Structural Consequences of Deproteinating the 50S Ribosome

Abstract

Ribosomes are complex ribonucleoprotein particles. Purified 50S ribosomes subjected to high-salt wash, removing a subset of ribosomal proteins (r-proteins), were shown as competent for in vitro assembly into functional 50S subunits. Here, we used cryo-EM to determine the structures of such LiCl core particles derived from E. coli 50S subunits. A wide range of complexes with large variations in the extent of the ordered 23S rRNA and the occupancy of r-proteins were resolved to between 2.8 Å and 9 Å resolution. Many of these particles showed high similarity to in vivo and in vitro assembly intermediates, supporting the inherent stability or metastability of these states. Similar to states in early ribosome assembly, the main class showed an ordered density for the particle base around the exit tunnel, with domain V and the 3′-half of domain IV disordered. In addition, smaller core particles were discovered, where either domain II or IV was unfolded. Our data support a multi-pathway in vitro disassembly process, similar but reverse to assembly. Dependencies between complex tertiary RNA structures and RNA-protein interactions were observed, where protein extensions dissociated before the globular domains. We observed the formation of a non-native RNA structure upon protein dissociation, demonstrating that r-proteins stabilize native RNA structures and prevent non-native interactions also after folding.

Full text

Citation: Larsson, D.S.D.; Kanchugal
P, S.; Selmer, M. Structural
Consequences of Deproteinating the
50S Ribosome. Biomolecules 2022,12,
1605. https://doi.org/10.3390/
biom12111605
Academic Editor: Leonard
B. Maggi, Jr.
Received: 30 September 2022
Accepted: 25 October 2022
Published: 31 October 2022
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biomolecules
Article
Structural Consequences of Deproteinating the 50S Ribosome
Daniel S. D. Larsson , Sandesh Kanchugal P †and Maria Selmer *
Department of Cell and Molecular Biology, Uppsala University, SE 751 24 Uppsala, Sweden
*Correspondence: maria.selmer@icm.uu.se; Tel.: +46-18-4714177
† Present address: MAX IV Laboratory, Fotongatan 2, SE 224 84 Lund, Sweden.
Abstract:
Ribosomes are complex ribonucleoprotein particles. Purified 50S ribosomes subjected to
high-salt wash, removing a subset of ribosomal proteins (r-proteins), were shown as competent for
in vitro
assembly into functional 50S subunits. Here, we used cryo-EM to determine the structures
of such LiCl core particles derived from E. coli 50S subunits. A wide range of complexes with large
variations in the extent of the ordered 23S rRNA and the occupancy of r-proteins were resolved to
between 2.8 Å and 9 Å resolution. Many of these particles showed high similarity to
in vivo
and
in vitro
assembly intermediates, supporting the inherent stability or metastability of these states.
Similar to states in early ribosome assembly, the main class showed an ordered density for the
particle base around the exit tunnel, with domain V and the 3
0
-half of domain IV disordered. In
addition, smaller core particles were discovered, where either domain II or IV was unfolded. Our data
support a multi-pathway
in vitro
disassembly process, similar but reverse to assembly. Dependencies
between complex tertiary RNA structures and RNA-protein interactions were observed, where
protein extensions dissociated before the globular domains. We observed the formation of a non-
native RNA structure upon protein dissociation, demonstrating that r-proteins stabilize native RNA
structures and prevent non-native interactions also after folding.
Keywords:
ribosome assembly; RNA structure; RNA folding; RNA-protein interactions; single-particle
cryo-EM
1. Introduction
The ribosome is a large macromolecular complex translating mRNA into protein
in all domains of life. Structural biology has elucidated the architecture of ribosomes,
where each subunit consists of structured ribosomal RNA (rRNA) stabilized by ribosomal
proteins
(r-proteins)
that use positively charged loops and tails linked to globular domains
to stabilize the tertiary structure of the rRNA. The intricate assembly of ribosomes is, in all
organisms, a tightly regulated process that has been fine-tuned by evolution.
The assembly of the 50S subunit from E. coli has been extensively studied
in vitro
,
and methods were early developed to reconstitute active subunits from components [
1
].
Another
in vitro
approach is to promote the dissociation of r-proteins through high-salt
wash of ribosomal particles. Proteins were shown to dissociate from the core at defined
concentrations of, e.g., LiCl [
2
] in a non-denatured state [
3
,
4
]. The observed order of
leaving r-proteins nearly agrees with the reverse of the
in vivo
assembly order followed
by quantitative mass spectrometry [
5
] as well as the
in vitro
reconstitution protocol [
6
]. A
subset of the ribosomal proteins that dissociate during salt wash also exchanges
in vivo
,
presumably as a mechanism by which damaged proteins can be continuously replaced [
7
].
In vivo
,E. coli makes use of a multitude of auxiliary RNA modification enzymes
and ribosome assembly factors to fold and assemble ribosomes from three rRNAs and
54 r-proteins (reviewed in [
8
,
9
]). Despite the complexity of the process, the assembly
time for a mature ribosome
in vivo
in exponentially dividing E. coli has been estimated to
only 2 min [
10
]. Our structural understanding of the folding and assembly of ribosomes
has greatly benefitted from progress in cryogenic electron microscopy (cryo-EM), where
Biomolecules 2022,12, 1605. https://doi.org/10.3390/biom12111605 https://www.mdpi.com/journal/biomolecules

Biomolecules 2022,12, 1605 2 of 20
classification schemes in single-particle reconstruction have proven essential to studying
these heterogeneous particle ensembles. To enrich ribosome assembly intermediates
in vivo
for these studies, bacteria have been perturbed by assembly inhibitors [
11
,
12
], the knock-
down of r-proteins or assembly factors [
13
–
15
], or subjected to pull-downs with assembly
factors as bait [16,17].
In one such study, a bL17 depletion strain was used to generate ribosomal 50S sub-
particles [
13
], allowing the identification of several cooperative folding blocks. In agreement
with previous studies [
12
], a model of three major routes of LSU folding upon bL17
perturbation was proposed, where the central protuberance (CP), the particle base, and the
L7/L12 stalk formed in different orders.
Structural studies of assembly intermediates from LSU
in vitro
reconstitution [
18
]
showed five distinct precursors that shared many features with in vivo assembly interme-
diates from the bL17-depleted strain. The later intermediates displayed an intricate variety
of folding states of the protuberances and the peptidyl transfer center (PTC).
Salt-washed LiCl core particles have been demonstrated to be assembly-competent,
allowing reconstitution into active 50S subunits [
1
,
6
]. They also work as
in vitro
substrates
for RlmF, an early rRNA modifying enzyme that lacks activity on naked RNA [
19
], sup-
porting at least local similarity to states occurring during ribosome assembly
in vivo
. To
date, there is no high-resolution structure of these particles, but early negative-stain elec-
tron microscopy showed a high degree of heterogeneity where approximately 40% of the
ribosomes remained as compact particles lacking the three characteristic protuberances of
the LSU [20].
Here, we set out to further characterize the structures of LiCl core particles using
single-particle cryo-EM, with the goal to characterize the high-resolution structure and
composition of these particles. We identified six major types of 50S sub-particles with a
wide range of sizes and a multitude of variants of each of these. The method allowed
observation of core particles smaller than the so-far identified 50S assembly intermediates
and the opportunity to examine the effects of removing particular r-proteins. A comparison
of these particles enabled us to identify stability dependencies between rRNA helices and
r-proteins, reconstruct possible disassembly and assembly pathways, and demonstrate
strong similarities between intermediates formed during disassembly and assembly.
2. Materials and Methods
2.1. Preparation of LiCl Core Particles
Frozen cell pellets of E. coli
∆
ybiN from the Keio collection (JW5107, [
21
]) (24 g, wet
weight) were resuspended in 100 mL of opening buffer (20 mM Tris-HCl pH 7.5,
100 mM
NH
4
Cl, 10 mM Mg(OAc)
2
, 0.5 mM EDTA and 3 mM
β
-mercaptoethanol) containing
5µg/mL
DNase I (Sigma-Aldrich, St. Louis, MI, USA) and cOmplete
™
EDTA-free protease
inhibitor cocktail (Roche, Basel, Switzerland) and the cells were opened using a CF Cell
Disrupter (Constant Systems Ltd., Daventry, UK). The lysate was cleared by centrifugation
(Sorvall SS-34 at 16,000 RPM for 45 min) and filtration (0.45
µ
m pore size). Crude ribosomes
were obtained by pelleting (Beckmann Type 45Ti at 40,000 RPM for 18 h) 25 mL fractions
of lysate through 27 mL cushions of sucrose buffer (20 mM Tris-HCl pH 7.5, 500 mM
NH
4
Cl, 10.5 mM Mg(OAc)
2
, 0.5 mM EDTA-Tris, 1.1 M sucrose, 3 mM
β
-mercaptoethanol).
The ribosome pellets were dissolved in wash buffer (20 mM Tris-HCl pH 7.5, 500 mM
NH
4
Cl, 10.5 mM Mg(OAc)
2
, 0.5 mM EDTA-Tris, 7 mM
β
-mercaptoethanol), and sedimented
through sucrose cushions a second time (25 mL of ribosomes and 1.5 mL of sucrose buffer
in a Beckmann SW32Ti at 24,800 RPM for 16 h). The crude ribosomes were separated on
pre-formed linear 15–30% sucrose gradients (Beckmann SW32Ti at 19,300 RPM for 17 h) in
TC buffer (20 mM Tris-HCl pH 7.5, 60 mM NH
4
Cl, 5.25 mM Mg(OAc)
2
, 0.25 mM EDTA-Tris,
3 mM
β
-mercaptoethanol), and the 70S peak was pooled and pelleted (Beckmann Type
45Ti at 45,000 RPM for 18 h). To obtain individual subunits, 70S pellets were resuspended
in SU buffer (TC buffer but with 3 mM Mg(OAc)2) and subunits separated on pre-formed
linear 15–30% sucrose gradients in SU buffer (Beckmann SW32Ti at 19,300 RPM for 16 h).

