ABSTRACT
 Title of Dissertation:                                
Dissertation Directed by:
 FROM RIBOSOMAL ROTATION TO AUTISM AND 
CANCER
 Sergey O. Sulima, Doctor of Philosophy, 2013
 Professor Jonathan D. Dinman 
Department of Cell Biology and Molecular Genetics 
The ribosome transits between two main conformational states ? non-rotated and 
rotated - as it progresses through the translation elongation cycle. How this transition is 
controlled is not known. Here, we present evidence that the essential ribosomal protein 
L10 regulates this process in yeast ribosomes through a flexible loop located within 13 ? 
of the peptidyltransferase center. While deletion of the entire loop is lethal, viable 
mutants in this region were found to promote opposing effects on the natural equilibrium 
between the two conformational states of the ribosome. Mutants causing rotational 
disequilibria show defects in essential ribosomal processes including ligand binding, 
peptidyltransfer, decoding, reading frame maintenance, and gene expression. Large-scale 
chemical modification analyses of rRNA of the mutant ribosomes identified networks of 
near-, medium-, and long-range allosteric interactions involved in coordinating 
intersubunit rotation. These allosteric pathways map from L10 in the core of the large 
subunit and propagate outward through both subunits, linking all of the functional centers 
of the ribosome.  Mutants of ribosomal protein L3 promoting opposing structural effects 
suppress the effects of the L10 mutants by re-establishing the correct rotational 
equilibrium. This loop is also involved in recruitment of the biogenesis factor Sdo1p as 
part of a quality control mechanism in which pre-60S subunits undergo a ?test drive? 
before maturation. This suggests that the correct rotational status is important for 
ensuring late-stage maturation of the large subunit, indicating that the L10 loop is a key
 regulator of ribosome function throughout the ribosomal life cycle. A model is presented 
describing how the unidirectionality of translation and ribosomal rotation are powered by 
this intrinsic mobile loop. Additionally, mutants in this element have been identified in a 
fraction of T-cell acute lymphoblastic leukemia patients, and these mutants were found to 
also favor the rotated state. This suggests that a rotational disequilibrium is at the heart 
of this disease, and a mechanistic model is presented that describes how a ribosomal 
mutation can lead to cancer, and identifies new treatments. Moreover, analysis of 
mutations in the essential C-terminal end of L10 linked with autism has provided
 functional insight into the mechanism of this disorder. 
FROM RIBOSOMAL ROTATION TO AUTISM AND CANCER
 By
 Sergey O. Sulima
 Dissertation submitted to the Faculty of the Graduate School of the
 University of Maryland, College Park, in partial fulfillment
 of the requirements for the degree of
 Doctor of Philosophy
 2013
 Advisory Committee:
 Professor Jonathan D. Dinman, Chair
 Associate Professor Volker Briken
 Professor James N. Culver 
Associate Professor Jason D. Kahn 
Associate Professor Douglas A. Julin, Dean?s Representative
? Copyright by
 Sergey O. Sulima
 2013
ii
 Preface
 There is poetry in the real world. Science is the poetry of reality. 
-Richard Dawkins 
iii
 Acknowledgements
 I would like to thank my advisor Dr. Dinman for years of mentorship, friendship, 
and particularly the insight to have a plastic bag in his car. A big thank you to all 
members of the Dinman lab, past and present, for a fun and cooperative work experience, 
especially the lab grandma. A special thank you goes to my parents for the nature and the 
nurture. 
This work was supported in part by a National Institute of Health grant to Dr. 
Jonathan D. Dinman and a National Institute of Health training grant in molecular and 
cell biology to Sergey O. Sulima.
iv
 Table of Contents
 Preface........................................................................................................................... ii
 Acknowledgements ...................................................................................................... iii
 Table of Contents......................................................................................................... iv
 List of Tables ............................................................................................................... vi
 List of Figures ............................................................................................................. vii
 List of Abbreviations ................................................................................................... ix
 Chapter 1 .......................................................................................................................1
 Introduction ...............................................................................................................1
 Ribosome History and Anatomy ..............................................................................2
 Ribosome Biogenesis and Disease .........................................................................10
 Ribosome Assembly Overview ..........................................................................10
 Ribosome Assembly Defects..............................................................................15
 Ribosome Function ................................................................................................18
 Translation.........................................................................................................18
 Initiation ............................................................................................................19
 Elongation..........................................................................................................25
 tRNA charging, selection and accommodation ...............................................25
 Peptidyltransfer ..............................................................................................30
 Translocation and ribosomal rotation..............................................................32
 Termination....................................................................................................36
 Antibiotics and Ribosome Inhibition ..................................................................39
 Translational Recoding ......................................................................................42
 Programmed -1 Ribosomal Frameshifting ......................................................42
 Programmed +1 Ribosomal Frameshifting .....................................................47
 Missense and Nonsense Suppression..............................................................48
 Scope of Work and Thesis Summary......................................................................49
 Chapter 2 .....................................................................................................................52
 Eukaryotic rpL10 drives ribosomal rotation..........................................................52
 Introduction ...........................................................................................................52
 Results ...................................................................................................................57
 Discussion .............................................................................................................88
 Chapter 3 .....................................................................................................................98
 Ribosomopathies: a new model for cancer biogenesis............................................98
 Introduction ...........................................................................................................98
 Results .................................................................................................................103
 Discussion ...........................................................................................................115
 Chapter 4 ...................................................................................................................119
 Work in progress: the rpL10 C-terminal tail and autism....................................119
 Introduction .........................................................................................................119
 Results .................................................................................................................122
 Discussion ...........................................................................................................127
 Chapter 5 ...................................................................................................................131
 Conclusion and future directions ..........................................................................131
 Summary and significance ...................................................................................131
v
 Medical relevance and future avenues..................................................................133
 Final words ..........................................................................................................135
 Chapter 6 ...................................................................................................................138
 Materials and Methods..........................................................................................138
 Media, strains, plasmids, and genetic manipulation. .............................................138
 Translational fidelity and polysome analyses. ......................................................139
 mRNA abundance and telomere length analyses. .................................................140
 Peptidyltransferase activity. .................................................................................140
 Sdo1p cloning, purification and labeling. .............................................................141
 Ribosome preparation. .........................................................................................142
 Ribosome/tRNA interactions................................................................................142
 Ribosome/protein interactions..............................................................................144
 Ribosome binding competition.............................................................................145
 Preparation of complexes for chemical probing....................................................145
 rRNA structure probing........................................................................................146
 Statistical analyses. ..............................................................................................147
 Acknowledgments ...............................................................................................147
 Appendix 1: Yeast strain list .....................................................................................148
 Appendix 2: Yeast plasmid list..................................................................................149
 Appendix 3: Oligonucleotide primer list...................................................................150
 References ..................................................................................................................152
vi
 List of Tables
 Table 1. Summary of genetic and functional analyses of L10 loop mutants....................59
 Table 2. Comparison of yeast rRNA chemical modification profiles with 
non-rotated and rotated E. coli standards........................................................................65
 Table 3. Establishing the rotational status of mutant ribosomes......................................71
 Table 4. Yeast strains generated...................................................................................148
 Table 5. Plasmids generated.........................................................................................149
 Table 6. Oligonucleotides used in directed mutagenesis...............................................150
 Table 7. Oligonucleotides used in telomere length assays. ...........................................151
vii
 List of Figures
 Figure 1. Electron microscopy images of bacterial ribosomes. .........................................3
 Figure 2. Yeast ribosome structure...................................................................................6
 Figure 3. Location of ribosomal intersubunit bridges. ......................................................7
 Figure 4. tRNA structure..................................................................................................9
 Figure 5. Position of tRNAs on the 70S ribosome. ...........................................................9
 Figure 6. Ribosomal subunit biogenesis summary..........................................................13
 Figure 7. Pathway of 60S maturation in the cytoplasm...................................................14
 Figure 8. Ribosome biogenesis and disease....................................................................17
 Figure 9. Electron micrograph of a polysome chain. ......................................................19
 Figure 10. Eukaryotic initiation summary. .....................................................................21
 Figure 11. The ?closed loop? mRNA structure...............................................................23
 Figure 12. Examples of viral and cellular IRES secondary structures.............................24
 Figure 13. Structure of a tRNA synthetase complex. ......................................................26
 Figure 14. Scheme of tRNA selection at the ribosomal A-site. .......................................29
 Figure 15. Mechanism of peptidyltransfer......................................................................31
 Figure 16. Positions of tRNAs during translation. ..........................................................33
 Figure 17. Ribosomal intersubunit rotation. ...................................................................34
 Figure 18. The elongation cycle of translation................................................................35
 Figure 19. Eukaryotic release factor is a structural tRNA mimic. ...................................37
 Figure 20. Mechanisms of peptidyltranfer and peptide release. ......................................38
 Figure 21. Steps of antibiotic inhibition of translation....................................................41
 Figure 22. Structure of a typical  -1 PRF signal..............................................................43
 Figure 23. The mechanism of -1 PRF.............................................................................46
 Figure 24. Effects of -1 PRF on viral propagation. .........................................................47
 Figure 25. The ribosome over 50 years. .........................................................................51
 Figure 26. rpL10 is strategically positioned in the core of the LSU. ...............................56
 Figure 27. rpL10 loop mutants have opposite effects on ligand binding to 
the A- and P-sites...........................................................................................................60
 Figure 28. Primary ligand binding data related to Figure 27...........................................61
 Figure 29. Visualization of chemical probing data by hSHAPE......................................63
 Figure 30. Biochemical verification of the rotational controls. .......................................66
 Figure 31. Structure of the B7a intersubunit bridge in the non-rotated and 
rotated states in E. coli ribosomes. .................................................................................68
 Figure 32. rpL10 loop mutants alter the rotational equilibrium of the 
ribosome........................................................................................................................69
 Figure 33. Structural probing analysis of 3? of the LSU. ................................................75
 Figure 34. Structural probing analysis of 5? of the LSU. ................................................76
 Figure 35. Structural probing analysis of the SSU..........................................................78
 Figure 36. 3D reactivity data of the PTC........................................................................79
 Figure 37. 3D reactivity data of the factor binding site...................................................79
 Figure 38. Points of rpL10 important for intersubunit rotation. ......................................81
 Figure 39. Primary data related to rotational mutants from Figure 38.............................82
 Figure 40. rpL10 loop mutants affect peptidyltransferase activity and 
translational fidelity. ......................................................................................................84
viii
 Figure 41. A mutant of rpL10 can be intrinsically suppressed with an rpL3 
mutant. ..........................................................................................................................86
 Figure 42. Primary data related to Figure 41. .................................................................87
 Figure 43. Models of rpL10 function: rpL10 is at the center of a cascade of 
allosteric communication pathways throughout the ribosome.........................................92
 Figure 44. Sdo1 structurally resembles tRNA. ...............................................................95
 Figure 45. Involvement of the rpL10 loop in ribosome rotation throughout 
the ribosomal life cycle..................................................................................................96
 Figure 46. Localization of rpL10 and the loop in the large ribosomal 
subunit.........................................................................................................................102
 Figure 47. Overexpression of NMD3 or co-expression of NMD3-Y379D
 suppresses the rpl10-R98S growth defect. ....................................................................103
 Figure 48. Overexpression of NMD3 or co-expression of NMD3-Y379D
 suppresses the rpl10-R98S biogenesis defect................................................................104
 Figure 49. Co-expression of NMD3-Y379D does not suppress the rpl10-
 R98S rotational defect..................................................................................................105
 Figure 50. Co-expression of NMD3-Y379D does not suppress the rpl10-
 R98S biochemical defects. ...........................................................................................107
 Figure 51. Primary ligand binding data related to Figure 50.........................................108
 Figure 52. Co-expression of NMD3-Y379D does not suppress the rpl10-
 R98S translational fidelity defects. ...............................................................................110
 Figure 53. Co-expression of TIF6-Y192F does not suppress the rpl10-
 S104D biochemical and translational fidelity defects. ..................................................111
 Figure 54. Sdo1p-V36A has a higher affinity to rpL10-S104D ribosomes....................112
 Figure 55. Co-expression of NMD3-Y379D does not suppress the rpl10-
 R98S telomere maintenance defects. ............................................................................114
 Figure 56. Co-expression of NMD3-Y379D does not suppress the rpl10-
 R98S telomere length defects. ......................................................................................114
 Figure 57. Model of T-ALL progression. .....................................................................117
 Figure 57. Location of L10 regions in the LSU. ...........................................................120
 Figure 58. Multiple sequence alignment of L10. ..........................................................121
 Figure 59. Genetic analysis of rpL10 ?body? and ?tail? mutants. .................................123
 Figure 60. Translation fidelity analysis of rpL10 ?body? and ?tail? 
mutants........................................................................................................................125
 Figure 61. Polysome profiling of rpL10 ?body? and ?tail? mutants. .............................126
 Figure 62. The rpL10 C-terminal tail and location of the L206 ?autism 
residue.? ......................................................................................................................130
 Figure 63. Ribosomopathies: proposed chain of causality. ...........................................137
ix
 List of Abbreviations
 1M7 1-methyl-7-nitroisatoic anhydride
 5-FOA 5-Fluoroorotic acid 
aa-tRNA Aminoacyl tRNA
 AF Assembly factor
 ARS Aminoacyl-tRNA synthetase 
ATP Adenosine triphosphate 
CP Central protuberance
 Cryo-EM Cryogenic electron microscopy 
DBA Diamond-Blackfan anemia 
DC Decoding center 
EF Elongation factor
 eEF Eukaryotic elongation factor 
eIF Eukaryotic initiation factor
 eRF Eukaryotic release factor 
GAC GTPase-associated center 
GDP Guanosine diphosphate 
GEF Guanine exchange factor 
GGQ Glycine-glycine-glutamine motif 
GTP Guanosine triphosphate 
HIV-1 Human immunodeficiency virus type 1
 IF Initiation factor
 IRES Internal ribosomal entry site 
x
 LSU Large ribosomal subunit 
m7G 7-methylguanosine 
mRNA Messenger RNA
 NGD No-go decay 
NMD Nonsense mediated decay 
NRD Non-functional ribosomal decay 
ORF Open reading frame 
PABP Poly-A binding protein 
Pol Polymerase 
Poly A Poly-adenosine
 PRF Programmed ribosomal frameshifting 
PTC Peptidyltransferase center & premature termination codon 
RF Release factor
 RP Ribosomal protein
 RRF Ribosomal recycling factor
 rRNA Ribosomal RNA
 SARS Severe acute respiratory syndrome 
Sec Selenocysteine
 SHAPE Selective 2?-hydroxyl acylation analyzed by primer extension 
snoRNAs Small nucleolar RNAs
 SRL Sarcin-ricin loop 
SSU Small ribosomal subunit
 T-ALL Acute T-cell lymphoblastic leukemia 
xi
 tRNA Transfer RNA
 YPAD Yeast media containing: yeast extract, peptone, adenine sulfate, dextrose  
1
 Chapter 1
 Introduction
 Faithful conversion of genomic information into functional proteins is central to 
life. In all domains of life, this is accomplished by the largest RNA-protein complex in 
the cell, the ribosome, in a process called ?translation?. The ribosome translates the 
information contained in messenger RNAs (mRNAs) into polypeptide chains with 
tremendous speed and accuracy, linking the nucleic acid and protein worlds. There are 
significant differences in both the assembly and structure of eukaryotic ribosomes as 
compared to their bacterial counterparts: these are the basis for approximately half of our 
current arsenal of antibiotics. However, given the rapid emergence of antibiotic 
resistance, increasing our understanding of ribosome mechanics is critical. From a 
mechanistic point of view, the ribosome can be viewed as a model ?nanomachine.? In 
addition to rational drug development, studies of the wiring and workings of the machine 
provide a platform for the understanding of macromolecular assembly and function.
 Defects associated with this nanomachine and canonical translation have broad impacts 
on human health including birth defects, cancer, heart and lung diseases, connective 
tissue disorders, and mental retardation. Viruses, immune to antibiotics and reliant on 
ribosomes, have evolved ways to subvert canonical translation strategies. Despite all 
this, ribosomal molecular mechanics and dynamics, as well as mechanisms of quality 
control of ribosome assembly, are not well understood. Understanding of these important 
areas will provide the foundation for the design of new classes of anti-bacterial and anti-
 viral drugs, nanodevices, and treatment of translation-linked diseases.  
2
 Ribosome History and Anatomy 
The ribosome is thought to have originated in the pre-biotic world where it 
evolved to bridge the realms of RNAs and proteins and was first detected approximately
 3.5 billion years later as a ?small particle? by Albert Claude in the early 1940s 1,2. It was 
not until the 1950s that these so-called ?microsomes?, operationally defined in terms of 
sedimentation, optical inspection, and chemical composition, became linked on 
experimental grounds to protein synthesis 3?6. Purification of microsomes was the major 
breakthrough in allowing development of cell-free protein synthesis systems in 1955. 
Around the same time, through the novel technique of transmission electron microscopy
 George Palade was able to obtain images of the microsome in the cytoplasm as well as on 
the endoplasmic reticulum, for which he won the Nobel Prize in 1974 7,8. These images 
provided the first insights into the structure of these small particles, and similar 
techniques were the basis for the very first strikingly accurate 3D models developed by 
James Lake and shown in Figure 1. After a few transient aliases including ?deoxycholate-
 insoluble? and ?ribonucleoprotein particle?, Howard Dintzis coined the term ?ribosome? 
for purified microsomes in 1958 based on the presumed role of the particle?s large RNA 
content (rRNA) 9. This was also the year of the first ribosome symposium. The following 
?golden age of translation? of the 1960s saw the discovery of mRNA, the dissection of 
the ribosome into its components, and the resolution of the translational process into 
partial functions. Much of this golden age, as well as the myriad breakthroughs in the 
field between the 1970s and the 1990s, will be referred to in later sections and chapters. 
3
 More recently, the major focus in the field has shifted to high resolution structural 
analyses. At the turn of the century the combined efforts of several groups resulted in the 
elucidation of eubacterial ribosomal structures at the atomic level, revealing in detail this 
complex macromolecule and its discrete functional centers 11,12,13. This work was 
awarded the Noble Prize in chemistry in 2009. Shortly after, ribosomal X-ray 
crystallographic structures at the atomic level became available for the yeast 
Saccharomyces cerevisiae 14,15. As of 2013, lower resolution structures obtained through 
cryogenic electron microscopy exist for complete fruit fly and human ribosomes 16, as 
well as for the small subunit of rabbit ribosomes 17.   The ribosome consists of two 
Figure 1. Electron microscopy images of bacterial ribosomes.
 Electron microscopy images from 1973 reveal the shape of the ribosomal subunits. These 
images were used by James Lake to construct the very first ribosome models on the right.
 Parts of the image obtained from Wittman 10. 
4
 subunits defined by the sedimentation coefficient S: the large subunit (LSU, 50S in 
prokaryotes and 60S in eukaryotes) and the small subunit (SSU, 30S in prokaryotes and 
40S in eukaryotes). These subunits have distinct functions, but must also interact with 
one another along with numerous trans-acting factors to form a holoenzyme that
 coordinates a complex series of events. The secondary and tertiary structures of the 
ribosomal core (the ?proto-ribosome?), including the active site, are well conserved 
across all domains of life. However, the eukaryotic ribosome is much larger and more 
complex, containing additional RNA in the form of so-called expansion segments as well 
as many additional proteins or protein extensions. As a result, its construction is 
fundamentally different in many key ways. Mitochondrial ribosomes, consisting of 28S 
and 39S subunits, structurally more closely resemble prokaryotic ribosomes ? one of the 
strongest supports for the endosymbiotic theory of cellular evolution 18,19. 
Compared with the 4500 nucleotides of rRNA and 54 proteins of the bacterial 70S 
ribosome, the yeast 80S ribosome contains 5500 nucleotides of rRNA (SSU: 18S rRNA; 
LSU: 5S, 5.8S, 25S rRNA) and 79 proteins (LSU and SSU proteins are prefixed with ?L? 
or ?S? respectively). The structural landscape of the 3.6 MDa yeast ribosome is shown in 
Figure 2. The SSU contains the mRNA decoding center (DC), a region responsible for 
reading the mRNA to ensure proper acceptance of transfer RNAs (tRNAs), and the tunnel 
through which mRNA moves. The most salient structural feature in the SSU is the 
flexible ?head? domain. The LSU harbors the catalytic peptidyltransferase center (PTC) 
at its core, the exit tunnel for the growing polypeptide chain, three tRNA binding pockets
 and a single binding site that must distinguish between elongation and release factors in 
response to specific circumstances. The main structural features of the 60S include the 
5
 stalk which consists of proteins L7 and L12 and is required for factor binding, and the 
central protuberance (CP) which consists of L11 and 5S rRNA.  
The yeast ribosome is an RNA-protein complex and has an approximate 1:1 RNA 
to protein mass ratio. It is the largest and most abundant natural ribozyme, and is also the 
only natural RNA-based polypeptide polymerase. Ribosomal RNA is highly modified, 
containing over 100 modifications including base methylations, ribose 2?-hydroxyl 
methylations, and the most common nucleotide modification ? pseudouridylations. Mice 
and humans with impaired pseudouridylation develop cancer and dyskeratosis congenita 
20,21. Moreover, out of the 25 non-canonical basepair types, 20 are present in the ribosome
 22. A balance between flexibility and rigidity is achieved through a plethora of structural 
RNA elements and motifs including tetraloops, E-loops, U- and K-turns, purine stacks, 
coaxial stacking, ribose zippers, and most commonly ? the A-minor motif 23.  Most 
ribosomal proteins contain globular portions that tend to localize to the solvent sides of 
the two subunits, as well as non-globular solvent-inaccessible and flexible extensions or 
?tails? interacting with the rRNA core. Historically, it was first thought that ribosomal 
proteins were the central players in ribosome function while rRNA was relegated to a 
minor, scaffolding function. As understanding of the ribosome progressed and it was 
established that catalysis occurs in an entirely RNA-based environment, these perceived 
roles were almost completely reversed. However, more recently ribosomal proteins have 
begun to be appreciated as being more than just the structural glue: it has been shown that 
over half are essential, and a small set of ribosomal proteins are even required for 
catalytic activity 24,25. Thus, the 1:1 RNA to protein mass ratio is likely also indicative of 
the contribution of each to ribosomal structure and function.
6
 Figure 2. Yeast ribosome structure. 
Subunits are shown from the intersubunit face view (top), side (middle) and in the 80S 
context (bottom). Proteins = blue/brown; rRNA = black/grey. Generated from 3 ?
 crystallography structures using PyMOL (PDB code: 3U5B/C/D/E) 26.
7
 The 80S ribosome must function as one unit. To this end, the interaction surface 
between the two subunits contains 17 connecting intersubunit bridges to provide 
communication pathways and structures 15. These bridges consist of RNA:RNA, 
protein:protein, and RNA:protein interactions and are shown in Figure 3.  Because the 
two subunits must dynamically interact with one another as the ribosome undergoes 
global conformational rearrangements during translation, some of these intersubunit 
bridges (such as B3) consist of strong permanent interactions, while others (such as B7a) 
are malleable in composition and can thus break and reform throughout translation. 
Figure 3. Location of ribosomal intersubunit bridges. 
Out of the 17 intersubunit bridges, B1b/c is the only protein:protein bridge, while all 
others are RNA:protein or RNA:RNA. Image from Ben-Shem 15. 
8
 Precise subunit interactions are necessary to guide tRNAs through the ribosome. 
