From The Royal Institute of Technology,
School of Technology and Health
SINGLE-PARTICLE CRYOELECTRON MICROSCOPY
OF MACROMOLECULAR
ASSEMBLIES
Kimberley Cheng
Stockholm 2009
All previously published papers were reproduced with permission from the publisher.
Published by The Royal Institute of Technology. Printed by Universitetsservice US-AB
© Kimberley Cheng, 2009
ISBN 978-91-7415-509-9
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ABSTRACT
In this thesis, single-particle cryo-electron microscopy (cryo-EM) was used to study the
structure of three macromolecular assemblies: the two hemocyanin isoforms from
Rapana thomasiana, the Pyrococcus furiosus chaperonin, and the ribosome from
Escherichia coli.
Hemocyanins are large respiratory proteins in arthropods and molluscs. Most
molluscan hemocyanins exist as two distinct isoforms composed of related
polypeptides. In most species the two isoforms differ in terms of their oligomeric
stability, and thus we set out to investigate the two Rapana thomasiana hemocyanins
(RtH) in order to explain this behaviour. Our findings showed that the two RtH
isoforms are identical at the experimental resolution. Furthermore, three previously
unreported connections that most likely contribute to the oligomeric stability were
identified.
Chaperonins are double-ring protein complexes that assist the folding process of
nascent, non-native polypeptide chains. The chaperonin from the hyperthermophilic
archaea Pyrococcus furiosus belongs to Group II chaperonins, and unlike most other
group II chaperonins it appears to be homo-oligomeric. The 3D reconstruction of the
Pyrococcus furiousus chaperonin revealed a di-octameric structure in a partially
closed/open state, something in between the closed folding-active state and the open
substrate-accepting state.
The ribosome is the molecular machine where protein synthesis takes place. In
bacteria there is a unique RNA molecule called transfer-messenger RNA (tmRNA) that
together with its helper protein SmpB rescues ribosomes trapped on defective
messenger RNAs (mRNAs) through a process called trans-translation. tmRNA is about
4 times the size of a normal tRNA, and it is composed of a tRNA-like domain (TLD)
that is connected to the mRNA-like domain (MLD) by several pseudoknots (PKs) and
RNA helices. During trans-translation, tmRNA utilize its TLD to receive the
incomplete polypeptide from the peptidyl-tRNA in the ribosomal P site of the stalled
ribosome. Subsequently, its MLD is used to tag the incomplete polypeptide with a
degradation signal. When tmRNA enters a stalled ribosome the MLD and pseudoknots
form a highly structured arc that encircles the beak of the small ribosomal subunit. By
utilizing maximum-likelihood based methods for heterogeneity analysis we could
observe the Escherichia coli ribosome in a number of different tmRNA·SmpB-bound
states. The cryo-EM map of the post-accommodated state revealed that the TLD·SmpB
part of the tmRNA·SmpB complex mimics native tRNAs in the A site of stalled
ribosomes. The density map also showed that the tmRNA arc remains well structured
and that it is still attached to the beak of the small ribosomal subunit. The
reconstructions of the double-translocation tmRNA-bound ribosome complex showed
that the pseudoknots of tmRNA still form an arc, and that they are located at positions
similar to the ones assigned for the pseudoknots in the post-accommodated state. In
addition, the tmRNA arc exists in two states; one stable and highly structured and
another more flexible and disorganized.
LIST OF PUBLICATIONS
I.
Cheng K., Koeck P.J.B., Elmlund H., Idakieva K., Parvanova K., Schwarz H.,
Ternström T. and Hebert H. Rapana thomasiana hemocyanin (RtH):
Comparison of the two isoforms, RtH1 and RtH2, at 19A and 16A resolution.
Micron (2006) 37:566-76
II.
Karlström M., Cheng K., Purhonen P., Ladenstein R., Hebert H. and Koeck
P.J.B. Low resolution structure and apparent melting temperature of the
chaperonin from Pyrococcus furiosus. (2009) Manuscript
III.
Cheng K., Ivanova N., Scheres S.H.W., Pavlov M.Y., Carazo JM., Hebert H.,
Ehrenberg M. and Lindahl M. tmRNA·SmpB complex mimics native
aminoacyl-tRNAs in the A site of stalled ribosomes. J. Struct Biol. 2009 (In
press)
IV.
Cheng K., Ivanova N., Scheres S.H.W., Pavlov M.Y., Carazo JM., Hebert H.,
Ehrenberg M. and Lindahl M. Structural analysis of double translocated
tmRNA on the 70S ribosome indicates flexibility of the tmRNA structure.
(2009) Manuscript
CONTENTS
Introduction.......................................................................................................... 1
Hemocyanin – An oxygen transporter ................................................................ 2
Structure of molluscan hemocyanins........................................................... 2
Medical Significance.................................................................................... 3
Rapana thomasiana hemocyanin................................................................. 3
Chaperonin – protein folding assistant................................................................ 6
Structure and mechanism............................................................................. 6
Ribosome – The workbench of translation ......................................................... 8
The message – mRNA......................................................................... 8
The adaptor molecule - tRNA............................................................. 9
The protein-synthesizing machine – The ribosome ......................... 10
The translation process............................................................................... 12
Initiation ........................................................................................... 12
Elongation ........................................................................................ 12
Termination ...................................................................................... 14
Trans-Translation ....................................................................................... 14
tmRNA and SmpB: structure and function ...................................... 15
The mechanism of trans-translation ................................................ 17
Physiological significance of trans-translation............................... 18
Structural studies of tmRNA·ribosome complexes .......................... 18
Single-Particle Cryo-Electron Microscopy....................................................... 22
Specimen preparation................................................................................. 22
Negative Staining.............................................................................. 22
Vitrification....................................................................................... 22
Data Processing .......................................................................................... 23
Heterogeneity analysis ............................................................................... 25
Conclusions........................................................................................................ 29
Acknowledgements ........................................................................................... 30
References.......................................................................................................... 32
LIST OF ABBREVIATIONS
3D
5S RNA
16S RNA
23S RNA
30S
50S
70S
aaRS
Ala-tmRNA
ASL
ATP
ATPase
Cs
CCA
CCT
Cryo-EM
CTF
DNA
E. coli
EF-G
EF-Tu
EM
FU
GDP
GroEL
GroES
GTP
GTPase
H. tuberculata
HtH
IF
KLH
M. crenulata
ML
MLD
mRNA
NMR
NpH
ORF
P. furiosus
Pf Cpn
PK
PTC
RF
Three-dimensional
Small RNA molecule in 50S
RNA molecule in 30S
Large RNA molecule in 50S
Small subunit in 70S
Large subunit in 70S
Bacterial ribosome
Aminoacyl-tRNA synthetase
tmRNA charged with alanine
Anticodon stem-loop
Adenosine triphosphate
Enzyme hydrolyzing ATP
Spherical aberration of the EM lens system
Nucleotide sequence at the 3’-end of all tRNA molecules
Eukaryotic chaperonin (Chaperonin containing TCP-1)
Cryo-electron microscopy
Contrast transfer function
Deoxyribonucleic acid
Escherichia coli
Elongation factor G
Elongation factor Tu
Electron microscopy
Functional unit
Guanosine diphosphate
Group I chaperonin in E. coli
Co-protein to GroEL
Guanosine triphosphate
Enzyme hydrolyzing GTP
Haliotis tuberculata
Haliotis tuberculata hemocyanin
Initiation factor
Keyhole limpet hemocyanin
Megathura crenulata
Maximum-likelihood
mRNA-like domain
Messenger RNA
Nuclear magnetic resonance
Nautilus pompilius hemocyanin
Open reading frame
Pyrococcus furiosus
Chaperonin from P. furiosus
Pseudoknot
Peptidyl transferase center
Release factor
RNA
RNase
RRF
rRNA
RtH
SmpB
smpB
SNR
ssrA
TEM
TLD
tmRNA
tRNA
Ribonucleic acid
Enzyme degrading RNA
Recycling factor
Ribosomal RNA
Rapana thomasiana hemocyanin
Small protein B
Gene encoding SmpB
Signal-to-noise ratio
Gene encoding tmRNA
Transmission electron microscopy
tRNA-like domain
Transfer-messenger RNA
Transfer RNA
INTRODUCTION
Many tasks in the living cell are performed by proteins. Since structure and function are
closely correlated, proteins must obtain a specific structure to function properly. While
many proteins function by themselves, some are multiply used to form large
macromolecular complexes resulting in an incredible large diversity of shapes and sizes
of these assemblies [1]. Similar to individual proteins, macromolecules are not static in
their native state. Instead they undergo functionally important conformational changes,
often triggered by ligand binding, in order to perform their specific biological activity.
Take hemoglobin for instance. Hemoglobin is a respiratory protein whose function is to
transport oxygen in the cells. It is composed of four protein molecules (called subunits
in the context of the complex). When hemoglobin go between the oxygenated and deoxygenated state it “breathes”, i.e. the subunits move in relation to each other. This
allows the oxygen-binding sites to communicate with each other to increase or decrease
the affinity towards oxygen. Thus, by performing structural studies of a macromolecule
complex in its different functional states, and characterizing the differences, a deeper
insight into its biological process can be obtained.
X-ray crystallography and Nuclear Magnetic Resonance (NMR) are the traditional
methods used to determine the three-dimensional (3D) structure of proteins at atomic
resolution. NMR has the advantage of analyzing proteins in solution, thus allowing the
protein to be in all its functional states. However, this technique is limited to studying
proteins smaller than 50kDa. X-ray crystallography on the other hand has been an
extremely useful tool for determining the structure of large individual proteins, but it
runs into difficulties with large multi-component complexes. In addition, to form
crystals the protein complex needs to be in one conformational state only. In many
cases this is not easy to achieve, especially when the complex has a flexible region.
Thus, often formation of crystals only takes place when the flexible part has been
removed. Furthermore, crystal packing may lead to steric constraints that limit the
number of functional states the complex can assume and thereby restrict the functional
states that can be studied.