Biomolecules 2022,12, 1605 3 of 20
The 50S subunit fractions were pooled and pelleted (Beckmann Type 45Ti at 45,000 RPM
for 18 h). The pellets were resuspended in LiCl-wash buffer (10 mM Hepes-KOH pH 7.5,
10 mM Mg(OAc)
2
, 3.5 M LiCl) and incubated for 5 h on ice. The 50S LiCl core particles
were pelleted (Beckmann Type 90Ti at 44,000 RPM for 4 h), resuspended in SU buffer, and
separated on pre-formed linear 15–30% sucrose gradients in SU buffer (Beckmann SW32Ti at
19,900 RPM for 16 h). The 50S LiCl core particle fractions were pooled, pelleted (Beckmann
Type 90Ti at 45,000 RPM for 17 h), and resuspended in storage buffer (
20 mM
Tris-HCl
pH 7.5, 50 mM NH
4
Cl, 5 mM Mg(OAc)
2
, 3 mM
β
-mercaptoethanol). The concentration of
50S LiCl core particles was measured by 260 nm absorbance (assuming 1 A
260
unit equals
38 nM) and aliquots were flash-frozen and stored at −80 ◦C.
2.2. In Vitro Methylation Assay
Assays were performed in duplicates. Each reaction contained 30 pmol 50S-LiCl parti-
cles in reaction buffer (50 mM HEPES-KOH pH 7.5, 20 mM NH
4
Cl, 100 mM KCl, 2.5 MgOAc,
8 mM
β
-mercaptoethanol). The addition of 400 pmol unlabeled S-Adenosyl-L-methionine
(Sigma-Aldrich, USA) doped with 16 pmol S-[methyl-
3
H]-adenosyl-L-methionine
(4 Ci/mmol; PerkinElmer, Waltham, MA, USA) was followed by the addition of 40 pmol
RlmF, or buffer (negative control) to a final volume of 30 µL. All reactions were incubated
at 37
◦
C for up to 30 min, quenched in 2 mL of ice-cold 10% trichloroacetic acid, and
incubated on ice for 10 min. The precipitated reactions were applied to BA85 nitrocellulose
filters (Whatman, Little Chalfont, UK) under vacuum and washed five times with 7 mL
of cold 10% trichloroacetic acid. The washed filters were placed in vials containing 5 mL
of Filter Safe scintillation cocktail (Schleicher & Schuell GmbH, Keene, NH, USA), shaken
for 30 min, and counted in an LC6500 scintillation counter (Beckman, Brea, CA, USA). For
calculation of the fraction of modified particles, the CPM per pmol of doped SAM mix was
measured. This allowed conversion of CPM per pmol of ribosomal particles to pmol of
modified particles.
2.3. Cryo-EM Sample Preparation
A 300-
µ
L reaction containing 5.2 nmol of C-terminally His-tagged RlmF and 1.2 nmol
of 50S LiCl core particles in imaging buffer (50 mM Hepes-KOH pH 7.5, 100 mM NH
4
Cl,
10 mM
Mg(OAc)
2
, 0.5 mM EDTA-Tris, 6 mM
β
-mercaptoethanol, 0.5 mM sinefungin
(Sigma-Aldrich, USA), 0.5 U/mL of RiboLock RNase inhibitor (Thermo Fisher Scientific,
Waltham, MA, USA)) was incubated with 200
µ
L of Ni-Sepharose (GE Healthcare, Uppsala,
Sweden) for 30 min at 25
◦
C under gentle agitation in a spin-filter and, subsequently,
centrifuged at 0.5 RFC for 3 min. The Ni-Sepharose beads were washed twice by incubation
for
10 min
in 500
µ
L of imaging buffer with 0.2 mM sinefungin, 0.1 U/mL of RiboLock
RNase inhibitor and 30 mM imidazole, and the ribosome-RlmF complex was eluted in
200 uL of imaging buffer with 0.2 mM sinefungin, 0.1 U/mL of RiboLock RNase inhibitor
and 300 mM imidazole. The eluate was sedimented in 50-
µ
L aliquots through 50
µ
L of
50% sucrose cushion in imaging buffer (Sorvall RC M150GX for 90 min at 80,000 RPM at
4
◦
C). The pellets were resuspended in 10
µ
L of imaging buffer and stored on ice before
being flash-frozen on cryo-EM grids. The protein content was verified by SDS-PAGE
and LC-MS/MS analysis of the lane cut from the SDS-PAGE gel. Sample preparation is
summarized in Figure S1.
For the LC-MS/MS analysis, the sample was digested by trypsin and the peptides
were subjected to reverse phase separation on a C18-column with a 90-min gradient and
electrosprayed to a Q Exactive Orbitrap mass spectrometer (Thermo Finnigan, San José, CA,
USA). Tandem mass spectrometry was performed by high-energy collision dissociation
fragmentation. The data were analyzed using Proteome Discoverer 1.4 (Thermo Fisher Sci-
entific) using the Sequest algorithm against a database of the E. coli K-12 BW25113 proteome
(the parental strain to the Keio collection). The search criteria for protein identification were
set to at least two matching peptides of 95% confidence level per protein.