The job of the 20 tRNAs, each 75-90 nucleotides long and carrying one specific amino 
acid, is to provide the ribosome with building blocks for protein synthesis. Transfer 
RNAs are the most modified RNA molecules: 80 modifications out of a total of 95 
reported have been observed in tRNAs 27. The secondary and tertiary tRNA structure is 
shown in Figure 4. Based on base complementarity, the secondary structure of a tRNA 
can be drawn as a cloverleaf containing 4 stems. Five regions of the tRNA are not base 
paired: the CCA acceptor stem, the D-loop and T-loop, the anticodon loop and the 
variable "extra arm". These structural elements result in an L-shaped molecule upon 
folding and allow tRNAs to interact with both the mRNA in the SSU (the anticodon) and 
the PTC in the LSU (3? CCA end). As tRNA molecules interact with the ribosome they 
move through a distinct path through functional sites on the ribosomes.  There are well 
defined tRNA binding sites along this path: the A- and P- sites, where the aminoacyl- and 
peptidyl-tRNAs reside before peptide bond formation, and the E-site, from which 
deacylated or ?used? tRNAs exit the ribosome 28. While high resolution structures of 
yeast ribosomes bound with tRNAs do not exist, such structures have been resolved in 
the bacterium T. thermorphilus, shown in Figure 5.  
9
 Figure 4. tRNA structure. 
Non base-paired regions form the 4 shown loops. The anticodon interacts with the 
decoding center in the SSU, while the acceptor stem is covalently linked to the 
appropriate amino acid and interacts with the PTC in the LSU. Image modified from 
Griffiths-Jones 29.
 Figure 5. Position of tRNAs on the 70S ribosome. 
tRNAs in A, P and E sites are orange, red and blue, respectively, and mRNA is  yellow. 
Image from Yusupov 28.
10
 Ribosome Biogenesis and Disease 
Healthy yeast cells contain approximately 200,000 ribosomes distributed 
throughout the cytoplasm as well as bound to the endoplasmic reticulum (the ?rough? 
part of it), where proteins of the intra-cellular transport system are translated 30. Up to 
2000 ribosomes are produced per minute per cell to keep up with such a demand. 
Impaired ribosome assembly is the underlying cause of the set of pathological conditions 
known as ribosomopathies. As such, accurate piece by piece ribosome assembly and 
subsequent quality check are essential components of ribosome biogenesis. 
Ribosome Assembly Overview 
Ribosomes must be assembled from rRNA and proteins in a complicated and 
energetically demanding process taking place in both the nucleus and cytoplasm of cells. 
In prokaryotes, in vitro reconstitution of functional ribosomal subunits from purified or in 
vitro transcribed rRNAs and ribosomal proteins has been demonstrated, and assembly 
maps for both subunits have been constructed 31?34. Ribosome assembly in eukaryotes is 
much more complex, and reconstitution such as seen in bacteria has never been achieved. 
Most of the knowledge pertaining to eukaryotic ribosome biogenesis comes from studies 
in yeast. Unlike in prokaryotes, in which ribosome biogenesis is determined and driven 
by the intrinsic properties of its rRNA and proteins and follows an established assembly 
gradient, yeast ribosome assembly involves transient association of ~200 essential 
accessory proteins, trans-acting assembly factors that do not form part of the mature 
ribosome structure 35,36. These proteins include GTPases, ATPases, kinases, helicases and 
chaperones and promote essential functions such as rRNA processing, modification and 
11
 folding, and assembly of ribosomal proteins. Eukaryotes also bear the added burden of 
exporting immature ribosomal subunits from the nucleus to the cytoplasm. A summary of 
the subunit processing pathway including known factors is given in Figure 6.
 In yeast, the assembly process begins in the nucleolus, a specialized 
subcompartment of the nucleus that is organized around the rDNA transcription units. 
18S, 25S and 5.8S rRNA are transcribed in the nucleolus as a 35S polycistronic precursor 
rRNA by RNA polymerase I. This pre-rRNA undergoes a series of endonucleolytic 
cleavages and exonucleolytic trimmings, along with extensive modifications by ~70 
small nucleolar RNAs and protein co-factors into the final 18S, 25S and 5.8S rRNAs, 
which serve as the catalytic core of the ribosome 37. The 5S rRNA is transcribed by RNA 
polymerase III in the nucleoplasm, undergoes maturation in the cytoplasm, and is 
transported to the nucleolus in complex with L5 and L11 38. Similarly, ribosomal protein
 mRNAs are transcribed by RNA polymerase II in the nucleoplasm, translated into mature 
proteins in the cytoplasm, after which they are transported back to the nucleolus for 
additional processing steps. These steps consist of the formation of numerous assembly 
intermediates, including a 90S pre-ribosome that begins to fold and associate with 
ribosomal proteins, cleavage steps resulting in 43S and 66S preribosomal particles, and 
their transport from the nucleolus into the nucleus and subsequently the cytoplasm for 
final rRNA processing and protein assembly events 39. Both subunits are exported with a 
small complement of non-ribosomal trans-acting factors, some of which (Nmd3) 
facilitate export while others (Tif6) prevent the premature interaction of the ribosomal 
subunits. These factors must be released in the cytoplasm and shuttled back to the 
nucleus for subsequent rounds of maturation and export. After this recycling step, the 
12
 fully assembled and mature subunits can interact to form translationally active 80S 
ribosomes.
 While the sequence of events constituting 40S cytoplasmic maturation is largely 
unknown, the recently established order of 60S cytoplasmic maturation steps is shown in 
Figure 7 40. Maturation is initiated by the ATPase Drg1 which facilitates the replacement 
of Rlp24 by Rpl24, which in turn recruits Rei1. Together with Jjj1 and Ssa1/Ssa2, Rei1 
enables the release of the export receptor Arx1, freeing up the polypeptide exit tunnel.
 Concurrently, Yvh1 enables replacement of Mrt4 with P0 to form the foundation upon 
which the ribosome stalk is constructed. At this point the last component that is part of 
the final mature subunit, ribosomal protein L10, is incorporated. This is followed by the 
recruitment of the Efl1:Sdo1 complex and the subsequent Sdo1-induced GTP hydrolysis 
by Efl1 to release the anti-association factor Tif6 from the subunit joining face of the 
subunit. The release of Tif6 leads to activation of Lsg1 to release the export adaptor 
Nmd3 from the joining face - the last step of 60S maturation. 
13
 Figure 6. Ribosomal subunit biogenesis summary. 
Red arrows = endonucleolytic steps; blue arrows = exonucleolytic steps; cyan = 
ATPases; green = GTPases; orange = kinases. Figure is from Strunk 41.
14
 Figure 7. Pathway of 60S maturation in the cytoplasm. 
The ribosomal stalk is assembled in the cytoplasm which, along with incorporation of 
L10, enables the recruitment of the G-protein Efl1 in complex with Sdo1. This results in 
the GTP hydrolysis induced dissociation of Tif6 and Nmd3 and a mature 60S subunit. 
Figure modified from Lo 40. 
15
 In addition to central roles in cell viability, ribosome biogenesis and protein 
synthesis are two of the most energy consuming processes in a growing cell. Thus it is 
critical for cells to have evolved a surveillance system for monitoring proper assembly of 
the translational machinery. It was recently demonstrated that pre-40S subunits undergo a 
proofreading step late during their cytoplasmic maturation 42. This involves a translation-
 like cycle, whereby mature 60S subunits join pre-40S subunits in an eIF5B (eukaryotic 
initiation factor 5B)-dependent manner. However, the resulting 80S-like ribosomes are 
not translationally active, as they do not contain mRNA or initiator tRNA. Rather, this 
translation-like cycle serves as a final quality control step in which major functions of the 
maturing small subunit, such as binding to 60S subunits and to translation factors, are 
tested. Along the same lines, a ?test drive? of immature 60S subunits has recently been 
reported by our lab and will be discussed in more detail in chapter 2.
 Ribosome Assembly Defects 
Given the essential role ribosomes play in the central dogma of biology, it is 
likely that defects in translation would result in wide-spread deleterious phenotypic 
effects. However it was not until recently that disorders of ribosome dysfunction, 
collectively called ribosomopathies, were described 43. In 1999, recurrent mutations in the
 ribosomal protein gene RPS19 were reported in patients with Diamond-Blackfan anemia 
(DBA), a rare congenital bone marrow failure syndrome 44. Since then, mutations in a 
number of ribosomal proteins have been identified in up to 50% of patients with DBA.
 Moreover, other congenital syndromes have been linked to non-RP related ribosomal 
16
 defects and include Schwachman-Diamond syndrome, X-linked dyskeratosis congenita, 
cartilage hair hypoplasia, and Treacher Collins syndrome 45. In addition, the 5q? 
syndrome, a subtype of myelodysplastic syndrome, is caused by a somatically acquired
 deletion of chromosome 5q, which leads to haploinsufficiency of the ribosomal protein 
S14 and an erythroid phenotype highly similar to Diamond-Blackfan anemia 46. The 
genetic abnormalities in all of these disorders cause defects in a step in ribosome 
biogenesis, shown in Figure 8, and lead to a higher incidence of cancer. 
A precise mechanism of ribosome biogenesis impairment has recently been 
described for Schwachman-Diamond syndrome (SDS), an inherited bone marrow failure 
syndrome 47. SDS has an incidence estimated at 1 in 50 000 births and is characterized by 
a strong predisposition to leukemia, with bone marrow transplantation as the only 
definitive therapy. Approximately 90% of SDS cases are caused by mutations in the 
SBDS (Shwachman-Bodian-Diamond syndrome) gene. The SBDS protein functions in a 
late step in the cytoplasmic maturation of 60S subunits by promoting the release of eIF6 
(eukaryotic initiation factor 6) from pre-60S subunits. This trans-acting factor keeps the 
nascent 60S subunit in a functionally inactive state during cytoplasmic 60S assembly, but 
needs to be released for final 60S maturation. Much like its yeast homolog Sdo1, SBDS 
appears to promote the release of eIF6 by stimulating the GTPase activity of Efl1. The 
mutant form of the protein in SDS patients is unable to release eIF6 as efficiently and 
hence stalls 60S maturation. 
The causes of the hypo-proliferative clinical symptoms of most ribosomopathies, 
such as anemia and dystosis, have recently been linked to the tumor-suppressor protein 
p53 48. This protein is known to play a fundamental role in the surveillance of protein 
17
 translation, and recent studies show that it can be activated by ribosome dysfunction. The 
HDM2 protein (Human Double Minute 2) can bind many ribosomal proteins and is a 
central regulator of p53, acting as a ubiquitin ligase that leads to p53 degradation, thereby 
providing a link between ribosome biogenesis and the p53 pathway: a disruption in 
ribosome biogenesis leads to an accumulation of free ribosomal proteins binding HDM2,
 the inhibition of HDM2 activity, and the consequent accumulation of p53 and apoptosis
 49. However, how clinically hypo-proliferative diseases can also cause hyper-proliferation 
(cancer) is unclear and will be addressed in Chapter 3. 
Figure 8. Ribosome biogenesis and disease. 
The step in ribosome biogenesis that is thought to be affected by each ribosomopathy is 
depicted. Image from Teng 50. 
18
 Ribosome Function
 Translation
 Once properly assembled, ribosomes can begin synthesizing polypeptides. Protein 
synthesis is remarkably fast: elongation of the polypeptide chain by one amino acid 
occurs in approximately 60 ms 51. It is also highly accurate, making approximately 1 error 
every 103 - 104 codon reads 52. Several ribosomes may translate a single mRNA molecule 
at the same time, resulting in a ?polysome? which is shown in Figure 9. As with other 
polymerization reactions, the translational process is divided into the phases of initiation, 
elongation, and termination and is assisted by a multitude of essential trans-acting
 factors. A fourth less understood phase comprises ribosome recycling. During initiation 
the 80S ribosome is assembled on the mRNA and is guided to a start codon, marking the 
beginning of the protein product. The bulk of translation happens during elongation, 
when new amino acids are continuously added to the growing polypeptide chain as the 
ribosome moves one triplet codon at a time along the mRNA. Upon reaching a stop 
codon, the full-length protein is released by specialized release factors during the 
termination phase of translation. The 80S complex may either be recycled back to the 
beginning of the message for another round of translation or dissociate to begin 
translation of another message. 
19
 Figure 9. Electron micrograph of a polysome chain. 
The beginning of the mRNA is to the right (at the arrow) and the end is to the left. The 
synthesized polypeptide can be seen growing longer and longer, the further along the
 ribosome is positioned along the message. Image obtained from the official website of the 
Nobel Prize Foundation 53. 
Initiation
 Initiation is a major rate-limiting step of translation, and is thus the principal 
target for regulation 54. In eukaryotes, this process involves a series of initiation factors 
(eIFs) and is depicted in Figure 10. The starting point for assembly of the initiation 
complex is the production of free small subunits. Binding of eIF1A and eIF3 to the SSU 
prevents premature association with 60S 55. The initiator methionine tRNA (Met-
 tRNAiMet) is delivered to the SSU in an initiation ternary complex containing eIF2 
coupled with GTP. The eIF1A and eIF3 stimulate binding of the ternary complex to form
20
 a 43S pre-initiation complex 56. Before mRNAs can be bound by this complex they must 
first interact with several factors. Particularly, the 3? poly-A tail of the message is 
recognized by the Poly-A Binding Protein (PABP), while the 5? 7-methylguanosine cap 
(m7G) is bound by eIF4E. The scaffold factor eIF4G interacts with both these 
components, linking the 5? and 3? ends of the mRNA thereby circularizing the message
 57. This results in a translationally competent ?closed loop? message which is able to 
interact with the 43S pre-initiation complex. The eIF4G facilitates 43S binding near the 
5? cap by interactions with eIF3, resulting in a 48S pre-initiation complex. Powered by 
ATP hydrolysis of eIF4A, this complex begins scanning the mRNA towards the 3? 
direction in search of a start codon.  Pairing of the anticodon of Met-tRNAiMet with the 
AUG start codon in the ribosomal P-site triggers GTP hydrolysis by eIF2, stimulated by 
the concerted action of the GTPase activating protein eIF5 and the 40S itself 58?62. All the 
initiation factors are subsequently released, and the 60S is able to associate with the 40S
 complex, with the Met-tRNAiMet stably occupying the ribosomal P-site. The eIF2-GDP is 
recycled to the eIF2-GTP by the guanine exchange factor (GEF) eIF2B to enable the 
reassembly of the ternary complex. Rare eukaryotic initiation from the ribosomal A-
 site has also been described, suggesting that the repertoire of translation initiation may be 
greater than anticipated 62. 
21
 Figure 10. Eukaryotic initiation summary. 
Known steps and factors are indicated. The GEF eIF2B is inhibited by the 
phosphorylation of eIF2 of its ? subunit by various kinases, activated by different kinds 
of stress. Image from Hinnenbusch 63. 
22
 As discussed above, successful initiation is reliant on the formation of the ?closed 
loop? structure that results from mRNA circularization, shown in Figure 11. Many 
viruses have evolved unique methods to fulfill the need for features of translationally 
competent eukaryotic mRNAs. For example, flu viruses can obtain their own 5? m7G 
caps through a cap-snatching endonuclease 64. This results in the capping and 
translational activation of viral mRNAs, while simultaneously rendering cellular mRNAs 
inactive.  In another approach, picornaviruses covalently attach a VPg (viral protein 
genome-linked) protein to the 5? ends of their mRNA. The VPg mimics a 5? cap and is 
able to interact with initiation factors, thereby fulfilling the function of the cap 65,66. 
Another successful viral initiation strategy bypasses the need for a 5? cap and other 
cellular mRNA features altogether. Instead, some viruses have evolved Internal 
Ribosomal Entry Site elements (IRES elements) 67,68. These are special secondary 
structures located at the 5? end of viral mRNAs, some mimicking tRNAs and interacting 
with initiation factors, which can recruit ribosomes and direct them to initiate internally. 
Many IRES elements of various shapes and sizes have been described, and some are 
shown in Figure 12. The key for viruses is to lure the ribosome onto its message and to 
overcome the rate-limiting step of initiation.  More recently, such IRES elements have 
also been described in eukaryotes 69,70,71. IRES-mediated cellular translation appears to be 
favored under circumstances when cap-dependent translation is compromised, such as 
starvation conditions 72.
23
 Figure 11. The ?closed loop? mRNA structure.  
Atomic force microscopy reveals complexes formed on circularized, translationally 
active mRNAs. Image from Mendez 73. 
24
 Figure 12. Examples of viral and cellular IRES secondary structures.
 IRES elements in Plautia stali intestine, Hepatitis C, Foot-and Mouth Disease, and 
Human Immunodefiency virus genes are shown, as well as in the human proto oncogene 
c-Myc.  Image from Filbin 74. 
25
 Elongation
 Once stably established on the mRNA during initiation, the ribosome elongates 
the polypeptide chain by reading one codon at a time and relying on tRNAs and 
elongation factors (eEFs) to supply the correct amino acid at each codon. This is when 
the bulk of protein synthesis occurs and is the fastest phase of translation.  
tRNA charging, selection and accommodation
 The first step in elongation is to bind a tRNA carrying an amino acid to the 
ribosomal A-site. Before this can happen, tRNAs need be activated or ?charged? -
 covalently linked with an amino acid by a class of enzymes called aminoacyl-tRNA 
synthetases (ARS). Although codon:anticodon is the key interaction directing translation, 
the correct recognition of and aminoacylation of the tRNA by its cognate synthetase is of 
comparable importance and can be regarded as a second genetic code. Mis-charging of 
tRNAs can lead to the incorporation of the wrong amino acid and production of an 
aberrant protein product. Every cell requires an ARS for each of the 20 amino acids of 
the genetic code. Correct aminoacylation depends on the selection of two appropriate 
substrates by the corresponding ARS: the tRNA and the amino acid. While tRNAs are 
large and specific and can be easily distinguished, amino acids must be distinguished 
solely by the nature of their side chains. The aminoacylation reaction which takes place at 
a site of the enzyme called ?the synthetic site? occurs in two steps 75. The amino acid is
 first activated by ATP-driven adenylation and is then transferred to the CCA end of the
 cognate tRNA. Steric exclusion of amino acids with larger side-chains and recognition of 
specific properties of each amino acid generally make this synthetic site specific enough 
26
 so that only the correct amino acid can be activated and transferred. However, amino 
acids having similar properties to and a smaller size than the cognate amino acid can be 
mis-activated at frequencies that are too high to maintain an unambiguous code. As a 
consequence, enzymes facing this problem have evolved a second active site for 
proofreading called ?the editing site?, where mis-activated amino acids or mis-acylated 
tRNAs are hydrolyzed 76?79. The presence of two catalytic sites with different activities 
led to the "double-sieve? model of fidelity and is shown below in Figure 13. 
Figure 13. Structure of a tRNA synthetase complex. 
The L-shaped structure of the tRNA allows the tRNA synthetase to bind both the 
acceptor stem and the anticodon loop. Charging of the cognate amino acid is further 
ensured through the proofreading editing site. Image from Berg 80. 
27
 Properly charged tRNAs are delivered to the ribosome in an elongation ternary 
complex with eEF1A and GTP. The interaction of this ternary complex and the ribosome 
follows distinct steps shown in Figure 14, and culminates in either acceptance 
(accommodation) or rejection of the tRNA 81?83. First, initial binding of the ternary 
complex results in a labile interaction with the A-site, in which the anticodon of the 
tRNA is positioned in the decoding center. Cognate and near-cognate ternary complexes 
are stabilized by the following codon-recognition step, which is governed by codon-
 anticodon Watson-Crick base-pairing stability. Non-cognate ternary complexes are 
quickly and efficiently rejected at this first selection stage. Upon accurate tRNA 
selection, yeast 18S bases A1755, A1756, and G577 (E. coli A1492, A1493, G530) flip 
out from a syn to an anti conformation stabilizing the mini-helix formed from cognate 
codon-anticodon interactions 84,85. This stabilization induces conformational changes in 
the ribosome that triggers the GTPase activity of eEF1A 86,87,88. Near-cognate tRNAs fail 
to stimulate rearrangement of the three 18S rRNA bases, and thus fail to stimulate GTP 
hydrolysis by eEF1A. The conformational change of eEF1A to the GDP-bound form 
results in loss of its affinity for the tRNA and the ribosome. Once released from eEF1A, 
the aa-tRNA relocates to the LSU portion of the A-site in the process of accommodation. 
This is also accompanied by the release of the tRNA from the E-site. 
The efficiency of the initial selection step is not sufficient to distinguish between 
the cognate and near-cognate ternary complexes, because the differences in the rate 
constants of the backward reaction (k?2 in Figure 14) and the forward reaction (k3 in 
Figure 14) between cognate and near-cognate complexes are not large enough 89?91. 
Therefore, most of the discrimination of near-cognate aa-tRNA is accomplished only 
28
 after GTP hydrolysis during the stage of selection known as ?proofreading.? This step is 
much less understood than the selection step during decoding. However, recent evidence 
suggests that recognition of the canonical geometry of the codon-anticodon helix in the 
DC induces stabilization of the entire anticodon loop by rRNA from both ribosomal 
subunits with involvement of magnesium ions 92. This triggers fixation of the anticodon 
stem through a cascade of rearrangements in 25S rRNA and ribosomal proteins. The 
latter strengthen the binding of cognate tRNA on the ribosome via their flexible tails 
while concurrently stimulating positioning of the acceptor end in the PTC. An induced-fit 
mechanism in such a closed system of interactions as in the ribosome, in which the 
accommodation step is accomplished by the surrounding of cognate tRNA by dynamic 
layers of rRNA and proteins from both subunits, seems entirely reasonable. The kinetic 
data indicate that cognate aa-tRNA is faster in A-site accommodation and peptide bond 
formation 93. As such, this ?proofreading? step is unlike analogous functions of other 
macromolecules such as tRNA synthetases discussed above or DNA polymerases. 
Rather, this is a ?kinetic proofreading? step. Together, the two selection strategies ensure 
that cognate aa-tRNAs are bound to the ribosome more stably, and are faster in 
?productive? forward reactions of GTP hydrolysis and A-site accommodation than near-
 and non-cognate tRNAs.
29
 Figure 14. Scheme of tRNA selection at the ribosomal A-site. 
The initial binding step in tRNA selection, governed by the rate constants k1 and k-1, is a 
codon-independent reaction between the ternary complex and the ribosome, while the 
following codon-recognition step is codon dependent. The active site of the elongation 
factor undergoes a conformational change during GTPase activation. This step is pivotal 
for establishing the irreversible step essential to proofreading and appears to limit the rate 
of GTP hydrolysis. During the proofreading stage after factor dissociation the tRNA 
either moves into the A-site (accommodation) for peptidyltransfer or dissociates from the 
ribosome (rejection). Accommodation is regulated by the rate constant k5 and depends on 
codon-anticodon interactions.
30
 Peptidyltransfer 
The catalytic core of the ribosome is composed entirely of rRNA, with the nearest 
protein component approximately 13 ? away 15. Peptidyltransfer is mediated solely by 
the LSU, and it alone can synthesize peptide bonds as rapidly as the full ribosome. Upon 
accommodation, the 3? CCA end of the aminoacyl-tRNA is stabilized by the 25S rRNA 
and positioned directly in the PTC resulting in almost instantaneous peptidyltransfer. The 
ribosome PTC catalyzes the aminolysis of an ester bond and is shown in Figure 15. The 
?-amino group of A-site aminoacyl tRNA nucleophilically attacks the P-site peptidyl 
tRNA at the carbonyl carbon of the ester bond that links the peptide to the tRNA, 
resulting in the deacylation of the P-site tRNA and the transfer of the growing 
polypeptide chain to the A-site tRNA. Amines react intrinsically fast with esters to form 
peptide bonds (?10?4 M?1 s?1 at room temperature), and the ribosome accelerates this 
reaction by ?106 -to107 -fold 94. This acceleration is achieved by lowering the entropy of 
activation, whereas the enthalpy of activation is the same for the reaction on the ribosome 
and in solution. Thus, the ribosome seems to rely on entropic catalysis as the mechanism 
of peptide-bond formation 94,95. Recent evidence suggests that the correct binding of 
substrates induces structural rearrangements in the PTC resulting in an orientation 
suitable for catalysis. In particular, this includes the involvement of conserved active site 
residues in substrate positioning, desolvation, and organization of an electrostatic 
environment via a hydrogen network that stabilizes the reaction intermediates and allows 
appropriate proton shuttling 96. 
31
 Figure 15. Mechanism of peptidyltransfer. 
