual synthesis of a translation- competent ribosome, followed by nuclear export and maturation in the cytoplasm, are long and complex processes and sensitive to many influences (Karbstein, 2011; Nierhaus, 1991; Wilson and Nierhaus, 2007). Their regulation controls the cell’s basic disposition to perform translation. Similarly crucial to the synthesis of ribosomes is the synthesis of factors that are involved in translation.

The actual accessibility of both ribosomes and factors is the next layer of control. For example, 80S monosomes were found to bind eEF2, but not to be translationally active (Khatter et al., 2015; Voorhees et al., 2014); the 40S subunit in these complexes is thus not accessible for translation initiation. Stability of an 80S•eEF2 complex has also been observed in vitro (Budkevich et al., 2014; Davydova et al., 1993). An example for the control of protein factors is the eukaryotic initiation factor eIF4E, which can be bound and thus inactivated by eIF4E-binding protein (4E-BP), whose affinity is regulated by post- translational phosphorylation; initiation is reduced then by the decreased availability of eIF4E (Bah et al., 2015; Marcotrigiano et al., 1999). Also eEF2 is known to be inactivated by phosphorylation (Ling and Ermolenko, 2016).

Besides such quite simple mechanisms that are based on the accessibility of substrates, there might exist a more complex machinery that is aimed to enhance the production of specific proteins and down-regulate others. Whereas in long-term, the proteome of a cell is controlled by transcription, dynamic, rapid adaption mechanisms could base on differential translation. How this works exactly is not clear; discussed is among others the existence of ‘specialized ribosomes’ (Xue and Barna, 2012). An indicator for a possible regulation mechanism is that ribosomal proteins can be modified post-translationally. The ribosomal protein with the longest history of investigations of its posttranslational modification is ribosomal protein eS6, which is phosphorylated and whose putative impact on translation is one of the questions raised in this thesis (see below). Also, the topology of translation in the cell and the composition and architecture of polyribosomes (see below) might be a dynamic mechanism to alter the translatome.

Actively translating ribosomes are organized into polysomes

It is the normal case that on one mRNA, multiple ribosomes are translating the message at the same time and that depending on the length of the mRNA, multiple initiation events happen before the first ribosome reaches the stop codon.

Such assemblies of multiple ribosomes on the same mRNA are called polyribosomes or polysomes. Each ribosome that is part of a polysome group is at some stage of translation (Figure 13). Elongation intermediates are the by far most often sampled states, because the elongation cycle must take place for each codon between start and stop in contrast to initation, which takes place only at the start codon, and termination, which takes place only at the stop codon.

By isolating polysomes ex vivo, Behrmann et al. were able to reconstruct ten different states of mammalian elongation (Behrmann et al., 2015). The differential population of the states reflects the distribution of functional states in polysomes. Although the time between purification of the polysomes used for analysis and the freezing of the sample might allow for changes, that work shows the in vivo energy landscape of translation as close to the native state as possible. Of special interest for studying the regulation of translation is the distribution of functional states (Figure 13). The captured states by far don’t represent the complete spectrum of possible conformational and functional states of the mammalian ribosome during elongation, but only local minima. Referring to the nomenclature of the ‘dynamics‘ section, the states represent fluctuations in the macro- to mesoscopic range (Munro et al., 2009).

The shapes of polysomal assemblies have been extensively studied (Afonina et al., 2013, 2014; Brandt et al., 2010; Myasnikov et al., 2014; Viero et al., 2015). The relative orientations of the ribosomes in a polysomal assembly raise the question of how ribosomal proteins or RNA on the solvent side might promote these orientations via specific inter-ribosomal contacts. Moreover, it was observed that the abundance of rectangular shapes is associated with a low translation activity (Myasnikov et al., 2014; Viero et al., 2015) and that it can be increased by treatment of cells with rapamycin or by serum- withdrawal (Viero et al., 2015). Change in the ratio of ribosomes to mRNA can also modify the pattern of proteins synthesized (Lodish, 1974). Polysome patterns after post mitochondrial supernatant (Duncan and McConkey, 1982) as well as investigations of the shapes of polysomes (Myasnikov et al., 2014; Viero et al., 2015), suggest that polysomes react to different conditions, like serum withdrawal, by changing their shape and/or composition.

In eukaryotic cells, polysomes can be additionally differentiated by the cellular compartment they are attached to. Essentially, there are cytosolic polysomes and endoplasmatic reticulum (ER)-bound polysomes. So far, it is agreed upon that every ribosome has the potential to join either class and that it is the signaling peptide encoded in the mRNA that is responsible for recruiting the ribosome to the ER membrane (Meister, 2011).

Phosphorylation of ribosomal protein eS6: a possible switch for translation regulation?

Among the many discussed possibilities of how translation might be regulated, there are post-translational modifications of ribosomal proteins (Xue and Barna, 2012). Some ribosomal proteins undergo phosphorylation, e.g. ribosomal protein eS6 (eS6), a eukaryote- specific protein of the 40S subunit located at the foot of the 40S body/platform (Figure 3A). eS6 is 249 residues long in humans and consists of a globular part and an alpha helical part with a flexible C-terminus (Figure 14).

eS6 is a necessary protein for ribosome biogenesis. Its conditional deletion has been shown to lead to a p53-dependent inhibition of cell cycle progression (Panić et al., 2006; Sulić et al., 2005; Volarevic, 2000) in multiple tissues.

The alpha helical, C-terminal part of eS6 reaches into the vicinity of expansion segment 6 of the 18S rRNA and displays five serine residues (S235, S236, S240, S244, S247) at its C-terminus, which are the substrates of four independent kinases (RSK, Protein kinase A, S6K and CK1) with partially overlapping specificities (Bandi et al., 1993; Krieg et al., 1988; Wettenhall and Morgan, 1984) (Figure 15).

Phosphorylation happens in an ordered fashion, such that first, the residues serine 235 and serine 236 are phorsphorylated, then serine 240 and serine 244 and last serine 247 (Martin-Pérez and Thomas, 1983; Wettenhall et al., 1992). Phosphorylation of serine 247 requires prior phosphorylation of serines 240/244, and phosphorylation of serine 247 in turn promotes/stabilizes phosphorylation of serines 240/244 (Hutchinson et al., 2011). The modification is counteracted by the action of phosphatase (PP) 1 which is responsible for dephosphorylation at all five sites (Barth-Baus et al., 2002; Belandia et al., 1994; Hutchinson et al., 2011; Li et al., 2012).

The phosphorylation of the five serine residues at the C-terminus is massively increased in regenerating liver cells by partial hepatectomy, which was the first model for studying eS6-phosphorylation (Gressner and Wool, 1974). In the context of growth, two pathways lead to eS6 phosphorylation: The PI3K/Akt/TCS/Rheb/mTORC1/ S6K pathway and the Ras/Raf/MEK/ERK/RSK pathway (Figure 16) (reviewed in (Meyuhas, 2008, 2015)). A large number of stimuli have been shown to induce eS6 phosphorylation via these pathways, including stimulation of serum-starved cells with serum (Meyuhas, 2008, 2015; Roux et al., 2007).

As to the consequences of this modification, many observations have been made that are thought to be associated with eS6 phosphorylation, but none of them can be explained mechanistically. In 2005, Ruvinsky and colleagues found that global protein synthesis is higher in knockin mouse embryonic stem cells (MEFs) (P-/-) compared to wild type MEFs, but these observations could not be reproduced by using S6K-deficient mice (S6K1 -/-, S6K2 -/-) (Chauvin et al., 2014; Mieulet et al., 2007; Ruvinsky et al., 2005). Many cell- types derived from eS6P-/- mice are significantly smaller than in the wild type, among them pancreatic beta cells, IL7-dependent cells from fetal livers, MEFs, and muscle myotubes (Granot et al., 2009; Ruvinsky et al., 2005, 2009). However, some cells have normal size, for example acinar cells from pancreas (Pende et al., 2004; Ruvinsky et al., 2005). eS6P-/- mice have impaired renal hypertrophy after unilateral nephrectomy (Xu, 2015). Interestingly, there seems to be a link to initiation, as phosphorylation of serine 247 promotes association with mRNA Cap-binding complex in vitro (Hutchinson et al., 2011), and phosphorylation of serines 240/244 is required for Cap binding (Roux et al., 2007). Further, by covariance analysis of a cryo-EM map of a 43S initiation complex, density variance in the eS6 region was shown to be related to density variance of initiation factors (Liao et al., 2015) (the sample analyzed was (Hashem et al., 2013)). Nevertheless, if there is a clear structural role of eS6 phosphorylation is not known.

Single particle cryo-EM as a tool in structural biology

In cryogenic transmission electron microscopy (cryo-EM), a two- dimensional (2D) projection is generated from a thin vitrified specimen through which an electron beam is sent. In order to reconstruct the three-dimensional (3D) volume, multiple viewing angles of the same object must be covered. To this end, the specimen can either be imaged from different angles by tilting it, as done in cryo-EM tomography, or a specimen that is imaged at a fixed angle of 0° must contain multiple copies of the object in different orientations. The latter approach is used in single particle cryo-EM, which will be discussed in the following paragraphs.