Biomolecules 2022,12, 1605 4 of 20
2.4. Cryo-EM Grid Freezing and Data Collection
The cryo-EM specimens were prepared on QuantiFoil R 2/2 grids with 2-nm contin-
uous carbon (QuantiFoil Micro Tools GmbH, Großlöbichau, Germany). The grids were
glow-discharged for 40 s at 20 mA and 0.38 mBar using an EasiGlow (Ted Pella, Inc.,
Redding, CA, USA). Plunge-freezing was done using a Vitrobot mark IV (Thermo Fisher
Scientific, USA) using 3
µ
L of sample at 160 nM ribosome concentration after 30 s of incuba-
tion on the grid at 100% humidity and 10
◦
C and blotting for 3.5 s. Data were collected on
two different occasions on the same Titan Krios microscope from two different grids from
the same batch. The microscope was equipped with a Falcon-III direct electron detector
used in integrating mode. The nominal magnification was 75,000
×
, corresponding to a
pixel size of 1.09 Å in the specimen plane, and the acceleration tension was 300 kV. On
the first occasion, movies were collected with 66.2 e
−
/pixel/second during 0.773 s for a
total dose of 41.0 e
−
/Å
2
, and the sample stage was set to either 0
◦
(3052 movies) or 15
◦
(2196 movies) tilt. In the second session, movies were collected with 68.4 e
−
/pixel/second
during 0.770 s for a total dose of 42.2 e
−
/Å
2
, and the sample stage was set to a 30
◦
tilt
(6120 movies). Data collection parameters are summarized in Table S1.
2.5. Cryo-EM Data Processing
The 3D single-particle reconstruction was done using RELION version 3.0 [
22
] fol-
lowing the conventional RELION workflow [
23
]. The 0
◦
, 15
◦
, and 30
◦
tilt micrographs
were initially processed separately and later combined. Movies were motion corrected
in 5
×
5 patches using the RELION implementation of the MotionCor2 algorithm [
24
].
Per-micrograph defocus parameters were estimated using gCTF version 1.18 [
25
]. Parti-
cle picking was done using the Laplacian-of-Gaussian algorithm in RELION. Local CTF
parameters were estimated for the tilted micrographs using gCTF. An ab initio 3D initial
model was generated from primary data to prevent model bias. Multiple rounds of 2D
and 3D classifications were performed to prune the data down to a set of 384,374 particles.
Automatic 3D refinement, per-particle refinement of CTF parameters, and particle polishing
were performed several times until no further improvement in resolution could be achieved.
RELION version 3.1 [
26
] was then used to estimate higher order aberrations and anisotropic
magnification, followed by further CTF-refinement and particle polishing, to produce the
final 2.84-Å consensus reconstruction. The mask for estimating resolution was generated
by masking the final reconstruction with a 155-Å spherical mask to remove artifacts at the
rim of the reconstruction followed by low-pass filtering to 15 Å, thresholding that map at a
sufficiently low value to encompass diffuse density associated with the particle map but
excluding solvent noise (0.012), expanding the mask first by 5 pixels, and finally adding a
5-pixel soft edge. Estimation of local resolution and local low-pass filtering was performed
using the RELION algorithm using a B-factor determined by fitting to an approximately
linear regime at a high resolution of the Guinier plot (
−
32.2 Å
2
). The efficiency of the orien-
tation distribution (E
od
) parameter was calculated using cryoEF [
27
] and the distribution
of Euler angles was plotted (Figure S2). All but one class had E
od
values in the range of
0.6–0.8 (Table S2), which can be expected for ribosome data sets [
27
]. To identify distinct
particle subsets, 3D classification without alignment was performed based on the consensus
reconstruction. The particles in the consensus reconstruction were classified into six classes,
chosen based on initial testing with more classes, and each class was refined separately
according to the above procedure. Subsequently, classes were further 3D-classified in the
same manner in tiers, each time requesting four classes. Data processing is summarized in
Tables S1 and S2.
2.6. Model Refinement in Cryo-EM Maps
A high-resolution crystal structure of the E. coli ribosome (PDB ID 4YBB [
28
]) was
rigid-body fitted into the consensus map using UCSF Chimera version 1.14 [
29
]. The pixel
size was refined to 1.07 Å by monitoring the map-model fit while varying the pixel size.
Large disordered regions were pruned from the model and refinement was performed

Biomolecules 2022,12, 1605 5 of 20
using real_space_refine in Phenix version 1.19 [
30
] with reference model restraints to PDB
ID 7K00 [
31
] and interactive rebuilding using Coot version 0.9 [
32
]. In the final deposited
model, all nucleotides not supported by the density were removed. No water oxygens
or ions were modeled. Figures were prepared using ChimeraX version 1.4 [
33
]. Model
refinement parameters are summarized in Table S1.
2.7. Occupancy of Structural Elements and Hierarchical Clustering
The atomic model of a mature ribosome (PDB ID 4YBB), as well as a model refined
against the consensus reconstruction, were rigid body fitted to each class reconstruction
using UCSF Chimera 1.14 [
29
], and the map values were calculated at the position of each
heavy atom (i.e. non-hydrogen) in the LSU using the phenix.map_value_at_point program.
For these calculations, unsharpened maps produced by Relion 3D refinement were used,
which are on an absolute gray scale. Maps were low-pass filtered to 5 Å to avoid spurious
results due to local misalignments between the maps and the models, and the pixel size
was adjusted to the calibrated pixel size.
The occupancy was calculated for each structural element (chain or 23S rRNA helix)
as the fraction of atoms with a map value above a threshold of 0.05. The threshold was
confirmed by manual inspection of the maps. The values were normalized according to [
13
],
i.e., by dividing by the value by the maximum observed value for a particular feature across
all classes or by the median values across all features and maps, whatever was larger.
Manual inspection to gauge occupancies of r-protein was performed by identifying
the map threshold where the strongest recognizable feature could be seen. Density for 23S
rRNA helices was classified as being near-native, slightly distorted, highly distorted, or
disordered, which were translated into numerical values 0.1, 0.085, 0.045, and 0, respectively,
for clustering and plotting purposes.
The boundaries for the 23S rRNA helices were taken from [
34
], specifically the seg-
mentation in the E. coli secondary structure map downloaded from the Ribosome Gallery
on 22 September 2020 (http://apollo.chemistry.gatech.edu/RibosomeGallery/bacteria/E%
20coli/LSU/E_coli_LSU_Helices_2.png), which differs slightly from the original publica-
tion. RNA helices not named in the map were given the number of the helix immediately
preceding with an additional letter or the next number if available, cf. Table S3.
Hierarchical clustering using the complete linkage method and the Euclidean met-
ric was calculated using the function scipy.cluster.hierarchy.linkage in the SciPy library
(https://www.scipy.org/, accessed on 10 August 2020), both for classes (columns) as well
as features (rows). R-proteins that could not be identified in any of the classes were ex-
cluded from the analysis. The clustering of features was rather sensitive to the chosen
parameters (e.g., reference model, threshold, low-pass filtering, occupancy normalization,
exclusion/inclusion of r-proteins, etc.), but the general trends were reproducible.
3. Results
3.1. Sample Preparation
LiCl core particles were prepared from active 50S subunits of E. coli
∆
ybiN, a strain
lacking the RlmF-mediated methylation of A1618, by salt wash and sucrose gradient
purification (Figure S1A). In an
in vitro
tritium-labeling assay, these LiCl core particles
could be methylated by RlmF [
19
], and methylation showed saturation after 10 min at
~20% modification (Figure S3). With the aim to determine the structure of RlmF bound to
a pre-ribosomal-like substrate, the sample was enriched for such particles by pull-down
with His-tagged RlmF in the presence of the SAM analog sinefungin (Figure S1B). The
pull-down yield was 4-fold higher in the presence of RlmF and sinefungin compared to the
negative control without both. In the resulting population, RlmF was by SDS-PAGE judged
to be present at approximately the same stoichiometry as uL2 (Figure S1C) and LC-MS/MS
analysis showed a similar or higher signal as for the stably bound r-proteins. However, no
distinct density for RlmF could be found in any of the particle classes after single-particle
reconstruction, indicating that it dissociated after the pull-down.