The ?-amino group of aminoacyl tRNA in the A site attacks the carbonyl carbon of the 
peptidyl-tRNA in the P site. The universally conserved bases in green, among others, 
promote electrostatic shielding and a concerted proton-shuttle mechanism resulting in 
stabilization of a six-membered transition state and facilitating catalysis. Figure modified 
from Rodnina 96,97. 
32
 The ribosomal reaction rate acceleration is much less efficient compared to many 
protein enzymes, which use chemical catalysis and accelerate reactions by up to 1023.
 Moreover, the intrinsic rate of peptidyltrasfer is estimated to be >300 s-1 and is much 
faster than the approximate 10 s-1 rate of accommodation 87,98. Thus it seems that 
evolution has favored the increase in speed and fidelity of the rate-limiting steps of 
protein synthesis which do not involve chemistry, such as ternary complex binding 
described above, rather than peptide-bond formation itself. As a result, the catalytic core 
has retained its structure and mechanism of action during the course of the evolution from 
a pre-biotic ribozyme into a modern ribosome, and is one of the most conserved elements 
in all of life 99,100.  
Translocation and ribosomal rotation 
In order for translation to continue after a single round of peptidyltransfer, the A-
 site must be emptied to allow binding of a new ternary complex. For this to happen, the 
polypeptide-containing tRNA in the A-site needs to move into the P-site, while the 
?empty? deacylated tRNA needs to move from the P-site into the E-site. This is fueled by 
GTP and eEF2 in a process called translocation and results in ligand repositioning shown 
in Figure 16. Immediately after peptidyltransfer, the classically positioned P/P- and A/A 
site tRNAs (nomenclature signifies the position of tRNAs in the SSU/LSU) adopt a 
?hybrid? P/E and A/P conformation 101,102,103. Adoption of this state is not possible before 
deacylation of the P-site tRNA, possibly due to the anchoring effect of the polypeptide 
chain in the exit tunnel 104. The movement of the P-site tRNA into the hybrid state 
involves large motions in the whole body of the molecule, while adoption of the hybrid 
33
 state of the A-site tRNA only involves the shifting of the CCA-end into the P-site. This 
hybrid state conformation can bind the ternary complex mimic eEF2, which interacts 
with both subunits to promote translocation through GTP hydrolysis and the transition 
back to a classical state 105,106. This is accompanied by a ribosomal shift in the 
3?direction by one codon and the transfer of the tRNAs from the SSU P- and A-sites to 
the E- and P-sites. 
Figure 16. Positions of tRNAs during translation. 
The relative position of tRNAs during classical, hybrid, and pre-accommodation states. 
Figure modified from Noller 107. 
34
 The transition between the classical and hybrid states above is accompanied by 
dramatic rearrangements in the structure of the whole ribosome. These are embodied by 
the rotation (previously referred to as ?ratcheting?) of the SSU by approximately 6? in a 
counterclockwise motion relative to the LSU, as shown in Figure 17 108?112. The hybrid 
state corresponds to the rotated conformation, while the classical state corresponds to a 
non-rotated conformation. In the rotating SSU, the head region is the most mobile 
element, rotating at the neck by 14? which translates to a movement of up to 25 ?. 
Corresponding structural changes in LSU are far less well known, and will be discussed 
in detail in the next chapter. The transition to a rotated state results in the disruption of 
several non-centrally located intersubunit bridges including B7a and B2b/c. 
Figure 17. Ribosomal intersubunit rotation.
 Subtle rotation of the SSU relative to the LSU during translation. Image modified from 
Schmeing 85. The ?head? domain of the SSU is indicated. 
35
 The recently adopted term ?rotation? in place of ?ratcheting? more accurately 
reflects the more continuous and fluid movements involved in the process: there may be 
up to 50 intermediate conformational states, with the fully rotated and fully non-rotated 
states representing the two extremes 101. As such, the ribosome transits between these two 
conformational states as it progresses through the translation elongation cycle. Recent 
evidence suggests that ternary complex binding and peptidyltransfer occur primarily in 
the context of a non-rotated ribosome, while eEF2 binding and translocation occur in a 
rotated ribosome 113,114,115. A summary of this process is shown in Figure 18.
 Figure 18. The elongation cycle of translation. 
Ribosomal intersubunit rotation drives the elongation cycle and ligand binding. Image 
modified from Schmeing 85. 
36
 Termination
 There are 3 stop codons, also known as nonsense codons (UGA, UAG, and 
UAA), which mark the end of the protein product. There are no cognate tRNAs for stop 
codons. Instead, protein factors called release factors (RFs) bind to the A-site when it is 
positioned over one of the stop codons and trigger a hydrolysis reaction in the ribosome 
active site, resulting in the release of the polypeptide chain from the P-site tRNA.
 Eukaryotes have one omnipotent class I RF (eRF1), which decodes all three stop codons. 
It associates with the class II RF (the GTPase eRF3) in the cytoplasm and binds to the 
ribosome as another ternary complex, resulting in GTP hydrolysis by eRF3 and 
subsequent activation of eRF1 116,117 . Because eRF1 and aminoacyl tRNAs interact with 
the same ribosomal structural elements of the A-site, this release factor may also act as a 
functional tRNA mimic 118 (Figure 19.)  Binding of release factors to the A site induces 
conformational changes similar to those for peptidyltransfer during translation 
elongation. Indeed, the mechanism of peptide release resembles the mechanism of 
peptide bond formation, differing only in the identity of the nucleophile and the 
requirement for protein factors. While an amine is the functional nucleophile during 
elongation, a water molecule plays that role in peptide release; the two mechanisms are 
compared in Figure 20. During elongation, a particular arrangement of the PTC excludes 
water from the active site to avoid premature peptide hydrolysis. During termination, the 
high conserved GGQ (glycine-glycine-glutamine) motif of eRF1 interacts with the PTC 
in a manner analogous to the 3? CCA ends of tRNAs, and helps position an attacking 
water molecule properly for reaction 119?121. 
37
 Figure 19. Eukaryotic release factor is a structural tRNA mimic.  
The similarity of the structures is shown by a side view (left) and a front view (right). The 
N-terminal (N) domain mimics the anticodon arm, the middle (M) domain is equivalent 
to the amino acid acceptor arm and the C-terminal (C) domain is responsible for 
interaction with eRF3. Image modified from Chavatte 122. 
Peptide release leaves the 80S ribosome still bound to the mRNA with deacylated 
tRNA in its P-site and eRF1 in its A-site, all of which need to be disassembled and 
released from the mRNA to allow further rounds of translation. While in bacteria subunit 
and factor dissociation is accomplished by the crowbar-shaped ribosome recycling factor 
(RRF), no such eukaryotic factors have been found 123.  However, though the mechanism 
of action remains enigmatic, it has recently been shown that the ATPase ABCE1 can 
promote ribosome splitting in combination with eRF1 124. The ABCE1 protein is also 
required to recycle stalled ribosomes in the eukaryotic mRNA surveillance pathways no-
 go decay (NGD) and no-stop decay (NSD). The protein Dom34 and its interaction partner 
the GTPase Hbs1 are paralogs of the eRF1-eRF3 system and are similarly recruited as a 
38
 ternary complex to the ribosomal A-site of stalled ribosomes. This assembly in turn 
recruits ABCE1, culminating in ribosome splitting and recycling 124,125. Another possible 
post-termination and ribosome recycling model includes eIF3 induced separation of the 
subunits 126. Lastly, a subunit dissociation step may not be needed at all. A linkage 
between termination and initiation is possible due to an interaction between eRF3 and 
PABP in the closed loop mRNA structure. As such, in some circumstances eRF3-guided
 ribosomes are able to move right back to the beginning of the message along the 
circularized mRNA. Translation is in fact strongly inhibited upon disruption of the eRF3-
 PABP interaction 127. 
Figure 20. Mechanisms of peptidyltranfer and peptide release. 
Both reactions occur in the same active site and begin with the same ground state 
complex, where the labile P-site ester linkage sits in the PTC. In eRF1 induced 
hydrolysis, the C-terminus of the polypeptide chain attached to the P-site tRNA 
undergoes nucleophilic attack at the ester carbon by a water molecule, stabilized by the 
glutamine of the GGQ motif, leading to release of the newly synthesized polypeptide.
39
 Antibiotics and Ribosome Inhibition 
Because of the central role of translation in the cell, as well as the structural 
differences between prokaryotic and eukaryotic ribosomes, protein synthesis inhibitors 
have been the most successful clinical antibiotics to date. Because the ribosome exceeds 
the size of an average antibiotic by four orders of magnitude, it provides a multitude of 
targets: approximately 50% of all existing antibiotics impair translation. The emergence 
of high resolution ribosomal structures in this decade has led to insights into the mode of 
antibiotic interactions and exact mechanisms of action 128?133 . 
There are several general classes of ribosomal antibiotics, most of which are 
obtained from natural producers such as Streptomyces, that can be grouped based on their 
structures and modes of action 134. Antibiotics interfering with almost every step of 
translation are clinically available, some shown in Figure 21. For example, 
orthosomycins and pactamycin bind to the 50S and 30S subunits, respectively, preventing
 formation of the initiation complex. Kasugamycin also prevents initiation by binding in 
the path of mRNA in the 30S subunit and inhibiting initiator tRNA interactions. 
Affecting elongation, sparsomycin blocks tRNA binding at the 60S A-site while 
tetracyclins prevent tRNA accommodation.  Aminoglycosides such as paromomycin can 
bind to the decoding center leading to stabilization of codon:anticodon mismatches and 
subsequent mistranslation. Pulvomycin prevents the binding of aa-tRNA to the eEF1A-
 homolog EF-Tu and hence ternary complex formation, while kirromycin prevents the 
conformational changes of EF-Tu required for GTP hydrolysis. Many types of antibiotics 
interfere with the catalytic activity in the PTC. Puromycin mimics the CCA end of an aa-
 tRNA, enters the ribosomal A-site and gets incorporated into the nascent polypeptide 
40
 chain triggering premature peptide release. Chloramphenicol blocks the peptide exit 
tunnel, leading to an obstruction referred to by Nobel Laureate Thomas Steitz as 
?molecular constipation?. Fusidic acid prevents conformational changes required for 
dissociation of the eEF2-homolog EF-G after GTP hydrolysis, thus trapping the factor on 
the ribosome and stalling translation at the translocation step. Ribotoxins and thiopeptides 
block the binding of EF-G and its structural homologs such as the ternary complex and 
release factors. 
The oxazolidinone class of synthetic antibiotics represents the first new class of 
drug to enter the antibiotic market in over 20 years 135. While their precise mode of action 
is not known, these antibiotics appear to interact with the 50S subunit and inhibit 
initiation or early stages of elongation 136. Interestingly, partial translation of full proteins
 is more detrimental to bacterial cells than complete translational shut down. As a result, 
more powerful antibiotics actually allow a fraction of proteins to escape inhibition. These 
?resistant? proteins stall biochemical pathways and result in toxic intermediates 137. 
Because of the increasingly alarming rate of the development of bacterial resistance and 
cross-resistance to antibiotics, advancing our understanding of the precise mechanisms of 
ribosomal function and drug interactions is crucial to outpace the rapidly evolving 
pathogenic strains.  
41
 Figure 21. Steps of antibiotic inhibition of translation. 
The prokaryotic life cycle of the ribosome is impaired by a variety of clinical drugs at the 
indicated steps. This image is from Sohmen 138. 
42
 Translational Recoding
 As discussed above, canonical translation initiates at a start codon, proceeds 
uninterrupted along an open reading frame (ORF) one triplet codon at a time, and 
terminates at a stop codon yielding a single protein product. However, translation can 
also proceed through a variety of non-canonical pathways, generically referred to as 
translational recoding. While recoding events are intrinsically rare (10-4 ? 10-5 events per 
codon) 139, they can be induced to occur at higher rates at specific sequences by cis-acting 
elements on mRNAs. Such exceptions to translational rules include shifting ribosomes 
into an alternate reading frame during elongation, directing ribosomes to alternate start 
sites during initiation, and suppression or recoding of stop codons 140. Although originally 
identified in viruses, many of such recoding signals have since been documented in both 
prokaryotic and eukaryotic organisms 141.
 Programmed -1 Ribosomal Frameshifting 
A programmed -1 ribosomal frameshifting (-1 PRF) event induces a particular 
fraction of elongating ribosomes to ?slip? back on the message, and out of the original
 ORF, by one nucleotide in the 5? direction. This results in the establishment of a new 
ORF and is followed by the continuation of elongation in the new reading-frame. Thus, a 
-1 PRF signals allows the synthesis of multiple proteins from a single, unaltered RNA 
template. This is a genome condensation strategy that allows viruses, all of which are 
limited by the size of their nucleocapsids, to maximize genomic coding space while 
maintaining genome integrity.  -1 PRF was first identified in the Rous Sarcoma Virus, 
and is utilized by a plethora of other RNA viruses including those causing Acquired 
43
 Immunodeficiency Syndrome (HIV-1) and Severe Acute Respiratory Syndrome (SARS
 CoV) 142?145. The large number of well-characterized viral -1 PRF signals has allowed the 
definition of the parameters and rules governing - 1 PRF efficiency in at least one class of 
signals, shown in Figure 22. A typical -1 PRF signal consists of 3 important elements: a 
?slippery site? of 7 nucleotides over which the ribosomal shift in frame actually takes 
place; a short spacer region of less than 12 nucleotides; and a strong stimulatory structure 
downstream, which is usually a pseudoknot 146?149.  
Figure 22. Structure of a typical  -1 PRF signal. 
The slippery site, where the A- and P-site tRNAs are positioned and the shift in frame 
occurs, is denoted by IUPAC notation (N=any 3 identical nucleotides, W=A or U, and 
H?G). 22 functional slippery sites are known. While a pseudoknot is the most common 
type of stimulatory element, other mRNA structures are capable of filling that role as 
well.
44
 A model of the mechanism of a -1 PRF event is shown in Figure 23. Generally it 
is thought that the function of the pseudoknot is to stall the ribosome and position it over 
the slippery site, making the ?slip? more likely and biologically relevant. The nature of 
the slippery sequence enables re-pairing of the non-wobble bases of both the aa- and 
peptidyl-tRNAs with the -1 frame codons. As this is occurring, the energetic barrier of 
the pseudoknot is overcome by the ribosome, and elongation can continue in the newly 
established reading frame. Frameshift ?attenuators? consisting of hairpin structures 
upstream of the slippery site have been shown to influence rates of frameshifting as well 
150. A number of models have been presented addressing the exact time point and 
mechanism of the shift in frame. More recently, the ?many pathways model? has unified 
previous competing models by demonstrating that -1 PRF products merely represents an 
endpoint accessible by at least three different kinetic pathways 151. 
In addition to enrichment of genomic content, -1 PRF is also used by viruses to 
ensure production of viral gene products in their correct stoichiometric ratios and is 
shown in Figure 24. For example, the L-A virus of yeast has a simple icosahedral 
structure of T=1. In Euclidian solid geometry, the simplest sphere (T=1) is composed of 
60 identical subunits and resembles a traditional soccer ball. Translating this structure to 
viruses, these subunits are the capsid proteins (also called Gag for ??Group specific 
antigen??) which typically self-assemble to form the viral capsid 152. The only other 
component required for L-A virus propagation is a single molecule of an enzyme capable 
of replicating the genome, the viral replicase, or polymerase. Thus, the ideal ratio of 
capsid to replicase is 60:1. The L-A viral -1 PRF signal has evolved to shift ribosomes 
from the Gag open reading frame to the Pol open reading frame at a rate of 1.8%, thus 
45
 producing the desired 60:1 ratio of capsid to replicase proteins 153. Changing this rate by 
mutations in the frameshift signal, mutations in the host translational apparatus, or with 
small molecule inhibitors alters this important stoichiometric ratio and proper virion 
assembly 154. This general concept applies to many other viruses as well, with specific -1 
PRF efficiencies ranging 1-10%. Because the correct frameshift structure and function 
are necessary for viral propagation, targeted altering of these parameters may open up 
new avenues for anti-viral treatments. 
Many functional -1 PRF signals have also been identified in eukaryotic systems 
where their role appears to be quite different from the viral context 155?158. Over 95% of 
??cellular?? -1 PRF events direct elongating ribosomes into premature termination codons, 
including in humans, leading to an inverse relationship between -1 PRF efficiency and 
mRNA half-lives. Studies using endogenous -1 PRF signals from yeast demonstrated that 
these can function as mRNA destabilizing elements through both the nonsense-mediated 
decay (NMD) and the No-Go decay pathways 159,160. In unpublished studies from our 
laboratory, we have shown that endogenous -1 PRF signals from human genes also 
function as mRNA destabilizing elements in mammalian cells through NMD, play an 
important role in regulation of the immune response, and that frameshifting efficiency 
can be regulated through miRNAs. In addition, it was recently demonstrated that levels of 
-1 PRF directly correlate with telomere length in yeast 161. Thus, it is now clear that -1 
PRF is much more widespread and versatile than originally thought, is employed by 
organisms representing every branch in the tree of life, and presents a powerful new 
method of gene regulation.
46
 Figure 23. The mechanism of -1 PRF. 
The SARS coronavirus -1 PRF signal is used as an example. An mRNA pseudoknot 
forces elongation ribosomes to pause over the slippery site which induces a shift by 1 
base in the 5? direction and the subsequent repairing of A- and P-site tRNAs with the new 
-1 frame codons. The pseudoknot is eventually denatured, and elongation in the new ORF 
continues. Image obtained from ViralZone 162. 
47
 Figure 24. Effects of -1 PRF on viral propagation. 
The correct ratio of structural to enzymatic proteins is achieved by a -1 PRF signal in the 
viral genome directing ribosomes into an alternate reading frame at a set rate, resulting in 
the production of the Gag-pol fusion protein. No shift in frame yields the Gag protein 
product. Increase/decrease in the frameshifting rate results in improper virion assembly 
and a non-functional virus. Image adapted from Dinman 154. 
Programmed +1 Ribosomal Frameshifting 
A less widespread and understood example of a recoding event is programmed +1 
ribosomal frameshifting (+1 PRF). Like -1 PRF, this mechanism utilizes cis-acting 
mRNA elements, is employed by viruses and transposable elements to regulate the 
synthesis of structural and enzymatic proteins, and has been found in human, mouse, 
bacteria, and yeast genomes 163?166. Unlike -1 PRF however, +1 PRF induces elongating 
ribosomes to bypass one nucleotide in the 3? direction. A slippery site is required for this, 
48
 while a downstream stimulatory structure is not. Rather, in bacteria ribosomal pausing is 
caused by a low abundance of a particular tRNA or release factor 2 (RF2) as the 
ribosome is positioned over the slippery site 167. While there is only one generally well-
 understood type of -1 PRF signal, +1 PRF signals appear to be case-specific, with 
different mechanisms for different signals. However, recent modeling of E. coli +1 PRF 
signals revealed that the mechanism is influenced by distinct kinetic parameters: 1) 
destabilization of deacylated  tRNA in the E-site, 2) rearrangement of peptidyl tRNA in 
the P-site, and 3) availability of cognate aminoacyl tRNA at the A-site. While all three 
parameters are important for efficient +1 PRF, a rate constant of ~1.9 sec-1 for slippage of 
the P-site tRNA by one downstream nucleotide appears to be the driving force of this 
mechanism 168. It is also greatly enhanced by the presence of a ?hungry codon? in the A-
 site decoding a low abundance tRNA or RF2. 
Missense and Nonsense Suppression 
In addition to reading frame maintenance, translational fidelity can also be 
assessed by missense suppression (the rate at which near- or non-cognate tRNAs are
 mistakenly accommodated), and nonsense suppression (the rate at which suppressing 
tRNAs are incorporated at stop codons, thereby reading through the codon). Nonsense 
suppression is found to be a translational strategy in a number of viruses, often used to 
produce gag-pol fusion proteins as a result of cis-acting elements on mRNAs not unlike 
in PRF 169?171. Similarly, the incorporation of the essential 21st amino acid selenocysteine 
(Sec), the major biological form of the element selenium, occurs through an induced 
nonsense suppression mechanism in all three kingdoms. This requires specific trans-
49
 acting factors and secondary structures in the mRNA to actively recode an in-frame UGA 
from a stop codon to Sec sense codon 172,173. While the human genome only encodes 25 
selenoproteins, variations in these Sec-containing proteins or their synthetic machinery is 
linked to a range of human disorders including cancer. The much more puzzling 22nd
 amino acid pyrrolysine, described in archaea and bacteria and present in only 1%  of 
sequenced genomes, is universally encoded by the stop codon UAG as well 174. 
All recoding strategies provide means for genomic expansion and gene regulation, 
and it is estimated that as many as one-third of inherited genetic disorders are caused by 
frameshift or nonsense mutations 154. In the context of this thesis, recoding mechanisms 
can be used as functional tools to characterize the ribosomal components involved in 
maintaining translational fidelity and function. 
Scope of Work and Thesis Summary 
The progress made over the last 50-60 years in visualizing and understanding the 
structural constituents of the ribosome is remarkable, as shown in Figure 25.  As 
described in this chapter, the general principles of ribosome function are also very well 
understood. However, many important questions still remain about ribosomal mechanics, 
in particular 1) how intersubunit rotation is coordinated, 2) how the different functional 
centers of the ribosome communicate with one another, 3) the nature of the quality 
control mechanisms employed to ensure that only functional subunits are allowed to enter 
the pool of active ribosomes, and 4) the nature and mechanisms of ribosome malfunction 
in disease. 
50
 Studies of the essential ribosomal protein L10 (rpL10) have shed significant light 
on all of these questions. rpL10 was initially identified in yeast in a screen for mutants 
that were synthetically lethal with a deletion of subunit 6 of cytochrome bc1 175. 
Subsequent genetic studies showed that QSR1, as it was originally named, was essential 
for protein synthesis and allelic to RPL10 176. This protein plays an essential role in 
ribosome biogenesis, and its incorporation into the LSU in the cytoplasm is one of the 
last steps of LSU maturation 177. In addition, rpL10 is strategically located near functional 
centers of the ribosome, including the elongation factor and tRNA binding sites. One 
segment of rpL10 also comes within 13 ? of the PTC. As such, it is the closest protein 
element approaching the catalytic RNA core of the ribosome, and the protein has 
consequently been shown to be involved in coordinating aa-tRNA movement near the 
PTC 178. The high level of sequence conservation of rpL10 in all organisms and the 
location close to regulatory and catalytic sites of the ribosome are indicative of its 
importance in ribosome function, and we have referred to it as the ?CPU of the LSU.? In 
additional support of this, recent studies have demonstrated that defects of rpL10 are 
found in patients suffering from autism and acute T-cell acute lymphoblastic leukemia
 (T-ALL) 179,180. 
The work presented in the following chapters further dissects the role of rpL10 in 
the ribosome while also addressing important questions regarding translation in general. 
Chapter 2 describes how a flexible loop of rpL10 near the PTC coordinates ribosomal
 rotation, that the rotational status is important throughout the ribosomal lifecycle, and 
offers a model explaining how pre-60S subunits are functionally monitored during 
ribosome biogenesis. Chapter 3 then demonstrates that malfunction of this loop 
51
 influences T-ALL, offers a mechanistic model of the disease, and proposes an 
explanation for the high incidences of cancer in patients with ribosomopathies. Chapter 4 
focuses on the eukaryotic-specific C-terminal ?tail? of rpL10, its pivotal role in 
recruitment of trans-acting factors and ribosome biochemistry, and how impairment of 
this domain can result in autism. Finally, Chapter 5 summarizes all these findings and 
discusses their significance and future implications.  
Figure 25. The ribosome over 50 years. 
The very first images of ?microsomes? and the modern ribosome at atomic resolution. 
EM image obtained from Kirsch 181, conceptual idea from Dinman 182. 