Construction of the transmission electron microscope

The interior of a transmission electron microscope (TEM) is evacuated. Electrons are emitted from the cathode, or electron gun (Figure 17A). Because electrons interact with an electromagnetic field, they can be focused using magnetic lenses. The condenser lens (Figure 17B) transforms the emitted electrons to a parallel beam. The following acceleration of the electrons in the column is directly coupled to a theoretical point resolution limit, as will be explained below. Nowadays, usually 120-300 kV microscopes are used for single particle cryo-EM. The object (Figure 17C) is inserted into the column via a vacuum lock. The incident wave is modified by the object such that it carries the information about the object’s structure. The ojective lens system (Figure 17D) focuses the scattered wave to form the real image. The focused beam forms the Fourier transform/diffraction pattern of the wave in the back focal plane (Figure 17E). A phase plate can be inserted here to shift the phases (Orlova and Saibil, 2011). As high frequency information is scattered at high angles, at this point the size of the objective aperture plays an important role (Figure 17F). The projector lens system contains several magnification lenses (Figure 17G). To exclude electrons of certain energies (e.g. those that lost energy and changed wavelength due to interactions with the sample), which would otherwise reduces image quality, an energy filter can be used. Finally, the electron wave reaches the detector (Figure 17H).

Cryogenic transmission electron microscopy allows imaging of biological specimens

The distinctive feature of cryo-EM versus other structural methods is that it allows imaging of biological samples at near-native conditions. The specimen is plunge frozen in liquid ethane, a process that happens so fast that the hydration state of the molecule and its native conformation are unaffected (Dubochet et al., 1988). The molecules are now embedded in a thin layer of amorphous ice and can be imaged in the microscope. Additionally to the basic elements of a TEM described above, the cryo-TEM must ensure that the specimen is steadily kept at a temperature of below -180° C. Therefore, the specimen is inserted into the column via a special cryo-holder where it is kept in liquid nitrogen. The microscope is cooled with liquid nitrogen or sometimes helium.

Interaction of the electrons with the sample

Electrons can interact with the specimen or pass through it without interaction. If the electron passes without interaction, no information about the specimen is gained, and it contributes to background. The changes that occur to the electron by interacting with the sample make it carry information about the sample. In case of interaction, two types are distinguished: elastic and inelastic scattering.

When scattered elastically, the electron is deflected by the Coulomb potential of the specimen atoms without loss of energy. The deflection is expressed in a change of phase of the electron wave (Figure 18). Elastic scattering is the main interaction of electrons with biological specimens, which consist of light atoms (H, C, N, O, P, S). Elastic scattering contributes to phase contrast.

In inelastic scattering, the passing electron transfers energy to the specimen and both energy and phase of the electron change (Figure 18).

The lower the kinetic energy of the electron (e.g., at low voltage), the higher is the probability of inelastic and elastic scattering events. The general probability of electron-sample interaction and thus signal-to-noise ratio can be enhanced by increasing the electron dose. However, transfer of energy from the electron to the sample is associated with sample damage (Egerton et al., 2004). Low dose conditions with mean exposure of 20-30 electrons per Å2 are therefore usually applied for cryo EM of biological samples to limit the damage. To obtain a meaningful signal nevertheless, cryo-EM single particle analysis uses many low dose measurements of copies of the sample, which are then summed up (Cheng et al., 2015; Frank et al., 1992).

Cryo-EM of biological specimens is dominated by phase contrast

Image detectors record the electrons that hit a pixel. Because of the negligible amplitude loss in biological material and under the necessary low dose conditions, there will be almost no contrast visible for an image taken by an ideal cryo-TEM in focus.

There are two ways to solve this problem:

1. To use heavy atoms to increase amplitude contrast. For example, in negative staining, uranylacetate is used to stain the outside of the particles, such that one obtains a ‘negative’ (as example, see Figure 45). This is often done for screening of a sample and is justified for determining the overall shape of a particle, but will not yield high resolution.

2. To optimize the (phase) contrast transfer function (CTF), which describes the relation between the original image and the one that is detected and contains microscope-specific information. It is the convolution of the sinus of the aberration function χ(f) (1) and an envelope function E(f) that represents the dampening of the signal (de Jong and Van Dyck, 1993) (2). The aberration function χ(f) depends on the spatial frequency f, the wavelength λ, the spherical aberration Cs, which is microscope-specific, and the defocus δ, which can be adjusted via the objective lens current (de Jong and Van Dyck, 1993) (1). The envelope function depends amongst others on the defocus and on the electron source.

Images are usually not taken in focus. Instead, a defocus (δ) is employed, which increases the phase shift of the scattered vs. the unscattered beam. The efficiency by which the CTF transfers information depends on the spatial frequency and is different for different defocus values (Figure 19). Some information is missing completely, namely at each zero crossing. To compensate for this selective loss of information, images are usually taken at different defocus values, e.g. in a range of -500 to -2500 nm. Later, the exact defocus is recalculated for each micrograph (sometimes, even for each particle) and used for correction of the images during processing.

Spherical aberration (Cs) means that the electron beam is not focused exactly by the objective lens; instead, different components meet the optic axis at different heights, contributing to phase contrast as well.

By adjusting the defocus, χ(f) can be optimized such that the transferred contrast is enhanced. The samples presented in this work were all imaged employing this approach of taking images in defocus for enhancing phase contrast.

Another way of enhancing phase contrast is borrowed from light microscopy and consists in the usage of a phase plate (the term ‘Volta phase plate’ is often used in cryo-EM, in light microscopy the term ‘λ/4-plate‘ is used). It delays part of the electron beam and thus enhances the phase shift such that it can be measured as intensity (Zernike, 1942). Phase plates are not essential for visualizing large molecular assemblies like the ribosome, but their optimization and introduction into the field of cryo-EM nowadays allows visualization of very small molecular complexes that can hardly be realized when employing defocus variation (Danev and Baumeister, 2017).

Direct electron detectors contributed a great deal to the ‘resolution revolution’

The establishment of direct electron detectors contributed a great deal to the improvement of resolution in cryo-EM. Single electrons can be detected by these cameras, which possess a very thin active layer from which individual diodes can read single pixels (Faruqi McMullan, 2010). The key is that the incoming electrons are recorded as change in the potential of the diodes and not first converted to light by scintillators as in old CMOS detectors. The point spread function (PSF) is therefore much smaller and the position of the incoming signal is recorded with a higher accuracy and a higher signal-to-noise ratio.

The current state of the art is that images are collected as multiple frames (movie mode). Thus, beam-induced motion of the sample can be compensated by realigning the movie frames, a process that is called motion correction (Cheng et al., 2015; Zheng et al., 2017) (Figure 20).

Speed is also an important factor: The K2 Summit camera (Gatan), for example, has a sampling rate of ~400 images per second, enabling the detection of single events (incoming electrons). Such event is recorded over the whole detector area, of which each pixel detects a different intensity. Additionally to an intensity maximum in a certain pixel, the surrounding (weaker) intensities are measured by the neighboring pixels and thus the position relative to the maximum inside the pixel itself (quadrant-wise) can be recovered. Superresolution mode takes advantage of this and leads to a theoretically halved pixel size (Figure 21).

Image processing leads from 2D projections to 3D volumes

The particle images that are recorded present 2D projections formed by sending a set of parallel beams through the specimen. The intensity recorded by each pixel can be interpreted as the sum or integral of the object’s intensities along the beam.

The information that a 2D projection image contains can be described by the sum of different wave functions. Low frequency information contributes to the overall shape of structures in the image, like the rough outline of a ribosomal particle, whereas high frequency information contributes tofine details, like the position of single residues.

The Radon transform (Radon, 1986 -originally published in 1917) describes the function that calculates the integrals of a 2D image at given angles and thus disassembles the 2D image in its projections. Importantly, if all angles are completely covered, an object can be completely reconstituted from its projections using the inverse Radon transform. The inverse Radon transform is the approach that in principle is used to obtain 3D reconstructions of the molecules imaged by cryo-TEM, only that the set of projections from a cryo- EM experiment is not 100% complete. Moreover, the data contains noise that might interfere with high-frequency information.

To carry out inverse Radon transform from experimental data, several methods can be used: 1) Filtered backprojection, where the 2D image is smeared along the projection axis in real space, 2) Fourier interpolation, where the Fourier transforms of the projection are interpolated in Fourier space, forming the Fourier transform of the reconstructed image, which is then inverse Fourier transformed. A high-pass filter (e.g., ramp filter) can be applied to the Fourier transforms to unblur the resulting image. 3) Algebraic reconstruction technique (ART), where the reconstruction problem is formulated as a large set of equations that are then iteratively solved (Kaczmarz, 1937).

However, for these methods, of which nowadays Fourier interpolation is most commonly used for cryo-EM reconstructions, the angles from which the projections were generated must be known. Therefore, a key procedure in single particle cryo-EM is the finding and optimization of the orientation parameters (see below).

Notably, a single projection image has a very low signal to noise ratio. Therefore, similar projections can be grouped to one image stack. Subsequent class averaging improves the signal-to-noise ratio. 2D classification of particle images is a good start when handling data of which the 3D structure is unknown. From these 2D classes or a selection, an ab initio reconstruction can be calculated by back- projecting the 2D images to guess a 3D volume, which over several rounds of comparison with the backprojections is improved.

In case that one has already an idea of the structure, this step can be skipped and one can proceed with refinement and sorting.

Refinement is the optimization of the orientation parameters. In the approach of projection matching, cross-correlation or maximum likelihood methods are used to compare the original projection images with computed projection images of the reconstructed volume/the reference.

Sorting means that one splits the dataset into subpopulations in case of heterogeneity of the sample. The ribosome is a good example for an object that gives quite heterogeneous datasets because of its intrinsic dynamic described above (subunit rotation, head swivel, etc.). Large differences can be sorted by adding a neutral reference, for example an empty, strongly filtered volume (Figure 22). Smaller, more local differences can be unveiled by masking the region of interest and comparing reference projections obtained through this mask (focused reassignment) (Penczek et al., 2006).