Biomolecules 2022,12, 1605 6 of 20
3.2. High-Resolution Cryo-EM Reconstruction of the 50S LiCl Core Particle
Cryo-EM imaging, single-particle reconstruction, and a multi-tiered classification
scheme were used to determine the structure of the LiCl core particle (Figure 1). Because
of strong preferred orientation [
20
], data collected at 15
◦
and 30
◦
tilts were used to im-
prove the angular sampling (Figure S2). The micrographs showed heterogeneous particles
(Figure S4a,d) and several of the 2D class averages had one disordered side (Figure S4g).
Biomolecules 2022, 12, x FOR PEER REVIEW 6 of 21
be methylated by RlmF [19], and methylation showed saturation after 10 min at ~20%
modification (Figure S3). With the aim to determine the structure of RlmF bound to a pre-
ribosomal-like substrate, the sample was enriched for such particles by pull-down with
His-tagged RlmF in the presence of the SAM analog sinefungin (Figure S1B). The pull-
down yield was 4-fold higher in the presence of RlmF and sinefungin compared to the
negative control without both. In the resulting population, RlmF was by SDS-PAGE
judged to be present at approximately the same stoichiometry as uL2 (Figure S1C) and
LC-MS/MS analysis showed a similar or higher signal as for the stably bound r-proteins.
However, no distinct density for RlmF could be found in any of the particle classes after
single-particle reconstruction, indicating that it dissociated after the pull-down.
3.2. High-Resolution Cryo-EM Reconstruction of the 50S LiCl Core Particle
Cryo-EM imaging, single-particle reconstruction, and a multi-tiered classification
scheme were used to determine the structure of the LiCl core particle (Figure 1). Because
of strong preferred orientation [20], data collected at 15° and 30° tilts were used to improve
the angular sampling (Figure S2). The micrographs showed heterogeneous particles (Fig-
ure S4a,d) and several of the 2D class averages had one disordered side (Figure S4g).
Figure 1. Single-particle reconstruction and hierarchical classification of LiCl core particles. The
widths of the branches in the diagram reflect the number of particle images (italic font) in each class
(bold font). Classes are sorted from left to right, approximately according to the amount of ordered
density. Images show the reconstructions in “crown view” with the L1 protuberance to the left and
the L7/L12 stalk to the right (cf. Figure S6 for additional views).
3.3. LiCl Washing of the LSU Produces a Range of Sub-Particles
A consensus reconstruction from ~384,000 particle images produced a map at 2.84 Å
resolution (Figures 2 and S5). The reconstructed density is dome-like with the ordered
part at the 50S solvent side, centered on the expanded nascent-chain exit tunnel (Figure
3a,b). None of the three characteristic protuberances of the LSU are ordered.
Figure 1.
Single-particle reconstruction and hierarchical classification of LiCl core particles. The
widths of the branches in the diagram reflect the number of particle images (italic font) in each class
(bold font). Classes are sorted from left to right, approximately according to the amount of ordered
density. Images show the reconstructions in “crown view” with the L1 protuberance to the left and
the L7/L12 stalk to the right (cf. Figure S6 for additional views).
3.3. LiCl Washing of the LSU Produces a Range of Sub-Particles
A consensus reconstruction from ~384,000 particle images produced a map at
2.84 Å
resolution (Figures 2and S5). The reconstructed density is dome-like with the ordered part
at the 50S solvent side, centered on the expanded nascent-chain exit tunnel (Figure 3a,b).
None of the three characteristic protuberances of the LSU are ordered.
Classification resulted in six major classes, numbered 1–6 according to size
(
Figures 1and S6
). Three levels of further classification generated several sub-classes,
where the largest sub-class 5-5 collected approximately 1/3 of the initial particles. Model
refinement (Figures 3a and S5d) and some of the analysis and figures are based on the
consensus map, which is very similar to class 5-5; importantly, all conclusions are also valid
for this particle.
Most classes could be resolved to better than 4.5 Å resolution, but the smallest and
largest particles could only be resolved to approximately 9 Å resolution
(Figures S7 and S8)
,
probably due to a low number of particles and, for the smallest particles, heterogeneity
in structure as well as the presence of r-proteins. All particles have resolutions that allow
interpretation on the level of presence or absence of double-helical RNA elements.

Biomolecules 2022,12, 1605 7 of 20
Biomolecules 2022, 12, x FOR PEER REVIEW 7 of 21
Classification resulted in six major classes, numbered 1–6 according to size (Figures
1 and S6). Three levels of further classification generated several sub-classes, where the
largest sub-class 5-5 collected approximately 1/3 of the initial particles. Model refinement
(Figures 3a and S5d) and some of the analysis and figures are based on the consensus map,
which is very similar to class 5-5; importantly, all conclusions are also valid for this parti-
cle.
Most classes could be resolved to better than 4.5 Å resolution, but the smallest and
largest particles could only be resolved to approximately 9 Å resolution (Figures S7 and
S8), probably due to a low number of particles and, for the smallest particles, heterogene-
ity in structure as well as the presence of r-proteins. All particles have resolutions that
allow interpretation on the level of presence or absence of double-helical RNA elements.
Figure 2. Consensus reconstruction of the 50S LiCl core particle. (a) The map (gray) in “crown view”,
with a rigid-body docked model of the mature 50S particle (PDB ID 4YBB) colored as in (b). (b)
Secondary-structure map of the LSU rRNA (adapted with permission (CC BY-SA 3.0) from Ref.
[34]). Folded parts in the consensus particle are indicated with filled circles and outlined with a
dashed line. Disordered helices, loops, or single nucleotides are shown as white dots and labeled.
The RNA helices are colored in orange, indigo, blue, purple, yellow, or red tones for domains 0–VI,
respectively. (c) Four different views of the map with the rRNA colored according to (b) and r-
proteins in white.
Figure 2.
Consensus reconstruction of the 50S LiCl core particle. (
a
) The map (gray) in “crown
view”, with a rigid-body docked model of the mature 50S particle (PDB ID 4YBB) colored as in
(
b
).
(b) Secondary
-structure map of the LSU rRNA (adapted with permission (CC BY-SA 3.0) from
Ref. [
34
]). Folded parts in the consensus particle are indicated with filled circles and outlined with a
dashed line. Disordered helices, loops, or single nucleotides are shown as white dots and labeled.
The RNA helices are colored in orange, indigo, blue, purple, yellow, or red tones for domains 0–VI,
respectively. (
c
) Four different views of the map with the rRNA colored according to (
b
) and r-proteins
in white.
3.4. The LSU Is the Most Stable at the Solvent-Side
In the majority of the particles, more than half of the 23S rRNA is folded, including
domains 0, I, III, VI, most of domain II, and the 5
0
half of domain IV (i.e., H62–H67, from now
on referred to as sub-domain IV
50
) (Figure 2b). The folded regions are at the solvent side
and particle base of the LSU and to a high degree correlate with the presence of r-proteins
(Figure S5b). Several rRNA helices appear as folded but flexible low-resolution features.
The three major protuberances and the subunit interface, including functionally important
regions, such as the PTC and binding sites for tRNAs and GTPases, are unstructured
(Figure 2). In addition, many single nucleotides involved in native tertiary contacts are
disordered (Figure 2b).

Biomolecules 2022,12, 1605 8 of 20
Biomolecules 2022, 12, x FOR PEER REVIEW 8 of 21
Figure 3. Local resolution and the expanded exit tunnel. (a) Consensus reconstruction seen from the
interface side, centered on the nascent-chain exit tunnel, colored according to local resolution, with
the refined atomic model colored according to Figure 2b. (b) Central slice through the map, rectan-
gle in (a), showing the expanded exit tunnel and the absence of the CP. The rigid-body fitted model
of the mature 50S particle (PDB ID 4YBB [28]) is colored as in Figure 2b. The solid black line indicates
the extent of the ordered density and the dashed line indicates the extent of the mature particle. (c,d)
Class 2-2 in the same view and model as in (a,b). In this particle, sub-domain IV5′ and parts of do-
mains II and III are disordered compared to the consensus structure, in particular, nucleotide A1618
in H49a. The thick black line in (d) indicates the extent of the density in class 2-2, while the thin and
dashed lines are the same as in (b).
3.4. The LSU Is the Most Stable at the Solvent-Side
In the majority of the particles, more than half of the 23S rRNA is folded, including
domains 0, I, III, VI, most of domain II, and the 5′ half of domain IV (i.e., H62–H67, from
now on referred to as sub-domain IV5′) (Figure 2b). The folded regions are at the solvent
side and particle base of the LSU and to a high degree correlate with the presence of r-
proteins (Figure S5b). Several rRNA helices appear as folded but flexible low-resolution
features. The three major protuberances and the subunit interface, including functionally
important regions, such as the PTC and binding sites for tRNAs and GTPases, are unstruc-
tured (Figure 2). In addition, many single nucleotides involved in native tertiary contacts
are disordered (Figure 2b).
3.5. The LiCl Core Particle Classes Are Analogous to In Vivo and In Vitro Assembly Intermedi-
ates
Classes 2, 5, and 6 show strong similarities to in vitro reconstitution intermediates
[18] and to in vivo assembly intermediates under bL17 depletion [13] (Figure S9), although
Figure 3.
Local resolution and the expanded exit tunnel. (
a
) Consensus reconstruction seen from
the interface side, centered on the nascent-chain exit tunnel, colored according to local resolution,
with the refined atomic model colored according to Figure 2b. (
b
) Central slice through the map,
rectangle in (
a
), showing the expanded exit tunnel and the absence of the CP. The rigid-body fitted
model of the mature 50S particle (PDB ID 4YBB [
28
]) is colored as in Figure 2b. The solid black line
indicates the extent of the ordered density and the dashed line indicates the extent of the mature
particle. (
c
,
d
) Class 2-2 in the same view and model as in (
a
,
b
). In this particle, sub-domain IV
50
and parts of domains II and III are disordered compared to the consensus structure, in particular,
nucleotide A1618 in H49a. The thick black line in (
d
) indicates the extent of the density in class 2-2,
while the thin and dashed lines are the same as in (b).
3.5. The LiCl Core Particle Classes Are Analogous to In Vivo and In Vitro Assembly Intermediates
Classes 2, 5, and 6 show strong similarities to
in vitro
reconstitution intermediates [
18
]
and to
in vivo
assembly intermediates under bL17 depletion [
13
] (Figure S9), although
they all include bL17. Major class 1 is smaller than any previously studied 50S assembly
intermediate and seems to represent a minimal stable core of the LSU.
To arrange the particles into a tree that allowed identification of cooperativity between
binding of r-proteins and folding of rRNA, automated (Figures S10 and S11) and manual
(Table S4) occupancy estimations were subjected to hierarchical clustering (Figure S12a–d).
The classes grouped into smaller (classes 1, 2, 3, and 5-1), medium-sized (classes 4 and
5, except 5-1 and 5-8), and larger particles (classes 5-8 and 6) (Figure S12i), analogous to
early, intermediate, and late 50S assembly intermediates (Figure S9). In the small particles,