52
 Chapter 2
 Eukaryotic rpL10 drives ribosomal rotation
 Introduction
 The ribosome is an essential and complex nanomachine that provides a model for 
understanding principles of macromolecular assembly and functional coordination. The 
eukaryotic yeast ribosome is a 3.6 megadalton RNA-protein complex, consisting of 79 
intrinsic ribosomal proteins and 4 ribosomal RNAs (rRNAs) 14. Different biochemical 
functions are spatially separated from one another in the two subunits of the ribosome.  
The small subunit (SSU) contains the mRNA decoding center, while the large subunit 
(LSU) harbors separate regions with distinct functions: the peptidyltransferase center,
 PTC (responsible for catalysis), the peptide exit tunnel, three transfer RNA (tRNA) 
binding pockets, and a single binding site that must distinguish between the two 
elongation factors and release factor in response to specific circumstances. The subunits 
must also interact with one another as a holoenzyme to coordinate a complex series of 
events, particularly allosteric movements that occur throughout the course of the 
elongation cycle.  The two extreme conformational states are termed ?rotated? (aka 
ratcheted or hybrid) and ?non-rotated? (aka classical). How information is exchanged 
over long distances between spatially distinct functional centers, and how these centers 
then work in concert to ensure timely rotation, proper ligand binding, and ultimately 
unidirectional and faithful translation remains largely unclear. Additionally, while the 
roles of the SSU (particularly the head) and tRNAs in ribosome rotation have been 
investigated, the involvement of the LSU in this process has not been explored.
53
 Ribosomal protein L10 [rpL10, aka L16 15] plays essential roles in ribosome 
biogenesis and translational fidelity. Incorporation of rpL10 into the LSU in the 
cytoplasm constitutes a late step of LSU assembly 183. In particular, rpL10 works in 
conjunction with Shwachman-Diamond protein Sdo1p and the eEF2-like GTPase Efl1p 
to promote the release of the anti-association factor Tif6p and the nuclear export adapter 
Nmd3p, the last steps of 60S maturation 47,184,185. Thus, large subunits lacking rpL10 are 
unable to join with the SSU. rpL10 is located near the corridor through which aminoacyl-
 transfer RNAs (aa-tRNAs) move during the process of accommodation and is involved in 
tRNA movement through this structure 178. It is also located near several other functional 
centers of the LSU, including the PTC, the A-site finger (H38), the elongation factor 
binding site, and the GTPase associated center (see Figure 26). The C-terminus of rpL10
 contacts 5S rRNA, which interacts with rpL5 and rpL11 at the head of the central 
protuberance. Thus, rpL10 is well positioned to act as a sensor of activity near the PTC, 
and transduce that information to other functional centers to coordinate ribosome 
function. 
An essential internal loop of rpL10 (aa 102-112, previously called the ?P-site 
loop?) that makes the closest approach to the PTC (13 ?) of any ribosomal protein  is 
crucial for Tif6p and Nmd3p release 186. Mutagenesis of the loop revealed two classes of 
mutants based on their effects on ribosome biogenesis 187. The current study employs the 
strongest representative mutants of these two classes: S104D (improper subunit joining 
and 60S biogenesis defect) and A106R (high 60S/40S ratio).   Here, we present evidence 
that these mutations promote opposing effects on the natural equilibrium between the two 
extreme conformational states of the ribosome. Structural and biochemical analyses of 
54
 empty ribosomes demonstrate that the S104D mutation drives the ribosome towards the 
rotated state, favoring binding of eEF2 and disfavoring binding of eEF1A/aa-tRNA/GTP 
(elongation ternary complex) and inhibiting peptidyltransfer.  In contrast, A106R causes 
empty ribosomes to distribute toward the non-rotated state, favoring elongation ternary 
complex binding over eEF2, and leaving peptidyltransferase activity unaffected. We also 
show that the non-rotated conformation favors Sdo1p binding, stimulating binding of 
tRNAs to the A-site. Further, Sdo1p competes with acetylated-aa-tRNA for binding to the 
P-site and inhibits peptidyltransfer, suggesting that Sdo1p stabilizes the non-rotated state 
through binding the P-site.  Large scale rRNA chemical modification analyses reveal 
distinct information transmission pathways originating from rpL10 in the heart of the 
LSU and emanating throughout the LSU and the SSU.  
These observations lead us to propose that the rpL10 loop plays a central role in 
much of the ribosomal life cycle by helping to set the conformational status of the LSU, 
coordinate intersubunit rotation, and communicate this information to the decoding center 
on the SSU.  During late LSU biogenesis, the loop senses Sdo1p recruitment to the P-site, 
initiating a ?test drive? to ensure the functionality of pre-60S subunits. After ensuring 
proper 80S assembly, the loop monitors the tRNA occupancy status of the PTC A-site: in 
the absence of A-site ligand (aa-tRNA) it can sample this space, while the introduction of 
ligand displaces it.  We suggest that the positioning of the rpL10 loop determines which 
state the LSU assumes in a process involving cascades of allosteric interactions that link 
functional centers in the LSU with those in the SSU. The downstream effects of 
rotational disequilibrium are wide-ranging, impacting translational reading frame 
maintenance, the ability to discriminate between cognate and near- and non-cognate 
55
 codons, and termination codon recognition. Mutants of ribosomal protein L3 that confer 
opposing effects on ribosome structure and function can suppress the structural, 
biochemical, and functional defects of the rpL10 loop mutants by re-establishing the 
normal rotational equilibrium.  In sum, we propose that the rpL10 loop is a master 
controller of ribosome structure and function, influencing critical steps in both ribosome 
assembly and biogenesis, and the protein-synthetic phase of elongation. We suggest that 
the unidirectionality of translation is aided by this intrinsic feature of the ribosome, and 
that the LSU alone has the ability to independently influence intersubunit rotation. 
56
 Figure 26. rpL10 is strategically positioned in the core of the LSU.
 a. The big picture: rpL10 in the context of the subunit interface of the LSU. b. A close-up 
view of rpL10 and the local environment. The hypothetical loop structure is circled and 
indicated by dashed red lines and the approximate positions of S104 and A106 are 
indicated. The protein is situated between Helices 38 and 89, and appears to be an 
extension of Helix 39. It is located in close proximity to several functional centers of the 
LSU including the PTC, the aa-tRNA accommodation corridor, and the elongation factor 
binding site. It is also positioned to communicate with the SSU through Helix 38 and the 
5S rRNA. 
57
 Results
 The S104D and A106R mutants promote opposing effects on ligand binding to the 
ribosomal A- and P-sites. Saturation mutagenesis of the rpL10 loop revealed two 
classes of mutants based on their ribosome biogenesis defects.  Class I mutants, 
exemplified by rpL10-S104D, displayed halfmer polysomes which are indicative of 
subunit joining defects on the mRNA. Class II mutants, exemplified by rpL10-A106R, 
exhibited higher 60S/40S subunit ratios 187. A summary of additional genetic and 
functional analyses of loop mutants displaying the strongest defective phenotypes is 
depicted in Table 1. Of these, rpL10-S104D and rpL10-A106R were chosen for further 
examination. While the rpL10 loop was not resolved by X-ray crystallography 15, cryo-
 EM studies suggested that the tip of this loop is in close proximity to the P-site tRNA
 188,189.  Thus, it was speculated that P-site ligand binding would be affected by these 
mutants.  However, steady state binding of acetylated-[14C]Phe-tRNAPhe to 80S 
ribosomes purified from isogenic strains expressing wild-type and mutant forms of rpL10 
showed no significant differences in binding of this model P-site ligand to the ribosomal 
P-site (Figures 27a and 28a). In contrast, the mutants showed significant changes in their
 ability to bind elongation ternary complex ([14C]Phe-tRNAPhe?eEF1A?GTP) to the 
ribosomal A-site: specifically the rpL10-S104D mutant promoted a 2-fold increase in KD
 for this ligand, while the rpL10-A106R mutant promoted a 2-fold decrease (Figures 27a 
and 28b).  To ascertain whether these differences were due to changes in affinity for the 
elongation factor or the aa-tRNA itself, the same experiment was performed using 
[14C]Phe-tRNAPhe alone (i.e. non-enzymatic binding).  Under these conditions, the aa-
 tRNA binding defect of the rpL10-S104D mutant was exacerbated (~6-fold increased KD
58
 relative to wild-type ribosomes), and this was partially ameliorated by addition of eEF1A
 and GTP (Figures 27b and 28d).  In contrast, the binding of [14C]Phe-tRNAPhe to wild-
 type or rpL10-A106R ribosomes was not influenced by  eEF1A. These data indicate that 
the tRNA binding defects are intrinsic to the ribosomal A-site and not to binding sites 
unique to eEF1A.   
Eukaryotic elongation factor 2 (eEF2), which drives translocation, is a structural 
mimic of the elongation ternary complex, and includes a tRNA-like domain (Domain IV) 
that also interacts with the ribosomal A-site of the decoding center 105. Steady-state 
binding assays of purified eEF2 to 80S ribosomes, as monitored by the extent of 
diphtheria toxin [14C]- ADP ribosylation, also revealed reciprocal changes in KD values 
for the two mutants (Figures 27a and 28c).  Importantly, while the S104D mutant 
ribosomes exhibited decreased affinity for aa-tRNA and elongation ternary complex, they 
displayed increased affinity for eEF2.  Conversely, A106R mutant ribosomes displayed 
increased affinity for aa-tRNA and elongation ternary complex and decreased affinity for 
eEF2.  
Sdo1p is required at a late step in 60S maturation, coupling the GTPase activity of 
the Efl1p to release of Tif6p 47.  Steady state binding assays using purified [32P]-labeled 
Sdo1p revealed that mutant ribosomes displayed defects in binding this ligand similar to 
those observed with elongation ternary complex, i.e. the rpL10-S104D mutant promoted 
decreased affinity for Sdo1p, while the rpL10-A106R mutant promoted increased affinity 
for it (Figures 27c and 28e). Sdo1p did not bind mRNA or tRNA alone, and the Sdo1p?N 
mutant lacking the N-terminal FYSH domain that is essential for protein function 190
 displayed negligible binding to wild-type ribosomes (Figure 28e). Scatchard plot analyses 
59
 demonstrated that Sdo1p binds to a single site on the ribosomes (Figure 28f). 
Competition assays in which wild-type ribosomes were pre-incubated with increasing 
amounts of Sdo1p revealed that this protein stimulated binding of aa-tRNA to ribosomes 
in a concentration dependent manner, while inhibiting both binding of Ac-aa-tRNA to the 
P-site and peptidyltransferase activity (Figure 27d). 
L103S S104D C105G A106R A106P A106L
 Growth Slow Very slow Very slow Slow
 Temperature 
sensitivity Yes
 Tif6 
suppression Yes Yes Yes
 Cycloheximide Hs S S
 Anisomycin Hs R R R
 Polysome 
profile
 ? 60S ? mer ? mer ? 60S ? mer ?80S
 Table 1. Summary of genetic and functional analyses of L10 loop mutants. 
Cell growth at various temperatures (30?C, 15?C, 37?C), the ability of Tif6 mutants to 
suppress growth defects when co-expressed in mutant yeast cells, and the sensitivity to 
antibiotics were monitored by 10-fold dilution spot plating. S = sensitive, Hs = 
hypersensitive, R = resistant. Sucrose gradient analyses of mutant cells revealed halfmer 
defects (1/2 mer), elevated 60S/40S ratios (? 60S), or reduced polysomes (?80S). Blank 
spaces = wild-type phenotype. The S104D and A106R mutants were chosen for further 
examination based on these results. 
60
 Figure 27. rpL10 loop mutants have opposite effects on ligand binding to the A- and 
P-sites.
 Panels a - c: Steady state binding of indicated ligands to ribosomes isolated from cells 
expressing wild-type rpl10 and the rpl10-S104D and rpl10-A106R mutants. a.
 Dissociation constants obtained from binding assays of Ac-aa-tRNA to the P-site and 
ternary complex to the A-site as monitored by filter binding, and eEF2 as monitored by 
extent of ribosylation of unbound protein. b. Titration of eEF1A into aa-tRNA binding 
reactions similar to ternary complex in panel a; 1 = amount for maximum activity. c.
 Dissociation constants obtained from assays of binding of Sdo1p as monitored by levels 
of radiolabel detection in ribosomes. d. Competition assays. Binding of aa-Phe-tRNAPhe
 or Ac-Phe-tRNAPhe at half maximal concentrations were monitored in the presence of 
increasing amounts of Sdo1p. Peptidyltransferase activity was similarly monitored.  Bars 
indicate s.e.m. (n=4 for a-b, n=3 for c), * P < 0.05, ** P < 0.01 (compared to wild type 
unless noted).   
61
 Figure 28. Primary ligand binding data related to Figure 27. 
Panels a - c: Single site binding isotherms for ribosomes and various ligands. Binding of 
Ac-aa-tRNA to the P-sites (panel a), ternary complex to the A-site (panel b), and eEF2  
(panel c) to ribosomes isolated from wild-type, rpl10-S104D  and rpl10-A106R  cells. d. 
Titration of eEF1A into aa-tRNA binding reactions similar to panel b. e. Single site 
binding isotherms for indicated ribosomes and Sdo1p.  Sdo1p?N is an N-terminal 
deletion mutant that cannot bind ribosomes. Sdo1p does not bind significantly to mRNA 
or tRNA alone.  f. Scatchard plot of Sdo1p binding to wild-type ribosomes indicates 
single site binding. 5 pmoles of ribosomes in 120 ?l total volume were used at each 
dilution throughout. Bars indicate standard error of the mean. 
62
 The A106R and S104D mutants have opposing effects on the rotational equilibrium 
of the ribosome. During the elongation cycle, the ribosome transits through a large 
number of conformations, characterized at the extremes by states described as non-
 rotated and rotated 15,101,104,109,110,112.  Recent single molecule experiments using E. coli
 ribosomes have shown that the aa-tRNA?EF-Tu?GTP elongation ternary complex has 
higher affinity for non-rotated ribosomes than rotated ribosomes, and that the converse is 
true for EF-G 191. Thus, the ligand binding data described above suggested that the 
A106R and S104D mutants drive the structural equlibria of ribosomes toward either the 
non-rotated (substrate for binding elongation ternary complex) or rotated (substrate for 
eEF2) states respectively.  While chemical modification profiles and atomic resolution 
structures are well-defined for non-rotated and rotated E. coli ribosomes 110,192?196, no 
equivalent information exists regarding yeast ribosomes.  Thus, in order to examine the 
rotational status of yeast ribosomes, it was first necessary to demonstrate that chemical 
protection patterns of well-defined E. coli and yeast ribosome complexes are similar.   
For E. coli, the standard for non-rotated ribosomes are those which are primed with 
polyU and contain N-Ac-PhetRNAPhe in the P-site, while rotated ribosomes are primed 
with polyU and contain deacylated tRNA in the P/E site and EF-G-GDPNP 195.  Similar 
complexes were prepared using yeast ribosomes (rotated yeast ribosomes contained eEF-
 2-GDPNP instead of EF-G-GDPNP). These two complexes, plus empty wild-type control 
ribosomes were chemically probed with 1M7 (or DMSO only controls), and base 
reactivities were assessed by hSHAPE and quantified using ShapeFinder 197,198.  
Representative complete electropherograms for these three samples are shown in Figure 
29.  
63
 Figure 29. Visualization of chemical probing data by hSHAPE. 
A representative electropherogram segment from chemical probing in H63 of the LSU for 
the indicated complexes showing differences in chemical reactivities (peak areas). The 
areas under the 1M7 reactivity peaks are integrated to generate quantitative data and 
enable statistical analyses (see Table 2). Ethanol is the negative control. 
64
 Table 2 shows that the chemical modification profiles of landmark E. coli rRNA 
bases are very closely matched by their yeast counterparts in the non-rotated and rotated 
states (compare columns 3 and 4, with columns 6 and 7).  Columns 8 - 10 of Table 2 also 
depict conversion of the statistically normalized reactivity data to a scale from 0 ? 4, with 
4 being most reactive 197,199. Occupation of the P-site by Ac-aa-tRNA (non-rotated 
ribosomes) resulted in nearly identical chemical modification patterns at equivalent 
rRNA bases in the P-sites of the small and large subunits, and in the large subunit E-sites 
(indicated by color coded boxes in Table 2). Similarly, the chemical modification patterns 
in the large subunit P- and E-sites of rotated yeast ribosomes closely matched published 
data for rotated E. coli ribosomes.  Additionally, the non-rotated control was verified 
biochemically by comparing the ternary complex and eEF2 steady state binding profiles 
(Figure 30). Note that the rotated control could not be biochemically assessed as the 
factor binding site was already occupied by eEF2-GDPNP.  Collectively, these data 
demonstrate that these two complexes represent non-rotated and rotated yeast ribosomes.  
65
 Table 2. Comparison of yeast rRNA chemical modification profiles with non-rotated 
and rotated E. coli standards.
 1. E. coli rRNA base number. 2. Corresponding S. cerevisiae rRNA base number. 3.
 Chemical modification profile comparing polyU primed E. coli ribosomes containing Ac-
 aa-tRNA in the P-site to empty ribosomes using nomenclature as described 195. 4.
 Chemical modification profile comparing polyU primed E. coli ribosomes containing 
deacylated tRNA in the P-site plus EF-G-GDPNP to empty ribosomes using 
nomenclature as described 195. 5. Intensity of normalized hSHAPE signals in empty wild-
 type yeast ribosomes obtained from integrated areas under electropherogram peaks in 
arbitrary units as described 197. 6. Intensity of normalized hSHAPE signals in polyU 
primed wild-type yeast ribosomes containing Ac-aa-tRNA in the P-site. 7. Intensity of 
normalized hSHAPE signals in polyU primed wild-type yeast ribosomes containing 
deacylated tRNA in the P-site plus eEF2-GDPNP. 8-10. Arbitrary units from columns 5-7 
converted to statistically determined reactivity values as described 199. Boxes are colored 
to highlight congruent results where the extent of base reactivity/protection is determined 
relative to empty wild-type ribosomes.
 E. coli 
nucleotide1
 Yeast 
nucleotide2
 E. coli P site 
occupied 
compared to empty 
70S3
 E. coli EF-G-GDPNP 
compared to empty 
70S4
 80S empty5
 80S P site 
occupied6
 80S eEF2-
 GDPNP7
 80S empty 
reactivity8
 80S P site 
occupied 
reactivity9
 80S eEF2-
 GDPNP 
reactivity10
 A532 A579 Weakly protected 29.209533 19.3633623 23.26285 3 2 2
 G693 G904 Protected 21.486294 2.30601785 5.4394358 3 1 1
 A794 A1005 Protected 5.0085212 0.63071889 2.8545354 1 0 0
 C795 C1006 Protected 0.3092358 0.22039727 0.0685841 1 0 0
 G926 G1150 Protected NI NI NI
 G966 U1191 Weakly protected NI NI NI
 G1338 G1575 Weakly protected 52.590355 37.046344 10.990177 3 2 2
 A1916 A2259 Weakly protected NI 1.46208226 0.7667143 NI 0 0
 A1918 G2261 Weakly protected 16.591722 12.9900386 6.2834256 2 1 2
 U1926 U2268 Weakly protected 237.04377 0.40327763 18.495792 4 0 2
 G2252 G2620 Protected Protected 7.3710908 2.34212702 3.3526858 2 1 1
 G2253 G2621 Protected Reactive / unchanged 5.7993268 2.77326129 5.8762237 2 1 2
 A2439 A2808 Protected Protected NI NI 1.1447785 0 0 1
 A2451 A2820 Protected Protected 70.682962 0.27775391 11.154668 4 0 2
 G2505 G2874 Weakly protected 1.3841216 1.58462719 1466.4545 1 1 4
 U2506 U2875 Protected 8.4806306 4.65375882 0 2 1 0
 U2584 U2953 Protected 1.8857207 0.57815281 0.9600475 1 0 0
 U2585 U2954 Protected 80.722973 NI 11.056566 4 NI 2
 G2112 A2456 Reactive / unchanged Protected 1.2368216 2.1285347 0.1310783 1 2 0
 G2116 U2460 Reactive / unchanged Protected 2.7718079 3.18348329 1.6371288 1 2 1
 C2394 C2764 Reactive / unchanged Protected 0.4569259 0.29872902 0.0868647 0 0 0
 Weakly protected
 Protected
 Reactive / unchanged
 NI=Not Interpretable 
SSU P-site
 LSU P-Site
 LSU E-site
66
 Figure 30. Biochemical verification of the rotational controls. 
Dissociation constants of ternary complex and eEF2 for empty wild-type and control non-
 rotated (+Ac-Phe-tRNAPhe) ribosomes. Bars indicate s.e.m. (n=4).
 Having established standards defining the chemical reactivity profiles of non-
 rotated and rotated yeast ribosomes, these were used to determine the rotational statuses 
of mutant ribosomes. To this end, the chemical protection profiles of ?landmark? basepair 
interactions located in several universally conserved intersubunit bridges 15,85,111,112 were 
examined using vacant ribosomes. Although FRET experiments showed that vacant 
bacterial ribosomes are not as structurally dynamic as pre-translocation ribosomes (i.e. 
containing deacylated tRNA in the P site) 108, we nonetheless chose to make comparisons 
using vacant ribosomes for two reasons: 1) to monitor the intrinsic influence of the L10 
internal loop on ribosome conformational states, i.e. in the absence of trans-acting factors 
(e.g. tRNAs), and 2) because ribosomes harboring deacylated tRNAs in the P-sites alone 
do not occur in physiological conditions.   
67
 The B7a intersubunit bridge undergoes dramatic rearrangements during ribosome 
rotation 111.  Specifically, when the ribosome is in the non-rotated state, E. coli A702 
(yeast G913) of the small subunit rRNA interacts with E. coli A1847 (yeast A2207) of 
the large subunit rRNA, protecting both bases from chemical attack. This interaction is 
disrupted when the ribosome assumes the rotated state, rendering both bases susceptible 
to chemical modification (Figures 31 and 32a). Kethoxal was used to probe G913, and 
1M7 was used to probe A2207, and the extent to which these bases were modified in 
purified empty isogenic wild-type and mutant ribosomes, and with control rotated and 
non-rotated ribosomes was quantitatively assessed and normalized using hSHAPE 199.  
Figure 32b and 32c show that both of these bases were reactive along a continuum, 
beginning with non-rotated wild-type control (least reactive) to rpL10-A106R, empty 
wild-type, rpL10-S104D, and finally rotated wild-type control (most reactive). The 
intermediate peak heights observed with empty wild-type ribosomes are consistent with 
the view that the 80S ribosome is free to transit between the two states in the absence of 
ligands 101. 
68
 Figure 31. Structure of the B7a intersubunit bridge in the non-rotated and rotated 
states in E. coli ribosomes. 
The interaction between A1847 in the LSU and A702 in the SSU is disrupted upon 
rotation, rendering both bases susceptible to chemical modification. Structures were 
generated in PyMOL using the most recent high resolution structures of bacterial 
ribosomes 109. 
69
 Figure 32. rpL10 loop mutants alter the rotational equilibrium of the ribosome.
 a. The yeast B7a intersubunit bridge. In the non-rotated state, A2207 (25S rRNA) forms 
an interaction with G913 (18S rRNA). In the rotated state, that interaction is disrupted, 
and the marked 2? OH-group on A2207 becomes accessible to modification by 1M7.  
Similarly, marked atoms on G913 become accessible to modification by kethoxal upon 
rotation. b. Reactivity peaks obtained by hSHAPE after probing of the landmark base 
A2207 (arrows) at the LSU side of the B7a intersubunit bridge with 1M7. c. Reactivity 
peaks obtained by hSHAPE after chemical probing of the landmark base G913 at the 
SSU side of the B7a intersubunit bridge with kethoxal. Data were assigned five levels of 
reactivity: 0 (for less than a trace?s median value), 1 (for a value between the median and 
the mean), 2 (for a value between the mean and the 1st standard deviation), 3, (for a value 
between the 1st and the 2nd standard deviation) and 4 (for values above the 2nd standard 
deviation). Mean, median and standard deviation were calculated on a per trace basis.  