Pitfalls to avoid when calculating a structure are reference bias and overrefinement. Reference bias means that noise can align to features of any given reference volume and reproduce it (example: Mao et al., 2013). Therefore, the reference must be chosen with care, optimally it is a volume coming from the dataset itself by ab initio reconstruction, and it must be low pass filtered. Filtering is also important for avoiding overrefinement, that means that noise in the images aligns to fine structure of the reference projections and thus distorts the overall result.

The final maps and resolution limits of cryo-EM

Both the hardware (the microscopes and the image detectors) and the software that is used to reconstruct the 3D volume from 2D projections have been optimized in the past years and the slogan ‘resolution revolution’ has become very popular to describe the improving quality of single particle cryo-EM structures (Kuhlbrandt, 2014).

In analogy to light microscopy, the theoretical resolution limit of a transmission electron microscope is directly coupled to the wave length. In a 300 kV microscope, the electron gains a kinetic energy of 300 keV, and its wavelength is 1.969 pm according to formula (3) (Relativistic formula for the calculation of velocity, where λ is the wavelength, h is the Planck constant, E is the energy, m0 is the rest mass of the electron, E0 is the rest energy of the electron (Reimer and Kohl, 2008)). One possible way of estimating the resolution limit of a (light) microscope is to use Abbe’s equation (4) (Lipson et al., 1995). For a microscope with a numerical aperture NA=0.01 and light at the wavelength λ=1.969 pm, the theoretical point resolution limit would be ~0.99 Å: atomic resolution.

This example of Abbe’s equation illustrates the potential of an EM compared to a light microscope due to the difference in wavelength. However, for estimating the resolution of a final map, the point resolution of the microscope does not play any role. Instead, statistical measures are used.

The final map’s resolution is based on self-consistency of two raw (unfiltered) half maps: In Fourier space, the Fourier transforms of the half maps are compared pixel by pixel using cross correlation along the radius (representing the spatial frequency) of the Fourier shells. As result, correlation values (FSC) are found for each spatial frequency. The spatial frequency that falls below the threshold of 0.143 FSC is defined as the last spatial frequency with a sufficient correlation value, and thus all information that is in a higher frequency range is considered unreliable because it contains more noise than signal (Rosenthal and Henderson, 2003). The reciprocal spatial frequency corresponds to the resolution (Examples in the results section, Figure 28, Figure 40, Figure 50, Figure 51).

In practice, the resolution and, importantly, the quality of the final map that is obtained of a biological object depends on many factors. During the first half of 2018 (01/01/2018-01/07/2018), there was only one structure released in the EM-database (http://emsearch.rutgers. edu) from single particle cryo-EM of less than 2 Å resolution. It was the structure of beta-galactosidase at 1.9 Å (Bartesaghi et al., 2018). 423 structures had a resolution between 2-5 Å, and the number of structures greater than 5 Å was 221. The structure with the currently best resolution deposited in the EM-database is reported to have 1.6 Å resolution (Danev R, Yanagisawa H, Kikkawa M Cryo-EM structure of mouse heavy-chain apoferritin at 1.62 Å). Usually, biological objects reach a high resolution when they are very symmetric and exhibit a low degree of flexibility.

Finally, it is important to note that the estimation of the resolution of a cryo-EM structure is based on conventions which not the entire community agrees upon (van Heel and Schatz, 2005, 2017) and which are susceptible to distortion by improper refinement (e.g. ‘overrefinement’ or model bias). One number is not enough to entirely and reliably assess the quality of a cryo-EM reconstruction. Also the local resolution is important, especially local resolution of factors or any regions that are important for answering the biological question asked when imaging the given molecule.

The technical advances of the recent years resulting in an improved quality of cryo-EM maps makes it possible to model structures at near atomic resolution. De novo atomic modeling, so far only possible in X-ray crystallography, can now be done using cryo-EM. That means that at its current state, cryo-EM can be used to actually solve a structure.

Aim 1

To visualize and mechanistically understand mammalian translocation.

Translocation is one of the least understood processes in protein biosynthesis. Its correct completion is crucial for the continuation of the translation elongation cycle, and ultimately for protein synthesis. As upon translocation, the contacts between the ribosome and the tRNA2•mRNA- module extensively rearrange, it is a process which requires utmost accuracy. Efficient translocation depends on the action of the specialized GTPase eEF2; however, detailed insights into the mechanism, by which eEF2 catalyzes translocation is lacking and opposing hypotheses are vividly discussed: The Brownian ratchet model, in which eEF2 is supporting intrinsic conformational changes that lead to translocation, and the power stroke model, according to which eEF2, being a motor protein, actively moves the tRNAs in the direction of translocation (Chen et al., 2016; Liu et al., 2014; Rodnina et al., 1997; Spirin, 2009).

Moreover, structural knowledge on tRNA translocation is dominated by studies carried out in the bacterial system, and structural data on mammalian tRNA translocation does not exist. Therefore, this work is dedicated to studying how translocation is performed in the mammalian system, using an in vitro reconstitution of a rabbit 80S•tRNA2•mRNA•eEF2•GMPPNP complex. The goal is to obtain structures of mammalian translocation intermediates to characterize mammalian translocation and compare it to translocation in the bacterial and yeast system.

Aim 2

To investigate the 80S•tRNA2•mRNA•eEF2•GDP complex that is observed upon addition of eEF2•GTP to a programmed PRE complex.

To understand why eEF2 can stably bind to ribosomes and two tRNAs in vitro (Budkevich et al., 2014), and how this fits to the model of translocation, I look at an in vitro reconstituted rabbit 80S•tRNA2•mRNA•eEF2•GDP complex.

Aim 3

To revisit the mammalian polysome landscape to find out if serum deprivation influences the distribution of states.

Cryo-EM of actively translating polysomes gives insight into the energy landscape of mammalian translation (Behrmann et al., 2015). Among other stimuli, serum deprivation and subsequent serum restimulation has been hypothesized to influence translation, e.g. via changing polysome patterns (Duncan and McConkey, 1982; Viero et al., 2015). I want to investigate if overnight serum starvation and 30 minutes of serum deprivation can change the energy landscape of translation.

Aim 4

To look at the phosphorylation site of ribosomal protein eS6 and its possible impact on its surrounding structures to find out the role of eS6 phosphorylation.

The investigation of structural changes induced by eS6 phosphorylation is the fourth aim of this thesis. eS6 is a eukaryote-specific protein of the 40S subunit. It undergoes phosphorylation in response to various stimuli, including serum deprivation/restimulation. Surprisingly, there are almost no works tackling the role of eS6 phosphorylation from a structural perspective. Since structural information on the eukaryotic ribosome from crystal structures and cryo-EM density maps have emerged in the last years, eS6 could be structurally characterized in yeast and recently also in mammalian ribosomes. However, the last residues of the C-terminus of eS6, including all five phosphorylatable serines, are missing in the available structures, and similarly the neighbouring expansion segments are not well resolved, such that the phosphorylation and its structural consequences are not characterized yet (Behrmann et al., 2015; Ben-Shem et al., 2011). This thesis aims to investigate the C-terminal region of eS6 and its surrounding.

List of Materials

References

Abeyrathne, P.D., Koh, C.S., Grant, T., Grigorieff, N., & Korostelev, A.A. (2016). Ensemble cryo-EM uncovers inchworm-like translocation of a viral IRES through the ribosome. eLife, 5, e14874.

Adams, P.D., Afonine, P.V., Bunkóczi, G., Chen, V.B., Davis, I.W., Echols, N., Headd, J.J., Hung, L.-W., Kapral, G.J., Grosse-Kunstleve, R.W., et al. (2010). PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. Sect. D Biol. Crystallogr., 66, 213-221.

Afonina, Z.A., Myasnikov, A.G., Khabibullina, N.F., Belorusova, A.Y., Menetret, J.-F., Vasiliev, V.D., Klaholz, B.P., Shirokov, V.A., & Spirin, A.S. (2013). Topology of mRNA chain in isolated eukaryotic double-row polyribosomes. Biochem., 78, 445-454.

Afonina, Z.A., Myasnikov, A.G., Shirokov, V.A., Klaholz, B.P., & Spirin, A.S. (2014). Formation of circular polyribosomes on eukaryotic mRNA without cap-structure and poly(A)-tail: a cryo electron tomography study. Nucleic Acids Res., 42, 9461-9469.

Agirrezabala, X., Lei, J., Brunelle, J.L., Ortiz-Meoz, R.F., Green, R., & Frank, J. (2008). Visualization of the Hybrid State of tRNA Binding Promoted by Spontaneous Ratcheting of the Ribosome. Mol. Cell, 32, 190-197.

Alejo, J.L., & Blanchard, S.C. (2017). Miscoding-induced stalling of substrate translocation on the bacterial ribosome. Proc. Natl. Acad. Sci., 114, 201707539.

Allen, G.S., Zavialov, A., Gursky, R., Ehrenberg, M., & Frank, J. (2005). The Cryo-EM Structure of a Translation Initiation Complex from Escherichia coli. Cell, 121, 703-712.

Amunts, A., Brown, A., Toots, J., Scheres, S.H.W., & Ramakrishnan, V. (2015). The structure of the human mitochondrial ribosome. Science, 348, 95-98.