Biomolecules 2022,12, 1605 9 of 20
most of 23S rRNA domains 0, I, III, VI, and a core of domain II are folded (Figure 4). In the
medium-sized particles, more of domain II, as well as sub-domain IV
50
and parts of domain
V are folded. The large particles are mature-like, but parts of sub-domain IV
30
and parts
of domain V are unfolded. The clustering of structural features (rows in Figure S12a–d)
approximately agrees with previously identified folding blocks [13].
Biomolecules 2022, 12, x FOR PEER REVIEW 9 of 21
they all include bL17. Major class 1 is smaller than any previously studied 50S assembly
intermediate and seems to represent a minimal stable core of the LSU.
To arrange the particles into a tree that allowed identification of cooperativity be-
tween binding of r-proteins and folding of rRNA, automated (Figures S10 and S11) and
manual (Table S4) occupancy estimations were subjected to hierarchical clustering (Figure
S12a–d). The classes grouped into smaller (classes 1, 2, 3, and 5-1), medium-sized (classes
4 and 5, except 5-1 and 5-8), and larger particles (classes 5-8 and 6) (Figure S12i), analogous
to early, intermediate, and late 50S assembly intermediates (Figure S9). In the small parti-
cles, most of 23S rRNA domains 0, I, III, VI, and a core of domain II are folded (Figure 4).
In the medium-sized particles, more of domain II, as well as sub-domain IV5′ and parts of
domain V are folded. The large particles are mature-like, but parts of sub-domain IV3′ and
parts of domain V are unfolded. The clustering of structural features (rows in Figure S12a–
d) approximately agrees with previously identified folding blocks [13].
Figure 4.
Ordered rRNA for all particle classes mapped on the secondary-structure map of the
LSU (adapted with permission (CC BY-SA 3.0) from Ref. [
34
]). Ordered RNA helices are colored
according to Figure 2b. Partially ordered helices have half-filled circles (lighter color) and disordered
helices are gray.

Biomolecules 2022,12, 1605 10 of 20
3.6. The Misfolded 30Strand of Helix H73
Non-native density at the base of the L7/L12 stalk shows up in several classes (1-1, 2-1,
3-1, 4, and 5-2), as a low-resolution protruding loop at low map thresholds (
Figures 5and S6
,
star). One arm of the loop consists of H97 (Figure 5d), slightly shifted compared to the
native particle. The other arm, with the appearance of an rRNA stem-loop, emerges outside
the particle between H1, H94, and H97. Intriguingly, H73 is natively positioned just inside
in most “non-star” particles, but absent in all of the “star” classes (Figure S13). Helix H73 is
at a 4-way junction, but the region of its 3
0
strand (Figure 5b) can, according to secondary
structure prediction, form an alternative stem-loop (Figure 5c). A predicted 3D model
of the stem-loop is compatible in size with the observed non-native density (Figure 5d).
Closing the loop, an unidentified protein seems to bind to the top of helices H97 and the
misfolded H73. Possibly, this could be uL6, which natively binds H97 and is detected at
low levels in the LC-MS/MS data (Table S4).
3.7. R-Proteins Leave the 50S According to In Vivo Assembly Groups
A number of proteins appear to be present across all classes (uL3, uL4, bL17, bL20,
bL21, uL22, uL23, uL29, and bL34), and in most cases also uL13 and uL24 (Table S4, Figure
S14). All of these, except bL32, have firm support in LC-MS/MS data (Table S4). These
proteins are all early binders
in vivo
, as identified by pulse-labeling coupled with MS
analysis [
5
] (Figure S14). Furthermore, four proteins that are only found in larger particles
(uL2, uL14, bL19, and bL32) bind in the subsequent step in vivo.
Fifteen proteins do not have clear density in any of the classes (uL1, uL6, bL9, uL10,
uL11, bL12, uL15, uL16, bL25, bL27, bL28, bL31, bL33, bL35, or bL36), although diffuse
density prevented unambiguous assessment of some proteins. Trace levels of uL1, uL5,
uL6, bL9, uL11, uL15, and bL31 were detected by LC-MS/MS (Table S4), suggesting that
some of these r-proteins remain bound to disordered rRNA.
The high solubility of the CP-associated r-proteins uL5, uL18, and bL25 destabilizes
the CP in the LiCl-washed ribosomes. The CP and the 5S rRNA are only discernable in the
largest particle classes, 6 and 5-8 (Figure S6), together with uL5 and uL18 at low occupancy
in class 6, and uL18 in class 5-8.
For the three largest r-proteins in the consensus reconstruction (uL2, uL3, and uL4),
the long loops and tails that in the native particle interact with the 23S rRNA are invisible
(Figure S5c). The interacting rRNA helices are in some cases unstructured and in other
cases folded but in a non-native position.
3.8. Comparison of Particle Classes Show Dependencies between Different Structural Elements
Based on the clustering analysis, structural consequences of the removal of certain
r-proteins and of unfolding of rRNA were analyzed by pair-wise comparison between
the particle classes. This revealed folding and stability dependencies that are mostly
additive (Figure 6).

Biomolecules 2022,12, 1605 11 of 20
Biomolecules 2022, 12, x FOR PEER REVIEW 10 of 21
Figure 4. Ordered rRNA for all particle classes mapped on the secondary-structure map of the LSU
(adapted with permission (CC BY-SA 3.0) from Ref. [34]). Ordered RNA helices are colored accord-
ing to Figure 2b. Partially ordered helices have half-filled circles (lighter color) and disordered hel-
ices are gray.
3.6. The Misfolded 3′ Strand of Helix H73
Non-native density at the base of the L7/L12 stalk shows up in several classes (1-1, 2-
1, 3-1, 4, and 5-2), as a low-resolution protruding loop at low map thresholds (Figures 5
and S6, star). One arm of the loop consists of H97 (Figure 5d), slightly shifted compared
to the native particle. The other arm, with the appearance of an rRNA stem-loop, emerges
outside the particle between H1, H94, and H97. Intriguingly, H73 is natively positioned
just inside in most “non-star” particles, but absent in all of the “star” classes (Figure S13).
Helix H73 is at a 4-way junction, but the region of its 3′ strand (Figure 5b) can, according
to secondary structure prediction, form an alternative stem-loop (Figure 5c). A predicted
3D model of the stem-loop is compatible in size with the observed non-native density
(Figure 5d). Closing the loop, an unidentified protein seems to bind to the top of helices
H97 and the misfolded H73. Possibly, this could be uL6, which natively binds H97 and is
detected at low levels in the LC-MS/MS data (Table S4).
Figure 5. Non-native density close to the base of the L7/L12 stalk. (a) In classes 1-1, 2-1, 3-1, and 4,
the density of H97 extends and loops back (asterisk) at lower map thresholds. In class 5-2, the
Figure 5.
Non-native density close to the base of the L7/L12 stalk. (
a
) In classes 1-1, 2-1, 3-1, and
4, the density of H97 extends and loops back (asterisk) at lower map thresholds. In class 5-2, the
density for the upper (black arrowhead) and lower (white arrowhead) arms are not connected.
Analogous classes 1-2, 2-2, 3-2, 5-1, and 5-3, respectively, do not show this extra density (not shown).
(
b
) Secondary structure map of the E. coli 23S rRNA shown for parts of domains 0, V, and VI (adapted
with permission (CC BY-SA 3.0) from Ref. [
34
]). The black line encloses the segment that was used
for the secondary-structure prediction in (
c
). (
c
) Predicted secondary structure of the 3
0
-strand of
H73 and neighboring nucleotides. Colors as in (
b
). (
d
) Loop density for class 4 (rectangle in (
a
))
with a rigid-body docked model of the 8-bp stem-loop in (
c
) predicted by 3dRNA [
35
] colored as in
(
c
). Models of H97 and H95 are shown for reference. The density connecting H97 and H73-3
0
could
represent an unidentified protein.