A technique for structural studies that has attracted growing interest over the past
decade is single-particle reconstruction. Single-particle reconstruction is an
experimental/computational technique that utilizes transmission electron microscopy
(TEM) to address problems which crystallographic methods are unable to cope with.
Combined with cryo-electron microscopy (cryo-EM) it is often the choice of method
for structural studies of macromolecular complexes.
In this thesis, single-particle cryo-EM was conducted on three different
macromolecular assemblies. The first paper describes a comparison of the two
hemocyanin isoforms from the marine snail Rapana thomasiana. Structural
characterization of the chaperonin from the hyperthermophilic archaea Pyrococcus
furiosus is carried out in paper II. In paper III and IV, two functional states of the
ribosome from Escherichia coli stalled at a truncated messenger RNA (mRNA) are
studied.
1
HEMOCYANIN – AN OXYGEN TRANSPORTER
Hemocyanins are giant copper-containing multi-subunit respiratory proteins, freely
dissolved in the hemolymph of many arthropod and mollusc species [2]. The oxygen
binding sites in the two phyla are very similar, both in the way they involve a pair of
copper atoms to how the copper is coordinated via histidine ligands (Figure 1) [3, 4].
Apart from these similarities the molluscan and arthropod hemocyanins represent two
individual classes of proteins with distinct tertiary and quaternary structure.
Figure 1. The oxygen-binding site in arthropod and molluscan hemocyanins. The copper atoms
are coordinated via histidine ligands. Coppers are brown and oxygens are red. Upon oxygenation the
copper-copper distance in molluscan hemocyanin increases [3, 5]. PDB code: 1JS8.
STRUCTURE OF MOLLUSCAN HEMOCYANINS
Molluscan hemocyanins are based on large polypeptide chains, each ranging from 350
kDa to 450 kDa, that are folded into 7 (cephalopods) or 8 (gastropods, chitons and
bivalves) globular functional units (FUs, termed a-g/h). The FUs, each containing one
oxygen-binding site, are connected by linker peptide strands thus yielding a subunit
structure that resembles a pearl-necklace (Figure 2A). Two subunits are paired in an
anti-parallel fashion as stable dimers (the repeating unit), and in the basic quaternary
structure of molluscan hemocyanins five such oblique dimers assemble into a hollow
cylindrical molecule (Figure 2B) with an internal collar complex. In cephalopods and
chitons, the decamer is the only structure found, whereas in gastropods and bivalves
these decamers form di-decamers or even higher oligomers [2, 6, 7].
Structural information of hemocyanins has come from EM (e.g. immunoelectron
microscopy [8, 9], cryo-EM [10-13]), X-ray crystallography [3, 5] and Small-angle
neutron scattering studies [14]. Several pathways for the subunits have been proposed
based on the immunoelectron microscopy studies [9], and recently one of them was
suggested by Gebauer and colleagues to be the most probable one [15].
It has been shown in some gastropods, e.g Megathura crenulata [16, 17] and Haliotis
tuberculata [18], that the hemocyanin exists in two structurally distinct isoforms and
that they are homo-oligomeric. In Helix pomatia, up to three different hemocyanin
isoforms are found [19]. Although the different isoforms in these species are composed
of closely related subunits, they differ in terms of their oligomeric stability [20-24],
carbohydrate content [25] and oxygen-binding properties [23, 26].
2
(A)
(B)
Figure 2. The basic structure of the molluscan hemocyanin. (A) Schematic of the polypeptide chain
(subunit) with its eight different functional units. (B) Schematic of two oblique anti-parallel subunits (the
repeating unit) arranged about a molecular 5-fold symmetrical axis within the cylindrical decamer. Note,
in order to simplify the description at this stage the two subunits in the dimer are displayed as linear. In
reality, the pathway of the subunit has still not been firmly established. However, it is widely
acknowledged that the two subunits within the dimer are anti-parallel.
MEDICAL SIGNIFICANCE
Over 30 years ago the keyhole limpet hemocyanin (KLH) from the marine gastropod
M. crenulata was found to possess immunostimulatory properties [27-30]. Most
hemocyanins are glycoproteins, and the immunological response of KLH has often
been related to its specific carbohydrate content and composition. For instance, the
success of using KLH for the treatment of bladder carcinoma (reviewed in [31]) is
probably due to a carbohydrate epitope that is cross-reactive with an equivalent epitope
on the bladder tumor cell surface [32].
KLH is also widely used as a hapten carrier [33]. Haptens are small molecules such
as drugs, hormones and peptides that, in general, require the assistance of a larger
carrier protein to stimulate the immune system for the production of antibodies. The
efficiency of KLH as a carrier protein is due to its large molecular size and its
abundance of lysine residues for coupling haptens. For other medical applications of
KLH, see [33].
The successful use of KLH in both clinical and biotechnological applications have
made it interesting to study hemocyanins from other species (mostly gastropods),
especially their structure and carbohydrate content. Recently it was shown that the
hemocyanin from Concholepas concholepas possess adjuvant immunostimulatory
properties, as well as immunotherapeutic effects against bladder cancer [34, 35].
Moreover, the hemocyanin from H. tuberculata has been considered to be a possible
substitute/complement for KLH in biotechnological purposes due to its higher
oligomeric stability under certain ionic conditions [22].
RAPANA THOMASIANA HEMOCYANIN
Rapana thomasiana (prosobranch gastropod) is a marine snail that was originally
found along the coast of Japan. However, fifty years ago it was discovered in
relatively large amounts along the west coast of the Black Sea. Considered that the
Black Sea is practically a closed sea with salinity less than half of what the Pacific
Ocean contains, it might be expected that some physiological properties of this
3
gastropod will have been affected to adapt to the new environmental conditions.
Similar to M. crenulata and H. tuberculata, its hemocyanin (RtH) is also present in
two independent homo-oligomeric isoforms [36]. RtH are composed of two
structurally distinct subunits, RtH1 and RtH2, with molecular masses of 420 kDa and
450 kDa, respectively. Recently it was shown that RtH possess immunostimulatory
properties similar to those of KLH [37, 38]. Previous EM studies of specimen
prepared from native RtH containing both isoforms showed that the complexes were
solitary di-decamers [36]. Apart from this, no additional structural studies were
performed.
The different hemocyanin isoforms in most mollusc species, including R. thomasiana,
exhibit differences in their re-association behaviour under different ionic conditions
[20-24, 36]. In paper I the structure of the two RtH isoforms were studied in order to
detect something that could, hopefully, provide an explanation to this behaviour. The
cryo-EM maps showed that the quaternary structure of RtH1 and RtH2 (Figure 3)
were identical at the resolution of the experiment. While the reconstructions did not
give any clues to why the two isoforms differ in their oligomeric stability, they did
reveal three previously unreported connections. Based on the subunit pathway
proposed by Gebauer et al. [15], one connection was identified to be within the
subunit dimer (Figure 3D, full arrow) while the other two were between two adjacent
subunit dimers (Figures 3E and 3F, full arrows). The former connection is most likely
involved in stabilizing the subunit dimer while the two latter connections are formed
during the pentamerization process to stabilize the decameric ring. Molluscan
hemocyanins require Ca2+ and/or Mg2+ ions to maintain their quaternary structure. A
decrease of these ions results in dissociation of the oligomeric complex into
decamers, subunit dimers or individual subunits depending on the ionic conditions
[22, 36, 39]. It is therefore possible that the aforementioned non-covalent connections
provide binding sites for Ca2+ and Mg2+.
After the publication of our work, the three non-covalent connections previously
mentioned were also observed in HtH1 (one of the hemocyanin isoforms from H.
tuberculata) [40], KLH1 [41] and also in the Nautilus pompilius (cephalopod)
hemocyanin (NpH) [40, 42]. However, compared to our work the connections were
designated differently due to the new subunit pathway proposed. The new pathway of
the subunits in NpH was determined by fitting homology models of the FUs into a 9Å
cryo-EM map [42]. Recently it was shown that this subunit arrangement is valid for
KLH1 as well [41], and thus it was proposed that it also applies to other molluscan
hemocyanins. The most remarkable with the new subunit pathway is the suggestion
that the two FUs in the arch (FU-g) are from two adjacent subunit dimers [42].
If the FUs in our cryo-EM maps are re-assigned according to the new subunit
pathway [42], the linker region between FU-fY (wall) and FU-gY (arch) will be
absent. The covalent linker regions are very long, and are thus presumed to be very
flexible. According to the new pathway, the FU-fY ĺ FU-gY linker region spans a
very long distance [42]. In addition, it is also highly exposed. It is therefore likely that
it shows variability in conformation during the cooperative oxygen binding process,
and as a result it is averaged out in our reconstructions. Thus, the absence of this
linker region in our cryo-EM maps is most likely due to the presence of different
hemocyanin conformers in the sample. In the recent study where a 9Å cryo-EM
density map of KLH1 was obtained, the sample was kept in a chamber containing
25% oxygen prior to grid preparation for the EM work [41]. This procedure was
carried out in order to ensure full oxygenation of the hemocyanin complexes, and is
probably what enabled the authors to define this linker region in their reconstruction.
4
(A)
Tiers
(B)
(C)
1st
2nd
3rd
(D)
(E)
(F)
Figure 3. 3D reconstruction of RtH2. FUs labeled with yellow (FUY) and white (FUW) colors
correspond to FUs within the front subunit dimer, while FUL and FUR refer to FUs from its left and right
neighboring subunit dimers, respectively. (A) Top view. (B) External side view. The three tiers of the
wall and the major groove (broken line) within the subunit dimer are marked out. (C) Internal side view
showing the arch (FU-g) and the collar elements (FU-h). (D) Fragment of the RtH2 map showing the
non-covalent intra-subunit dimer connection between FU-gY and the first tier of the wall (full arrow), and
the covalent linker region (broken arrow). (E) Same fragment as in (D) rotated about 90° to reveal the
linker region (broken arrow), and the non-covalent inter-subunit dimer connection (full arrow) to FU-gW.