Andersen, C.B.F., Becker, T., Blau, M., Anand, M., Halic, M., Balar, B., Mielke, T., Boesen, T., Pedersen, J.S., Spahn, C.M.T., et al. (2006). Structure of eEF3 and the mechanism of transfer RNA release from the E-site. Nature, 443, 663-668.

Antoun, A., Pavlov, M.Y., Andersson, K., Tenson, T., & Ehrenberg, M. (2003). The roles of initiation factor 2 and guanosine triphosphate in initiation of protein synthesis. EMBO J., 22, 5593-5601.

Archer, S.K., Shirokikh, N.E., Beilharz, T.H., & Preiss, T. (2016). Dynamics of ribosome scanning and recycling revealed by translation complex profiling. Nature, 535, 570-574.

Bah, A., Vernon, R.M., Siddiqui, Z., Krzeminski, M., Muhandiram, R., Zhao, C., Sonenberg, N., Kay, L.E., & Forman-Kay, J.D. (2015). Folding of an intrinsically disordered protein by phosphorylation as a regulatory switch. Nature, 519, 106-109.

Ban, N., Beckmann, R., Cate, J.H.D., Dinman, J.D., Dragon, F., Ellis, S.R., Lafontaine, D.L.J., Lindahl, L., Liljas, A., Lipton, J.M., et al. (2014). A new system for naming ribosomal proteins. Curr. Opin. Struct. Biol., 24, 165-169.

Bandi, H.R., Ferrari, S., Krieg, J., Meyer, H.E., & Thomas, G. (1993). Identification of 40 S ribosomal protein S6 phosphorylation sites in Swiss mouse 3T3 fibroblasts stimulated with serum. J. Biol. Chem., 268, 4530-4533.

Bartesaghi, A., Aguerrebere, C., Falconieri, V., Banerjee, S., Earl, L.A., Zhu, X., Grigorieff, N., Milne, J.L.S., Sapiro, G., Wu, X., et al. (2018). Atomic Resolution Cryo-EM Structure of β-Galactosidase. Structure, 26, 848-856.e3.

Barth-Baus, D., Stratton, C.A., Parrott, L., Myerson, H., Meyuhas, O., Templeton, D.J., Landreth, G.E., & Hensold, J.O. (2002). S6 phosphorylation-independent pathways regulate translation of 5’-terminal oligopyrimidine tract-containing mRNAs in differentiating hematopoietic cells. Nucleic Acids Res., 30, 1919-1928.

Becker, T., Armache, J.-P., Jarasch, A., Anger, A.M., Villa, E., Sieber, H., Motaal, B.A., Mielke, T., Berninghausen, O., & Beckmann, R. (2011). Structure of the no-go mRNA decay complex Dom34–Hbs1 bound to a stalled 80S ribosome. Nat. Struct. Mol. Biol., 18, 715-720.

Behrmann, E., Loerke, J., Budkevich, T.V., Yamamoto, K., Schmidt, A., Penczek, P.A., Vos, M.R., Bürger, J., Mielke, T., Scheerer, P., et al. (2015). Structural Snapshots of Actively Translating Human Ribosomes. Cell, 161, 845-857.

Belandia, B., Brautigan, D., & Martín-Pérez, J. (1994). Attenuation of ribosomal protein S6 phosphatase activity in chicken embryo fibroblasts transformed by Rous sarcoma virus. Mol. Cell. Biol., 14, 200-206.

Belardinelli, R., Sharma, H., Peske, F., Wintermeyer, W., & Rodnina, M.V. (2016). Translocation as continuous movement through the ribosome. RNA Biol., 13, 1-7.

Ben-Shem, A., Jenner, L., Yusupova, G., & Yusupov, M. (2010). Crystal Structure of the Eukaryotic Ribosome. Science, 330, 1203-1209.

Ben-Shem, A., Garreau de Loubresse, N., Melnikov, S., Jenner, L., Yusupova, G., & Yusupov, M. (2011). The structure of the eukaryotic ribosome at 3.0 Å resolution. Science, 334, 1524-1529.

Berman, H.M. (2000). The Protein Data Bank. Nucleic Acids Res., 28, 235-242.

Bhat, M., Robichaud, N., Hulea, L., Sonenberg, N., Pelletier, J., & Topisirovic, I. (2015). Targeting the translation machinery in cancer. Nat. Rev. Drug Discov., 14, 261-278.

Blanchard, S.C., Kim, H.D., Gonzalez, R.L., Puglisi, J.D., & Chu, S. (2004). tRNA dynamics on the ribosome during translation. Proc. Natl. Acad. Sci., 101, 12893-12898.

Bommer, U., Burkhardt, N., Junemann, R., Spahn, C.M.T., Triana-Alonso, & F. Nierhaus, K.H. (1997). Ribosomes and polysomes. In Subcellular Fractionation: A Practical Approach, J. Graham and D. Rickwood, Eds., (Washington, DC: IRL Press), pp. 271-301.

Bourne, H.R., Sanders, D.A., & McCormick, F. (1991). The GTPase superfamily: conserved structure and molecular mechanism. Nature, 349, 117-127.

Brandt, F., Carlson, L.-A.A., Hartl, F.U., Baumeister, W., & Grünewald, K. (2010). The Three-Dimensional Organization of Polyribosomes in Intact Human Cells. Mol. Cell, 39, 560-569.

Brilot, A.F., Korostelev, A.A., Ermolenko, D.N., & Grigorieff, N. (2013). Structure of the ribosome with elongation factor G trapped in the pretranslocation state. Proc. Natl. Acad. Sci., 110, 20994-20999.

Budkevich, T., Giesebrecht, J., Altman, R.B., Munro, J.B., Mielke, T., Nierhaus, K.H., Blanchard, S.C., & Spahn, C.M.T. (2011). Structure and Dynamics of the Mammalian Ribosomal Pretranslocation Complex. Mol. Cell, 44, 214-224.

Budkevich, T.V., El’skaya, A.V., & Nierhaus, K.H. (2008). Features of 80S mammalian ribosome and its subunits. Nucleic Acids Res., 36, 4736-4744.

Budkevich, T.V., Giesebrecht, J., Behrmann, E., Loerke, J., Ramrath, D.J.F., Mielke, T., Ismer, J., Hildebrand, P.W., Tung, C.S., Nierhaus, K.H., et al. (2014). Regulation of the mammalian elongation cycle by subunit rolling: A eukaryotic-specific ribosome rearrangement. Cell, 158, 121-131.

Carter, A.P., Clemons, W.M., Brodersen, D.E., Morgan-Warren, R.J., Hartsch, T., Wimberly, B.T., & Ramakrishnan, V. (2001). Crystal structure of an initiation factor bound to the 30S ribosomal subunit. Science, 291, 498-501.

Chaisuparat, R., Rojanawatsirivej, S., & Yodsanga, S. (2013). Ribosomal Protein S6 Phosphorylation is Associated with Epithelial Dysplasia and Squamous Cell Carcinoma of the Oral Cavity. Pathol. Oncol. Res., 19, 189-193.

Chauvin, C., Koka, V., Nouschi, A., Mieulet, V., Hoareau-Aveilla, C., Dreazen, A., Cagnard, N., Carpentier, W., Kiss, T., Meyuhas, O., et al. (2014). Ribosomal protein S6 kinase activity controls the ribosome biogenesis transcriptional program. Oncogene, 33, 474-483.

Chen, J.Z., & Grigorieff, N. (2007). SIGNATURE: A single-particle selection system for molecular electron microscopy. J. Struct. Biol., 157, 168-173.

Chen, C., Cui, X., Beausang, J.F., Zhang, H., Farrell, I., Cooperman, B.S., & Goldman, Y.E. (2016). Elongation factor G initiates translocation through a power stroke. Proc. Natl. Acad. Sci., 113, 7515-7520.

Chen, J., Petrov, A., Tsai, A., O’Leary, S.E., & Puglisi, J.D. (2013). Coordinated conformational and compositional dynamics drive ribosome translocation. Nat. Struct. Mol. Biol., 20, 718-727.

Chen, V.B., Arendall, W.B., Headd, J.J., Keedy, D.A., Immormino, R.M., Kapral, G.J., Murray, L.W., Richardson, J.S., & Richardson, D.C. (2010). MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. Sect. D Biol. Crystallogr., 66, 12-21.

Cheng, Y., Grigorieff, N., Penczek, P.A., & Walz, T. (2015). A Primer to Single-Particle Cryo-Electron Microscopy. Cell, 161, 438-449.

Chou, F.-C., Sripakdeevong, P., Dibrov, S.M., Hermann, T., & Das, R. (2012). Correcting pervasive errors in RNA crystallography through enumerative structure prediction. Nat. Methods, 10, 74-76.

Connell, S.R., Takemoto, C., Wilson, D.N., Wang, H., Murayama, K., Terada, T., Shirouzu, M., Rost, M., Schüler, M., Giesebrecht, J., et al. (2007). Structural Basis for Interaction of the Ribosome with the Switch Regions of GTP-Bound Elongation Factors. Mol. Cell, 25, 751-764.

Cornish, P.V., Ermolenko, D.N., Noller, H.F., & Ha, T. (2008). Spontaneous Intersubunit Rotation in Single Ribosomes. Mol. Cell, 30, 578-588.

Crick, F. (1970). Central Dogma of Molecular Biology. Nature, 227, 561-563.

Cukras, A.R., Southworth, D.R., Brunelle, J.L., Culver, G.M., & Green, R. (2003). Ribosomal proteins S12 and S13 function as control elements for translocation of the mRNA:tRNA complex. Mol. Cell, 12, 321-328.