Biomolecules 2022,12, 1605 12 of 20
Biomolecules 2022, 12, x FOR PEER REVIEW 12 of 21
Figure 6. Relations between particle classes. Similar particles are connected by lines, representing
possible disassembly paths during salt wash. The right side contains particles with stronger density
for domain V, while the left-side particles have stronger densities for uL2, H34–35, and sub-domain
IV5′. The left side-shoot consists of particles with misfolded H73 (asterisks, Figure 5). The particles
are shown as in Figure S6. Orange and blue designations are similar classes in Davis et al. [13] and
Nikolay et al. [18] (see also Figure S9).
3.8.1. Sub-Domain IIItail Can Stabilize Sub-Domain IV5′ in Absence of uL2
Class 1-1, the smallest reconstructed particle, only consists of domains I (except H25),
III, VI, 0 (H26, H26a, and H61) and IV5′ (particularly strong density for H63) (Figure 4).
This is remarkable since the folding of sub-domain IV5′ in other classes coincides with the
presence of uL2 (see below). Class 1-2 has additional density for the core of domain II, a
more mature-like domain 0 and H25 of domain I, but less density for domain IV and the
distal half of domain III (helices H54–59, hereafter called sub-domain IIItail [36]).
Based on these structures, we hypothesize that in lieu of uL2, sub-domain IIItail can
stabilize sub-domain IV5′ through contacts between H57 and H58 in domain III and H63.
The same correlation between the folding of sub-domains IV5′ and IIItail was observed
Figure 6.
Relations between particle classes. Similar particles are connected by lines, representing
possible disassembly paths during salt wash. The right side contains particles with stronger density
for domain V, while the left-side particles have stronger densities for uL2, H34–35, and sub-domain
IV
50
. The left side-shoot consists of particles with misfolded H73 (asterisks, Figure 5). The particles
are shown as in Figure S6. Orange and blue designations are similar classes in Davis et al. [
13
] and
Nikolay et al. [18] (see also Figure S9).
3.8.1. Sub-Domain IIItail Can Stabilize Sub-Domain IV50in Absence of uL2
Class 1-1, the smallest reconstructed particle, only consists of domains I (except H25),
III, VI, 0 (H26, H26a, and H61) and IV
50
(particularly strong density for H63) (Figure 4).
This is remarkable since the folding of sub-domain IV
50
in other classes coincides with the
presence of uL2 (see below). Class 1-2 has additional density for the core of domain II, a
more mature-like domain 0 and H25 of domain I, but less density for domain IV and the
distal half of domain III (helices H54–59, hereafter called sub-domain IIItail [36]).
Based on these structures, we hypothesize that in lieu of uL2, sub-domain III
tail
can
stabilize sub-domain IV
50
through contacts between H57 and H58 in domain III and H63.

Biomolecules 2022,12, 1605 13 of 20
The same correlation between the folding of sub-domains IV
50
and III
tail
was observed
under bL17 depletion [
13
], and domain III was shown to fold independently of r-proteins
or the rest of the 23S rRNA [
37
]. Sub-domain III
tail
is mostly folded in all of the particles,
independently of domain IV and uL2, suggesting that it stabilizes domain IV rRNA at the
early stages of ribosome biogenesis.
In class 2 particles, the RNA structure is stabilized by additional r-proteins. Unlike
in class 1, there is no density for sub-domain IV
50
(Figure 7). In class 2-1, the whole flank
opposite to the CP (sub-domain III
tail
and helices in domains 0 and VI) is shifted 20 Å away
from the subunit interface compared to the native particle, whereas in class 2-2, this lobe is
instead in near-native position.
Biomolecules 2022, 12, x FOR PEER REVIEW 13 of 21
under bL17 depletion [13], and domain III was shown to fold independently of r-proteins
or the rest of the 23S rRNA [37]. Sub-domain IIItail is mostly folded in all of the particles,
independently of domain IV and uL2, suggesting that it stabilizes domain IV rRNA at the
early stages of ribosome biogenesis.
In class 2 particles, the RNA structure is stabilized by additional r-proteins. Unlike in
class 1, there is no density for sub-domain IV5′ (Figure 7). In class 2-1, the whole flank
opposite to the CP (sub-domain IIItail and helices in domains 0 and VI) is shifted 20 Å away
from the subunit interface compared to the native particle, whereas in class 2-2, this lobe
is instead in near-native position.
Figure 7. Side-by-side comparisons of particles of different sizes. RNA is colored as in Figure 2b and
proteins are shown in white.
The exit tunnel is expanded in the smaller particles, in particular in class 2-2 (Figure
3c,d). This would allow access of RlmF to its modification site A1618 in classes 1, 2, and 3-
1. These constitute 17% of the total number of particles, in reasonable agreement with 20%
of the particles being labeled after in vitro methylation.
Class 3-1 is rather similar to 2-1 but H63 is more pronounced and there is more diffuse
density in the H33–H35a region. Correlated to this, domain IIItail is closer to its native po-
sition, stabilized by H63 and possibly by the higher occupancy of uL3 and bL17.
In class 3-2, domain II is in a more native position compared to 3-1. It is the smallest
particle with unambiguous density for H34–H35 (Figure 7), correlated with improved po-
sitioning of nearby domain III (e.g., H58) and H96 and H101 in domain VI.
Figure 7.
Side-by-side comparisons of particles of different sizes. RNA is colored as in Figure 2b and
proteins are shown in white.
The exittunnel is expanded in the smaller particles, in particular in class 2-2 (
Figure 3c,d
).
This would allow access of RlmF to its modification site A1618 in classes 1, 2, and 3-1. These
constitute 17% of the total number of particles, in reasonable agreement with 20% of the
particles being labeled after in vitro methylation.
Class 3-1 is rather similar to 2-1 but H63 is more pronounced and there is more diffuse
density in the H33–H35a region. Correlated to this, domain III
tail
is closer to its native
position, stabilized by H63 and possibly by the higher occupancy of uL3 and bL17.
In class 3-2, domain II is in a more native position compared to 3-1. It is the smallest
particle with unambiguous density for H34–H35 (Figure 7), correlated with improved
positioning of nearby domain III (e.g., H58) and H96 and H101 in domain VI.
3.8.2. R-Protein uL3 Is Important for the Stability of Domains 0 and VI
Class 5-1 shares attributes with classes 2-2 and 3-2, but H63 is close-to-natively folded
(Figure 7). Similar to in larger class 5 particles, diffuse density extends from H73 across the
particle to H22 at lower thresholds (see below).

Biomolecules 2022,12, 1605 14 of 20
Higher occupancy of uL3 in 5-1 than in 3-2 seems to cause stronger density for domains
0 and VI (except H96). The same trend is also observed between 2-2 and 2-1. In the mature
particle, the 128–153 loop of r-protein uL3 directly interacts with several of these helices
(Figure S13b), supporting the importance of uL3 in maintaining the native H73 structure,
preventing its misfolding (see above).
3.8.3. R-Protein uL2 Stabilizes Sub-Domain IV50
Class 4 has many similarities to 5-1, but the misfolded H73 dislocates a lobe at the
solvent side close to the base of the stalk, including H1, H41, uL13, bL20, and bL21. Class 4
also has significantly stronger density for sub-domain IV
50
and more mature-like H33–H35a.
The differences in domains II and IV seem correlated with increased occupancy of uL2, also
seen in classes 5-2, 5-3, 5-5, and 5-6 and in the clustering analysis (Figure S12a,b, rows 85
and 89, respectively).
Class 5-2 is highly similar to class 4 (Figure 4), but with stronger density for sub-
domain IV50.
3.8.4. Helices H79 and H88 Anchor the Core of Domain V
Class 5-3 combines features from 3-2, 5-1, and 5-2 (Figure 6). Class 5-3 has density
for r-protein uL19 and low levels for neighboring uL14, similar to the first particle along
folding pathway C in [13] (Figure S9).
Class 5-4 is quite similar to class 5-3, but with slightly weaker sub-domain IV
50
and
uL2. Domain V is on the other hand more well-defined, in particular H79, H88, and the
proximal end of H89.
Class 5-5 shows less distinct density for domain V compared to 5-4, although H79
seems ordered. Instead, the density for uL2 and sub-domain IV
50
is stronger (even more
than in 5-3), placing it in a different path in Figure 6. The distal part of helix H67 is
well-defined, but the 30-half of domain IV is disordered (Figure 7).
3.8.5. Stability of Sub-Domain IV30Depends on Domain V
In class 5-6, all the elements that are visible in either of the smaller particles in class 5
are ordered (Figure 6).
Class 5-7 shows stronger and more well-defined density for domain V than the smaller
particles in class 5. There is clear density for helices H74, H75, H76, H79, and H88, albeit
shifted from their native positions (Figure 7). Similarly, the folding of domain V along
the C/E paths starts with H79 to expand towards the two protuberances under bL17
depletion [13]. The higher occupancy of uL13 in class 5-7 seems to stabilize H1 and H41.
Class 5-8 shows more native structure for domain V, which is partly held in place by
sub-domain IV
30
(Figure 7), but the distal ends of H68 and H69 are flexible. There is also
weak density for the rest of the 3
0
end of domain V, including H90–93 and the proximal
end of H89. Native positioning of H89, however, requires uL16 [
18
]. Finally, the H82–H87
branch of domain V, which forms part of the CP, shows partial order, and very weak density
can be discerned for additional large CP elements.
3.8.6. The CP Is Severely Destabilized in the LiCl Core Particles
Class 6 is the only particle where most of the CP rRNA, including 5S rRNA, is folded,
but otherwise it is similar to class 5-8. The CP is highly depleted of r-proteins but contains
low levels of uL5, uL18, and uL30. Relative to the mature particle, the entire CP has shifted
away from the subunit–interface side.
3.9. Folding of H49a and H35 Demonstrate Analogies between Disassembly and Assembly
Step-by-step comparison of the smaller to larger particles allows the folding of parts
of the LSU to be modeled as an intricate sequential addition of structural layers onto
a pre-existing core. As an example, we analyzed the structure of helices around H49a
(modification site of RlmF) and H35, in a region of tertiary interactions between domains 0,