(F) The non-covalent collar-wall connection between individual subunit dimers (arrow).
5
CHAPERONIN – PROTEIN FOLDING ASSISTANT
Chaperonins are a specific class of molecular chaperones that assist the folding process
of proteins in the crowded environment of the cell (reviewed in [43-45]). Their general
structure of a multi-subunit double-ring assembly with two central cavities allows them
to encapsulate protein substrates for ATP-driven protein folding (Figure 4).
Chaperonins are found in all organisms studied to date (with the exception of a few
Mycoplasmas), and they are essential for cell viability.
There are two main chaperonin groups (reviewed in [46, 47]): Group I (reviewed in
[48]) consist of eubacterial chaperonins (exemplified by E. coli GroEL) and
homologous proteins in mitochondria and chloroplasts, and Group II (reviewed in [49])
which the eukaryotic chaperonin CCT and chaperonins from archaea belong to. GroEL1
is composed of 14 identical subunits that form the 7-fold rotational symmetric doublering complex [50]. The composition for group II chaperonins is more complex. CCT is
composed of eight distinct subunits [51], whereas archaeal chaperonins consist of one,
two or three different subunits [52, 53].
STRUCTURE AND MECHANISM
All chaperonin subunits have a similar overall structure consisting of three domains:
apical, intermediate and equatorial [54, 55]. Within the double-ring assembly the
subunits are arranged so that the equatorial domains form the inter-ring and intersubunit contacts, and the apical domains, which contain the binding sites for the protein
substrates, are located at each end of the cylinder (Figure 4). The intermediate domains
connect the apical and equatorial domains. The major structural difference between the
two chaperonin groups is that complexes belonging to Group II have an extra helical
extension of the apical domain serving as a built-in lid that closes the central cavity
[56]. GroEL, on the other hand, use the co-protein GroES for that function (Figure 4).
Figure 4. Schematic of chaperonin structures. Side view of Group I (left) and Group II (right)
chaperonin in cross-section. The equatorial domains are represented by dark blue rectangles; the light
blue ovals are the intermediate domains and the apical domains are represented by green ellipses. A coprotein (orange) is utilized by Group I to seal the folding chamber, whereas in Group II a helical
protrusion (lighter green extension) serves as a built-in lid structure to close the cavity. Surfaces on the
apical domain for substrate (grey, left only) binding are marked by brown patches. (Taken and modified
from [57]).
GroEL is the most extensively studied Group I chaperonin, and a great deal is known
about its reaction cycle (reviewed in [47, 58]). Briefly, the mechanism starts with
1
For simplicity, GroEL will be used to represent Group I chaperonins.
6
binding of the unfolded protein substrate to the apical domains in one ring (the cis
ring). Subsequent ATP binding to the equatorial domains increases the affinity for the
co-protein GroES, which seals the folding chamber and induces the release of the
substrate into the cavity. The substrate is encapsulated in the cavity until ATP binds to
the opposite ring (the trans ring), which cannot occur until ATP in the cis ring has
hydrolyzed. Substrate and ATP binding to the trans ring triggers the release of GroES
and the folded substrate from the cis ring, and a new cycle of protein folding takes
place on the trans ring.
In contrast to the situation for GroEL, the functional cycle for Group II chaperonins
is not well understood. X-ray crystallography and cryo-EM studies have identified
several conformations of archaeal and eukaryotic chaperonins [55, 59-62] that show
resemblance to some functional states observed for GroEL [54, 63-65], but it is still
unclear how they are coupled to the ATPase cycle. One factor that contributes to limit
the amount of information in this matter is the hetero-oligomeric nature of Group II
chaperonins, which impose difficulties to understand the properties of the distinct
subunits and how these are related to the mechanism.
Pyrococcus furiosus is a hyperthermophilic archaea that grows optimally at
approximately 100°C. The P. furiosus genome contain only one chaperonin gene,
suggesting a homo-oligomeric chaperonin complex [52, 66]. Due to its minimal
oligomeric complexity, the P. furiosus chaperonin is an attractive model system for
studying the structure-function relationship of Group II chaperonins. The chaperonin
from the closely related organism Pyrococcus horikoshii has also been studied [67], but
structural characterization is still unavailable for any Pyrococcus chaperonins.
In paper II, structural characterization of the chaperonin from P. furiosus (Pf Cpn)
was carried out. Expression and purification of Pf Cpn was performed according to the
protocol outlined in paper II, and electron micrographs of frozen-hydrated Pf Cpn
samples revealed ring shaped structures that are characteristic for chaperonins. Because
the end-view was the strongly preferred orientation, not all projection images were
selected. A total of 674 projection images, selected from 22 micrographs, were used in
the 3D reconstruction process. The resulting cryo-EM map revealed a di-octameric
structure (Figure 5), consistent with the number of subunits per ring predicted by
Archibald et al. [52] for Pyrococcus chaperonin. Moreover, the double-ring assembly
had 8-fold rotational symmetry. Pyrococcus species contain only one chaperonin gene
per genome [52], with a high sequence identity between the chaperonin genes. Hence,
it is very likely that all chaperonins within Pyrococcus species exist as 8-fold
symmetrical complexes. Comparison of our density map with previously determined
functional states of Group II chaperonins suggest that the 16-mer assembly is in a
partially closed/open state [59, 60, 68].
Figure 5. 3D reconstruction of Pf Cpn.
Top view (left) and side view (right).
7
RIBOSOME – THE WORKBENCH OF TRANSLATION
When a protein is synthesized the genetic information stored in DNA must be
accurately transferred to a polypeptide chain. This flow of information passes through
an RNA molecule (Figure 6). First, the information encoded in DNA is transcribed into
messenger RNA (mRNA). Subsequently, mRNA is used as a template when the
genetic message is translated into the 20 different amino acids. The code words in
mRNA are deciphered by transfer RNAs (tRNAs), small adaptor molecules that
understand the two languages nucleic acids and amino acids. The entire translation
process takes place on the ribosome, a large molecular machine found in all cells.
During translation, the ribosome interacts with mRNA, tRNA and other molecules that
are necessary for the synthesis of the polypeptide.
Ribosomes and the other key components essential for translation have been
extensively studied over the years, which in turn have resulted in a large number of
publications. The purpose of this chapter is to give a brief background of protein
synthesis and the components involved; hence, only a fraction of the publications will
be covered. For a more detailed coverage the work of A. Liljas [69] is recommended. In
the next few paragraphs his disposition will be followed.
Figure 6. Transfer of information from DNA to RNA to protein.
The message – mRNA
The mRNAs are a class of RNA molecules that carries genetic information, copied
from DNA, to the ribosomes for translation into protein. Figure 7 shows the genetic
code, which is the set of rules that dictates how the genetic message is translated. The
code words are triplets of nucleotides called codons [70-72]. Since there are four
different nucleotides in mRNA (A, U, C, G), 64 (or 43) different triplet combinations
are possible. Out of these 64 codons, 61 specify individual amino acids. Thus, the
genetic code is degenerate, meaning that some amino acids are encoded by more than
one codon. Only methionine and tryptophan have one codon. On the other hand
leucine, serine and arginine are each specified by up to six codons (Figure 7). Codons
encoding one specific amino acid usually differ in their third base. For example, the
codons for alanine are GCU, GCC, GCA, and GCG. In addition to encoding individual
amino acids, some codons also signal the beginning and end of the translation process.
Generally the start (initiation) codon is AUG, which is the same as the methionine
codon. The three codons UAA, UGA, and UAG do not encode amino acids; instead
they are stop (termination) codons that signal the end of the polypeptide chain
synthesis. The sequence of codons in mRNA that is between the start and stop signal is
termed a reading frame.
8
U
C
A
G
U
C
A
G
Phe
Phe
Leu
Leu
Leu
Leu
Leu
Leu
Ile
Ile
Ile
Met
Val
Val
Val
Val
Ser
Ser
Ser
Ser
Pro
Pro
Pro
Pro
Thr
Thr
Thr
Thr
Ala
Ala
Ala
Ala
Tyr
Tyr
STOP
STOP
His
His
Gln
Gln
Asn
Asn
Lys
Lys
Asp
Asp
Glu
Glu
Cys
Cys
STOP
Trp
Arg
Arg
Arg
Arg
Ser
Ser
Arg
Arg
Gly
Gly
Gly
Gly
U
C
A
G
U
C
A
G
U
C
A
G
U
C
A
G
3rd base in codon
1st base in codon
2nd base in codon
Figure 7. The genetic code. The codons (three-base code words) are translated into the 20 amino acids,
which are given in their three letter codes. The three termination codons, as well as the initiation codon,
are in bold.
The adaptor molecule - tRNA
The decoding process where the nucleotide sequence of mRNA is converted into the
amino acid sequence of the protein is performed by tRNA molecules. Each amino acid
has its own tRNA it is attached to. When a codon in the mRNA calls for a particular
amino acid, the tRNA carrying that amino acid will recognize the codon and thus
deliver its amino acid to the growing polypeptide chain.
tRNAs consist of a single stranded RNA molecule, about 75 nucleotides long,
folded into a stem-loop arrangement that resembles a cloverleaf when drawn in two
dimensions (Figure 8A). The structure contains four stems that are stabilized by
Watson-Crick base pairing. Three of the stems have loops. The middle loop carries the
anticodon, the nucleotide triplet that base-pair to the corresponding codon in mRNA.
The other two loops, which contain several modified bases, are part of the D and T
arms. The fourth stem is composed of the 3’- and 5’-ends of the chain. All tRNAs has a
conserved CCA sequence at their 3’-end that is not base-paired. The free 2’-hydroxyl or
3’-hydroxyl of the adenosine is covalently attached to the proper amino acid, thereby
yielding an aminoacyl-tRNA (aa-tRNA). Thus, this unlooped stem is referred to as the
acceptor or aminoacyl stem.