Czworkowski, J., & Moore, P.B. (1997). The conformational properties of elongation factor G and the mechanism of translocation. Biochemistry, 36, 10327-10334.

Czworkowski, J., Wang, J., Steitz, T.A., & Moore, P.B. (1994). The crystal structure of elongation factor G complexed with GDP, at 2.7 A resolution. EMBO J., 13, 3661-3668.

Danev, R., & Baumeister, W. (2017). Expanding the boundaries of cryo-EM with phase plates. Curr. Opin. Struct. Biol., 46, 87-94.

Dauden, M.I., Kosinski, J., Kolaj‐Robin, O., Desfosses, A., Ori, A., Faux, C., Hoffmann, N.A., Onuma, O.F., Breunig, K.D., Beck, M., et al. (2017). Architecture of the yeast Elongator complex. EMBO Rep., 18, 264-279.

Davydova, E.K., & Ovchinnikov, L.P. (1990). ADP-ribosylated elongation factor 2 (ADP-ribosyl-EF-2) is unable to promote translocation within the ribosome. FEBS Lett., 261, 350-352.

Davydova, E.K., Malinin, N.L., & Ovchinnikov, L.P. (1993). Ribosomes terminated in vitro are in a tight association with non-phosphorylated elongation factor 2 (eEF-2) and GDP. Eur. J. Biochem., 215, 291-296.

Dubochet, J., Adrian, M., Chang, J.J., Homo, J.C., Lepault, J., McDowall, A.W., & Schultz, P. (1988). Cryo-electron microscopy of vitrified specimens. Q. Rev. Biophys., 21, 129-228.

Duncan, R., & McConkey, E.H. (1982). Preferential Utilization of Phosphorylated 40‐S Ribosomal Subunits during Initiation Complex Formation. Eur. J. Biochem., 123, 535-538.

Dunkle, J.A., Wang, L., Feldman, M.B., Pulk, A., Chen, V.B., Kapral, G.J., Noeske, J., Richardson, J.S., Blanchard, S.C., & Cate, J.H.D. (2011). Structures of the Bacterial Ribosome in Classical and Hybrid States of tRNA Binding. Science, 332, 981-984.

Egerton, R.F., Li, P., & Malac, M. (2004). Radiation damage in the TEM and SEM. Micron, 35, 399-409.

Eliseev, B., Yeramala, L., Leitner, A., Karuppasamy, M., Raimondeau, E., Huard, K., Alkalaeva, E., Aebersold, R., & Schaffitzel, C. (2018). Structure of a human cap-dependent 48S translation pre-initiation complex. Nucleic Acids Res., 46, 2678-2689.

Emsley, P., & Cowtan, K. (2004). Coot: Model-building tools for molecular graphics. Acta Crystallogr. Sect. D Biol. Crystallogr., 60, 2126-2132.

Fasken, M.B., & Corbett, A.H. (2009). Mechanisms of nuclear mRNA quality control. RNA Biol., 6, 237-241.

Ferguson, A., Wang, L., Altman, R.B., Terry, D.S., Juette, M.F., Burnett, B.J., Alejo, J.L., Dass, R.A., Parks, M.M., Vincent, C.T., et al. (2015). Functional Dynamics within the Human Ribosome Regulate the Rate of Active Protein Synthesis. Mol. Cell, 60, 475-486.

Flis, J., Holm, M., Rundlet, E.J., Loerke, J., Hilal, T., Dabrowski, M., Bürger, J., Mielke, T., Blanchard, S.C., Spahn, C.M.T., et al. (2018). tRNA Translocation by the Eukaryotic 80S Ribosome and the Impact of GTP Hydrolysis. Cell Rep., 25, 2676-2688.e7.

Frank, J., & Agrawal, R.K. (2000). A ratchet-like inter-subunit reorganization of the ribosome during translocation. Nature, 406, 318-322.

Frank, J., Penczek, P., & Liu, W. (1992). Alignment, classification, and three-dimensional reconstruction of single particles embedded in ice. Scanning Microsc. Suppl., 6, 11-20; discussion 20-22.

Frank, J., Radermacher, M., Penczek, P., Zhu, J., Li, Y., Ladjadj, M., & Leith, A. (1996). SPIDER and WEB: Processing and Visualization of Images in 3D Electron Microscopy and Related Fields. J. Struct. Biol., 116, 190-199.

Frauenfelder, H., Sligar, S.G., & Wolynes, P.G. (1991). The energy landscapes and motions of proteins. Science, 254, 1598-1603.

Gao, H., Zhou, Z., Rawat, U., Huang, C., Bouakaz, L., Wang, C., Cheng, Z., Liu, Y., Zavialov, A., Gursky, R., et al. (2007). RF3 Induces Ribosomal Conformational Changes Responsible for Dissociation of Class I Release Factors. Cell, 129, 929-941.

Gao, Y.-G., Selmer, M., Dunham, C.M., Weixlbaumer, A., Kelley, A.C., & Ramakrishnan, V. (2009). The Structure of the Ribosome with Elongation Factor G Trapped in the Posttranslocational State. Science, 326, 694-699.

Gavrilova, L.P., & Spirin, A.S. (1971). Stimulation of non-enzymic translocation in ribosomes by p-chloromercuribenzoate. FEBS Lett., 17, 324-326.

Gavrilova, L.P., Kostiashkina, O.E., Koteliansky, V.E., Rutkevitch, N.M., & Spirin, A.S. (1976). Factor-free and factor-dependent systems of translation of polyuridylic acid by Escherichia coli ribosomes. J. Mol. Biol., 101, 537-552.

Gesteland, R.F., Cech, T.R., & Atkins, J.F. (1999). The RNA World (Cold Spring Harbor, New York). Cold Spring Harbor Laboratory Press.

Granot, Z., Swisa, A., Magenheim, J., Stolovich-Rain, M., Fujimoto, W., Manduchi, E., Miki, T., Lennerz, J.K., Stoeckert, C.J., Meyuhas, O., et al. (2009). LKB1 regulates pancreatic beta cell size, polarity, and function. Cell Metab., 10, 296-308.

Greber, B.J., Bieri, P., Leibundgut, M., Leitner, A., Aebersold, R., Boehringer, D., & Ban, N. (2015). The complete structure of the 55S mammalian mitochondrial ribosome. Science, 348, 303-308.

Gressner, A.M., & Wool, I.G. (1974). The stimulation of the phosphorylation of ribosomal protein S6 by cycloheximide and puromycin. Biochem. Biophys. Res. Commun., 60, 1482-1490.

Hallinan, T., Eden, E., & North, R. (1962). The Structure and Composition of Rat Reticulocytes. Blood, 20, 50-63.

Hansen, J.L., Schmeing, T.M., Moore, P.B., & Steitz, T.A. (2002). Structural insights into peptide bond formation. Proc. Natl. Acad. Sci. U. S. A., 99, 11670-11675.

Harauz, G., & van Heel, M. (1986). Exact Filters for General Geometry Three Dimensional Reconstruction. Optik (Stuttg)., 74, 146-156.

Hartman, H., & Smith, T.F. (2010). GTPases and the origin of the ribosome. Biol. Direct, 5, 36.

Hashem, Y., & Frank, J. (2018). The Jigsaw Puzzle of mRNA Translation Initiation in Eukaryotes: A Decade of Structures Unraveling the Mechanics of the Process. Annu. Rev. Biophys., 47, 125-151.

Hashem, Y., Des Georges, A., Dhote, V., Langlois, R., Liao, H.Y., Grassucci, R.A., Hellen, C.U.T., Pestova, T.V., & Frank, J. (2013). Structure of the mammalian ribosomal 43S preinitiation complex bound to the scanning factor DHX29. Cell, 153, 1108-1119.

van Heel, M., & Schatz, M. (2005). Fourier shell correlation threshold criteria. J. Struct. Biol., 151, 250-262.

Hilal, T., Yamamoto, H., Loerke, J., Bürger, J., Mielke, T., & Spahn, C.M.T. (2016). Structural insights into ribosomal rescue by Dom34 and Hbs1 at near-atomic resolution. Nat. Commun., 7, 13521.

Hirashima, A., & Kaji, A. (1970). Factor dependent breakdown of polysomes. Biochem. Biophys. Res. Commun., 41, 877-883.

Horan, L.H., & Noller, H.F. (2007). Intersubunit movement is required for ribosomal translocation. Proc. Natl. Acad. Sci. U. S. A., 104, 4881-4885.

Hutchinson, J.A., Shanware, N.P., Chang, H., & Tibbetts, R.S. (2011). Regulation of ribosomal protein S6 phosphorylation by casein kinase 1 and protein phosphatase 1. J. Biol. Chem., 286, 8688-8696.

Inoue-Yokosawa, N., & Kaziro, Y. (1974). The role of guanosine triphosphate in translocation reaction catalyzed by elongation factor G. Biochem. Biophys. Res. Commun., 60, 4321-4323.

Isken, O., & Maquat, L.E. (2007). Quality control of eukaryotic mRNA: Safeguarding cells from abnormal mRNA function. Genes Dev., 21, 1833-1856.

Jackson, R.J., Hellen, C.U.T., & Pestova, T.V. (2010). The mechanism of eukaryotic translation initiation and principles of its regulation. Nat. Rev. Mol. Cell Biol., 11, 113-127.

Jackson, R.J., Hellen, C.U.T., & Pestova, T.V. (2012). Termination and post-termination events in eukaryotic translation. Adv. Protein Chem. Struct. Biol., 86, 45-93.