70
 Additional key rRNA bases known to undergo structural changes during 
intersubunit rotation were also probed. For example, the B2a intersubunit bridge, formed 
between the distal loop of H69 of the LSU and h44 of the SSU is important for substrate 
selection on the ribosome 200, and bases comprising this bridge undergo significant 
rearrangement during subunit rotation, albeit not as dramatic as the B7a bridge 112. Table 
3 shows that bases involved in the B2 bridge are less reactive in non-rotated control and 
rpL10-A106R mutant ribosomes than their rotated control and rpL10-S104D 
counterparts. The B3 bridge was utilized as an internal control because it is not disrupted 
during intersubunit rotation, and is thought to be the pivot around which the subunits 
rotate.  Consistent with this, no significant changes in B3 base reactivities were observed.  
The aa-tRNA accommodation corridor (AC) closes upon ribosome rotation, rendering the 
?gate bases? less reactive 200?202. This is reflected as increased chemical reactivities of 
these bases (U2860, U2924 and U2926) in non-rotated control and A106R mutant 
ribosomes as compared to S104D and rotated controls. Additionally, specific bases in the 
Sarcin/Ricin loop are reactive in non-rotated E. coli ribosomes 195.  The same protection 
patterns are observed at the analogous yeast rRNA bases (U3023 and A3027).  
Collectively, the analyses shown in Table 3 support the hypothesis that the S104D mutant 
shifts the structural equilibrium of 80S ribosomes toward the rotated state and that the 
A106R mutant shifts it toward the non-rotated state.  
71
 Region rRNA base Non-rotated A106R S104D Rotated
 B7a A2207 2 2 4 4
 G913 (SSU) 1 2 4 4
 B2a U2258 2 2 3 4
 A2262 0 0 1 2
 C1644 (SSU) 1 1 2 2
 G1645 (SSU) 0 0 2 1
 B3 U2301 0 0 1 1
 G2302 0 0 0 0
 A1655 (SSU) 1 2 2 2
 U1656 (SSU) 2 2 2 2
 AC U2860 1 2 0 0
 U2924 4 2 0 1
 A2926 4 2 1 1
 SRL U3023 1 1 0 0
 A3027 2 3 0 0
 Table 3. Establishing the rotational status of mutant ribosomes.  
Non-rotated control yeast ribosomes were primed with polyU and contained Ac-Phe-
 tRNAPhe in the P-site.  Rotated control ribosomes were primed with polyU, loaded with 
deacylated Phe-tRNA, and incubated with eEF2-GDPNP. These complexes, along with 
salt-washed empty rpL10-A106R and rpL10-S104D ribosomes were chemically probed 
with 1M7 or kethoxal and analyzed by hSHAPE. B7, B2a and B3 denote the probed
 intersubunit bridges.  AC: Accommodation Corridor. SRL: Sarcin/Ricin Loop. Numerical 
values reflect statistically normalized reactivities on a scale of 0 - 4, with 4 being most 
reactive 197,199.
72
 The S104D and A106R mutants define local and long distance changes in rRNA 
structure corresponding to ribosome rotational status.  Having established the 
rotational status of the mutant ribosomes, 1M7 and hSHAPE were used to quantitatively 
assess the chemical reactivities of approximately half (~2000 nt) of the rRNA content of 
salt-washed empty 80S ribosomes purified from isogenic cells expressing wild-type and 
the mutant forms of rpL10.  The highly reproducible quantitative data enabled subtraction 
analyses for each base probed.  Figures 33-37 employ a heat map based approach in 
which the reactivity data obtained from A106R mutant ribosomes was subtracted from 
data obtained from S104D mutant ribosomes.  This analysis yields maps highlighting the 
differences in rRNA base reactivities between the conformational states of the two 
mutants.  In these maps, warmer colors signify higher reactivity, and implicit higher 
flexibility of rRNA bases in S104D relative to A106R, while cooler colors correspond to 
lower reactivity, and implicit higher structure and constraint. Reactivity scale numbers 
denote the extent of the differences with each step in color as statistically determined 199.  
These are mapped onto 2-dimensional flat rRNA maps in Figures 33-35, onto 3-
 dimensional atomic resolution structures in Figures 36-37, and are summarized in Figure
 43c.  
Significant differences between the two mutants were observed in the core of the 
PTC, the rRNA structure most proximal to the rpL10 loop (Figure 33). Three of the four 
bases proposed to play central roles in the induced fit model of peptidyltransfer 203, i.e. 
G2922 (E. coli G2553), U2924 (E. coli U2555), and U2954 (E. coli U2584) were less 
reactive in the rpL10-S104D mutant as compared to rpL10-A106R suggesting a role for 
yeast rpL10 in this process. The universally conserved A2819 (E. coli A2450), G2874 (E. 
73
 coli G2505), and U2875 (E. coli U2506) that constitute the ?entrance? to the PTC 202 also 
showed dramatic differences in chemical reactivities (Figures 33 and 36). 
Moving outward from the loop, rpL10 is framed by Helices 38 (the A-site finger), 
39, 43 and 89.  Significant differences in chemical reactivities between the two mutants 
are seen in all of these structures (Figures 33 and 34), suggesting that structural 
rearrangements involving the flexible loop may be transduced through the body of rpL10 
to neighboring rRNA structural elements. Focusing on these elements in more detail, 
Helices 89 and 90 - 92 (Figure 33) form the ?accommodation corridor? through which 3? 
ends of aa-tRNAs transit as they enter the PTC 202.  In this structure, U2860 in H89 (E. 
coli U2491) and U2924 in H92 (E. coli U2555) interact to ?close? the accommodation 
corridor in the rotated state.  Conversely, they do not interact in the non-rotated state and 
should be more extensively chemically modified when the accommodation corridor is 
open. In support of our model, the difference map shown in Figure 33 and summarized in
 Table 2 demonstrates that these bases are more protected from chemical attack in S104D 
mutant ribosomes as compared to A106R mutants. 
rpL10 also interacts directly with Helix 39.  Examination of Figure 34 reveals 
large scale structural differences between the two mutant ribosomes extending from this 
helix to Helix 44 (PTC-distal).  Importantly, these include the GTPase activating center 
(GAC) and the H43-44 structure, upon which the P0/P1/P2 (E. coli L7/L12) stalk is 
assembled 204.  This region of the LSU interacts with both the elongation ternary complex 
and eEF2 at different stages during the elongation cycle.  As discussed below, we suggest 
that the structural differences identified here trace an information transmission pathway 
that helps the ribosome distinguish between different trans-acting factors at different 
74
 points in its lifecycle. One such transmission pathway is shown in Figure 37: here, the 
reactivity data from Figure 34 is plotted onto the yeast ribosome high resolution 3D 
structure to illustrate a network of rRNA helices connecting the PTC-proximal and PTC-
 distal portions of rpL10. 
The three site allosteric model posits that structural changes in the ribosomal A 
and E sites are linked 205, and studies from our laboratory have identified an extensive 
network of rRNA structural changes extending along the entire path taken by tRNAs as 
they transit the ribosome 178,201,206,207. Examination of the chemical modification patterns 
in Helices 82 ? 88 (Figure 33), which trace the path of tRNAs from the A- to the E-sites 
in the LSU, reveals extensive differences in rRNA base reactivities between the two 
mutants consistent with linkage between the A-and E-sites.  
75
 Figure 33. Structural probing analysis of 3? of the LSU. 
Chemical probing analysis using 1M7 and hSHAPE, showing S104D ? A106R difference 
map spanning Helices 61 to the 3? end of 25S rRNA. PTC: peptidyltransferase center. 
SRL: sarcin ricin loop. The scale at right indicates the extent of differences in reactivities 
with each number corresponding to one standard deviation from the mean reactivity as 
previously described 199. 
76
 Figure 34. Structural probing analysis of 5? of the LSU. 
Heat maps showing differences in 1M7 reactivities between S104D and A106R (left) and 
G81D and A106R (right) spanning Helices 38 ? 44.  GAC: GTPase-associated center.
77
 The distal tip of H38 forms the LSU partner of the B1a intersubunit bridge.  
Although no differences in base reactivities were observed in this structure (Figure 34), 
presumably because it is intrinsically highly mobile 15,193,208, changes were observed in its 
SSU partner in the distal loop of h33, part of the 3? major or ?head domain? of the SSU 
(Figure 35). A block of changes in rRNA base reactivities were also observed along the 
universally conserved helices 32 ? 35 in the 3? major domain of the 18S rRNA.  These 
map to the mRNA entrance tunnel on the SSU opposite the decoding center (see Figure 
43c).  In the 3? minor domain, A1755 (E. coli A1492), which plays a central role in 
stabilizing cognate codon:anticodon interactions in the decoding center 84, is much more 
protected from chemical modification in rpL10-S104D compared to rpL10-A106R 
ribosomes (Figure 35).  The head of the SSU undergoes a dramatic series of 
rearrangements during intersubunit rotation 109,209. Multiple differences in rRNA base 
reactivities were also noted throughout the head domain (h38 ? h43), providing evidence 
that the rpL10 loop may influence intersubunit rotation through the B1b/c bridge in a 
pathway involving 5S rRNA and rpL11.  This is supported by pronounced differences in 
rRNA chemical protection patterns at the tip of H88 (Figure 33), which interacts with 
rpL11 as a monitor of P-site tRNA occupancy 206. This pathway is shown in Figure 43c.
78
 Figure 35. Structural probing analysis of the SSU. 
S104D ? A106R difference map of SSU rRNA encompassing the 3?-major, and 3?-minor 
domains. 
79
 Figure 36. 3D reactivity data of the PTC. 
Reactivity data from Figure 33 plotted onto 3D structures, showing the critical bases 
A2819, G2874 and U2875 forming a triangle in the core of the PTC.
 Figure 37. 3D reactivity data of the factor binding site. 
Superimposition of data from Figure 33 onto the yeast ribosome high resolution structure. 
80
 Identification of important points of contact through which structural changes in 
rpL10 are communicated to the surrounding rRNA.  For structural changes in the 
loop of rpL10 to be broadly communicated to both subunits, information needs to be 
transduced through points of contact between rpL10 and local rRNA structural elements. 
For example, structural changes at amino acid R7, located in the N-terminal ?hook? of 
rpL10, are involved in opening and closing the accommodation corridor 178. Examination 
of mutants identified in a previous genetic screen of rpl10 mutants suggested two 
additional candidates: F94 and G81 (Figure 38). F94 interacts with H38, and the F94I 
mutant was chosen for deeper analyses because of its strong resistance to anisomycin (not 
shown), a competitive inhibitor of aa-tRNA binding to the A-site.  Replacement of the 
aromatic phenylalanine with isoleucine (F94I) produced ribosomes with higher affinity 
for elongation ternary complex and decreased affinity for eEF2, i.e. distributed toward 
the non-rotated state similar to the A106R mutant (Figure 39a). hSHAPE analysis 
revealed that rRNA bases proximal to F94 (Figure 39d, circled) were deprotected in the 
F94I mutant, presumably due to loss of interactions involving the aromatic ring of 
phenylalanine. Interestingly, bases further down the H38 structure (Figure 39d, boxed) 
that interact with the aa-tRNA D-loop showed decreased reactivities similar to an rRNA 
mutant of these bases that also displayed increased affinity for elongation ternary 
complex 210.  G81 is located at the opposite end of the body of rpL10, closer to the 
solvent exposed (PTC-distal) side of the protein, and is one of the amino acid residues 
closest to the elongation factor binding site (H43).  The G81D mutant was chosen 
because of its anisomycin hypersensitivity (not shown), and it promoted the opposite 
effects, i.e. decreased affinity for elongation ternary complex and increased affinity for 
81
 eEF2 (Figure 39a-c).   hSHAPE analysis also revealed that rRNA bases comprising the 
eEF2 binding site and the GTPase-associated center in G81D underwent changes in 
reactivity similar to those observed with S104D (Figure 34). These findings suggest that, 
like S104D, G81D distributes ribosomes toward the rotated state. 
Figure 38. Points of rpL10 important for intersubunit rotation. 
Important bridges between L10 and the surrounding rRNA found to be involved in inter-
 subunit rotation are shown in green. U2860, the H89-side of a closed accommodation 
corridor, is shown in blue. Bases with increased reactivities in H39 are shown in orange. 
Bases with decreased reactivities in H43 are shown in purple, with the base proposed to 
contact G81 starred. The bases in H38 that interact with the phenyl rings of F94 are 
shown in cyan.  
82
 Figure 39. Primary data related to rotational mutants from Figure 38.
 a. Dissociation constants of RPL10, rpl10-F94I and rpl10-G81D ribosomes with 
ternary complex and eEF2. Bars indicate s.e.m. (n=4, * P < 0.05). b and c. Single site 
binding isotherms for data shown in panel a.    d. hSHAPE difference map of F94I 
minus S104D spanning Helix 38 ? Helix 40.
83
 Effects of rpL10 loop mutants on peptidyltransferase activity and translational 
fidelity. A single round, puromycin-based assay of peptidyltransferase activity 211
 revealed that S104D mutant ribosomes promoted approximately 60% of 
peptidyltransferase activity relative to wild-type and A106R mutant ribosomes (Figures 
40a-b).  As peptidyltransfer occurs in the context of the non-rotated state, this further 
supports the hypothesis that the rpL10-S104D mutant drives the equilibrium of ribosomes 
toward the rotated state. Programmed -1 ribosomal frameshifting (-1 PRF) directed by L-
 A virus-derived sequence requires slippage of both A- and P-site tRNAs 154, while Ty1
 directed +1 PRF only requires slippage of P-site tRNA 166.  Both mutants promoted 
enhanced -1 PRF but had no effects on +1 PRF (Figure 40c). To more completely 
understand how the structural changes induced by the S104D and A106R mutations 
impact the ability of the ribosome to accurately decode mRNAs, we employed dual-
 luciferase translational fidelity assays designed to measure rates of misreading of near-
 and non-cognate tRNAs 212, and misreading of a UAA termination codon 213.  The S104D 
mutant was generally more inaccurate than wild-type, the magnitude of which became 
greater as the extent of codon/anticodon mismatches increased (Figure 40d). In contrast, 
decoding was largely unaffected in cells expressing the A106R mutant, although 
decoding of near-cognate tRNAs was slightly more accurate. 
84
 Figure 40. rpL10 loop mutants affect peptidyltransferase activity and translational 
fidelity.
 a. Apparent rates of peptidyltransfer from single turnover peptidylpuromycin reactions 
for indicated ribosomes. b. Single round peptidyltransferase reaction data for wild-
 type and the rpl10-S104D and rpl10-A106R mutants. c. Programmed -1 and +1 
ribosomal frameshifting values obtained using dual-luciferase reporters. d.
 Misincorporation of a stop codon, and near- and-noncognate amino acids in mutants 
compared to wild-type levels as monitored using dual-luciferase reporters.  Bars 
indicate s.e.m. (n=4), * P < 0.05, ** P < 0.01. 
85
 An intra-ribosomal suppressor of the S104D mutant reveals the importance of LSU 
allostery in function of the mature ribosome.   If the rpL10 mutants affect the 
distribution of ribosomes between the non-rotated and rotated states by altering the 
distribution of allosteric structures intrinsic to the ribosome, then mutants of other 
ribosomal components that promote opposing effects may be able to suppress the defects 
exhibited by the rpl10 mutants.  The L3-W255C mutant, which lies in a flexible loop (the 
W-finger) on the opposite side of the aa-tRNA accommodation corridor from rpL10 (Fig. 
43c) promotes increased affinity of ternary complex to the A-site and decreased affinity 
for eEF2 211.  A high copy plasmid expressing L3-W255C was able to almost completely 
suppress the biochemical (binding of ternary complex and Sdo1p to the A-site), and 
translational fidelity (-1 PRF) defects of the rpL10-S104D mutant (Figure 41a-c and 
Figure 42a-b). The ability of this mutant to correct the changes in rRNA structure at the 
B7a bridge promoted by the rpL10-S104D mutant (Figure 41d) demonstrates that the 
proper rotational equilibrium was re-established.  The control experiment, co-expression 
of L3-W255C with the wild-type RPL3, did not affect any of these parameters. 
Interestingly, this mutant did not suppress the rpL10-S104D slow growth defect (data not 
shown) or the ribosome biogenesis defect of rpl10-S104D, as assessed by polysome 
profiling (Figure 42c).  No mutants of L3 able to suppress the rpL10-A106R mutant were 
identified. 
86
 Figure 41. A mutant of rpL10 can be intrinsically suppressed with an rpL3 mutant.
 Panels a-b. Binding ternary complex and Sdo1p to indicated ribosomes. c. Frameshifting 
analyses. d. Structural probing of the landmark base A2207 (arrows) at the LSU side of 
the B7a intersubunit bridge with 1M7. Bars indicate s.e.m. (n=4 for a and c, n=3 for b), 
* P < 0.05, ** P < 0.01. 
87
 Figure 42. Primary data related to Figure 41.
 Single site binding isotherms for wild-type and rpL10-S104D ribosomes isolated from 
cells expressing the rpl3-W255C mutant or empty vector.  a. Binding of ternary complex 
to the A-site.  b. Binding of Sdo1p. 5 pmoles of ribosomes in 120 ?l total volume were 
used at each dilution. Bars indicate standard error of the mean.  c. Sucrose density 
gradient analysis. Cultures of AJY2104 (GAL1:RPL10) expressing RPL10 or rpl10-
 S104D and L3 or L3-W255C were grown in glucose to repress genomic RPL10 for 6 hrs 
before cells were harvested. Extracts were prepared and 9 A260 units were sedimented 
through 7?47% sucrose gradients.
88
 Discussion
 Eukaryotic rpL10 and its prokaryotic homologs L16 in bacteria and L10e in 
archaea contain a conserved internal loop that approaches the catalytic center of the large 
subunit. This loop is not resolved in atomic resolution structures of vacant yeast 15 or 
prokaryotic ribosomes 214, suggesting that this structure is dynamic. However, the internal 
loop is resolved in high-resolution crystal structures of bacterial ribosomes containing P-
 site tRNA 189,215 and in cryo-EM imaging of translating yeast ribosomes 188, suggesting 
that it is stabilized by tRNA binding. While the loop is shorter in the bacterial and 
archaeal homologs, the N-terminus of the L27 proteins from these kingdoms appear to 
provide the structural mimic for the tip of this loop 216.  Here, we have shown that 
mutations of this loop affect the rotational status of the ribosome, altering the ribosome?s 
affinity for Sdo1p at the P-site, for aa-tRNAs and eEF2 at the A-site, and globally 
affecting ribosome function from biogenesis through translational fidelity.
 Current models posit that ribosomal rotational status is solely determined by 
binding of different tRNA species and trans-acting GTPases 96,106,217. However, the 
findings presented here suggest that control of rotation is an intrinsic property of the 
ribosome.  We suggest that when the A-site is unoccupied by ligand, the flexible rpL10 
loop can sample this space: we call this the ?flipped in? conformation.  In contrast, 
occupation of the A-site by tRNAs displaces, or ?flips out? this loop (Figure 43a-b). The 
effects of the two rpL10 mutants on A-site associated functions and on ribosome 
structure in the absence of ligands suggest that this loop has an intrinsic, central role in 
establishing the rotational status of the ribosome.  The finding that non-enzymatic 
binding of aa-tRNA to the A-site exacerbates the low affinity defect and that 
89
 peptidyltransferase activity is diminished in rpL10-S104D provide additional support for 
this idea. Addition of the positively charged arginine residue may create new charge-
 charge interactions with the A-site to stabilize the ?flipped in? state making it more 
difficult to displace by aa-tRNA (Figure 43a).  Conversely, addition of a negatively 
charged residue in the S104D mutation may interfere with the ability of the loop to stably 
occupy the A-site due to charge repulsion effects with this rRNA-rich environment, thus 
stabilizing the ?flipped out? conformation (Figure 43b).  
Multiple lines of evidence (visualized in Figure 38) support the rpL10 lateral 
displacement model shown in Figure 43a-b. Chemical probing with 1M7 revealed 
increased reactivity of the tip of H39 near the PTC-proximal end of rpL10 in the S104D 
and G81D mutants (Figure 36), suggesting that rpL10 loses contact with this element in 
the rotated state.   This is accompanied by increased protection of bases in H43 
(elongation factor binding site) on the opposite (PTC-distal) end of the body of rpL10, 
suggesting that this side of the protein moves closer to H43 (the eEF binding site in 
Figure 43). Notably, 25S rRNA G1249 is even more protected from chemical 
modification in rpL10-G81D relative to rpL10-S104D mutant ribosomes, suggesting that 
the longer sidechain at this residue increases the extent to which it interacts with the 
nucleobase. The converse rRNA chemical reactivity patterns are observed with the 
A106R and F94I mutants, which favor the non-rotated state.  Along the ?top? of rpL10, 
the F94I mutant promoted increased reactivity of bases in H38 that normally interact with 
this amino acid residue, indicating that this contact was broken by loss of the aromatic 
sidechain.  These findings suggest that the phenylalanine sidechain intercalates into H38, 
pulling this structure along with it as rpL10 is laterally displaced, as evidenced by 
90
 changes in the reactivity of the small subunit side of the B1a intersubunit bridge (Figure 
35).  The observation that there are no differences in the reactivities of these bases 
(A1006 and U1007) in rpL10-S104D compared to rpL10-A106R ribosomes (Figure 34) 
indicates that F94 remains associated with this region of H38 throughout this proposed 
lateral movement. On the ?bottom? side of rpL10, decreased reactivity of bases in the 
accommodation corridor in rotated control ribosomes and the S104D mutant (Table 2) is 
consistent with prior observations that another feature of rpL10 called the N-terminal 
?hook? releases H89 to close the corridor in the rotated state 178. 
How can a change in the position of a small peptide loop affect the conformation 
of a large and complex macromolecule like the ribosome? We suggest that this small 
conformational change is amplified through a complex series of allosteric networks.  Like 
the gas pedal in an automobile, the rpL10 loop is physically linked to multiple functional 
centers of the machine.  The chemical structural probing studies reveal allosteric 
information exchange networks propagating out from rpL10 to all of the functional 
centers of the ribosome (diagrammed in Figure 43c).  These pathways completely frame 
the path through which tRNA moves through the ribosome during translation, including 
the accommodation corridor and decoding center, the PTC, and the E-site.  They also 
encompass the critical intersubunit bridges involved in rotation through the elongation 
cycle and the elongation factor binding site. Previous studies provide evidence that 
flexible peptide elements provide mobile platforms upon which allosteric switches can be 
built and are widely used by the ribosome as transducers of information, supporting the 
notion that structural elements intrinsic to the ribosome play important roles in 
coordinating ribosomal conformations.  These include the N-terminal hook of rpL10 178, 
91
 the W-finger, N-terminal extension, and Basic Thumb elements of rpL3 211,218,219, an 
internal loop located in rpL5 220, and the P-site loop of rpL11 206. Importantly, there is a 
considerable degree of overlap between the allosteric networks identified in these and 
other studies 200,201,207,210,221, indicating that they are all components a single network of 
?switches? and ?wires? that coordinate ribosome structure with function.   This is further 
supported by the observation that a mutant in one switch, rpL3-W255C, can suppress the 
biochemical and translational fidelity defects of a second, i.e. the rpL10-S104D loop 
mutant. What distinguishes the rpL10 loop from other elements are the broad effects of 
loop mutants on ribosome biogenesis, elongation and termination.  These effects argue 
that the rpL10 loop is the central control element in this complex wiring scheme. It 
should be noted that while we interpret our results as evidence for the influence of rpL10 
on ribosome equilibria, alternative explanations are also possible.  For example, these 
results could be interpreted as evidence for changes in rRNA folding and the structure of
 the large subunit. However, incorporation of L10 occurs very late in the process of 
eukaryotic ribosome biogenesis after the pre-60S subunit has been exported to the 
cytoplasm, well after the core of the large subunit has been folded. Thus, the effects of
 these single amino acid mutants of rpL10 on rRNA folding are likely to be minimal.