Over 35 years ago the first 3D structures of tRNA were determined [73, 74]. The
structures revealed that the cloverleaf is folded into an L shape, with the anticodon loop
and acceptor stem forming the ends of the two arms while the D- and T-loops form the
elbow region (Figure 8B). In solution, the L shape is the basic structure for all tRNAs.
On the ribosome, however, the tRNA can adopt several conformations.
9
(A)
(B)
Figure 8. Structure of tRNA. (A) The cloverleaf conformation of yeast alanine tRNA (tRNAAla), the
first nucleic acid sequence of tRNA to be determined [75]. Modified nucleotides: D = dihydrouridine, I =
inosine, T = thymine, ȥ = pseudouridine and m = methyl group. The D arm nearly always contains a
dihydrouridine. Likewise, the T or TȥCG arm almost always contains thymidylate and pseudouridylate.
(B) The 3D structure of tRNA. The color coding is the same as in (A). The 3D structure of tRNA is
reproduced with the kind permission of N.R. Voss.
The anticodon of a tRNA read the codon of mRNA by forming Watson-Crick base
pairing. However, only the first and second bases of a codon form standard WatsonCrick base pairs with the third and second bases of the corresponding anticodon. The
third base in the codon, the so-called “wobble” position, can form nonstandard pairing
with the first base of the tRNA anticodon [76]. For this reason, a tRNA with a certain
anticodon can recognize several codons.
Attachment of a particular amino acid to a tRNA is performed by the enzymes
aminoacyl-tRNA synthetases (aaRS). Each of these 20 enzymes is specific for one
amino acid, and they use different means to identify the correct tRNA. In general, the
identity elements of the individual tRNAs are localized in the anticodon and acceptor
stems [77]. For glutamine tRNA (tRNAGln) the aaRS recognizes the correct anticodon
[78], while the single G·U base pair in the acceptor arm of alanine tRNA (tRNAAla) is
sufficient as an identity element (reviewed in [79]). Once a tRNA has been charged
with an amino acid, the cognate codon-anticodon interactions will guide the tRNA to
deliver its amino acid to the growing polypeptide chain. For this reason, it is crucial for
the fidelity of translation that the synthetases charge the tRNAs with their specific
amino acid [80].
The protein-synthesizing machine – The ribosome
A ribosome consists of two subunits with unequal size, both composed of ribosomal
RNA (rRNA) molecules and proteins. The size of the ribosome and its subunits, as well
as some of the steps in the translation process, varies between different species. Thus,
for simplicity only the bacterial ribosome will be covered.
10
In bacterial ribosomes (70S2) the two subunits are called the 30S (small) and the
50S (large) subunit. These ribosomal subunits have irregular shape, but together they
form a fairly globular complex with a cleft between them through which the mRNA
and tRNA molecules transits (Figure 9). Various conformational changes takes place in
the ribosome during translation (reviewed in [81]); where large-scale movements
undergone by the 30S subunit is mainly in the head region whereas in the 50S subunit it
is the two side protuberances that show flexibility. Rotation of the two subunits with
regard to each other is also observed.
Ribosomes contain about 65% rRNA and 35% protein, with the core of the
ribosomal subunits formed by the rRNA molecules. The 30S subunit has one large
rRNA molecule (16S RNA) while the 50S subunit has two rRNA molecules, one large
(23S RNA) and one small (5S RNA). In both subunits it is the large rRNA molecule
that provides the binding sites for the ribosomal proteins. Most of the ribosomal
proteins are located at the external surface (solvent side) of the subunits [82-85]. The
interface between the ribosomal subunits is mainly composed of rRNA. It is also here
the functional sites of the ribosome are found; the decoding site on the 30S subunit [83,
86-89] and the peptidyl transferase center (PTC) on the 50S subunit [84, 85, 90, 91].
tRNA molecules are able to concurrently contact the decoding site on the 30S subunit
and the PTC on the 50S subunit because of their L-shaped structure where the
anticodon is at one side and the CCA tail is at the opposite side (Figure 8B), about 75 Å
apart.
Figure 9. The 70S ribosome. L1: stalk of ribosomal protein L1; CP: Central protuberance; L7/L12:
stalk of L7/L12; h: head; sh: shoulder; b: beak; sp: spur. Arrow (right) indicates the inter-subunit space.
There are three classical binding sites for tRNA molecules on the ribosome. These are
the A site (for incoming aminoacyl-tRNA), the P site (for peptidyl-tRNA that carries
the growing polypeptide chain) and the E site (for deacylated tRNA before it
dissociates from the ribosome) [92-95]. The A site is located at the side of the L7/L12stalk whereas the E site is at the L1-side of the ribosome. A fourth ribosomal binding
site exists, which is where tRNA molecules are bound with EF-Tu prior to entry into
the A site [96, 97]. This is called the A/T site and it is here the selection between
cognate and non-cognate tRNA molecules takes place. tRNAs bound to this site are in a
intermediate or hybrid state called the A/T state [97]. During their transit through the
ribosome, tRNA molecules passes through two additional hybrid states, the A/P and the
P/E state [98-100].
2
Naming of the ribosome and its subunits, as well as the rRNAs, is based on their sedimentation rates.
11
THE TRANSLATION PROCESS
Protein synthesis is divided into three stages; initiation, elongation and termination.
Each step is catalyzed by specific translation factors. These factors, together with all the
other components necessary for translation, are listed in Table 1.
Table 1. Components required for the translation process in E. coli.
Stage
Necessary components
Initiation
mRNA
fMet-tRNAiMet (initiator tRNA)
Inititaiton/start codon (AUG) in mRNA
30S ribosomal subunit
50S ribosomal subunit
Initiation factors (IF1, IF2, IF3)
GTP
Elongation
Initiation complex (70S·mRNA· fMet-tRNAiMet in P site)
Aminoacyl-tRNAs
Elongation factors (EF-Tu, EF-G)
GTP
Termination
and recycling
Termination codon in mRNA
Release factors (RF1, RF2, RF3)
Recycling factor (RRF)
Initiation
Protein synthesis is initiated when an mRNA molecule binds to the free 30S subunit.
Correct initiation requires that the start codon (generally AUG) is positioned in the
ribosomal P site. In bacteria, the 30S subunit identifies the start codon by forming
interactions between its 16S RNA molecule and a nucleotide segment in mRNA that is
located upstream of the AUG start codon. This mRNA sequence, called the ShineDalgarno sequence [101], is complementary to the 3’-end of the 16S RNA. After the
correct placement of the start codon in the P site of the 30S subunit, the initiation
factors IF1 and IF3 bind to the A and E sites, respectively [102-104], and guides the
initiator tRNA (fMet-tRNAiMet) into the P site. Subsequently, binding of IF2·GTP,
release of IF3 and recruitment of the 50S subunit occurs. Lastly, GTP on IF2 is
hydrolyzed and together with IF1 this factor leaves the ribosome. The result is the
initiation complex composed of the 70S ribosome with fMet-tRNAiMet in the P site that
is ready for elongation.
Elongation
At the start of each elongation cycle, the 70S ribosome has a peptidyl-tRNA or the
initiator tRNA (fMet-tRNAiMet) bound in the P site. Elongation begins with aminoacyltRNA, elongation factor EF-Tu and GTP forming a ternary complex (aminoacyltRNA·EF-Tu·GTP) that binds to the A/T site on the ribosome. In the A/T site the
12
anticodon arm of the aminoacyl-tRNA molecule has a conformation that differs from
those seen in normal L-shaped structures (Figure 10) [97, 100, 105, 106]. This modified
tRNA structure allows codon-anticodon pairing in the decoding site on the 30S subunit
while the aminoacyl end is bound to EF-Tu far from the PTC on the 50S subunit [100].
If the codon does not match the anticodon, the ternary complex leaves the ribosome and
a new ternary complex is bound to the A/T site. If there is cognate codon recognition,
EF-Tu will hydrolyze its bound GTP and undergo a conformational change. The EFTu·GDP complex has a low affinity for the aminoacyl-tRNA and the ribosome, thus it
dissociates [107]. This will free the aminoacyl end of tRNA and allow it to swing into
the PTC on the 50S subunit, which results in the tRNA molecule regaining its normal
L-shaped structure. During this motion the position of the anticodon stem-loop (ASL)
is maintained, i.e. the ASL in the A-site tRNA and A/T-site tRNA has a similar
orientation (Figure 10) [100]. With a peptidyl-tRNA in P site and an aminoacyl-tRNA
in A site, the peptide can be transferred. This leads to the pre-translocation ribosome
complex with a peptidyl-tRNA in A site and a deacylated tRNA in P site.
Figure 10. A- and A/T-site tRNA. The A/T-site
tRNA (blue) has a bent anticodon arm, compared to
the A-site tRNA (red), due to a kink between the
anticodon stem and the D stem. PDB id: 1QZA for
A/T-site tRNA and 1QZB for A-site tRNA.
The final step of elongation is translocation. The end result of this procedure is the
post-translocation ribosome complex where the next codon of mRNA is exposed in the
A site and the peptidyl-tRNA in A site and the deacylated tRNA in P site has moved to
the P and E site, respectively. This reaction is catalyzed by the elongation factor G (EFG). During the course of translocation the two ribosomal subunits rotate with regard to
each other in a ratchet-like motion [108, 109]. In addition, the peptidyl-tRNA and the
deacylated tRNA passes through the hybrid A/P3 and P/E states, respectively [98]. The
molecular mechanism for translocation is not well understood. However, current
findings suggest the following model for the EF-G dependent translocation process
(Figure 11) (reviewed in [110]). After peptidyl transfer, EF-G·GTP binds to the pretranslocation ribosome complex (a), with peptidyl-tRNA in A site and deacylated
tRNA in P site, and hydrolyzes its GTP (b). Next, the two ribosomal subunits rotate
with regard to each other (ratchet-like motion) and the P-site tRNA enters the hybrid
P/E state (c). Subsequently, A-site tRNA shift to the A/P state (d). Then, structural rearrangement of the ribosome allows the two tRNA molecules to enter their classical P
and E sites (e). Finally, EF-G·GDP dissociates and the ribosome undergoes the second
step of the ratchet-like movement and returns to its normal conformation where it is
ready for a new cycle of elongation (f).