Biomolecules 2022,12, 1605 15 of 20
II, III, IV, and V that is stabilized by the positively charged extensions of r-proteins uL2,
uL22, and bL34 (Figures 8and S15).
Biomolecules 2022, 12, x FOR PEER REVIEW 16 of 21
Figure 8. Structure of the H49a-H35 region in different particles suggests a local order of assembly.
(a–f) Maps showing varying degrees of folded structure in this region. Lines indicate densities that
differ between maps (the dashed lines indicate weak/diffuse densities). The maps are not sharpened,
or low-pass filtered, and are shown at a level of 0.05. Figure S15 shows the same view for all particle
classes. (a) The overview shows the consensus map and a mature model, PDB ID 4YBB, colored as
in Figure 2b. (b) In class 2-2, H48 is folded, but not H49a. (c) Densities for H34, H35, and H35a
outside of H49a in class 5-1. (d,e) Occupancy for H64, H65, and uL2 is partial in class 5-3, but close
to full in 5-5. (f) Helices H90, H93, and H74 are folded into class 5-8. (g) Schematic view of stabilizing
interactions in the H49a-H35 region. Tertiary contacts for H49a and H35 are indicated with solid
lines and numbered 1–10 in the approximate assembly order, as deduced from the maps (see sec-
tions 3.9.1-3.9.2). Dashed lines indicate contacts to r-proteins. (h) Structure of helices and r-proteins
in (g) in the mature 50S (PDB ID 4YBB), tertiary contacts are numbered 1-10 as in (g).
3.9.2. Folding and Tertiary Interactions of H35
Helix H35, a stem-loop in domain II, packs outside H49a, contacting domains 0 (H73),
IV (H64 and H65), and V (H93) (Figures 3f and 8g). H35 is disordered in most small clas-
ses, such as 2-2 (Figure 8b), and only vaguely discernable in the other small particles. The
stability of H35 shows a correlation to the presence of r-protein uL2 (Figure 8c–e). uL2
interacts with H35a, which packs outside of H35 (Figure 8c), and in turn contacts both
H35 (contact 6) and H65 (Figure 8d). An A-minor contact interlocks H64 and H35 (contact
Figure 8.
Structure of the H49a-H35 region in different particles suggests a local order of assembly.
(
a–f
) Maps showing varying degrees of folded structure in this region. Lines indicate densities that
differ between maps (the dashed lines indicate weak/diffuse densities). The maps are not sharpened,
or low-pass filtered, and are shown at a level of 0.05. Figure S15 shows the same view for all particle
classes. (
a
) The overview shows the consensus map and a mature model, PDB ID 4YBB, colored
as in Figure 2b. (
b
) In class 2-2, H48 is folded, but not H49a. (
c
) Densities for H34, H35, and H35a
outside of H49a in class 5-1. (
d
,
e
) Occupancy for H64, H65, and uL2 is partial in class 5-3, but
close to full in 5-5. (
f
) Helices H90, H93, and H74 are folded into class 5-8. (
g
) Schematic view of
stabilizing interactions in the H49a-H35 region. Tertiary contacts for H49a and H35 are indicated
with solid lines and numbered 1–10 in the approximate assembly order, as deduced from the maps
(
see Sections 3.9.1 and 3.9.2
). Dashed lines indicate contacts to r-proteins. (
h
) Structure of helices and
r-proteins in (g) in the mature 50S (PDB ID 4YBB), tertiary contacts are numbered 1-10 as in (g).

Biomolecules 2022,12, 1605 16 of 20
3.9.1. Folding and Tertiary Interactions of H49a
Helix H49a is a stem-loop in domain III that bridges to domain II, close to the nascent-
chain exit tunnel (Figure 3c,d). In the mature particle, the stem of H49a packs against
helices H49b and H50, and the loop forms interactions with H50, H47, and H35 (Figure 8g).
Based on this region of the different particles (Figure S15), we reconstructed the sequential
assembly of these interactions (Figure 8b–f). In class 2-2 (Figure 8b), H48 and H49b are
folded but H49a is completely disordered. In other early classes (2-1 and 3-1), H49a is
folded but points away from H50 at different angles. In classes 5-1 (Figure 8c) and 3-2,
H49a packs against H49b (contacts 1 and 2 in Figure 8g). Although the loop of H49a lacks
most of the native tertiary contacts, there is weak density for A1616 connecting to H50
(contact 3). Class 4 is similar but also shows density for the 5
0
-end of H47 (contact 4) and
H35. In classes 5-3 (Figure 8d) and 5-2, H49a is in a close-to-native conformation. Contact 5
is formed through the stacking between A1618 of H49a and A749 of H35 observed in class
5-4, and the triple-A stack is completed by A1272 of H47 as observed in class 5-5 (Figure 8e).
The stack would be further stabilized by the N6 methylation of central A1618 [
38
]. Classes
5-6 to 5-8 (Figure 8f) show only a slight distortion of the H49a loop at the flipped-out base
of A1614 compared to the native structure where it forms an interaction with the extended
loop of uL22, with rather weak density for the tip in all classes.
3.9.2. Folding and Tertiary Interactions of H35
Helix H35, a stem-loop in domain II, packs outside H49a, contacting domains 0 (H73),
IV (H64 and H65), and V (H93) (Figures 3f and 8g). H35 is disordered in most small classes,
such as 2-2 (Figure 8b), and only vaguely discernable in the other small particles. The
stability of H35 shows a correlation to the presence of r-protein uL2 (Figure 8c–e). uL2
interacts with H35a, which packs outside of H35 (Figure 8c), and in turn contacts both H35
(contact 6) and H65 (Figure 8d). An A-minor contact interlocks H64 and H35 (contact 7),
observed in the more mature-like classes 5-3 and 5-5 (Figure 8d,e), but absent in e.g., class
5-1. The further interactions of H65 with the loop of H35 (contact 8) and its loop to H73
and H93 (contacts 9 and 10) and uL22 are only present in the largest particles, such as class
5-8 (Figure 8f), completing the native structure.
4. Discussion
Core particles of the LSU, where the most loosely attached ribosomal proteins and
the 5S RNA have been removed using high-salt wash protocols, were first used in studies
of ribosome assembly more than half a century ago [
2
,
39
]. Here, our results provide a
structural understanding of what these particles are, and of their similarities and differences
to structures occurring during in vivo and in vitro ribosome assembly.
The removal of loosely associated r-proteins with a high-salt wash allows access
to the smallest stable cores of the large subunit. These particles show density for only
4–8 r-proteins
, but additional ones are possibly bound to disordered rRNA regions. With
the exception of the smallest particles, the reconstructed cores show strong similarity to in-
termediates isolated from bL17-deficient cells or from
in vitro
reconstitution (Figure S9) and
their protein content agrees with intermediates formed during
in vivo
assembly [
5
]. Similar
to results from the purification of
in vivo
30S sub-particles at different salt concentrations,
“exposure to the high salt buffer does not destroy the known binding interdependences
between the ribosomal proteins” [
40
]. The similarities between states obtained during
in vitro
disassembly and different variants of assembly strongly support the innate propen-
sity of the rRNA and the r-proteins to adopt distinct, meta-stable complexes. Evolutionary,
this inherent property is likely important for the robustness of ribosome assembly upon
changing or challenging conditions, such as disabling mutations of an r-protein [
13
]. Still,
local minima in the energy landscape can lead to the trapping of misfolded structures. One
such example is the strong non-native secondary structures observed to form by the 3
0
strand of helix H73 (Figure 5). This manifests the need for ribosomal proteins not only to
facilitate formation of the active RNA structure during co-transcriptional
in vivo
assembly