92
 Figure 43. Models of rpL10 function: rpL10 is at the center of a cascade of allosteric 
communication pathways throughout the ribosome.
 a. The rpL10 loop ?flipped in? conformation is the substrate for ternary complex and 
Sdo1p. P: PTC-proximal, D: PTC-distal. b. The ?flipped out? loop conformation, 
substrate for eEF2. Binding of an aa-tRNA (indicated by the red ?t?) causes displacement 
of the loop from the A-site, precipitating structural rearrangements in rpL10. These 
include lateral displacement of the main body of the protein (dashed black arrow) and 
H38 toward the elongation factor binding site, creating the binding platform for eEF2.  
Release of the N-terminal hook of rpL10 from H89 enables closing of the aa-tRNA 
accommodation corridor. These movements also initiate allosteric transmission of 
information through the communication pathways shown in panel c to distantly located 
functional centers of the ribosome to set the stage for the next phase of elongation. These 
include rearrangements in the E-site in preparation for release of deacylated tRNA, and 
interactions with the decoding center and small subunit to initiate subunit rotation. 
c. Summary of chemical probing experiments mapping the allosteric information 
exchange pathways emanating from rpL10 to all the functional centers of the ribosome to 
influence intersubunit rotation. Intersubunit bridges B1a, B1b/c B2a/c, and B7a and 
ribosomal proteins L3, L10, L11, S15 and L18 are labeled. CP: Central Protuberance of 
the LSU. AC: Accommodation Corridor. PTC: Peptidyltransferase Center.  SRL: 
Sarcin/Ricin Loop. DC: Decoding Center.
93
 The rotational status of the 80S ribosome is an important determinant for binding 
of trans-acting factors, optimizing PTC activity and translational fidelity. Both mutants 
promoted increased rates of -1 PRF (Figure 40c), which requires slippage of both A- and 
P-site tRNAs 222, but did not affect +1 PRF, which only requires slippage of P-site tRNA
 166. This is consistent with the observation that the rpL10 loop mutants only affected A-
 site tRNA binding. Increased utilization of near- and non-cognate tRNAs, and 
termination codon readthrough by the rpL10-S104D mutant (Figure 40d) may be 
explained by the observation that A1755 (E. coli A1492) in h44 was hyperprotected from 
chemical modification (Figure 35). Flexibility of this base is critical for accurate 
decoding by stabilizing a mini-helix between cognate codon:anticodon interactions 84,223.  
Concurrently, LSU A2256 (E. coli A1913) in H69 was hyper-reactive in rpL10-S104D 
ribosomes (Figure 33). This base also plays a central role in mRNA decoding where it is 
paired with A1492 in the non-rotated state, but is unpaired and flexible in the rotated state
 92.  We suggest that the propensity of rpL10-S104D ribosomes to assume the rotated 
stated limits the ability of these bases to participate in mini-helix formation, leading to 
increased utilization of non-cognate ligands. 
Considering the mechanisms in place to ensure fidelity of gene expression we 
expect that safeguards have evolved to ensure that only correctly functioning ribosomes 
are utilized in translation. Ribosome assembly culminates in cytoplasmic maturation 
where essential ribosomal proteins are added and trans-acting factors are released 40. A 
critical step in this pathway is the release of the anti-association factor Tif6p by the 
concerted action of Sdo1p and the eEF2 paralog Efl1p. The observation that suppressing 
mutations in Efl1p appear to facilitate a conformational change in the protein akin to the 
94
 conformational change observed in eEF2 engendered the proposal that the LSU 
undergoes a ?test drive? in which the integrity of the P-site is assessed through activation 
of the GTPase of Efl1p 187. Notably, similar functional checkpoints governing pre-40S 
maturation have recently been proposed 224,42. The findings presented in the current study 
add further support and detail to this model. Specifically,  the ability of Sdo1p to compete 
for Ac-aa-tRNA binding in the P-site, inhibit peptidyltransferase activity, and stimulate 
aa-tRNA binding to the A-site [similar to the stimulatory effect of P-site bound Ac-aa-
 tRNA on aa-tRNA binding to the A-site 225] all suggest that Sdo1p interacts with the P-
 site, stabilizing a pseudo-non-rotated state of the LSU alone. 
The structure of Sdo1p has been likened to tRNA (Figure 44) 226 , and we propose 
that Sdo1p is a mimic for a P-site ligand that couples the GTPase activity of the 
elongation factor mimic Efl1p 47 to drive a pseudo-translocation event on the LSU
 (Figure 45). Because Efl1p and Sdo1p appear to act on 60S independent of 40S, we 
suggest that the rpL10-S104D and ?A106R mutants promote conformational changes 
within the 60S subunit alone that are analogous to those in the 60S subunit in the context 
of rotated and non-rotated 80S ribosomes.  The 60S subunit ?test drive? appears to 
provide the primary quality control check on assembly and function of the LSU before it 
is released into the pool of actively translating ribosomes. In particular, the ability to bind 
biogenesis ligands that structurally mimic elongation factors as well as the ability to 
undergo GTP hydrolysis is being functionally tested during this quality control point. 
Failure at any of these steps leads to the inability of Tif6p and Nmd3p to dissociate, and 
the subsequent degradation of such improperly assembled subunits through the non-
 functional ribosomal decay pathway 227. Sdo1p binding defects in the ?pseudo-rotated? 
95
 S104D mutant can be intrinsically suppressed with the rpL3-W255C mutant, likely due to 
its ability to re-establish the correct pre-LSU conformation. However, the failure of this 
mutant to rescue the ribosome biogenesis defect suggests that quality control involves 
more than mere monitoring of ligand binding.  
Figure 44. Sdo1 structurally resembles tRNA.
 Sdo1 and tRNA appear to be structural mimics that function at distinct time points during 
the ribosomal life cycle. Image modified from Ng 226. 
96
 Figure 45. Involvement of the rpL10 loop in ribosome rotation throughout the 
ribosomal life cycle.
 a. The elongation cycle of translation: rpL10 loop positioning and ligand binding. 
The elongation cycle begins at left with the ribosome in the non-rotated state where the 
E-site contains a deacylated tRNA, the P-site is occupied by peptidyl tRNA, and the un-
 occupied A-site can be sampled by the rpL10 loop, i.e. the ?flipped in? conformation. 
Following elongation ternary complex binding, aa-tRNA accommodation and 
peptidyltransfer, tRNAs assume the hybrid states and the loop assumes the ?flipped out? 
conformation, signaling the ribosome to assume the rotated state. eEF2 binds to rotated 
ribosomes, resulting in translocation, and the elongation cycle begins anew. 
b. The ?test drive.? Sdo1p and Efl1p interact with the pre-60S subunit in the pseudo-
 rotated state.  Efl1p-mediated pseudo-translocation drives Sdo1p into the P-site, 
stabilizing the pseudo-non-rotated state with the rpL10 loop in the ?flipped in? 
conformation.  This is followed by release of the anti-association factor Tif6p and Sdo1p, 
promoting the final steps of 60S maturation. 
97
 The biomedical importance of the rpL10 loop was recently highlighted by the 
discovery that a significant fraction of T-cell acute lymphoblastic leukemia patients 
harbor mutations of R98, and in one patient at Q123, both of which lie at the base of the 
loop 179.  Remarkably, these mutations prevent the release of Tif6p and Nmd3p, but can 
be suppressed by mutations in Nmd3p, indicating that like the S104D mutation, R98 and 
Q123 mutations in human Rpl10 cause a failure during the ?test drive? of the 60S 
subunit. These findings suggest that defects in ribosome biogenesis and/or in translational 
fidelity may be drivers of this neoplastic disease. Investigations are currently underway to 
address these questions, and some of these findings are presented in the following 
chapter. 
98
 Chapter 3
 Ribosomopathies: a new model for cancer biogenesis 
Introduction
 It has become clear that deregulation of a process as essential as cellular protein 
translation may play a central role in cancer development. Key cancer oncogenes and 
tumor suppressors act as master regulators of the translational apparatus. For instance, 
MYC expression in cancer cells causes elevated expression of several ribosomal proteins 
and translation factors and PTEN inactivation results in mTOR activation and 
phosphorylation of ribosomal protein S6, eventually leading to increased translation of 
ribosomal proteins, elongation factors and ribosome biogenesis factors 228. Over the last 
decade several congenital disease syndromes have become linked to genetic defects in 
ribosomal proteins or ribosome biogenesis factors. These syndromes, collectively called 
?ribosomopathies?, are characterized by pleiotropic abnormalities including birth defects, 
heart and lung diseases, connective tissue disorders, anemia and ataxia, and mental 
retardation 43. A common feature is that patients suffering from these diseases have a 
significantly elevated risk of developing cancer 229. Thus, ribosomopathies present an 
intriguing paradox: while patients initially present with hypo-proliferative disorders such 
as anemias, those who survive to middle age often develop hyper-proliferative diseases, 
i.e. cancers.   This appears to be a general phenomenon: loss of function mutations in 
several ribosomal proteins have been shown to cause tumor development in zebrafish 230. 
Although there is thus clear indication for a role for deregulated translation in cancer 
99
 cells, it is currently unclear how alterations that affect a vital cellular function as protein
 translation may promote the hypo-to-hyper-proliferation switch and carcinogenesis.
 Acute T-cell lymphoblastic leukemia (T-ALL) is a hematopoietic malignancy 
caused by accumulation of genetic lesions in developing T-cells that has recently been 
linked to ribosome dysfunction. T-ALL accounts for 15% of all pediatric leukemias and 
20-30% of pediatric patients do not achieve long-term remission 231. Exome sequencing 
on 67 T-ALL patients revealed that 9.8% of pediatric and 7.2% of adult T-ALL patients 
had somatically acquired mutations in the large ribosomal subunit protein genes RPL10, 
RPL5 and RPL22 179. Mutations in RPL10 were the most frequent ribosome mutations 
identified in this study and were almost exclusively in residue arginine 98 (R98) with the 
exception of one patient harboring the Q123P mutation which lies adjacent to R98 within 
the rpL10 3D structure. Both residues are at the base of a flexible loop in rpL10 that 
closely approaches the peptidyltransferase center in the catalytic core in the ribosome 
(Figure 46). The rpL10 protein is highly conserved across species: the yeast and human 
proteins are interchangeable and residue 98 is invariantly an arginine 180. As described in 
the previous chapter, rpL10 plays an important role in the late stages of 60S ribosomal 
subunit biogenesis. Both immature subunits are functionally inactive as they enter the 
cytoplasm and require additional maturation events and association with each other 
before they can engage in protein synthesis 40. A critical step in 60S maturation is the 
release of the anti-association factor Tif6, followed by release of Nmd3, the primary 
export adaptor for the pre-60S subunit in yeast and in humans 232,231. Tif6 release requires 
Sdo1 and the GTPase Efl1, a paralog of translation elongation factor 2. As detailed in the 
previous chapter, we have proposed that an essential internal loop of rpL10 initiates a 
100
 ?test-drive? of the 60S subunit involving these factors as a quality control checkpoint in 
ribosome assembly, ensuring that only properly functioning subunits are allowed to 
progress into the translationally active pool of ribosomes. Defective ribosomes carrying 
mutations in rpL10 specifically fail in this ?test drive? leading to their degradation 
through the non-functional ribosomal decay pathway 227. 
To test the impact of the T-ALL-associated mutations, the rpl10-R98S and rpl10-
 R98C mutants were expressed in yeast cells as the sole forms of RPL10. These mutations 
impaired cellular proliferation and polysome profiling revealed modest changes in the
 ratios of free 60S vs. 40S subunits, markedly reduced polysomes, and the presence of 
halfmers (initiation complexes waiting to join with 60S subunits), suggesting defects in 
both ribosome biogenesis and subunit joining. Examination of Tif6 and Nmd3 recycling 
in these strains revealed that both proteins accumulated in the cytoplasm, indicating a 
defect in their release 179. Thus, the rpl10-R98S and rpl10-R98C mutations appeared to 
arrest 60S biogenesis in the Efl1-dependent quality control step. Consistent with the 
yeast-based observations, mouse lymphoid cells expressing rpl10-R98S had lower 
proliferation rates than cells expressing wild type RPL10 and showed defective polysome 
profiles 179. 
In the studies presented here, the rpl10-R98S mutant was used to elucidate the 
molecular mechanisms underlying the translational defects that may lead to 
carcinogenesis. Trans-acting factors were identified that suppress the biogenesis defect of 
the rpl10-R98S mutant, thereby re-establishing ribosome production and cell 
proliferation. However, we show that suppression of the biogenesis and growth 
impairment defects fail to suppress the profound molecular and functional defects of 
101
 rpL10-R98S ribosomes. These ribosomes display rotational disequilibrium and resulting 
ligand binding defects. Such defects in turn promote elevated rates of programmed 
ribosomal frameshifting and destabilization of - 1 PRF signal containing messages. One 
class of -1 PRF mRNAs plays a critical role in yeast telomere maintenance 161.  In rpl10-
 R98S cells, the steady-state abundance of these mRNAs is decreased, resulting in short 
telomeres. These results suggest that suppression of the growth defect results from 
bypassing the ?test drive? and the non-functional ribosome decay (NRD) apparatus, a 
quality control system that enables cells to identify and eliminate sub-optimal ribosomes 
227. The suppressed cells then continue to actively produce and use mutant ribosomes that 
have specific biochemical and translational fidelity defects. These findings lead us to
 propose two different but not mutually exclusive models for ribosome mutations 
promoting cancer: 1) mutant ribosomes drive an altered gene expression program,
 promoting T-ALL; 2) the suppressing mutations these cells acquire to overcome the 
growth defects are, themselves, drivers of T-ALL. This work defines novel mechanisms
 by which acquired mutations in a ribosomal protein can promote dysregulation of gene 
expression and cancer.   
102
 Figure 46. Localization of rpL10 and the loop in the large ribosomal subunit.
 a. rpL10 in the context of the crown view of the LSU. b. Close-up of rpL10 and the local 
environment. The hypothetical loop structure is indicated by dashed red lines and the 
position of R98 is indicated. rpL10 is situated between Helices 38 and 89 and located in 
close proximity to several functional centers of the LSU including the PTC, the aa-tRNA 
accommodation corridor, and the elongation factor binding site. PTC: peptidyltransferase 
center.  Images were generated using PyMOL. 
103
 Results 
The rpl10-R98S growth and biogenesis defects are suppressed by the NMD3-Y379D
 mutant. Despite the proliferation impairment in in vitro lymphoid cell cultures 
expressing rpl10-R98S, the mutations in the patients were somatically acquired, were 
present in 100% of tumor cells in most patients, and were recurrent in 6.5% of pediatric 
patients 179.  These observations strongly suggest that these mutations must have been 
selected for in pre-cancerous cells because they imparted some particular advantage. We 
have noted that yeast carrying the rpl10-R98S mutation readily acquire suppressing 
mutations that compensate for the growth defect. Sequencing of candidate suppressing 
genes identified a mutation in the export adapter NMD3, NMD3-Y379D, which 
suppresses the growth defect (Figure 47) and the ribosome biogenesis defect (Figure 48) 
of rpl10-R98S cells. Whole genome sequencing of an additional mutant identified
 aneuploidy for chromosome VIII bearing NMD3 (data not shown). Indeed, increased 
gene dosage of NMD3 also suppresses the rpl10-R98S defects (Figures 47 and 48). 
Figure 47. Overexpression of NMD3 or co-expression of NMD3-Y379D suppresses 
the rpl10-R98S growth defect. 
Ten-fold dilution spot assay of isogenic strains demonstrating that the rpl10-R98S growth 
defect is suppressed by ectopic expression of NMD3 and by the NMD3-Y379D mutation.
104
 Figure 48. Overexpression of NMD3 or co-expression of NMD3-Y379D suppresses 
the rpl10-R98S biogenesis defect. 
Sucrose density gradient analysis. Indicated strains were grown in glucose to repress 
genomic RPL10 for 6 hrs before cells were harvested. Extracts were prepared and 9 A260 
units were sedimented through 7?47% sucrose gradients.
105
 The rpl10-R98S mutation alters ribosome rotational status, and this defect is not 
suppressed by co-expression of NMD3-Y379D.  As shown in the previous chapter, the 
internal loop of rpL10 is a central driver of ribosomal rotation. Thus, we examined the 
previously established landmark of ribosomal rotation (the B7a intersubunit bridge, see 
Chapter 2) to assess the rotational status of the mutant ribosomes.  1M7 was used to 
probe A2207 of the LSU-side of the B7a intersubunit bridge, and the extent to which 
these bases were modified in purified empty isogenic wild-type, mutant, and control 
ribosomes was quantitatively assessed and normalized using hSHAPE.  Reactivity 
profiles in Figure 49 show that rpL10-R98S ribosomes are in a structural disequilibrium 
favoring the rotated state. Importantly, this defect is not suppressed by co-expression of 
NMD3-Y379. 
Figure 49. Co-expression of NMD3-Y379D does not suppress the rpl10-R98S
 rotational defect. 
Reactivity peaks obtained by hSHAPE after probing of the landmark base A2207 
(arrows) at the LSU side of the B7a intersubunit bridge with 1M7. This demonstrates that 
the R98S mutant pushes ribosomes towards the rotated state, and this defect is not 
suppressed by NMD3-Y379.
106
 The rpl10-R98S mutation alters ribosomal affinity for specific ligands, and this 
defect is not suppressed by co-expression of NMD3-Y379D.  Characterization of other 
yeast rpl10 mutants has revealed the role of this protein in recruitment of trans-acting 
factors to the ribosome, which depend on the correct rotational state (see Chapter 2). 
Given the rotational defect of rpL10-R98S ribosomes, their ability to bind several ligands 
was assayed. Steady-state binding experiments using ribosomes purified from isogenic 
yeast strains revealed that rpL10-R98S ribosomes promoted ~3-fold lower affinity for 
ternary complex and aminoacyl-tRNA (aa-tRNA), and ~3-fold increased affinity for 
eEF2, both of which are ligands that specifically interact with the ribosomal A-site 
(Figures 50 and 51).  Similar to other rpL10 loop mutants, rpL10-R98S ribosomes did not 
affect binding of acetylated-aa-tRNA (Ac-aa-tRNA) to the ribosomal P-site. Also similar 
to other rpl10 mutants, R98S mutant ribosomes had ~2.5-fold lower affinity for Sdo1p.  
Notably, none of the biochemical defects were suppressed by expression of NMD3-
 Y379D (Figure 50).  This indicates that, while NMD3-Y379D can suppress the ribosome 
biogenesis defect, it does not correct the underlying biochemical defect conferred by the 
rpl10-R98S mutation.  This allows us to separate the effects of R98S on biogenesis from
 those on translation fidelity.
107
 Figure 50. Co-expression of NMD3-Y379D does not suppress the rpl10-R98S
 biochemical defects. 
Steady state binding of indicated ligands to ribosomes isolated from cells expressing 
wild-type, rpl10-R98S, and rpl10-R98S + NMD3-Y379D. Dissociation constants were 
obtained from binding assays of Ac-aa-tRNA to the P-site and ternary complex to the 
A-site as monitored by filter binding, eEF2 as monitored by extent of ribosylation of 
unbound protein, and Sdo1p as monitored by levels of radiolabel detection in 
ribosomes. Bars indicate s.e.m. (n=4), * P < 0.05, ** P < 0.01. 
108
 Figure 51. Primary ligand binding data related to Figure 50. 
Single site binding isotherms for ribosomes and various ligands. Binding of Ac-aa-tRNA 
to the P-site (panel a), ternary complex the A-site (panel b), and eEF2 (panel c) and 
Sdo1p (panel d) to ribosomes isolated from cells expressing wild-type, rpl10-R98S, and
 rpl10-R98S + NMD3-Y379D. 5 pmoles of ribosomes in 120 ?l total volume were used at 
each dilution throughout. Bars indicated standard error of the mean (n=4). 
109
 Translational fidelity defects of the rpl10-R98S mutant are not suppressed by 
NMD3-Y379D.  In general, ribosomes must maintain translational reading frame in order 
to faithfully translate the genetic code.  However, there are a growing number of 
examples of cis-acting mRNA sequences that direct elongating ribosomes to shift reading 
frame: this is called Programmed Ribosomal Frameshifting (PRF).  Different sequences 
direct ribosomes to slip by 1 base in either the 5? (-) or 3? (+) direction, i.e. -1 PRF and 
+1 PRF, respectively. Numerous studies have revealed mechanistic differences between 
the two: most -1 PRF events require slippage of both A- and P-site tRNA 151,222, while 
only the P-site tRNA is involved in +1 PRF, for example, in translation of the 
retrotransposon Ty1 from yeast 166.  Consistent with the biochemical data indicating a 
large subunit A-site specific defect, the rpl10-R98S mutation stimulated -1 PRF by ~2.6-
 fold while +1 PRF remained unchanged (Figure 52). Similarly, rpl10-R98S cells were 
defective for recognition of a termination codon (~2-fold increased readthrough) which 
requires efficient recruitment of the eRF1/eRF3 complex to the ribosomal A-site 117. 
Discrimination between sense and missense tRNAs during elongation, a process that 
occurs in the A-site of the small subunit, was only slightly affected in rpl10-R98S mutant 
cells. Consistent with the structural and biochemical findings, co-expression of NMD3-
 Y379D did not suppress either the -1 PRF or termination codon misreading defects.  To 
determine whether or not the genetic and biochemical trends we observed were unique to 
the R98S mutant, we assayed the effects of a second trans-acting suppressor of a 
different rpl10 mutant that exhibits similar phenotypic, biochemical and functional 
defects.  Similar to the results described above, the TIF6-Y192F suppressor of the growth 
and biogenesis defect of rpl10-S104D mutant cells 187 (Chapter 2) did not suppress the 
110
 ternary complex binding or -1 PRF defects (Figure 53). Additionally SDO1-V36A, also 
previously shown to suppress the growth defect of rpl10-S104D, exhibited a 2-fold
 higher binding affinity to ribosomes isolated from rpl10-S104D cells as compared to 
wild-type (Figure 54). This indicates that the suppressing activity of trans-acting 
biogenesis factors is likely driven by their increased (i.e. Sdo1) or decreased (i.e. 
Tif6/Nmd3) affinity to ribosomes. 
Figure 52. Co-expression of NMD3-Y379D does not suppress the rpl10-R98S
 translational fidelity defects. 
a. Programmed -1 and +1 ribosomal frameshifting values obtained using dual-
 luciferase reporters. b. Misincorporation of a UAA stop codon and a near-cognate 
amino acid (missense) in mutants compared to wild-type levels as monitored using 
dual-luciferase reporters.  ND = not determined. Bars indicate s.e.m. (n=4), * P < 0.05, 
** P < 0.01. 
111
 Figure 53. Co-expression of TIF6-Y192F does not suppress the rpl10-S104D
 biochemical and translational fidelity defects.
 a. Steady state binding of indicated ligands for ribosomes.  Dissociation constants 
obtained from assays of binding of ternary complex to the A-site and Ac-aa-tRNA to the 
P-site in ribosomes isolated from cells expressing wild-type, rpl10-S104D, and rpl10-
 S104D + TIF6-Y192F as monitored by filter binding. b. Programmed -1 and +1 
ribosomal frameshifting values obtained using dual-luciferase reporters. Bars indicate 
s.e.m. (n=4), * P < 0.05, ** P < 0.01.
112
 Figure 54. Sdo1p-V36A has a higher affinity to rpL10-S104D ribosomes.  
Dissociation constants obtained from assays of binding of Sdo1p and Sdo1p-V36A to 
ribosomes isolated from cells expressing wild-type or rpl10-S104D. Bars indicate s.e.m. 
(n=3), * P < 0.05, ** P < 0.01.
113
 Yeast telomere maintenance and length defects promoted by the rpl10-R98S mutant 
are not suppressed by NMD3-Y379D. Computational analyses suggest that ~10% of 
eukaryotic cellular mRNAs harbor operational -1 PRF signals. Over 95% of these are 
predicted to function as mRNA destabilizing elements through the nonsense-mediated 
mRNA decay pathway by directing translating ribosomes to premature termination 
codons 155.  More recently, -1 PRF was implicated in telomere maintenance in yeast 161.  