3
The first and second letter in A/P and P/E denotes the position of the ASL and the acceptor end,
respectively. For instance, a tRNA in the A/P state means that its ASL is still bound in the 30S A site
while its acceptor end has moved to the 50S P site.
13
Selection
Accommodation
Peptidyl transfer
50S
30S
P
A/T
P
P
A
A
EF-Tu·GDP
(a)
EF-G·GTP
EF-Tu·GTP
P
E
P
EF-G·GDP
A
(b)
(f)
(e)
E
P
(d)
P/E A/P
(c)
P/E
A
P
A
Figure 11. Summary of the elongation cycle. (Taken and modified from [69])
Termination
Protein synthesis is terminated when one of the three stop codons UAA, UAG and
UGA is exposed in the A site. In bacteria there are three release factors (RF1, RF2 and
RF3) that work in concert to terminate the translation process. RF1 and RF2 recognize
the stop codons UAG/UAA and UGA/UAA, respectively [111], and hydrolyze the
polypeptide chain from the peptidyl-tRNA. RF3 is a GTPase, and its function is to
remove RF1 and RF2 from the ribosome [112]. RF3 binds to the ribosome in complex
with GDP [113]. Provided that RF1 or RF2 is bound to the A site, and the P site is
occupied by a deacylated tRNA, RF3 exchange its GDP to GTP and subsequently
releases RF1 or RF2 from the ribosome. Lastly, RF3 hydrolyze its GTP to GDP and
dissociates from the ribosome.
After the termination step and the peptide have been released, the ribosome has to be
prepared for a new round of translation. This task is performed by the ribosomerecycling factor (RRF) which, together with EF-G, promotes the separation of the two
subunits and dissociation of the mRNA and the deacylated tRNA.
TRANS-TRANSLATION
If the ribosome reach the 3’-end of the mRNA molecule without encountering an inframe stop codon, it leads to two undesirable consequences. First, it causes a significant
14
loss of translational efficiency since the ribosome stalled on the defective mRNA
molecule will be unavailable for new rounds of translation. Secondly, the incomplete
polypeptide produced may be toxic to the cell. In bacteria, both these problems are
solved by the translation control system termed trans-translation (reviewed in [114116]). The main components in trans-translation are an RNA molecule called transfermessenger RNA (tmRNA) and its helper protein small protein B (SmpB). The
SmpB·tmRNA system recognizes and rescues stalled ribosomes and simultaneously
adds a C-terminal proteolysis tag to the incomplete polypeptide chain. Furthermore, the
system also promotes degradation of the defective mRNA molecule, thus preventing
any future ribosome stalling [117, 118].
tmRNA and SmpB: structure and function
As the name indicates, tmRNA is an RNA molecule that functions both as messenger
and transfer RNA [119-122]. A diagram over the secondary structure of tmRNA is
depicted in Figure 12. The 5’- and 3’-ends of the RNA molecule fold into the tRNAlike domain (TLD), which contains an acceptor arm, a T-arm and a D-loop. tmRNA
contains a G·U wobble base pair in its acceptor arm, the identity element recognized by
Alanyl-tRNA synthetase (Ala-RS). Hence, it is charged with Alanine by Ala-RS [121,
123, 124]. Instead of an anticodon arm, a long stem links the TLD to the other two
domains in the tmRNA molecule, the mRNA-like domain (MLD) and the pseudoknotrich domain consisting of four RNA pseudoknots (designated PK1 to PK4). The MLD
contains the open reading frame (ORF) that encodes the proteolysis peptide tag
(ANDENYALAA in E. coli).
MLD
Figure 12. Secondary structure diagram of tmRNA from Thermus thermophilus. The main three
domains are the tRNA-like domain (TLD, boxed out), the mRNA-like domain (MLD) and the four
pseudoknots (PK). Reprinted (and modified) with permission from [125]. Copyright © 2006 National
Academy of Sciences, U.S.A.
15
The four pseudoknots are the largest features of tmRNA, encompassing about twothirds of the tmRNA molecule (Figure 12). While the importance of the TLD and MLD
for the tmRNA activity has been firmly established, the specific function of the
pseudoknots is still unclear. PK2 to PK4 have been suggested to be important for the
proper folding of tmRNA [126], while PK1 may have a functional role for the tmRNA
activity [123, 127, 128]. PK1 is only 11 nucleotides upstream of the resume codon, and
although the reading frame selection is determined by the nucleotides immediately
upstream of the first codon [129, 130], this pseudoknot could be involved in positioning
the tmRNA ORF into the A site.
The most significant protein partner of tmRNA is SmpB. This protein binds tmRNA
with high affinity and specificity [131, 132], and it is essential for all known tmRNA
functions. As previously described, during normal translation aminoacyl-tRNA binds
the ribosome in a ternary complex with EF-Tu·GTP, and cognate codon-anticodon
interactions trigger GTP hydrolysis on EF-Tu followed by the release of EF-Tu·GDP
and accommodation of the tRNA acceptor stem into the PTC on the 50S subunit (see
‘elongation’ section). Likewise, Ala-tmRNA too is delivered to the ribosome in
complex with EF-Tu and GTP. However, SmpB is also required since tmRNA·EFTu·GTP alone does not associate stably with stalled ribosomes [131, 133]. Since
tmRNA lacks an ASL (Figure 12), accommodation of Ala-tmRNA into the ribosomal
A site do not require codon-anticodon interactions. It is not clear how the
SmpB·tmRNA mediated mechanism can bypass the decoding process and enter the
stalled ribosome. However, numerous studies suggest that SmpB can substitute for the
missing ASL, and that it does so with its highly conserved C-terminal tail [131, 134138]. Sundermeier et al. [134] showed that SmpB proteins with a mutated or truncated
C-terminal tail cannot support addition of the proteolysis tag, despite being able to bind
tmRNA and promote association of the SmpB·tmRNA complex with stalled ribosomes.
This indicates that the C-terminal tail of SmpB is required for a tmRNA function
following the recruitment of the SmpB·tmRNA complex to the ribosome, but prior to
trans-peptidation where the tmRNA-linked alanine is added to the incomplete
polypeptide. Consistent with these findings, chemical probing experiments revealed
that SmpB interact with the three key conserved nucleotides A1492, A1493 and G530
of the 16S RNA that form the 30S subunit decoding center, and that G530 in particular
interact with the C-terminal end of SmpB [137]. Further support comes from X-ray
crystallography studies of SmpB in complex with the TLD of tmRNA [135, 136].
SmpB binds to the elbow region of TLD (Figure 13) and it has been suggested that the
protein substitutes for the anticodon arm and the D stem that is present in all native
tRNAs but missing in the TLD of tmRNA. Furthermore, biochemical experiments
demonstrated the necessity of SmpB on tmRNA for the ribosome-dependent GTPase
activation of EF-Tu in the Ala-tmRNA·SmpB·EF-Tu·GTP quaternary complex [138].
Altogether, these and other related studies indicate that SmpB functions as an ASL
mimic during the initial stages of trans-translation.
Figure 13. The TLD·SmpB
complex versus tRNA. Left,
TLD is in red and SmpB is
in blue. Right, tRNA. The
relative orientation of the
TLD·SmpB complex and the
tRNA is the same.
PDB codes: 2CZJ for the
TLD·SmpB complex, 1GIX
for tRNA.
16
The mechanism of trans-translation
The SmpB·tmRNA mediated mechanism is depicted in Figure 14, and the process can
be described as follows [114, 139]. tmRNA forms a complex with SmpB that is
alanylated by Ala-RS. Subsequently, the Ala-tmRNA·SmpB complex is recognized by
EF-Tu·GTP (box 1). The Ala-tmRNA·SmpB·EF-Tu·GTP quaternary complex bind
ribosomes stalled at the 3’-end of an mRNA. The lack of an anticodon stem in tmRNA
is compensated for by SmpB, which presumably interact with the ribosome near the
decoding center and thus triggers GTP hydrolysis on EF-Tu.
Figure 14. Schematics of the trans-translation process.
Reprinted, with permission, from the Annual Review of Microbiology, Volume 62 Copyright © 2008 by
Annual Reviews. www.annualreviws.org
After GTP hydrolysis, the TLD of Ala-tmRNA accommodates into the ribosomal A
site and the incomplete polypeptide is transferred to Ala-tmRNA (trans-peptidation).
17
Next, the TLD moves to P site and the ribosome switch template from the defective
mRNA to the ORF of tmRNA. Unlike normal translation, the resume codon is not
specified by initiation factors. Instead, the reading frame of the tmRNA-encoded tag
sequence is determined by bases upstream of the resume codon. After removal from the
ribosome, the defective mRNA molecule is degraded by RNase R (box 2). Translation
of the tmRNA ORF continues until a stop codon at the end of the reading frame is
encountered. This promotes translation termination, ribosome recycling and
polypeptide release. The released nascent polypeptide is recognized by several cellular
proteases, due to the 11 amino acid degradation tag at its C-terminus, and is thus
degraded (box 3). The end result is a rescued ribosome that is now free to engage in a
new round of translation, while the defective mRNA and the incomplete polypeptide
has been eliminated and will not be to any harm for the cell.
Physiological significance of trans-translation
To date, all sequenced bacterial genomes contain the genes encoding SmpB and
tmRNA (smpB and ssrA, respectively) [114, 140, 141]. For most bacterial species,
where the exceptions are some pathogenic bacteria (such as Neisseria gonorrheae,
Mycoplasma genitalium and Mycoplasma pneumoniae [142, 143]), the tmRNA·SmpB
system is non-essential under ideal growth conditions. However, tmRNA activity is
frequently important when cells respond to stress, differentiate or during pathogenesis
[144]. For instance, E. coli strains lacking ssrA and smpB exhibit slower growth at
high temperature [145], and they also show increased sensitivity towards amino acid
starvation [146] and to antibiotics that promote translational frameshifting, readthrough and stalling [147]. Similarly, Bacillus subtilis deleted of ssrA and smpB show
reduced growth rate at both high and low temperature and an increased level of
tmRNA and SmpB during temperature stress [148, 149]. Furthermore, an eightfold
increase of tmRNA levels has also been observed for Thermotoga maritime during
antibiotic challenge [150].