Jagannathan, S., Nwosu, C., & Nicchitta, C.V. (2011). Analyzing mRNA Localization to the Endoplasmic Reticulum via Cell Fractionation. In Methods in Molecular Biology (Humana Press, Totowa, NJ), pp. 301-321.

de Jong, A.F., & Van Dyck, D. (1993). Ultimate resolution and information in electron microscopy II. The information limit of transmission electron microscopes. Ultramicroscopy, 49, 66-80.

Juhling, F., Morl, M., Hartmann, R.K., Sprinzl, M., Stadler, P.F., & Putz, J. (2009). tRNAdb 2009: compilation of tRNA sequences and tRNA genes. Nucleic Acids Res., 37, D159-D162.

Karbstein, K. (2011). Inside the 40S ribosome assembly machinery. Curr. Opin. Chem. Biol., 15, 657-663.

Khatter, H., Myasnikov, A.G., Natchiar, S.K., & Klaholz, B.P. (2015). Structure of the human 80S ribosome. Nature, 520, 640-645.

Kim, S.-H., Jang, Y.H., Chau, G.C., Pyo, S., & Um, S.H. (2013). Prognostic significance and function of phosphorylated ribosomal protein S6 in esophageal squamous cell carcinoma. Int. J. Cancer, 132, 19-30.

Kolupaeva, V.G., Unbehaun, A., Lomakin, I.B., Hellen, C.U.T., & Pestova, T.V. (2005). Binding of eukaryotic initiation factor 3 to ribosomal 40S subunits and its role in ribosomal dissociation and anti-association. RNA, 11, 470-486.

Kozak, M. (1999). Initiation of translation in prokaryotes and eukaryotes. Gene, 234, 187-208.

Kuhlbrandt, W. (2014). The Resolution Revolution. Science, 343, 1443-1444.

Kumar, P., Hellen, C.U.T., & Pestova, T.V. (2016). Toward the mechanism of eIF4F-mediated ribosomal attachment to mammalian capped mRNAs. Genes Dev., 30, 1573-1588.

Lee, A.S.Y., Kranzusch, P.J., Doudna, J.A., & Cate, J.H.D. (2016). eIF3d is an mRNA cap-binding protein that is required for specialized translation initiation. Nature, 536, 96-99.

Li, X., Mooney, P., Zheng, S., Booth, C.R., Braunfeld, M.B., Gubbens, S., Agard, D.A., & Cheng, Y. (2013). Electron counting and beam-induced motion correction enable near-atomic-resolution single-particle cryo-EM. Nat. Methods, 10, 584-590.

Li, Y., Mitsuhashi, S., Ikejo, M., Miura, N., Kawamura, T., Hamakubo, T., & Ubukata, M. (2012). Relationship between ATM and Ribosomal Protein S6 Revealed by the Chemical Inhibition of Ser/Thr Protein Phosphatase Type 1. Biosci. Biotechnol. Biochem., 76, 486-494.

Liao, H.Y., Hashem, Y., & Frank, J. (2015). Efficient Estimation of Three-Dimensional Covariance and its Application in the Analysis of Heterogeneous Samples in Cryo-Electron Microscopy. Structure, 23, 1129-1137.

Liljas, A., Ehrenberg, M., & Åqvist, J. (2011). Comment on “The mechanism for activation of GTP hydrolysis on the ribosome”. Science, 333, 37; author reply 37.

Ling, C., & Ermolenko, D.N. (2016). Structural insights into ribosome translocation. Wiley Interdiscip. Rev. RNA, 7, 620-636.

Liu, T., Kaplan, A., Alexander, L., Yan, S., Wen, J. Der, Lancaster, L., Wickersham, C.E., Fredrick, K., Fredrik, K., Noller, H., et al. (2014). Direct measurement of the mechanical work during translocation by the ribosome. Elife, 3, e03406.

Lodish, H.F. (1974). Model for the regulation of mRNA translation applied to haemoglobin synthesis. Nature, 251, 365-448.

Loerke, J., Giesebrecht, J., & Spahn, C.M.T. (2010). Multiparticle Cryo-EM of Ribosomes. Methods Enzymol., 483, 161-177.

Lomakin, I.B., & Steitz, T.A. (2013). The initiation of mammalian protein synthesis and mRNA scanning mechanism. Nature, 500, 307-311.

Mao, Y., Wang, L., Gu, C., Herschhorn, A., Désormeaux, A., Finzi, A., Xiang, S.-H., & Sodroski, J.G. (2013). Molecular architecture of the uncleaved HIV-1 envelope glycoprotein trimer. Proc. Natl. Acad. Sci. U.S.A., 110, 12438-12443.

Maracci, C., & Rodnina, M.V. (2016). Review: Translational GTPases. Biopolymers, 105, 463-475.

Marcotrigiano, J., Gingras, A.C., Sonenberg, N., & Burley, S.K. (1999). Cap-dependent translation initiation in eukaryotes is regulated by a molecular mimic of eIF4G. Mol. Cell, 3, 707-716.

Márquez, V., Wilson, D.N., Tate, W.P., Triana-Alonso, F., & Nierhaus, K.H. (2004). Maintaining the ribosomal reading frame: The influence of the E site during translational regulation of release factor 2. Cell, 118, 45-55.

Martin-Pérez, J., & Thomas, G. (1983). Ordered phosphorylation of 40S ribosomal protein S6 after serum stimulation of quiescent 3T3 cells. Proc. Natl. Acad. Sci. U. S. A., 80, 926-930.

Mastronarde, D.N. (2005). Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol., 152, 36-51.

Mazmanian, S.K., Liu, G., Ton-That, H., & Schneewind, O. (1999). Staphylococcus aureus sortase, an enzyme that anchors surface proteins to the cell wall. Science, 285, 760-763.

Meister, G. (2011). RNA biology: an introduction. Wiley-VCH.

Melnikov, S., Ben-Shem, A., Garreau de Loubresse, N., Jenner, L., Yusupova, G., & Yusupov, M. (2012). One core, two shells: bacterial and eukaryotic ribosomes. Nat. Struct. Mol. Biol., 19, 560-567.

Meyuhas, O. (2008). Chapter 1 Physiological Roles of Ribosomal Protein S6: One of Its Kind. Int. Rev. Cell Mol. Biol., 268, 1-37.

Meyuhas, O. (2015). Ribosomal Protein S6 Phosphorylation: Four Decades of Research. In International Review of Cell and Molecular Biology, (Elsevier Ltd), pp. 41-73.

Mieulet, V., Roceri, M., Espeillac, C., Sotiropoulos, A., Ohanna, M., Oorschot, V., Klumperman, J., Sandri, M., & Pende, M. (2007). S6 kinase inactivation impairs growth and translational target phosphorylation in muscle cells maintaining proper regulation of protein turnover. Am. J. Physiol. Physiol., 293, C712-C722.

Milon, P., Konevega, A.L., Gualerzi, C.O., & Rodnina, M.V. (2008). Kinetic checkpoint at a late step in translation initiation. Mol. Cell, 30, 712-720.

Mirande, M. (2010). Processivity of translation in the eukaryote cell: Role of aminoacyl-tRNA synthetases. FEBS Lett., 584, 443-447.

Mitchell, S.F., & Parker, R. (2014). Principles and Properties of Eukaryotic mRNPs. Mol. Cell, 54, 547-558.

Moazed, D., & Noller, H.F. (1989). Intermediate states in the movement of transfer RNA in the ribosome. Nature, 342, 142-148.

Mohan, S., & Noller, H.F. (2017). Recurring RNA structural motifs underlie the mechanics of L1 stalk movement. Nat. Commun., 8, 14285.

Munro, J.B., Altman, R.B., O’Connor, N., & Blanchard, S.C. (2007). Identification of Two Distinct Hybrid State Intermediates on the Ribosome. Mol. Cell, 25, 505-517.

Munro, J.B., Sanbonmatsu, K.Y., Spahn, C.M.T., & Blanchard, S.C. (2009). Navigating the ribosome’s metastable energy landscape. Trends Biochem. Sci., 34, 390-400.

Munro, J.B., Altman, R.B., Tung, C.-S., Sanbonmatsu, K.Y., & Blanchard, S.C. (2010). A fast dynamic mode of the EF-G-bound ribosome. EMBO J., 29, 770-781.

Murray, J., Savva, C.G., Shin, B.S., Dever, T.E., Ramakrishnan, V., & Fernández, I.S. (2016). Structural characterization of ribosome recruitment and translocation by type IV IRES. Elife, 5, e14690.

Myasnikov, A.G., Afonina, Z.A., Ménétret, J.-F., Shirokov, V.A., Spirin, A.S., & Klaholz, B.P. (2014). The molecular structure of the left-handed supra-molecular helix of eukaryotic polyribosomes. Nat. Commun., 5, 5294.

Nguyen, K., & Whitford, P.C. (2016). Steric interactions lead to collective tilting motion in the ribosome during mRNA-tRNA translocation. Nat. Commun., 7, 10586.

Nielsen, P.J., Duncan, R., & McConkey, E.H. (1981). Phosphorylation of Ribosomal Protein S6 Relationship to Protein Synthesis in HeLa Cells. Eur. J. Biochem., 120, 523-527.

Nierhaus, K.H. (1991). The assembly of prokaryotic ribosomes. Biochimie, 73, 739-755.

Nierhaus, K.H. (1993). Solution of the ribosome riddle: how the ribosome selects the correct aminoacyl-tRNA out of 41 similar contestants. Mol. Microbiol., 9, 661-669.