To test if rpl10-R98S-promoted changes in -1 PRF also lead to changes in gene 
expression, we used dual luciferase reporter-based assays of -1 PRF directed by elements 
located in the yeast EST1, EST2, STN1 and CDC13 mRNAs. The rpl10-R98S mutant 
stimulated -1 PRF promoted by cis-acting elements located in these mRNAs by 2 to 3-
 fold as compared to isogenic wild-type controls. Consistent with the biochemical 
findings, NMD3-Y379D did not suppress these -1 PRF defects (Figure 55a). Similar to 
results observed with other yeast mutants that promoted changes in -1 PRF 159,161, the 
steady-state abundances of endogenous EST1, EST2, STN1 and CDC13 mRNAs were 
significantly reduced in rpl10-R98S cells, as monitored by qRT-PCR (Figure 55b). 
Consistent with previous data, the NMD3-Y379D mutant did not affect the abundance of 
these mRNAs in rpl10-R98S mutant cells.  Furthermore, rpl10-R98S cells exhibited 
significantly shortened telomeres as assessed by PCR, and this defect was not 
ameliorated by co-expression of NMD3-Y379D (Figure 56).  
114
 Figure 55. Co-expression of NMD3-Y379D does not suppress the rpl10-R98S
 telomere maintenance defects. 
a. Programmed -1 ribosomal frameshifting directed by sequences in the following yeast 
genes: EST1 (signal beginning at nt 1272); EST2 (signal beginning at nt 1251); STN1
 (signal beginning at nt 1203); and CDC13 (signal beginning at nt 1272). b. Expression of 
endogenous EST1, EST2, STN1 and CDC13 mRNAs was monitored by qRT-PCR. Bars 
indicate s.e.m. (n=4), * P < 0.05, ** P < 0.01.
 Figure 56. Co-expression of NMD3-Y379D does not suppress the rpl10-R98S
 telomere length defects.
 Abundance of telomere repeat sequences were quantified by PCR, with the single copy 
reference gene SGS1 as the loading control. Bars indicate s.e.m. (n=9), ** P < 0.01.
115
 Discussion 
It is currently unclear how and by what mechanisms defects in a crucial cellular 
process like translation can lead to carcinogenesis. In particular, how a mutation that 
negatively impacts cell growth can promote a hyper-proliferative disease presents a 
paradox. While this is not the first time that defects in the translation machinery have 
been linked to cancer, the molecular mechanism of how a mutation in a specific 
ribosomal protein can promote a specific type of cancer is highly novel. Furthermore, the 
evidence presented above suggests that the acquisition of defects in ribosomal proteins 
throughout life may represent yet another unexplored mechanism to deregulate the 
translation machinery in cancer cells.
 Our findings reveal that the malfunction of the rpL10 internal loop is at the heart 
of T-ALL. In particular, while the mutations described in Chapter 2 were located at the 
tip of the loop (much like fingers on a hand), thereby impairing its ?sensitivity?, the T-
 ALL mutations localize to the base of the loop (wrist of the hand), thereby possibly
 changing the general flexibility and dynamic nature of the structure and its ability to 
influence ribosomal rotational status. NMD3-Y379D suppresses the ribosome biogenesis 
defects but not the intrinsic biochemical defects conferred by the rpl10-R98S mutant, the 
most frequent acquired mutation in T-ALL patients. An identical trend applies to a 
different suppressor, TIF6-Y192F, which suppresses the ribosome biogenesis defects of 
the rpl10-S104D mutant but not the downstream translational functions. The observed 
suppression is possibly achieved due to a lower affinity of such mutant biogenesis factors 
to the ribosome, resulting higher rates of dissociation and hence passing of the pre-60S 
quality check point. These observations lead us to propose a novel model for 
116
 carcinogenesis offering an explanation for the apparent link between hypo-and hyper-
 proliferation seen in ribosomopathies (depicted in Figure 57). Given the importance of 
rpL10 in ribosome biogenesis, mutations that arise in this protein lead to defects in 
ribosome assembly. As the mutant ribosomes are prevented from entering the active pool 
of ribosomes because of a failed ?test drive?, the translational capacity of the cell is 
greatly diminished, resulting in a severe proliferation defect (hypo-proliferation). Such 
mutants are under selective pressure to select for mutations that suppress the biogenesis 
defects, which is sensed by the NRD apparatus. Once acquired, these suppressing 
mutations can subvert the quality control process and allow utilization of defective 
ribosomes. However, the continued use of defective ribosomes in translation ultimately 
drives altered gene expression patterns, evidenced by decreased abundances of -1 PRF 
signal containing mRNAs, including those implicated in telomere length maintenance 
(Figures 55 and 56). Thus, we suggest that suppression of the ribosome biogenesis 
defects represents a critical event in the progression of these cells to T-ALL (hyper-
 proliferation). As such, while providing a temporary fix by bypassing the primary quality 
check point in ribosome assembly, suppressing mutations are ultimately themselves the 
drivers of disease.
117
 Figure 57. Model of T-ALL progression. 
A mutation in L10 results in the inability of the pre-60S subunits to pass the quality 
control checkpoint, leading to decreased ribosome assembly and proliferation. A 
ribosome biogenesis suppressor can arise due to selective pressure allowing the bypass 
the quality ?test drive? thereby boosting production of defective ribosomes. Continued 
defective translation ultimately leads to an altered gene expression profile and the onset 
of T-ALL. 
Mutant ribosomes such as described here represent an attractive novel drug target. 
As detailed in Chapter 1, many clinically used antibiotics inhibit the proliferation of 
pathogenic bacteria by binding their ribosomes and interfering with translation. Although 
these drugs specifically target the prokaryotic ribosome, the presence of a ribosome 
mutation and resulting impaired ribosome function in eukaryotic cells may sensitize these 
cells to the effect of such antibiotics directly or in a synthetically lethal way. As such, 
clinically approved drugs could serve a second life as anti-cancer agents for ribosome 
defective cancer. 
118
 Although this study employed yeast as a model system to probe the molecular 
defects of T-ALL associated ribosome mutations, the function of the ribosome and the 
pathway of its assembly are highly conserved between yeast and humans. In fact, human 
rpL10 and Nmd3 can functionally replace their yeast counterparts 180,234. Consequently, 
the mechanisms of bypassing these defects are also likely to be conserved. Using the 
yeast work described in this chapter as a foundation, we are currently performing similar 
studies in mouse and human cells, including T-ALL patient T-lymphoblasts. While this 
work was focused on one suppressor, identifying additional suppressing mutations in 
yeast should uncover the pathways that cancer cells exploit to avoid the growth inhibition 
of mutations in RPL10. This knowledge will identify new cellular targets for novel drug 
therapies. In addition, a more complete mechanistic understanding of the ribosome 
defects contributing to T-ALL will direct the search for additional genes that are mutated 
in pediatric T-ALL to improve molecular diagnostics and will also improve prognostics 
for this heterogeneous disease. Importantly, because patients with congenital ribosome 
defects develop various tumor types 235, the acquired ribosome mutations described in T-
 ALL may only represent ?the tip of the iceberg?: acquired ribosome defects may represent 
a novel general concept involved in pathogenesis of many tumor types.
119
 Chapter 4
 Work in progress: the rpL10 C-terminal tail and autism
 Introduction
 The previous two chapters describe the importance of the ribosomal protein L10 
internal loop, which approaches the peptidyltransferase center (PTC) and acts as the 
primary sensor of activity in this ribosomal core. Here, we explore the remaining regions 
of the protein and their functions. These regions comprise the ?body? of L10 which lies 
in a plane along the face of the protein just above the tRNA accommodation corridor, and 
the C-terminal ?tail? which is unique to eukaryotes (Figure 57).  
The C-terminal tail of rpL10 is essential in yeast, as even small truncations of this 
structure are lethal. Mutations in the tail, as well as at the tip of the body of rpL10,
 resulted in profound defects in general growth, sensitivity to temperatures and antibiotics, 
and ribosome biogenesis. The locations of these mutations face and closely approach the 
factor binding site, suggesting that these regions of rpL10 play an important role in ligand 
binding.  This hypothesis is currently being tested through binding experiments assaying 
the affinity of several trans-acting factors to ribosomes containing mutations in the C-
 terminus of L10. 
Importantly, both mutations previously identified in rpL10 (L206M and H213Q) 
in children affected by autism in two independent families are located in the C-terminus 
of the protein 180. One of these mutations, L206M, affects the last amino acid that is 
conserved between yeast and humans, and almost completely conserved in the whole 
eukaryotic tree (Figure 58). We introduced this mutation, as well as additional amino acid 
120
 substitutions at the rpL10-L206 position covering a range of chemical properties, into
 yeast to assay their effects on ribosome structure and function. 
Figure 57. Location of L10 regions in the LSU. 
The crown view is shown, and the path taken by tRNAs through the ribosome and the 
three binding pockets are indicated. L10 is red, with the loop sensing ligand arrival at the 
ribosomal A-site indicated by dashed lines. Other ribosomal proteins are shown in brown, 
and rRNA is grey. 
121
 Figure 58. Multiple sequence alignment of L10. 
Alignment of RPL10/RPL16 protein sequences. Sequences are abbreviated as follows: hs 
Homo sapiens; bt Bos taurus; mm Mus musculus; gg Gallus gallus; xl Xenopus laevis; dr 
Danio rerio; pm Petromyzon marinus; bb Branchiostoma belcheri; dm Drosophila 
melanogaster; ce Caenorhabditis elegans; ha Hydra attenuate; sc Saccharomyces 
cerevisiae; eg Euglena gracilis; pf Plasmodium falciparum; at Arabidopsis thaliana af 
Archaeoglobus fulgidus; bs Bacillus subtilis. Exon 7 is representing the C-terminal 
domain spanning aa 165?214 of human RPL10. Red arrows: position of the highly 
conserved aa 206 changed through mutations (L206M) in families with autism. The 
figure display is shown with clustalW colors. Image modified from Klauck 180. 
122
 Results
 The ?body? and ?tail? mutants of rpL10 show general growth defects, and 
sensitivity to temperature and antibiotics.  As ribosome-associated defects commonly 
result in growth deficiencies, 10-fold dilution spot assays were performed on previously 
generated mutants having lost the ?killer? phenotype 236 on rich medium at the optimal 
temperature of 30?C (Figure 59). Many mutant cells assayed displayed significant growth 
defects, particularly those in the C-terminal tail (shown in blue in Fig. 59) or at the tip of 
the rpL10 body near the C-terminal tail (G81D, N144D, K145E, K145R). Because 
changes in ribosome structure and function also alter the growth characteristics of cells at 
various non-optimal temperatures, the same mutants were assayed with regard to their 
sensitivity or resistance to low (15?C) or high (37?C) temperatures.  Almost all of the 
mutants with growth defects at 30?C also showed slow growth phenotypes on both of 
these restrictive temperatures. Wild type and mutant cells were also spotted onto medium 
containing small molecule protein synthesis inhibitors. Anisomycin is a competitive 
inhibitor at the ribosomal A-site 237, while paromomycin induces the utilization of near-
 cognate codons due to the stabilization of the mini helix required for GTP hydrolysis by
 eEF1A 238. Sensitivity or resistance to these antibiotics suggests defects in the ribosomal 
A-site and the decoding center, respectively. As seen in Figure 59, several mutants 
displayed strong resistance to anisomycin, while others showed sensitivity to 
paromomycin. Notably, the most profound growth defects under all these conditions were 
seen in mutants of the C-terminal tail and also at the tip of the rpL10 body. One such 
mutant in the body of rpL10, G81D, was explored in more detail in Chapter 2 and was 
123
 shown to contact the elongation factor binding site and to participate in the required 
allosteric communication pathways initiated by rpL10 during ribosomal rotation. 
Figure 59. Genetic analysis of rpL10 ?body? and ?tail? mutants. 
Dilution spot assays at indicated temperature and in the presence of the antibiotic 
anisomycin or paromomycin. Mutants of the C-terminal tail are highlighted in blue. 
Figure modified from the thesis of Alexey Petrov 236. 
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 The ?body? and ?tail? mutants of rpL10 show defects in translational fidelity and 
ribosome biogenesis. As described in the previous chapters, defects in the translational 
apparatus frequently result in changes in -1 PRF efficiencies. These changes in turn can 
interfere with virus propagation by altering the ratio of structural to enzymatic proteins 
available for viral particle assembly, possibly leading to the observed loss the ?killer? 
phenotype of the rpl10 mutants. The dual luciferase assay was used to quantitatively 
assess rates of -1 PRF and the inability of ribosomes to correctly recognize termination 
codons (nonsense suppression). Consistent with the genetic analysis above, mutants in 
the C-terminal tail and at the tip of the body of rpL10 generally displayed elevated levels 
of -1 PRF and stop codon read-through, with several mutants displaying lower levels of -
 1 PRF as compared to wild-type (Figure 60). Five of these mutants also displayed 
profound defects in subunit biogenesis as assayed by polysome profiling (Figure 61). Of 
these, three (G81D, K145E, K145R) are located at the tip of the rpL10 body, and two 
(K40M, L206V) are located in the C-terminal tail. All these mutants displayed strong 
?halfmer? defects in the polysome profiles, indicative of defects in 40S and 60S subunit 
joining on the mRNA. In addition, all of these polysome profiles are also marked by a 
significantly elevated 40S/60S ratio, indicative of defects in 60S maturation.
 Importantly, the mutation at the position previously shown to play a role in autism 
(L206V) results in significant changes in both -1 PRF efficiency and ribosome 
biogenesis. Mutants in the rest of the rpL10 body did not show any significant phenotypic 
or functional defects. 
125
 Figure 60. Translation fidelity analysis of rpL10 ?body? and ?tail? mutants. 
Programmed -1 ribosomal frameshifting (L-A) and stop codon misincorporation (UAG)
 values compared to wild-type levels as monitored using dual-luciferase reporters. Wild-
 type level of -1 PRF: 8.1% +/- 0.17%. Wild-type level of nonsense suppression: 0.22% 
+/- 0.01%. Bars indicate s.e.m. (n=4). Mutants of the C-terminal tail are highlighted in 
blue.
126
 Figure 61. Polysome profiling of rpL10 ?body? and ?tail? mutants.
 Sucrose density gradient analysis. Indicated strains of L10 were grown in glucose to 
repress genomic RPL10 for 6 hrs before cells were harvested. Extracts were prepared and 
9 A260 units were sedimented through 7?47% sucrose gradients.
127
 Discussion
 The studies presented here reveal that, in addition to the loop of ribosomal protein 
L10 described previously, the eukaryotic C-terminal domain of the protein is an 
additional important ribosomal control element. Mutations in this element at the L206 
position have been reported in autism patients 180. Deletion of the entire C-terminus (aa 
170-221) or just the tip of the structure (aa 200-221) which interacts with the 5S rRNA 
(Figure 62), is lethal (data not shown).  This suggests that its interaction with 5S rRNA 
plays an important role in ribosome function. In particular, examination of the location of 
the L206 and S201 residues of rpL10 in high resolution structures reveals an apparent 
interaction with A65 of the 5S rRNA (Figure 62). 5S rRNA is known to be important in 
translational fidelity in yeast 239, and mutants of A65 are always lethal 240. Additionally, 
5S rRNA is hypothesized to act as a physical transducer of information 241: it is 
strategically positioned to link various functional centers of the ribosome and provides a 
connection to the small subunit. Thus, we hypothesize that the rpL10-L206/S201/5S-A65 
?sandwich? interaction is an important component of the allosteric communication 
pathway initiated by the rpL10 loop in response to ligand binding at the A-site (Chapter 
2). Disruption of this interaction likely results in an ?uncoordinated? ribosome and 
translational fidelity defects. This is supported by the fact that these two mutations, 
particularly S201F, display some of the more profound growth and function defects 
(Figures 59-61). To test this hypothesis, we have generated additional mutations at the 
L206 position spanning a range of side-chain properties (L206A/C/E/F/K/M/T). Notably 
this includes the exact L206M mutation found in autistic patients. As of the time of this 
writing, shuttle vectors containing these mutants have been generated, and the 
128
 corresponding yeast strains are being prepared. It is likely that most of these substitutions 
will be lethal: very significant changes in phenotype and fidelity seen in rpl10-L206V
 (Figures 59-61) despite minimal changes in side chain chemistry suggests that a very 
particular delicate type of interaction is needed at that position. 
Biochemical, translational fidelity, and gene expression analyses of the viable 
mutant strains will be performed and will provide insight into their hypothesized 
modulating effects on ribosome function, which might constitute a leading cause of the
 distinct neurodevelopmental syndrome autism. This disorder is a most often associated 
with mental retardation, and is caused mainly by genetic factors with at least three and 
possibly up to 100 genes 242. However, more than a hundred candidate genes have been 
investigated without conclusive evidence for involvement in autistic disorders 243, and 
functional insight into the mechanism of the disorder is missing. Because there is a high 
sequence conservation in rpL10 between humans and yeast (65% protein identity), and 
the amino acid at position 206 is invariably a leucine, these results should provide a solid 
foundation for future studies in regards to ribosome dysfunction and the onset of autism. 
Additionally, biochemical and structural analysis of the rpl10-L206 mutants will 
provide a more complete view of how the protein is able to sense ligand binding at the 
ribosomal A-site, and then communicate this information to distally located functional 
centers. Mutants in the body region of L10 show far smaller phenotypic and functional 
defects, except those at the very tip most closely approaching the factor binding site, and 
most C-terminal mutants show similar profound defects (Figure 62). Because the more 
deleterious mutations face the factor binding site, it is likely that they also influence 
ligand binding in this region. In particular, non-enzymatic binding experiments using 
129
 elongation ternary complex will reveal the role of the C-terminal end in biochemical 
function: for instance, amelioration of binding defects at the A-site seen in non-enzymatic 
binding would indicate that the C-terminal tail is a structure that is required for binding 
the protein component of the ternary complex. This would point to rpL10 participating in 
the interaction and coordination of the entire ternary complex: the loop senses the tRNA 
arrival at the A-site, while the C-terminal may act as a docking site for the elongation 
factor. Such results would likely indicate that the body of rpL10 provides a platform on 
which the two more dynamic structures, the loop and C-terminus, are attached. In this 
model, the body of rpL10 functions akin to a piston that is physically linked to multiple 
functional centers of the machine. The dynamic structures are involved in ligand binding 
and transduction of information and are sensitive to mutations as a result, while the
 robust ?-sheet foundation is much more tolerant to single amino acid substitutions.  
Additionally, chemical probing of 5S RNA in L206 mutants will unravel the role of the 
L206/S201/A65 ?sandwich? or ?hook? and the structural changes that need to take place 
at this position to ensure proper allosteric communication within the ribosome.  
130
 Figure 62. The rpL10 C-terminal tail and location of the L206 ?autism residue.? 
The location of L10 in the ?crown view? of the LSU is shown on the top right. Indicated 
rotation of this structure and zooming in yields the view on the left, showing the backside 
of L10 composed of the C-terminal tail and the tip of the body of the protein. Location of 
some of the mutations causing profound growth, fidelity, and biogenesis defects are 
shown in yellow. Further indicated rotation and zooming in yields the structure on the 
bottom right. Location of L206 is indicated in blue, A65 of the 5S rRNA is shown in 
green. The first amino acid that is part of the C-terminal truncation (?aa200-221) in the
 lethal mutant is indicated with a black arrow. 
131
 Chapter 5
 Conclusion and future directions
 At the fundamental level, this thesis endeavors to answer a basic biological 
question: how do ribosomes work? Particularly, how are unidirectionality, order, and 
quality control achieved during the process of translation? Chapters 2-5 describe the 
discovery of a central ribosomal control element participating in the coordination of all 
these parameters. Malfunction of this element contributes to promoting the neoplastic 
disease acute lymphoblastic T-cell leukemia (T-ALL). 
Summary and significance 
A flexible loop of the essential ribosomal protein L10 is strategically positioned 
near the ribosomal peptidyltransferase center, and is the first sensor of arrival of ligands 
at the ribosomal A-site. The ligand occupancy information is first transduced locally by 
movement of the protein platform as a result of dynamic changes in loop positioning, to 
ultimately all important functional centers of the ribosome through rRNA networks,
 setting the ribosome rotational mechanism in motion. While many other ribosomal 
proteins are also essential, L10 is a primary sensor and thus an essential initiator of 
allosteric communication pathways necessary for ribosome dynamics. As a result, any
 defects in this initial step get amplified through the subsequent series of outward 
radiating allosteric networks, ultimately leading to biologically significant ribosome 
dysfunction. This is a very significant finding in the field: contrary to the current view 
132
 that rotation can only be achieved through GTP hydrolysis of trans-acting factors, this 
shows that the ribosome alone can actively influence its own rotational status through this 
mobile loop element. Throughout this thesis and in the field in general, the ribosome is 
frequently referred to as a ?molecular machine?. And while translation does require 
tremendous sophistication and accuracy, this terminology perhaps stereotypes the view of 
the ribosome as a passive participant. The studies of the L10 loop and its involvement in 
ribosome structure and function throughout its lifecycle have shown that the ribosome is 
a more capable ?nanomachine? than previously thought, and such knowledge can be 
applied in nanotechnology. Concerns of machine design center around how structure of 
each component contributes to function, and how these components interact dynamically 
to ensure proper function. As one of the primary goals of nanotechnology is to create 
increasingly smaller machines that operate at the nanometer range, studies as this provide 
the foundation for the design of new classes of nanodevices, and are crucial complements 
to the structural work that has been revolutionizing our knowledge of the ribosome.
 While these studies offer a detailed description of the L10 loop function, they ultimately 
also provide a wealth of new information to the efforts to form a ?big picture? regarding 
information exchange in the ribosome. For example, this work for the first time identifies 
landmarks of rotated and non-rotated yeast ribosomes at single nucleotide resolution, as 
well as several long-range interactions between distally located functional centers -
 information that will be tremendously useful to future researchers of ribosome 
mechanics. 
Despite more than a half-century of intensive research on the ribosome, the exact 
roles of many ribosomal components in its biogenesis, and the link between ribosome 
133
 biogenesis and translation, are still poorly understood. The findings presented in this 
project improve our understanding of this link between ribosome biogenesis and 
translational fidelity. The L10 flexible loop was found to also be involved in a 60S 
quality control point during ribosome biogenesis, generating the first model of a ?test 
drive? of the large subunit. Indeed, bypassing of this test drive can promote suppression 
of slow-growing phenotypes. 
Medical relevance and future avenues 
Because of the essential broad roles played by L10 in biogenesis and translation, 
defects in the loop are linked to ribosome dysfunction and disease as exemplified by T-
 ALL. Analysis of such defective ribosomes and cells led to a possible description of the 
molecular mechanism of this disease, from the underlying structural changes to the 
biochemical and functional defects, and out to biological function. This illuminates a 
pathway from genotype to phenotype and in so doing identifies critical control nodes that 
can be targeted for therapeutic intervention (Figure 63). For example, this knowledge 
coupled with future small-drug screens could be used to identify new drugs and drug 
targets that can be exploited to specifically eliminate cells with acquired ribosome 
defects. Such drugs should cause minimal side effects, because they will leave the 
healthy cells without ribosome defects untouched. -1 PRF also represents an attractive 
drug target, as unpublished work from our lab has revealed that levels of frameshifting 
can be attenuated with miRNAs. Nonsense-mediated decay (NMD) could be altered by 
knock-down of components of the NMD machinery, though effects of such a treatment 
are unknown and could be severe, and knockout of NMD components is lethal in 
134
 embryonic mice 244. However, siRNA knockdown levels of specific NMD components 
could be controlled which could significantly decrease the degradation of target messages 
of interest. Alternatively, NMD could be controlled by miRNA-mediated stabilization of 
specific -1 PRF containing messages, leaving global NMD intact. 