Virulence in some pathogenic bacteria is tmRNA·SmpB-dependent. Yersinia
pseudotuberculosis and Salmonella enteric are two pathogens that exhibit defects in
virulence when they lack tmRNA activity [151-153]. Studies have shown that Y.
pseudotuberculosis cells lacking ssrA or smpB cannot cause lethal disease in a mouse
infection model [151]. The mutant strains where also unable to proliferate in
macrophages. Part of this virulence deficiency has been ascribed to defects in
expression and delivery of the virulence effector proteins known as Yops. Production
and secretion of Yops is regulated by the transcription factor VirF, and absence of
tmRNA activity leads to misregulation of VirF and a delayed Yop secretion. In
addition, Y. Pseudotuberculosis cells lacking the tmRNA·SmpB system are
nonmotile, have increased sensitivity to antibiotics, and they are also temperature
sensitive [151]. Similar to Y. Pseudotuberculosis, S. enteric also require tmRNA
activity to proliferate in macrophages, and for virulence in mice [152, 153].
Altogether, these findings emphasize the importance of the SmpB-tmRNA system for
survival under adverse conditions and in virulence of some pathogenic bacteria.
Structural studies of tmRNA·ribosome complexes
To date, the structural information available of tmRNA-bound ribosome complexes has
come from cryo-EM studies, and the complex most extensively studied has been with
tmRNA in the pre-accommodated state [125, 154]. This tmRNA·ribosome complex can
be obtained with the antibiotic kirromycin (kir). Kirromycin permits GTP hydrolysis on
EF-Tu but prevents EF-Tu·GDP from leaving the ribosome, thus allowing the
18
tmRNA·ribosome complex to be visualized in a state after GTP hydrolysis but prior to
accommodation of tmRNA into the A site. The initial cryo-EM structure of the preaccommodation tmRNA·ribosome complex revealed three major findings [154]; First,
the TLD of tmRNA was found to bind EF-Tu in a manner comparable to an aminoacyltRNA. Second, the remaining density of tmRNA was highly structured and formed a
large spiral that enclosed the beak of the 30S subunit in such a way that the ORF of the
MLD was positioned close to the mRNA entrance channel. Third, one SmpB was
found to bridge the TLD and the GTPase-associated center (GAC) on the 50S subunit.
The latter was corrected for in a more recent cryo-EM study when the presence of two
SmpB molecules in the pre-accommodation tmRNA·ribosome complex was observed
[125]. In this density map one SmpB (SmpB-1) was found to bind near the site where it
was previously observed [154], and the other (SmpB-2) was located close to the
decoding site on the 30S ribosomal subunit. Both SmpB molecules interact with the
TLD, where the complex formed between SmpB-2 and the TLD is similar to the
TLD·SmpB complex observed by X-ray crystallography [135, 136].
One of the main questions regarding tmRNA is the fate of its structure during the
course of trans-translation. As previously mentioned, the tmRNA molecule is highly
structured when it enters the stalled ribosome [125, 154]. Thus, the question that arises
is whether it stays structured or whether it unfolds. Wower et al. [155] recently
proposed that PK2 to PK4 of tmRNA unwinds as trans-translation progresses.
However, other studies suggest that the pseudoknots of tmRNA, PK3 in particular,
remain structured and that they stay outside the ribosome [156, 157].
In order to increase the structural information about trans-translation, we decided to
study tmRNA·ribosome complexes captured at stages subsequent to the preaccommodated state. The first is the post-accommodation complex (paper III) where
EF-Tu·GDP has dissociated and tmRNA has accommodated into the ribosomal A site.
The cryo-EM map shows the 70S ribosome with a density in the A site that is
connected to the remaining tmRNA density (Figure 15A). Fitting of the recently
published crystal structure of the T. thermophilus TLD·SmpB complex [135] into the
A-site density revealed the presence of one SmpB molecule (corresponding to SmpB-2
in the pre-accommodated state [125]) in the density map. This is in agreement with
previous biochemical studies [157, 158] where it was suggested that the SmpB
molecule bound to the 50S subunit (SmpB-1 in [125]) dissociates upon accommodation
of tmRNA into the A site. The fitting also revealed that the TLD·SmpB part of the
tmRNA·SmpB complex is positioned in the A site similarly to a canonical A-site tRNA
(Figure 15B), in line with the suggestion that SmpB mimics the anticodon arm and the
D-stem of native tRNAs [135-137]. Accordingly, during accommodation the SmpB
molecule moves deeper into the A-site pocket on the 30S subunit while the acceptor
stem of the TLD of the tmRNA·SmpB complex rotates away from its EF-Tu-bound
position and moves into the A site of the 50S subunit (Figure 15C). The distance the
acceptor arm of the TLD moves during accommodation is about 50 Å, which is very
similar to the distance of 46 Å that the acceptor arm of a canonical aminoacyl-tRNA
move during accommodation [159]. Altogether, our findings support the model in
which the TLD·SmpB complex is suggested to functionally mimic tRNA during the
first steps of trans-translation [135-137, 160, 161].
19
Figure 15. Reconstruction of the post-accommodation tmRNA·SmpB·ribosome complex. (A)
tmRNA·SmpB·70S complex in the post-accommodated state. (B) Left, TLD·SmpB complex fitted into
the A-site density. Right, the relative positions of the A-site tRNA and the A-site density. (C) Movement
of the TLD·SmpB complex during accommodation. The ribosomal orientation for (B) and (C) is
indicated by the corresponding thumbnails. tmRNA·SmpB is colored red, 30S and 50S ribosomal
subunits is in yellow and blue, respectively. CP, central protuberance; L1, stalk of protein L1; L7/L12,
stalk of proteins L7/L12; b, beak; sh, shoulder; sp, spur; dc, decoding center; post, post-accommodated
state; pre, pre-accommodated state [125]. PDB codes: 2CZJ for TLD·SmpB, 1GIX for tRNA.
Similarly to previously reported cryo-EM maps of tmRNA·ribosome complexes [125,
154, 162], the tmRNA in our density map was highly structured and formed an arc
around the beak of the 30S ribosomal subunit (Figure 15A). Comparison with the
tmRNA structure in the pre-accommodation ribosomal complex [125] shows that a
conformational change takes place in the arc when tmRNA goes from the preaccommodated to the post-accommodated state. The most significant change is the
downward flip undergone by PK1, which result in the MLD of tmRNA being
positioned closer to the A-site entry. It is likely that the re-arrangement of the tmRNA
structure may be caused by, or coordinated with, the rotation of the TLD·SmpB
complex in order to facilitate the entry of the MLD into the decoding center of the 30S
subunit.
These findings are in contradiction to the previously reported cryo-EM study of the
post-accommodation complex [125] where no movement in TLD or SmpB was
observed during tmRNA accommodation, and the arc density corresponding to PK2PK4 was absent in the density map. We ascribe these contradictory findings to low
occupancy of tmRNA in their post-accommodation complex [125].
In paper IV we studied the tmRNA·ribosome complex after two rounds of elongation,
where the P and E site should be occupied by a peptidyl-tRNAAla and the deacylated
tmRNA, respectively. The six reconstructions generated from the heterogeneity
analysis showed strong density in all three ribosomal sites, consistent with the double
translocation complex containing a deacylated tRNAAsp in the A site. Density for the
20
tmRNA arc composed of the pseudoknots was also observed in all cryo-EM maps, and
it was in a position somewhat similar to the arc in the post-accommodation ribosomal
complex. However, the arc exists in two states, one stable and highly structured and
another much more flexible and disorganized (Figure 16). The absence of density in the
fragmented arc structures are mostly in the hinge regions between the individual
pseudoknots, indicative of high flexibility in these parts of the structure that causes
them to become averaged out in our reconstructions. Thus, our findings suggest that the
pseudoknots in the double-translocation tmRNA·ribosome complex are still located
outside the ribosome, and that the individual tmRNA elements are highly dynamic
around the hinge regions.
Figure 16. Cryo-EM maps of double-translocation tmRNA-bound ribosome complexes. The
reconstructions are displayed with the arc density in red. The top left side of each class shows the atomic
coordinates of tmRNA in the post-accommodated state fitted into the isolated arc density for the
corresponding cryo-EM maps. The resolution and the number of particles contributing to each class are
indicated below each class. Cyan, PK1; black, linker between PK1 and ORF; magenta, ORF; yellow,
helix 5; purple, PK2, orange, PK3; gray, PK4. PDB code: 2OB7 for tmRNA coordinates.
21
SINGLE-PARTICLE CRYO-ELECTRON MICROSCOPY
Transmission electron microscopy (TEM) is an extremely flexible imaging technique
for a scientist. It can be utilized for 3D structural studies of protein molecules or
macromolecular complexes, as well as for studying the organization of organelles
within cells. Cryo-electron microscopy (cryo-EM) in combination with single-particle
reconstruction is a widely used technique to study macromolecular complexes, as these
assemblies are often either too large or too flexible for crystallographic methods [163].
The single-particle method is based on recording a large number of TEM images
(projections) of the complex, determine the angular orientation of each image, and
subsequently combine the images to recover the 3D structure of the complex. The main
advantage with this technique over crystallographic methods is that the complexes are
in a non-crystalline close-to-native state; hence, no steric constraints are present that
will limit the functional states they can assume. Single-particle cryo-EM has enabled
3D structures of macromolecular complexes to be obtained at subnanometer resolution
(reviewed in [164, 165]). Recently, near-atomic resolution has also been achieved
([166-169], reviewed in [170]).