Nilsson, L., & Nygård, O. (1992). Reduced puromycin sensitivity of translocated polysomes after the addition of elongation factor 2 and non-hydrolysable GTP analogues. FEBS Lett., 309, 89-91.

Nissen, P., Hansen, J., Ban, N., Moore, P.B., & Steitz, T.A. (2000). The structural basis of ribosome activity in peptide bond synthesis. Science, 289, 920-930.

Ogle, J.M., Murphy IV, F.V., Tarry, M.J., & Ramakrishnan, V. (2002). Selection of tRNA by the ribosome requires a transition from an open to a closed form. Cell, 111, 721-732.

Oppenheimer, N.J., & Bodley, J.W. (1981). Diphtheria toxin. Site and configuration of ADP-ribosylation of diphthamide in elongation factor 2. J. Biol. Chem., 256, 8579-8581.

Orlova, E.V., & Saibil, H.R. (2011). Structural Analysis of Macromolecular Assemblies by Electron Microscopy. Chem. Rev., 111, 7710-7748.

Panić, L., Tamarut, S., Sticker-Jantscheff, M., Barkić, M., Solter, D., Uzelac, M., Grabusić, K., & Volarević, S. (2006). Ribosomal protein S6 gene haploinsufficiency is associated with activation of a p53-dependent checkpoint during gastrulation. Mol. Cell. Biol., 26, 8880-8891.

Pelletier, J., & Sonenberg, N. (1988). Internal initiation of translation of eukaryotic mRNA directed by a sequence derived from poliovirus RNA. Nature, 334, 320-325.

Penczek, P.A., Frank, J., & Spahn, C.M.T. (2006). A method of focused classification, based on the bootstrap 3D variance analysis, and its application to EF-G-dependent translocation. J. Struct. Biol., 154, 184-194.

Pende, M., Um, S.H., Mieulet, V., Sticker, M., Goss, V.L., Mestan, J., Mueller, M., Fumagalli, S., Kozma, S.C., & Thomas, G. (2004). S6K1(-/-)/S6K2(-/-) mice exhibit perinatal lethality and rapamycin-sensitive 5’-terminal oligopyrimidine mRNA translation and reveal a mitogen-activated protein kinase-dependent S6 kinase pathway. Mol. Cell. Biol., 24, 3112-3124.

Pestova, T.V., & Hellen, C.U.T. (2003). Translation elongation after assembly of ribosomes on the Cricket paralysis virus internal ribosomal entry site without initiation factors or initiator tRNA. Genes Dev., 17, 181-186.

Pettersen, E.F., Goddard, T.D., Huang, C.C., Couch, G.S., Greenblatt, D.M., Meng, E.C., & Ferrin, T.E. (2004). UCSF Chimera—A visualization system for exploratory research and analysis. J. Comput. Chem., 25, 1605-1612.

Pisarev, A.V., Skabkin, M.A., Pisareva, V.P., Skabkina, O.V., Rakotondrafara, A.M., Hentze, M.W., Hellen, C.U.T., & Pestova, T.V. (2010). The role of ABCE1 in eukaryotic posttermination ribosomal recycling. Mol. Cell, 37, 196-210.

Pulk, A., & Cate, J.H.D. (2013). Control of Ribosomal Subunit Rotation by Elongation Factor G. Science, 340, 1235970.

Punjani, A., Rubinstein, J.L., Fleet, D.J., & Brubaker, M.A. (2017). cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination. Nat. Methods, 14, 290-296.

Radon, J. (1986). On the determination of functions from their integral values along certain manifolds. IEEE Trans. Med. Imaging, 5, 170-176.

Ramesh, M., & Woolford, J.L. (2016). Eukaryote-specific rRNA expansion segments function in ribosome biogenesis. RNA, 22, 1153-1162.

Ramrath, D.J.F., Yamamoto, H., Rother, K., Wittek, D., Pech, M., Mielke, T., Loerke, J., Scheerer, P., Ivanov, P., Teraoka, Y., et al. (2012). The complex of tmRNA-SmpB and EF-G on translocating ribosomes. Nature, 485, 526-529.

Ramrath, D.J.F., Lancaster, L., Sprink, T., Mielke, T., Loerke, J., Noller, H.F., & Spahn, C.M.T. (2013). Visualization of two transfer RNAs trapped in transit during elongation factor G-mediated translocation. Proc. Natl. Acad. Sci., 110, 20964-20969.

Ramrath, D.J.F., Niemann, M., Leibundgut, M., Bieri, P., Prange, C., Horn, E.K., Leitner, A., Boehringer, D., Schneider, A., & Ban, N. (2018). Evolutionary shift toward protein-based architecture in trypanosomal mitochondrial ribosomes. Science, 362, eaau7735.

Ratje, A.H., Loerke, J., Mikolajka, A., Brünner, M., Hildebrand, P.W., Starosta, A.L., Dönhöfer, A., Connell, S.R., Fucini, P., Mielke, T., et al. (2010). Head swivel on the ribosome facilitates translocation by means of intra-subunit tRNA hybrid sites. Nature, 468, 713-716.

Reimer, L., & Kohl, H. (2008). Transmission electron microscopy: physics of image formation. Springer, 5th edition.

Rodnina, M.V., Savelsbergh, A., Katunin, V.I., & Wintermeyer, W. (1997). Hydrolysis of GTP by elongation factor G drives tRNA movement on the ribosome. Nature, 385, 37-41.

Rogg, H., Wehrli, W., & Staehelin, M. (1969). Isolation of mammalian transfer RNA. BBA Sect. Nucleic Acids Protein Synth., 195, 13-15.

Rohou, A., & Grigorieff, N. (2015). CTFFIND4: Fast and accurate defocus estimation from electron micrographs. J. Struct. Biol., 192, 216-221.

Rosenthal, P.B., & Henderson, R. (2003). Optimal determination of particle orientation, absolute hand, and contrast loss in single-particle electron cryomicroscopy. J. Mol. Biol., 333, 721-745.

Roux, P.P., Shahbazian, D., Vu, H., Holz, M.K., Cohen, M.S., Taunton, J., Sonenberg, N., & Blenis, J. (2007). RAS/ERK Signaling Promotes Site-specific Ribosomal Protein S6 Phosphorylation via RSK and Stimulates Cap-dependent Translation. J. Biol. Chem., 282, 14056-14064.

Roy, A., Kucukural, A., & Zhang, Y. (2010). I-TASSER: a unified platform for automated protein structure and function prediction. Nat. Protoc., 5, 725-738.

Rubinstein, J.L., & Brubaker, M.A. (2015). Alignment of cryo-EM movies of individual particles by optimization of image translations. J. Struct. Biol., 192, 188-195.

Ruvinsky, I., Sharon, N., Lerer, T., Cohen, H., Stolovich-Rain, M., Nir, T., Dor, Y., Zisman, P., & Meyuhas, O. (2005). Ribosomal protein S6 phosphorylation is a determinant of cell size and glucose homeostasis. Genes Dev., 19, 2199-2211.

Ruvinsky, I., Katz, M., Dreazen, A., Gielchinsky, Y., Saada, A., Freedman, N., Mishani, E., Zimmerman, G., Kasir, J., & Meyuhas, O. (2009). Mice Deficient in Ribosomal Protein S6 Phosphorylation Suffer from Muscle Weakness that Reflects a Growth Defect and Energy Deficit. PLoS One, 4, e5618.

Scheres, S.H.W. (2012). RELION: Implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol., 180, 519-530.

Schmeing, T.M., & Ramakrishnan, V. (2009). What recent ribosome structures have revealed about the mechanism of translation. Nature, 461, 1234-1242.

Schuette, J.-C., Murphy, F.V., Kelley, A.C., Weir, J.R., Giesebrecht, J., Connell, S.R., Loerke, J., Mielke, T., Zhang, W., Penczek, P.A., et al. (2009). GTPase activation of elongation factor EF-Tu by the ribosome during decoding. EMBO J., 28, 755-765.

Schuwirth, B.S., Borovinskaya, M.A., Hau, C.W., Zhang, W., Vila-Sanjurjo, A., Holton, J.M., & Cate, J.H.D. (2005). Structures of the Bacterial Ribosome at 3.5 A Resolution. Science, 310, 827-834.

Schweins, T., Geyer, M., Scheffzek, K., Warshel, A., Kalbitzer, H.R., & Wittinghofer, A. (1995). Substrate-assisted catalysis as a mechanism for GTP hydrolysis of p21ras and other GTP-binding proteins. Nat. Struct. Biol., 2, 36-44.

Selmer, M., Dunham, C.M., Murphy, F.V., Weixlbaumer, A., Petry, S., Kelley, A.C., Weir, J.R., & Ramakrishnan, V. (2006). Structure of the 70S ribosome complexed with mRNA and tRNA. Science, 313, 1935-1942.

Semenkov, Y.P., Rodnina, M.V., & Wintermeyer, W. (1996). The “allosteric three-site model” of elongation cannot be confirmed in a well-defined ribosome system from Escherichia coli. Proc. Natl. Acad. Sci. U. S. A., 93, 12183-12188.

Shine, J., & Dalgarno, L. (1974). The 3’-Terminal Sequence of Escherichia coli 16S Ribosomal RNA: Complementarity to Nonsense Triplets and Ribosome Binding Sites. Proc. Natl. Acad. Sci., 71, 1342-1346.