Biomolecules 2022,12, 1605 17 of 20
but also to maintain the native structure in a partly folded structural context. During
in vivo
assembly, RNA helicases allow remodeling of such misfolded structures [
9
] and
during
in vitro
assembly, the optimized conditions of temperature and magnesium [
1
,
41
]
may prevent their formation. Still, also
in vivo
perturbation by depletion of ribosomal
proteins or assembly factors may produce unstable particles that are prone to degradation
or misfolding.
The r-protein occupancy of our LiCl core particles largely agrees with previous charac-
terizations [
2
,
6
,
20
,
42
]. The spontaneously exchangeable r-proteins
in vitro
and
in vivo
[
7
]
are the most loosely associated ones, a small subset of the ones that can be dissociated
with salt. There is no indication that their dissociation and re-association would cause
major structural changes, which seems evolutionary important to maintain the active
pool of ribosomes.
The wide range of LSU sub-particles that we observe allows analysis of the conse-
quences of removal of specific r-proteins. For some r-proteins, e.g., uL3, a primary binder,
and uL2, a later binder [
5
] there is a strong link between their presence and observation of
the native rRNA structure in their close surrounding. The central protuberance shows high
sensitivity to deproteination and unfolds as proteins leave. In contrast, it can fold early
during ribosome biogenesis as demonstrated in the bL17-depletion model [
13
] and during
LSU
in vitro
reconstitution [
18
]. The structural stability of domain III seems less dependent
on r-proteins.
To what extent does ribosome disassembly
in vitro
reverse the process of ribosome
assembly
in vivo
or
in vitro
? Apart from the CP-region proteins uL5, uL15, and uL18, the
r-proteins that bind first during assembly are also the ones that stay bound during high-salt
wash. The disassembly process is sterically constrained by outer assembly layers of the
ribosome, and thereby more similar to
in vitro
assembly where the structure forms starting
from the core and continuing outwards. During the co-transcriptional
in vivo
process,
there is an additional directionality, where at a given time only part of the rRNA can be
available for folding and tertiary interactions (reviewed in [
9
]). However, since functional
ribosomes can be assembled on circularly permutated 23S RNA linked to 16S rRNA, no
directionality can be required for functional 50S assembly [
43
]. The 5
0
and 3
0
ends of 23S
RNA together form H99, which remains folded in all our particles (Figure S12f), and the
ordered elements of the smallest particles are scattered in several blocks along the 23S rRNA
sequence, showing no correlation between directionality and stability of RNA structure.
In addition, the correlation between the occupancy of r-proteins in our core particles and
early binding during
in vivo
assembly [
5
] suggests that directionality is not a main factor
in r-protein affinity.
The reconstructed series of particles subjected to r-protein dissociation corroborates
the multi-pathway of assembly [
13
,
44
,
45
] by demonstrating the semi-independent stabil-
ity of folding blocks of rRNA and r-proteins. In particular, since parts of domains IV
and V can remain structured while the other unfolds, we can from the LiCl core parti-
cles derive
two distinct
pathways where either of these blocks independently remains
folded (Figure 6).
Compared to the well-studied early, co-transcriptional 30S assembly process [
46
], little
is known about the corresponding steps of 50S assembly. However, this study provides
examples of concepts that have been described for the 30S, such as the sequential reduction
of structural variability of RNA by binding of r-proteins, specifically in regions that require
long-range interactions to reach their native structure. There is a clear similarity between
how r-proteins during assembly recognize and bind to structured regions and further
contribute to the maturation of the nearby structure, and how the dissociation of r-proteins
seems to start with the disordering of their extensions and loops coupled to loosening
of rRNA structure to be followed by dissociation of the folded r-protein domains and
unfolding of the respective rRNA binding sites. The native packing of the RNA structure
sometimes requires such r-protein tails or loops, as an example, H49a does not adopt its
fully native conformation in absence of the uL22 loop (Figure 8f), the deletion of which

Biomolecules 2022,12, 1605 18 of 20
has been shown to lead to the accumulation of immature LSU particles [
47
]. Moreover, the
back sides of both subunits show higher stability towards high salt than the interface sides
(this study and [
40
]), in line with preventing the premature association of the subunits. We
observe a multi-pathway disassembly process, in agreement with the complex “assembly
landscape” of the 30S subunit that was proposed as analogous to the energy landscape
in protein folding [
48
]. These observations support the fact that core particles, produced
by high-salt wash, in some respects can be used as mimics of transient early assembly
intermediates, for example, in studies of maturation factors.
In conclusion, the
in vitro
-generated 50S LiCl core particles represent a collection
of intrinsically stable or meta-stable complexes, from which we can learn about the in-
herent structural properties of ribonucleoprotein particles, and specifically about states
that are likely to occur during 50S ribosome assembly. Recent developments in the clas-
sification and analysis of heterogeneous particle populations [
49
,
50
] will be crucial in
further such studies.
Supplementary Materials:
The following supporting information can be downloaded at: https:
//www.mdpi.com/article/10.3390/biom12111605/s1, Figure S1: Sample preparation for cryo-EM
imaging; Figure S2: Euler angles from single-particle reconstructions; Figure S3:
In vitro
methy-
lation assay; Figure S4: Examples of raw micrographs, particle picking and 2D classification;
Figure S5: Consensus
reconstruction and model refinement; Figure S6: Non-redundant major and
minor reconstruction classes; Figure S7: Fourier-shell correlation between independent half-set;
Figure S8: Guinier plots; Figure S9: Comparison of 50S sub-particles formed during assembly and dis-
assembly; Figure S10: Correlation between map and model for the 23S rRNA;
Figure S11: Correlation
between map and model for 5S rRNA and r-proteins in the LSU; Figure S12: Occupancy and clustering
of structural elements in the reconstruction classes; Figure S13: Fold of helix H73;
Figure S14: In vivo
assembly groups; Figure S15: Folding of the H49a-H35 region; Table S1: Cryo-EM data collec-
tion, reconstruction and modeling statistics; Table S2: Details about cryo-EM reconstruction classes;
Table S3: Naming
of “non-standardized” 23S secondary structure elements;
Table S4: R-protein
com-
position of LiCl core particles.
Author Contributions:
Conceptualization, M.S. and D.S.D.L.; methodology, D.S.D.L.; validation,
D.S.D.L. and M.S.; investigation, D.S.D.L. and S.K.P.; resources, M.S.; data curation, D.S.D.L.;
writing—original
draft preparation, D.S.D.L. and M.S.; writing—review and editing, D.S.D.L. and
M.S.; visualization, D.S.D.L.; supervision, M.S.; funding acquisition, M.S. All authors have read and
agreed to the published version of the manuscript.
Funding:
This research was funded by the Swedish Research Council, grants 2016-06264 and
2017-03827 to M.S.
Data Availability Statement:
Atomic coordinates for the consensus structure have been deposited
in the Protein Data Bank under accession code 7ODE. All maps have been deposited in the Elec-
tron Microscopy Data Bank with accession codes EMD-12826 for the consensus reconstruction and
EMD-12828 through EMD-12841, EMD-12843, and EMD-12844 for the different classes.
Acknowledgments:
The cryo-EM data were collected with the aid of Julian Conrad and Marta
Carroni at the Solna node of the Cryo-EM Swedish National Facility funded by the Knut and Alice
Wallenberg, Family Erling Persson, and Kempe Foundations, SciLifeLab, Stockholm University, and
Umeå University.
Conflicts of Interest:
The authors declare no conflict of interest. The funders had no role in the design
of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or
in the decision to publish the results.
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