SPECIMEN PREPARATION
Specimens observed in the EM are subjected to a high vacuum [171]; therefore they
must be prepared in such a way that the structure of the initially hydrated molecules is
preserved when the specimen is positioned in the vacuum. In addition, they must also
be protected from the radiation damage caused by the electron beam. Two of the most
commonly used specimen preparation methods will be covered in this section, namely
negative staining and vitrification, with emphasize on the latter technique.
Negative Staining
The negative staining method developed by Brenner and Horne [172] is based on
utilizing a heavy metal salt (e.g., uranyl acetate) to reduce the radiation damage and
protect the molecules from collapsing, while at the same time it also leads to an
increase in the contrast. However, since it is the heavy metal atoms distributed around
the molecules that are visualized, the structural information is limited to the shape of
the molecule. Furthermore, the structure of the molecules is only preserved to some
extent, deformation of the molecules occurs as a result of air drying [173]. Therefore,
negative stained specimens are limited in resolution [174, 175]. Nevertheless, negative
staining is technically easy to perform and is often used to evaluate whether a molecule
is suitable or not for high-resolution EM studies.
Vitrification
The technique of vitrifying samples for cryo-EM studies was developed more than 30
years ago (reviewed in [176], [177, 178]), and it revolutionized the field of biological
EM as it became possible to observe macromolecules under close-to-native conditions.
The trick with this technique is the amorphous ice (or vitreous ice) the sample is
embedded in, which enables the molecules to be fully hydrated despite the requirement
for vacuum conditions inside the EM. Thus, the molecule is preserved in a biologically
relevant conformation. Amorphous ice is produced when water is cooled extremely fast
22
(cooling rate ~ 105 K/s) [171], thus preventing formation of ice crystals that could
damage the specimen. This is done by plunging the grid into a cryogen, often liquid
ethane cooled by liquid nitrogen. The preparation steps are as follows: the sample is
applied to a grid. The grid is then blotted with filter paper to remove excess liquid,
leaving only a thin layer of the sample. Immediately after blotting, the grid is plunged
into liquid ethane.
The advantage with this technique is the reduction of radiation damage at low
temperatures [179] and that specimen collapse is avoided. Another advantage is that the
contrast observed is related to the biological molecule itself rather than some
contrasting agent, such as the heavy metal atoms in negative staining. However, the
latter case is also what leads to the low contrast in EM images of vitrified specimens,
which causes difficulties for the subsequent image processing procedures. In addition,
the electron dose needs to be limited in order to reduce the radiation damage; the
projection images produced are therefore very noisy with a low signal-to-noise ratio
(SNR), thus further imposing problems when the images are analyzed. Despite these
drawbacks, cryo-EM has still enabled structures of macromolecular complexes to be
determined with high accuracy. A significant factor contributing to this success is the
progress that has been made in developing more powerful software platforms, as well
as the increased computer power, to deal with the noisy EM images.
During imaging, the electron beam can eject electrons (called ‘secondary electrons’)
from the specimen. This leads to a buildup of positive charge that can distort the
recorded image through an effect termed charging. Charging can be reduced by the
carbon film that coats the EM grids, an effect that increases if the carbon support grids
have been pre-irradiated under the electron beam prior to specimen preparation [180].
Pre-irradiated carbon support grids are used mostly for studies of 2D crystals. However,
due to severe charging problems with the ribosome specimens (paper III and IV), these
grids were tested. The result was a significant reduction of the charging effect.
The pre-irradiated grids used were ‘holey carbon grids’ coated with an extra
(continuous) carbon film. Holey carbon grids are basically what the name implies, i.e.
grids coated with holey carbon. When the specimen is vitrified, in cases where regular
holey carbon grids are used, the macromolecules are dispersed in the holes away from
the carbon film. This is to avoid any orientation preference that can occur when the
molecules adsorb to the carbon film. However, it has been shown that the carbon
actually has an opposite effect on the ribosome [163]. Hence, not only did the extra
carbon film applied to the holey carbon grids reduce the charging effect, but it also
generated a more random orientation distribution of the ribosome complexes.
DATA PROCESSING
Once a set of electron micrographs has been collected, the individual projection images
of the molecules needs to be selected for the subsequent image processing procedure.
This can be done manually or automatically. The automatic particle selection methods
available are usually based on locating the particles by cross-correlating rotationally
averaged reference images with the entire micrograph. The user manually selects
several particles, which are then rotationally averaged and used as templates. A
combination of the two methods was used for the hemocyanin and the ribosome
projects, where boxer from the EMAN package [181] was first used to select all
23
particles in the right size range as the molecule, and then the gallery of particle
candidates was visually checked. Due to the strong preference of the chaperonin
complex to be in the end-orientation, particles for the chaperonin dataset were selected
manually. First all side-views were selected, and subsequently a reasonable amount of
images representing the end-orientation of the molecule was added to the dataset.
The next step is to correct the contrast transfer function (CTF) of the projection images
[163]. The rationale for this correction is that a number of factors (predominantly the
amount of defocus and the spherical aberration constant of the lens system (Cs))
contribute to affect the acquired images. The effect of the CTF is best described in
Fourier space, where it will act as a multiplicative function. It has the shape of a sine
wave with the period compressed at high resolution. The CTF for a certain micrograph,
and accordingly also for all the particles in the micrograph, can be obtained by
calculating Fourier transforms of images selected from this micrograph that contain
carbon film. By fitting the theoretical expression of CTF to the CTF of the carbon film,
defocus and CTF parameters for each micrograph can be determined. The Cs value is
taken to be a known constant for a specific microscope. With these values known,
correction of the sign changes (and possibly damping) caused by the CTF can be
performed on the individual projection images that originate from a certain micrograph.
CTFit from the EMAN package [181] was used for defocus determination, as well as
the CTF phase correction of the datasets from all projects. CTF amplitude correction
was only carried out on the hemocyanin (paper I) and the chaperonin (paper II)
datasets, and it was performed as part of the reconstruction procedure (see below).
Once the projection images have been selected and CTF correction (in our case, only
the phases) has been performed, the angular orientation of each projection needs to be
determined (angular assignment) so the dataset can be combined to recover the 3D
structure of the molecule (3D reconstruction). There are several software packages
available that provide the tools to process the projection images, and they all use
different approaches. For instance, SPIDER [182], FREALIGN [183] and STRUL
[184] define the orientation angles by aligning the experimental projections against 3D
reference volumes. For cases where no initial 3D volumes are available, 2D class
averages are generated from the individual projections and taken to represent SNR
enhanced projections. These can then be assigned Euler angles by maximizing the
common lines correlation coefficient, a technique employed by IMAGIC [185] and
RAD [186].
For the hemocyanin and chaperonin projects, the EMAN package [181] was used.
EMAN use projection matching to determine the orientation angles of the projection
images. However, instead of combining the individual projections to recover the 3D
volumes, ‘refined’ class averages are used. The reconstruction procedure utilized by
EMAN (figure 17) is as follows: First, the individual particles are selected from the
electron micrographs. Subsequently, the CTF parameters are determined and the phases
are corrected. 2D reference projections (so called re-projections) are then generated
from the preliminary 3D volume and subsequently used to classify the projection
images. Once the projections have been grouped into individual classes, ‘refined’ class
averages representing the different projection views are generated. In parallel, CTF
amplitude correction is also carried out. Re-assignment of Euler angles for the class
24
averages are then performed with projection matching. Finally, a new 3D model for the
next round of refinement is generated from the class averages.
EMAN [181] was also used to generate the initial models of hemocyanin and
chaperonin, which utilize a technique based on the rotational symmetry of the complex.
First, a search for projections with the best n-fold rotational symmetry (‘top’ views),
and projections with the best mirror or pseudo-mirror symmetry and the poorest n-fold
rotational symmetry (‘side’ views) is performed. Class averages of the top and side
views of the complex are then generated and subsequently used to construct a
preliminary 3D model. D5 and D8 symmetry was applied for the hemocyanin and
chaperonin complexes, respectively.
Refinement Loop
Particle Selection
Select
Particles
Determine CTF
Parameters and
Correct Phases
Generate
Preliminary
3D Model
Classify
by Projection
Matching
Align and
Average Particles
in Each Class
CTF Amplitude
Correction
Re-assign Euler
Angles to Class
Averages by
Projection Matching
Projections
of Model
Build 3D
Model
Initial Model
Final
Model
Figure 17. Schematics of the reconstruction process in EMAN. (Taken and modified from [181])
HETEROGENEITY ANALYSIS
In single-particle reconstruction, the underlying assumption is that the projection
images represent randomly oriented views of the same structure. If the sample contains
a mixture of conformational states, e.g. different conformers or different ligand binding
states, the dataset collected will be structurally heterogeneous. The 3D volume
recovered from such a dataset does not reflect any of the co-existing structures.
Consequently, the only way to generate an interpretable structure is to divide the
heterogeneous dataset into homogeneous subsets, which can then be used to recover the
individual 3D volumes. There are two approaches that can, in principle, perform this
task. These are known as supervised and unsupervised classification. The former
approach is a reference based classification method, where the heterogeneous dataset is
divided according to resemblance between the projection images and certain 3D
template volumes that represent different conformational states of the molecule. These
3D templates can be obtained from other experiments [187] or created artificially [188].
25
Two main unsupervised 3D classification methods have been described. The first one,
introduced by Penczek et al. [189], is based on focusing the classification to regions
that show high variance. These high variance regions are identified and localized using
3D variance maps. The procedure for focused classification begins with the calculation
of a 3D variance map [190]. A 3D spherical mask is then placed in the site
corresponding to the high variance region, and subsequently projected into a set of 2D
masks. Next, the experimental projections are sorted into angular sets according to the
closest projection direction as the 2D masks. Classification of each set is then
performed based on the regions outlined by the respective masks. In the case the
structures differs with regard to ligand or no ligand bound, high pixel values indicate
the presence of ligand and low pixel values indicate the absence of ligand. Two groups
will therefore be generated from the classification, one with classes containing images
with high pixel values inside the masked regions and another with low pixel values
inside the masked regions. These two groups are used to reconstruct two 3D volumes
that are subsequently used in a multi-reference 3D projection alignment procedure (i.e.
supervised classification) against all projection images in order to refine the angles.