The presence of acquired ribosome defects in T-ALL cancer cells is a novel 
finding and it is likely that this mechanism is of general relevance in cancer. For 
example, cancer risks have been documented in much detail in patients with the 
ribosomopathy Diamond-Blackfan anemia (DBA). Although DBA patients have a 5-fold 
higher incidence to develop cancer in general, the cancers they are most predisposed to 
are colon cancer (36-fold higher incidence than in the general population), osteosarcoma 
(33-fold higher) and acute myeloid leukemia (28-fold higher) 235. It is thus not unlikely 
that acquired ribosome defects may also be present in these cancer types. Therefore, 
public datasets could be used to analyze these particular tumor types first. Similar
 analyses can be extended to all other possible tumor types for which data are available
 afterwards. Such work should pave the way towards the development of diagnostic tests 
allowing the early identification of patients with acquired ribosome defects. Over the 
longer term, it may also lead to the development of innovative diagnostic tests for cancers 
associated with acquired ribosome defects. Development of this class of tests will enable 
earlier intervention using therapies tailored to specifically target the translation defect 
specific to the individual patient - personalized medicine.
 While mutations in several ribosomal proteins can lead to the onset of DBA, 
patients also exhibit tissue-specific defects including limb defects, cleft palate, 
abnormalities in heart development, growth failure and a predisposition for cancer 245. 
135
 Interestingly, mutations in different ribosomal proteins also give rise to different types of 
birth defects. For instance, most patients with rpL5 mutations have a cleft palate, whereas 
most individuals with rpL11 mutations do not have any craniofacial defects 246. As such, 
patients with genetic disorders that are linked to mutations in ribosomal proteins show 
remarkably specific phenotypes, suggesting that ribosomal proteins have unique 
functions in different tissues. Studying the molecular mechanism of these defective 
ribosomes can contribute to the exciting emerging field of ?specialized ribosomes? ?
 ribosomes whose specificity may be controlled and fine-tuned by tissue-specific and 
developmentally regulated signals 247. The hypothesis that despite the constitutive nature 
of the ribosome, several components could also give rise to divergent regulation of the 
molecular machine can be compared to the combustion engine: many subtle variations of 
the engine exist which are fine-tuned to a diversity of machines and applications, but the 
fundamental underlying mechanism remains the same. Direct proof supporting the 
existence of such specialized ribosomes would surely impact the field in a manner similar 
to the first ribosomal electron microscopy images of the 1970s and the first high 
resolution structures of the early 2000s. 
Final words 
The above examples illustrate that there are multiple exciting avenues for future 
studies on L10 as well as other ribosomal proteins in both yeast and mammals. Similar to 
scientists who have come before me, particularly Dr. Alexey Petrov who left behind a 
great platform to initiate my own PhD project, I am grateful and happy to have provided 
136
 a solid foundation for future graduate students. To borrow from the thesis title of the 
former lab member Dr. Michael Rhodin 248 ? we are all ?cogs in the nanomachine.? 
137
 Figure 63. Ribosomopathies: proposed chain of causality. 
An acquired mutation in L10 causes ribosomes to distribute more towards a rotated state. 
This affects the binding of ligands, which in turn drives elevated levels of - 1PRF and the 
resulting change in mRNA abundances. A changed protein expression profile induces 
cellular stress, which can lead to three possible mutually non-exclusive pathways. 
138
 Chapter 6
 Materials and Methods
 Media, strains, plasmids, and genetic manipulation. 
E. coli DH5? was used to amplify plasmid DNA. Transformation of E. coli and 
yeast, and preparation of yeast growth media (YPAD and synthetic drop out medium) 
were as reported earlier 249. Restriction enzymes were obtained from Promega 
(Madison, WI, USA) and Roche Applied Science (Indianapolis, IN, USA). DNA 
sequencing was performed by Genewiz (Germantown, MD, USA). 
The haploid S. cerevisiae strain AJY1437 (MAT? rpl10::Kan lys?0 met15?0 
leu2?0 ura3?0 his3?0 pAJ392) containing wild-type RPL10 on a centromeric URA3
 vector (pAJ392) has previously been described 187. In AJY3222, the wild type vector 
was replaced by wild type RPL10 on a centromeric LEU2 vector (pAJ2522) through 
standard 5-fluoroorotic acid (5-FOA) shuffling techniques 250.  AJY3209 harbors 
pAJ2609, a centromeric LEU2 vector expressing the rpl10-S104D allele.  Similarly, 
AJY3212 contains pAJ2612, which expresses the rpl10-A106R mutant from a 
centromeric LEU2 vector; AJY2784 contains pAJ2726, which expresses the rpl10-
 R98S mutant from a centromeric LEU2 vector. Generation of the rpl10-F94I and 
rpl10-G81D expressing strains, JD1308.F94I (MAT? rpl10::Kan met15?0 leu2?0 
ura3?0 his3?0 pJD589.F94I.HIS) and JD1308.G81D (MAT? rpl10::Kan met15?0 
leu2?0 ura3?0 his3?0 pJD589.G81D.HIS), was previously described 178. 
Identification of suppressing factors has previously been described 187. In suppression 
studies, AJY3209 was transformed with pAJ2240, which expresses TIF6-Y192F from 
139
 a centromeric URA3 vector; AJY2784 was transformed with pAJ2805, which 
expresses NMD3-Y379D from a centromeric URA3 vector. AJY2104 186 was 
transformed with pAJ2522 or pAJ2609 and RPL3 or rpl3-W255C expressed from 2 
micron TRP1 vectors. Standard molecular biology techniques were used to subclone 
RPL3 and rpl3-W255C into the HIS3 selectable pRS423 251. AJY3209 was 
subsequently transformed with pRS423-RPL3 and pRS423-rpl3-W255C.
 Mutations of the RPL10 C-terminal domain were generated using the mega-
 oligo mutagenesis method as previously described 252 (primers listed in Table 6) using 
pJD589 as the template. The generated plasmids (see Appendix 2) were transformed 
into JD1293 expressing RPL10 from a 2 micron URA3 maintenance plasmid, grown 
on selective (- his) medium, after which viable cells having lost the wild-type RPL10-
 containing vector were selected by replica plating on 5-FOA. 
Dilution spot assays were performed by growing yeast to mid-log growth 
phase in liquid culture, and spotting them in tenfold serial dilutions from 105 to 101
 colony-forming units per spot on appropriate media, followed by a two day incubation 
at 30?C or as appropriate. 
Translational fidelity and polysome analyses.  
The dual luciferase reporter plasmids pYDL-control, pYDL-LA, pYDL-Ty1, 
pYDL-UAA 213, and pYDL-AGC218 and pYDL-TCT218 212 were employed to monitor 
programmed -1 ribosomal frameshifting, programmed +1 ribosomal frameshifting, 
suppression of UAA, and suppression of an AGC near-cognate serine codon and a 
TCT non-cognate serine codon in place of the cognate AGA codon in the firefly 
140
 luciferase catalytic site, respectively. The reporters were expressed from high-copy 
URA3/LEU2-based plasmids (pJD375, pJD376, pJD376, pJD431, pJD642, pJD643).  
Assays were performed as previously described 253.  Sample readings were collected 
using a GloMax Multi-Microplate luminometer (Promega). All assays were repeated 4 
times. Sucrose density gradient analysis was carried out as described 187.
 mRNA abundance and telomere length analyses. 
qRT-PCR experiments to assay mRNA abundance were carried out as 
described 161. Similar methods were used to quantify telomere length in yeast cells. 
Genomic DNA was isolated from mid-logarithmic cell cultures using the ?Smash & 
Grab? yeast DNA preparation method as described 254. Each DNA sample was diluted 
serially. The telomere PCR (T) and the single copy reference gene SGS1 PCR (S) were 
performed using the Bio-Rad iTaqTM Universal SYBR Green system using primer 
pairs complementary to telomere repeats (listed in Table 7), and cycle threshold (Ct) 
values were determined. The T/S ratios were calculated from three experimental 
replicates at each of three DNA concentrations (100, 200 and 400 ng).
 Peptidyltransferase activity. 
Single turnover peptidylpuromycin reactions were performed to assay apparent 
rates of peptidyltransfer as described 255 with the following modification. Ribosomes 
pre-bound with Ac-[14C]Phe-tRNAPhe and polyuridylic acid (polyU) were loaded onto 
pre-wetted Millipore HA (0.45 micron) filters and washed with binding buffer. The 
141
 filters were placed into 15 mL scintillation vials and 2.4 mL of binding buffer + 0.05% 
Zwittergent (EMD BioScience), followed by a 30 min incubation on a rocker at 4?C. 
Aliquots (1 mL) were taken from each vial, incubated for 5 min at 30?C to activate 
ribosomes, reactions were initiated by adding pH-neutralized puromycin to 10 mM 
final concentrations, and the procedure completed as described 255. Reactions were 
repeated 4 times. 
Sdo1p cloning, purification and labeling. 
Sdo1p was cloned into a modified pET-21a, expressing Sdo1p with a C-
 terminal 6xhistidine tag and a phosphorylatable kemptide (the resulting amino acid
 sequence beyond the last residue of Sdo1: LEHHHHHHLRRASLG. The S gets 
phosphorylated). The protein was expressed in Codon Plus bacteria (Stratagene) and 
purified by Ni-NTA (Invitrogen) chromatography followed by gel filtration on 
sephacryl S200 (GE Healthcare).  Labeling reactions containing 10 ?g of Sdo1p, 5x 
molar excess of [32P]-?-ATP, 1 ?l PKA (NEB) in 100 ?l 1x kinase buffer (NEB) were 
incubated for 15 minutes at 30?C, passed through G25 columns (GE) to remove 
unincorporated ATP, and flow-through was measured by  scintillation counting. Flow-
 through values from control mixtures lacking PKA were subtracted from values of the 
labeling reactions to determine the specific activity of 32P-labeled Sdo1p, yielding 40-
 60% label incorporation. 
142
 Ribosome preparation. 
Purification of active 80S ribosomes using cysteine-charged sulfolink columns 
was performed as described 256, with the following modifications: after elution from 
the column, ribosomes were treated with 1 mM final concentration GTP and 1 mM 
final concentration pH-neutralized puromycin at 30?C for 30 min to remove 
endogenous tRNAs. After a 100,000 x g 16 -20 hr spin through a high salt glycerol 
cushion (20 mM HEPES-KOH pH 7.6, 60 mM NH4Cl, 500mM KCl, 10 mM 
Mg(OAc)2, 2 mM DTT, 25% glycerol), ribosomes were resuspended in elution buffer 
and passed through a low salt cushion (20 mM HEPES-KOH pH 7.6, 50 mM NH4Cl,  
5 mM Mg(OAc)2, 1 mM DTT, 25% glycerol) to increase purity. 
Ribosome/tRNA interactions. 
eEF1A preparation,  purification of aminoacyl-tRNA synthetases, charging of 
tRNAPhe (Sigma) with [14C]-phenylalanine (PerkinElmer) and purification of 
aminoacyl-tRNA and acetylated aminoacyl-tRNA were carried out as described 218,257. 
To assay steady-state dissociation rates (KD) of aa-tRNA to the ribosomal A-site, 2 
sets of reactions were set up in parallel. A mix containing 100 ?g of polyU, 50 pmoles 
of ribosomes, a 4-fold molar excess of tRNAPhe, all in binding buffer (80 mM Tris-
 HCl, pH 7.4 at 30o C, 160 mM NH4Cl, 15 mM Mg(CH3COOH)2, 2 mM spermidine, 
0.5 mM spermine, 6 mM ?-mercaptoethanol) in 150 ?l total volume was prepared and 
incubated for 30 min at 30?C to block the P-site. To prepare the ternary complex 
([14C]Phe-tRNAPhe?eEF1A?GTP), 100 ?g of soluble protein factors, 1 mM final 
concentration of GTP, 125 pmoles [14C]-Phe-tRNAPhe were mixed in in 50 ?l total 
143
 volume binding buffer and incubated for 30 min at 30?C. After incubation, serial 2-
 fold dilutions of the ternary complex reaction mix were prepared, resulting in 8 
fractions containing decreasing amounts of ternary complex (62.5 - 1 pmoles), in 105 
?l each. An equal amount of the ribosome mix (5 pmoles of ribosomes, 15 ?l) was 
added to each dilution, followed by incubation for 30 min at 30oC. The mixtures were 
applied onto pre-wetted nitrocellulose Millipore HA (0.45 micron) filters, washed with 
binding buffer, and radioactivity was measured via scintillation counting. Background 
control reactions without ribosomes were performed at each ligand dilution and 
subtracted from experimental ones. KD values were calculated using GraphPad Prism
 software fitted to single binding site with ligand depletion models:
 Y = (-b+?(b*b-4*a*c))/(2*a)
 Bmax initial Ymax value set to 1.0; a = -1; b = KD + X + Bmax; c = -1*X*Bmax 
X is total ligand added. Y is total binding (X and Y must be expressed in the same 
units).
 Non-enzymatic binding studies were performed likewise, but without eEF1A 
or by using 50 ?g of soluble protein factors (half maximum activity). To test binding 
of tRNA to the P-site, Ac-[14C]-Phe-tRNAPhe was used as the ligand, there was no 
need to pre-block with tRNAPhe, and 11 mM Mg(CH3COOH)2  was used in the binding 
buffer. All reactions were repeated 4 times.  
144
 Ribosome/protein interactions. 
6x-His-tagged eEF2 was purified from TKY675 yeast cells (a generous gift 
from Dr. T. Kinzy) as described 258.  Aliquots containing ribosomes (2 pmoles) were 
first pre-incubated with increasing concentrations of eEF2 (0.25 ? 32 pmoles), 100 ?g 
of polyU, 0.1 mM final concentration GDPNP (Sigma), and 4x molar excess of [14C] 
NAD (PerkinElmer) over ribosomes in 50 ?l total volume binding buffer at 30?C for 
20 minutes. Diphtheria toxin (0.2 ?g, Sigma) was added, and reactions were incubated 
for 30 min at 30?C. After precipitation with TCA (final concentration 15%) and 15 
min incubation on ice, reaction mixtures were applied onto GF/C filters, washed with 
5% TCA, and the amount of [14C]-ADP ribosylated eEF2 was determined by 
scintillation counting. Readings reflect unbound eEF2 and were subtracted from total 
eEF2 to obtain values bound.  Sdo1p binding assays were performed by incubating 
ribosomes (2 pmoles) with increasing concentrations of Sdo1p (0.25 ? 32 pmoles) and 
100 ?g of polyU in binding buffer (50 mM Tris-(OAc)2 pH 7.5 RT, 50 mM 
NH4(OAc)2, 10 mM Mg(OAc)2, 2 mM DTT)  in 50 ?l total volume at 30?C for 20 
minutes. The reaction mixtures were then applied onto pre-equilibrated 1 mL 
polyethylene filter spin columns (Pierce) containing 0.3 mL cysteine-charged sulfolink 
resin and incubated on ice for 5 minutes. The columns were spun and flow-through 
measured via scintillation counting.  Readings reflect unbound Sdo1p and were 
subtracted from total loaded to obtain Sdo1p bound. Background control reactions 
without ribosomes were performed at each ligand dilution. eEF2 assays were repeated 
4 times, Sdo1p assays were performed in triplicate.
145
 Ribosome binding competition. 
To monitor effects of Sdo1p on tRNA binding to the A-site, wild-type 
ribosomes (270 pmoles) primed with polyU (0.35 mg), were first mixed  with a 4-fold 
molar excess of uncharged tRNAPhe, 2.8 nmoles GTP, and 5 ?g of soluble protein 
factors (including eEF1A)  in 700 ?l total volume of ribosome binding buffer. After 
10 min incubation at 30?C, ribosome/tRNA complexes (50 ?l aliquots) were added to 
10 ?l aliquots of 2-fold dilutions of purified Sdo1p (500 pmol ? 31 pmol, plus a no-
 Sdo1p control) and incubated at 30?C for 10 min.  Subsequently, 25 pmol of [14C]Phe-
 tRNA was added to each ribosome/Sdo1p complex, and incubated at 30?C for 10 min. 
The reaction mixtures were applied onto pre-wetted nitrocellulose Millipore HA (0.45 
micron) filters, washed with binding buffer, and radioactivity was measured via 
scintillation counting. Background control reaction values from samples without 
ribosomes were subtracted from experimental ones. To monitor the ability of Sdo1p to 
compete for binding at the P-site, the same conditions were employed except that 
binding reactions were performed in 11 mM magnesium, and [14C]Ac-Phe-tRNAPhe
 was used.  Assays of peptidyltransferase activity were performed as described above 
in the presence of Sdo1p. All assays were performed twice in triplicate.
 Preparation of complexes for chemical probing. 
Fifty pmoles 80S ribosomes isolated from isogenic strains were incubated with 
100 ?g polyU in binding buffer (80 mM HEPES pH 7.5, 50 mM NaCl, 11 mM 
Mg(OAc)2, 6 mM ?-mercaptoethanol) at 30?C for 10 minutes. One set of ribosomes so 
prepared (?empty 80S? wild-type, rpL10-S104D, rpL10-A106R) was employed for 
146
 chemical protection assays.  To prepare control ?non-rotated? ribosomes, 200 pmoles 
N-acetyl-phenylalanyl tRNAPhe were added and incubations continued for 20 minutes. 
To prepare ?rotated? wild-type ribosomes, empty 80S ribosomes were first incubated 
with 200 pmoles of deacylated tRNAPhe.  eEF2 (400 pmoles) and GDPNP (final 
concentration of 1 mM) were then added to the deacylated tRNAPhe - ribosome 
mixture and incubated for an additional 20 minutes. These conditions were based on 
results of binding assays as above. 
rRNA structure probing. 
hSHAPE of rRNA with 1M7 was performed as described 199 using the 
following substrates: empty ribosomes isolated from isogenic wild type cells, cells 
expressing mutant ribosomes, and wild-type ribosomes containing acetylated-aa-
 PhetRNAPhe in the P-site (non-rotated control), and deacylated-tRNAPhe + eEF2-
 GDPNP (rotated control). The following primers were employed: 969 and 1780 in the 
SSU; 25-2, 1466, 2632, 2836, 25-7, and 3225 in the LSU. Data were analyzed using 
SHAPEFinder 197. For kethoxal studies, 25 pmoles of ribosomes in a 50 ?l volume 
were treated with 1 ?l of a 4% kethoxal solution (in pure ethanol), or 1 ?l of ethanol as 
control, and incubated for 10 min at 30?C. Reactions were stopped by addition of one 
half volume of stop solution (150 mM sodium acetate, 250 mM potassium borate), 
followed by analysis as above using primer 969. 
147
 Statistical analyses. 
Student?s t test for two-tailed p-value calculations was used throughout. Data 
analysis of dual-luciferase assays and rRNA probing was carried out as described
 199,253. After generation of rRNA chemical modification data using ShapeFinder 197 the 
median reactivities of each primer region were used to normalize the raw integrated 
peak values.
 Acknowledgments 
We wish to thank our collaborators for technical and intellectual assistance: 
The Arlen Johnson lab at the University of Texas at Austin and the Kim De 
Keersmaecker lab at the Vlaams Instituut voor Biotechnologie in Leuven, Belgium. 
This work was supported by grants to Jonathan D. Dinman from the Public Health 
service (2 R01 GM058859-11, 3 R01GM053655-15S1), and to Arlen W. Johnson (2 
RO1 GM53655 and 3 R01 GM053655-15S1). Sergey O. Sulima was partially 
supported by an NIH training grant T32GM080201.
148
 Appendix 1: Yeast strain list 
Table 4. Yeast strains generated. 
* 1293: (aka AJY1437) MAT? rpl10::Kan met15?0 leu2?0 ura3?0 his3?0 RPL10.URA3.2?, K+
 Strain 
Name Description
 JD1586 JD1293*; ?200-221 in L10
 JD1587 JD1293; ?170-221 in L10
 JD1588 JD1293; L206C in L10
 JD1589 JD1293; L206E in L10
 JD1590 JD1293; L206F in L10
 JD1591 JD1293; L206K in L10
 JD1592 JD1293; L206M in L10
 JD1593 JD1293; L206T in L10
149
 Appendix 2: Yeast plasmid list
 Table 5. Plasmids generated. 
Plasmid 
Name
 Backbone
 Plasmid Insertion
 Bacterial
 Marker
 Yeast
 Marker
 pJD1932 pRS423 Wild type L3 AmpR HIS3
 pJD1933 pRS423 L3-W255C AmpR HIS3
 pJD1934 pJD589 L10-?200-221 AmpR HIS3
 pJD1935 pJD589 L10-?170-221 AmpR HIS3
 pJD1936 pJD589 L10-L206C AmpR HIS3
 pJD1937 pJD589 L10-L206E AmpR HIS3
 pJD1938 pJD589 L10-L206F AmpR HIS3
 pJD1939 pJD589 L10-L206K AmpR HIS3
 pJD1940 pJD589 L10-L206M AmpR HIS3
 pJD1941 pJD589 L10-L206T AmpR HIS3
150
 Appendix 3: Oligonucleotide primer list 
Table 6. Oligonucleotides used in directed mutagenesis. 
* Bases at the 5? end of the deleted region are capitalized; bases at the 3? end of the deletion 
region are written in lower case. 
Strain Mutation Forward and Reverse 5? to 3? Oligonucleotide sequence
 JD1586 L10-?200-221
 CGGTGCTTTCGTTAAGTTCTTGTCCAAGAAGGTgttcttt
 tcaaacatttgaactaac * (forward)
 CAGGAAACAGCTATGAC (reverse)
 JD1587 L10-?170-221
 CCCAGGTCAACAAAAGATTATTTTGTCTAAGAAGTG
 GGGgttcttttcaaacatttgaactaac *
 CAGGAAACAGCTATGAC
 JD1588 L10-L206C
 CCAAGAAGGGTTCTGTGAAAACAACATCAGAGAAT
 TCCCAGAATACTTTGC
 CCTGCGTTATCCCCTGATTCTGTGGATAACCGTATT
 ACCGCC
 JD1589 L10-L206E
 CCAAGAAGGGTTCGAAGAAAACAACATCAGAGAAT
 TCCCAGAATACTTTGC
 CCTGCGTTATCCCCTGATTCTGTGGATAACCGTATT
 ACCGCC
 JD1590 L10-L206F
 CCAAGAAGGGTTCTTTGAAAACAACATCAGAGAAT
 TCCCAGAATACTTTGCTGC
 CCTGCGTTATCCCCTGATTCTGTGGATAACCGTATT
 ACCGCC
 JD1591 L10-L206K
 CCAAGAAGGGTTCAAAGAAAACAACATCAGAGAAT
 TCCCAGAATACTTTGCTGC
 CCTGCGTTATCCCCTGATTCTGTGGATAACCGTATT
 ACCGCC
 JD1592 L10-L206M
 CCAAGAAGGGTTCTATGGAAAACAACATCAGAGAA
 TTCCCAGAATACTTTGC
 CCTGCGTTATCCCCTGATTCTGTGGATAACCGTATT
 ACCGCC
 JD1593 L10-L206T
 CCAAGAAGGGTTCACTGAAAACAACATCAGAGAAT
 TCCCAGAATACTTTGC
 CCTGCGTTATCCCCTGATTCTGTGGATAACCGTATT
 ACCGCC
151
 Table 7. Oligonucleotides used in telomere length assays.
 Oligonucleotide 
purpose
 Oligonucleotide 
name 5? to 3? Oligonucleotide sequence
 Yeast telomere 
repeat probe
 yTel_Fwd CAGTGGTGTGGGTGTGCATGGTGGTGTGGGTGTGTGGAC
 yTel_Rev GCCCACAACCACACCCAACACATCCCA CACCACCTA
 Single copy 
reference gene 
probe
 ySGS1_Fwd GGCTCTCGTACACTGCCACATCGAATC AATATGCTGACGTACCC
 ySGS1_Rev CATGTGGAGATGCGGGAATGCGGGAGAATGTGGAGC
152
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