Shirokikh, N.E., & Preiss, T. (2018). Translation initiation by cap-dependent ribosome recruitment: Recent insights and open questions. Wiley Interdiscip. Rev. RNA, 9, e1473.

Shirokikh, N.E., Archer, S.K., Beilharz, T.H., Powell, D., & Preiss, T. (2017). Translation complex profile sequencing to study the in vivo dynamics of mRNA-ribosome interactions during translation initiation, elongation and termination. Nat. Protoc., 12, 697-731.

Shoemaker, C.J., Eyler, D.E., & Green, R. (2010). Dom34:Hbs1 promotes subunit dissociation and peptidyl-tRNA drop-off to initiate no-go decay. Science, 330, 369-372.

Shoji, S., Walker, S.E., & Fredrick, K. (2009). Ribosomal Translocation: One Step Closer to the Molecular Mechanism. ACS Chem. Biol., 4, 93-107.

Simonetti, A., Marzi, S., Myasnikov, A.G., Fabbretti, A., Yusupov, M., Gualerzi, C.O., & Klaholz, B.P. (2008). Structure of the 30S translation initiation complex. Nature, 455, 416-420.

Skabkin, M.A., Skabkina, O.V., Hellen, C.U.T., & Pestova, T.V. (2013). Reinitiation and other unconventional posttermination events during eukaryotic translation. Mol. Cell, 51, 249-264.

Spahn, C.M., Penczek, P.A., Leith, A., & Frank, J. (2000). A method for differentiating proteins from nucleic acids in intermediate-resolution density maps: cryo-electron microscopy defines the quaternary structure of the Escherichia coli 70S ribosome. Structure, 8, 937-948.

Spahn, C.M., Gomez-Lorenzo, M.G., Grassucci, R.A., Jørgensen, R., Andersen, G.R., Beckmann, R., Penczek, P.A., Ballesta, J.P., & Frank, J. (2004). Domain movements of elongation factor eEF2 and the eukaryotic 80S ribosome facilitate tRNA translocation. EMBO J., 23, 1008-1019.

Spahn, C.M.T., Jan, E., Mulder, A., Grassucci, R.A., Sarnow, P., & Frank, J. (2004). Cryo-EM visualization of a viral internal ribosome entry site bound to human ribosomes: the IRES functions as an RNA-based translation factor. Cell, 118, 465-475.

Spirin, A.S. (2009). The Ribosome as a Conveying Thermal Ratchet Machine. J. Biol. Chem., 284, 21103-21119.

Stapulionis, R., & Deutscher, M.P. (1995). A channeled tRNA cycle during mammalian protein synthesis. Proc. Natl. Acad. Sci. U. S. A., 92, 7158-7161.

Sulić, S., Panić, L., Barkić, M., Merćep, M., Uzelac, M., & Volarević, S. (2005). Inactivation of S6 ribosomal protein gene in T lymphocytes activates a p53-dependent checkpoint response. Genes Dev., 19, 3070-3082.

Susorov, D., Zakharov, N., Shuvalova, E., Ivanov, A., Egorova, T., Shuvalov, A., Shatsky, I.N., & Alkalaeva, E. (2018). Eukaryotic translation elongation factor 2 (eEF2) catalyzes reverse translocation of the eukaryotic ribosome. J. Biol. Chem., 293, 5220-5229.

Tang, G., Peng, L., Baldwin, P.R., Mann, D.S., Jiang, W., Rees, I., & Ludtke, S.J. (2007). EMAN2: An extensible image processing suite for electron microscopy. J. Struct. Biol., 157, 38-46.

Tourigny, D.S., Fernandez, I.S., Kelley, A.C., & Ramakrishnan, V. (2013). Elongation factor G bound to the ribosome in an intermediate state of translocation. Science, 340, 1235490.

Triana, F., Nierhaus, K.H., & Chakraburtty, K. (1994). Transfer RNA binding to 80S ribosomes from yeast: evidence for three sites. Biochem Mol Biol Int., 33, 909-915.

Tsuboi, T., Kuroha, K., Kudo, K., Makino, S., Inoue, E., Kashima, I., & Inada, T. (2012). Dom34:Hbs1 Plays a General Role in Quality-Control Systems by Dissociation of a Stalled Ribosome at the 3′ End of Aberrant mRNA. Mol. Cell, 46, 518-529.

UniProt Consortium, T.U. (2008). The universal protein resource (UniProt). Nucleic Acids Res., 36, D190-D195.

Valle, M., Zavialov, A., Sengupta, J., Rawat, U., Ehrenberg, M., & Frank, J. (2003). Locking and unlocking of ribosomal motions. Cell, 114, 123-134.

Viero, G., Lunelli, L., Passerini, A., Bianchini, P., Gilbert, R.J., Bernabò, P., Tebaldi, T., Diaspro, A., Pederzolli, C., & Quattrone, A. (2015). Three distinct ribosome assemblies modulated by translation are the building blocks of polysomes. J. Cell Biol., 208, 581-596.

Volarevic, S. (2000). Proliferation, But Not Growth, Blocked by Conditional Deletion of 40S Ribosomal Protein S6. Science, 288, 2045-2047.

Voorhees, R.M., Schmeing, T.M., Kelley, A.C., & Ramakrishnan, V. (2010). The Mechanism for Activation of GTP Hydrolysis on the Ribosome. Science, 330, 835-838.

Voorhees, R.M., Fernández, I.S., Scheres, S.H.W., & Hegde, R.S. (2014). Structure of the Mammalian Ribosome-Sec61 Complex to 3.4 Å Resolution. Cell, 157, 1632-1643.

Wasserman, M.R., Alejo, J.L., Altman, R.B., & Blanchard, S.C. (2016). Multiperspective smFRET reveals rate-determining late intermediates of ribosomal translocation. Nat. Struct. Mol. Biol., 23, 333-341.

Wettenhall, R.E.H., & Morgan, F.J. (1984). Phosphorylation of hepatic ribosomal protein S6 and 80 and 40 S ribosomes. Primary structure of S6 in the region of the major phosphorylation sites for cAMP-dependent protein kinases. J. Biol. Chem., 259, 2084-2091.

Wettenhall, R.E.H., Erikson, E., & Maller, J.L. (1992). Ordered multisite phosphorylation of Xenopus ribosomal protein S6 by S6 kinase II. J. Biol. Chem., 267, 9021-9027.

Whitford, P.C., Ahmed, A., Yu, Y., Hennelly, S.P., Tama, F., Spahn, C.M.T., Onuchic, J.N., & Sanbonmatsu, K.Y. (2011). Excited states of ribosome translocation revealed through integrative molecular modeling. Proc. Natl. Acad. Sci. U. S. A., 108, 18943-18948.

Wilson, D.N., & Nierhaus, K.H. (2007). The Weird and Wonderful World of Bacterial Ribosome Regulation. Crit. Rev. Biochem. Mol. Biol., 42, 187-219.

Wittinghofer, A., & Vetter, I.R. (2011). Structure-Function Relationships of the G Domain, a Canonical Switch Motif. Annu. Rev. Biochem., 80, 943-971.

Wool, I.G. (1979). The Structure and Function of Eukaryotic Ribosomes. Annu. Rev. Biochem., 48, 719-754.

Xue, S., & Barna, M. (2012). Specialized ribosomes: a new frontier in gene regulation and organismal biology. Nat. Rev. Mol. Cell Biol., 13, 355-369.

Yamamoto, H., Unbehaun, A., & Spahn, C.M.T. (2017). Ribosomal Chamber Music: Toward an Understanding of IRES Mechanisms. Trends Biochem. Sci., 42, 655-668.

Yang, J., Yan, R., Roy, A., Xu, D., Poisson, J., & Zhang, Y. (2015). The I-TASSER Suite: protein structure and function prediction. Nat. Methods, 12, 7-8.

Yusupova, G., & Yusupov, M. (2017). Crystal structure of eukaryotic ribosome and its complexes with inhibitors. Philos. Trans. R. Soc. B Biol. Sci., 372, 20160184.

Zernike, F. (1942). Phase contrast, a new method for the microscopic observation of transparent objects, Part I. Physica, 9, 974-986.

Zhang, K. (2016). Gctf: Real-time CTF determination and correction. J. Struct. Biol., 193, 1-12.

Zheng, S.Q., Palovcak, E., Armache, J.-P., Verba, K.A., Cheng, Y., & Agard, D.A. (2017). MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods, 14, 331-332.

Zhou, J., Lancaster, L., Donohue, J.P., & Noller, H.F. (2013). Crystal Structures of EF-G – Ribosome States of Translocation. Science, 340, 1235490.

Zhou, J., Lancaster, L., Donohue, J.P., & Noller, H.F. (2014). How the ribosome hands the A-site tRNA to the P site during EF-G-catalyzed translocation. Science, 345, 1188-1191.

Anatomy of the mammalian ribosome.

Bacterial translocation intermediates.

Electron-specimen interaction.

Elements of the transmission electron microscope.

Elongation cycle.

Features of the G-domain.

Messenger RNA (mRNA).

Model of ribosomal protein eS6

Motion correction.

Overview of the processes leading to translation initiation.

Overview of the translation cycle.

Pathways that lead to eS6 phosphorylation.

Ribosome dynamics and the energy landscape.

Ribosomes from different domains of life

Sites of ribosomal protein eS6 phosphorylation and dephosphorylation.

Structure of the bacterial translocase EF-G.

Superresolution mode of the K2 Summit camera (Gatan).

The CTF depends on the defocus.

The G-domain.

The human cytoplasmic 80S ribosome.

Transfer RNA.