The unsupervised 3D classification method used for the ribosome project (paper III and
IV) is based on the maximum-likelihood (ML) principle. The underlying idea in the
ML method is to find the statistical model that maximizes the probability to observe the
experimental data. Mathematically this is achieved by optimizing the following loglikelihood function [191]
L (4)
I
K
¦ ln ¦ ³ P( X
i 1
i
| k , M , 4 ) P ( k , M | 4 ) dM
k 1M
where the model Ĭ consists of K number of 3D structures and some parameters that
describe the probability to observe different projections through these 3D structures.
The first factor P ( X i | k , M , 4) represents the probability of observing image X i when
viewing class k in orientation M . The second factor P (k , M | 4) is the probability of
observing class k in projection direction M . The optimization of the log-likelihood
function is performed iteratively by introducing a weighting factor, which describes the
probability of viewing class k in projection direction M when presented to image X i .
This weighting factor, which is updated for each image in every iteration, determines
the contribution each projection image makes to every class and projection direction.
The XMIPP software package [192, 193] was used for the ML classification, and the
procedure is as follows: Projections are generated from the K different reference
structures. Subsequently, all the experimental images are compared with these
reference projections. Based on the resemblance, probabilities are calculated for each
image in all the different projection directions. These probabilities are then used to
generate new probability-weighted 2D averages. Finally the 2D averages are combined
into K different 3D reconstruction, which will be used in the next iteration. In addition,
the standard deviation for e.g. P( X i | k , M , 4) , as well as the probabilities for the
different projection directions, are also updated for the next iteration.
To generate the initial reference maps, a single iteration of the ML classification for
K randomly split subsets of the experimental data is performed with a low-resolution
volume that show the general features of the average structure as a starting model. For
26
paper III and IV, a ribosome structure low-pass filtered to 80 Å was used as the starting
model [191].
Figure 18 shows the first two ML classifications performed on the ribosome dataset
from paper III. In the first ML3D run [194], the 50S subunit particles and the intact 70S
ribosome particles were separated. Subsequently, a second classification was performed
using only the intact 70S ribosome particles. Likewise, in paper IV the 50S and 70S
particles were also separated before continuing with the intact 70S ribosome particles,
but in this case the first classification was carried out in Fourier space (MLF3D) [195].
The classes from paper III were refined separately by projection matching. Due to the
limited number of projections in the individual classes, in combination with poor SNR
of the projection images, no refinement was performed in paper IV.
27
Figure 18. Flow-chart showing the first two ML classifications of the tmRNA·SmpB·70S dataset
from paper III. The number of particles in each class is indicated below the corresponding
reconstructions.
28
CONCLUSIONS
Paper I – A comparison of the two R. thomasiana hemocyanin isoforms revealed that
up to a resolution of 19 Å, the two assemblies show no structural differences. Three
previously unreported connections were identified in both complexes, one within the
subunit dimer and two between adjacent subunit dimers, which could contain binding
sites for the ions Ca2+ and Mg2+.
My contributions: Design of research (together with PJBK), specimen preparation for
electron microscopy, data collection (together with TT), analysis of results (together
with PJBK and HE), writing of paper.
Paper II – Electron micrographs of vitrified P. furiosus chaperonin samples revealed
ring-shaped particles, and the density map obtained from subsequent image processing
suggest that the 16-mer assembly with an 8-fold symmetry is in a partially closed/open
state.
My contributions: Design of research (together with PJBK), protein purification
(together with MK), specimen preparation for electron microscopy (together with PP),
data collection (together with PP), analysis of results (together with PJBK and MK),
writing of paper (together with MK and PJBK).
Paper III – The structure of the tmRNA·SmpB-bound ribosome complex in the postaccommodated state showed that the TLD·SmpB part of the tmRNA·SmpB complex
mimics native tRNAs in the A site of stalled ribosomes. In addition, the tmRNA arc
formed by the pseudoknots remains highly structured after the structural rearrangements that occur during accommodation.
My contributions: Design of research (together with ML), specimen preparation for
electron microscopy, data collection, analysis of results (together with SHWS, ML and
HH), writing of paper (together with MP, ME, SHWS, ML, HH).
Paper IV – The pseudoknots of tmRNA in the double-translocation tmRNA·SmpB·70S
complex is located outside the ribosome in positions similar to the ones assigned for the
pseudoknots in the post-accommodated state. Furthermore, the maps show indications
of dynamic behavior around the hinge regions connecting the individual pseudoknots.
My contributions: Design of research (together with ML), specimen preparation for
electron microscopy, data collection, analysis of results (together with SHWS, MP, HH
and ML), writing paper (together with MP).
29
ACKNOWLEDGEMENTS
At last I have made it to the end of my PhD studies. It has not been easy and I could not
have pulled it off without all the people who supported and believed in me. So now I
would like to thank them, in special:
Hans Hebert for making me part of your lab these years. I would also like to thank you
for your encouragement and support, especially at times when I doubted myself.
Martin Lindahl for sharing the ribosome project with me, and for supervising me
during the second part of my PhD studies. I am also thankful for your patience with all
my questions. Not only were they never-ending, but they were also repeated several
times.
Philip Koeck, also for supervising me and for your patience with my questions. But
mostly I am grateful that you asked me to join the EM-group.
Past and present members of the EM-group, especially; Anna-Karin - thank you so
much for being such a good friend and colleague during these years! I could not have
asked for a better companion along this winding road. Caroline for being the glue in the
group that keeps things together. Pasi for all the help with the microscope. Hans E and
Dominika for your peptalk at the end when things were a bit rocky. Urban, for always
helping me more than you needed to. Ulrika, Priya, Karina, Peter, Tomas, Björn,
Ronny, Sipra and Qie.
My collaborators: Krassimira Idakieva for providing me with hemocyanin samples.
Rudolf Ladenstein and Mikael Karlström for letting me be part of the chaperonin
project which gave me the chance to do some protein purification work. Måns
Ehrenberg, Misha Pavlov and Natalia Ivanova for a nice collaboration on the ribosome
project. José María Carazo and Sjors Scheres, also for the nice collaboration on the
ribosome project and for introducing me to your awesome software package XMIPP!
Yoshi – thank you so much for welcoming me to your lab! That month really was one
of the best times of my life! I also want to thank Mie for all the help with the paperwork
and accommodation, and for the nice morning chats. Chris for preventing many lost in
translation scenarios. Tomohiro for teaching me all there is to know about how to preirradiate grids and also about mountain climbing.
Kurt Berndt for your great lectures at Södertörns Högskola. Thank you for opening my
eyes to structural biology.
Other colleagues at NOVUM; Tobias for being a good friend that is sweet, warm,
funny and smart - all at the same time! Sofia, Boel, Katarina and Linda for brightening
up our corner of level 7. Sunny, for our nice talks and for attending all my dance
classes. Patrick, also for our nice talks and for teaching me that stress is just a waste of
time and energy. Merita and Birgitta for always being happy and cheerful whenever we
30
run into each other in the hallways. Johan, Jianxin, Xiaofeng, Ivan, Joelle, Luca,
Magnus.
My students at Södertörns Högskola, best wishes to all of you!
All the staff at NOVUM, especially; Inger M for teaching me that sometimes it’s not
me, it’s ‘them’. Anders and Erik for coming to my rescue every time my computer was
giving me a hard time. Kristina for taking such good care of us PhD students!
Karl-Erik, tusen tack för all hjälp med det administrativa inför disputationen! Det hade
inte gått lika smidigt utan dig.
Mina kära vänner; Emelie, för att du är en toppenvän! Din omtänksamhet har ingen
begränsning! 2010 får bli mitt år istället. - Tobbe, för att du alltid ställer upp oavsett
om man behöver prata eller flytthjälp. Kristian, för att du lärde mig använda min dark
force på ett optimalt sätt. Micke, för alla härliga uppdateringsmail från Tyskland samt
dina ”bra” förslag på felsökningar då mikroskopet inte fungerar. Muna, för den gamla
goda tiden då vi var labkompisar på Södertörns högskola. Mimmi, för att du vägrar
kalla mig Kimberley. Är glad att ha en vän som dig! Atif - min gulliga (men inte söta)
kompis från högskoletiden. Sara A, Louise, Hanna, Sara B, Johanna, Nikolina och
Anders A - mina härliga danskompisar! Ni är en utav anledningarna till varför jag
älskar att dansa. Bara tanken på er får mig att le. Daniel D – min vän och
favoritdanslärare. Du och dina dansklasser har betytt så mycket mer för mig än du anar.
Agneta, för att du fick mig att inse hur jag vill att mitt liv ska vara.
Pia för att du är en fantastisk vän samt den bästa kusinen man kan önska sig! Din
självständighet, glädje samt intellekt är något jag strävar efter att uppnå. Din humor har
jag redan…. Min syster Grace - tack för att du inte gav upp hoppet om mig! Också en stor kram för
att du alltid fanns där och delade alla upp- och nedgångar i mitt liv. Lovar att inte klona
en människa!
Min bror Alex - du är en outtömlig källa av positiv energi! Ditt sätt att se allt från den
ljusa sidan är verkligen inspirerande.
Mamma och pappa, för ert stöd samt alla visdomsord ni delar med er.
Min svåger Magnus, min svägerska Veronika samt syskonbarnen Kevin, Viggo och
Axel, ni är verkligen ett härligt tillskott till min smått galna och vilda familj.
Anders – gigantisk puss och kram för ditt stöd, den har varit ovärderlig! Utan dig hade
jag aldrig klarat av att slutföra det hela. Tack för att du finns i mitt liv!
31
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