post-transcriptional regulation by rnases in streptococcus...
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Post-transcriptional regulation
by RNases
in Streptococcus pyogenes
Anne-Laure Lécrivain
Department of Molecular Biology
Umeå Center for Microbial Research (UCMR)
Laboratory for Molecular Infection Medicine Sweden (MIMS)
Um 2018
This work is protected by the Swedish Copyright Legislation (Act 1960:729)
Dissertation for PhD
ISBN: 978-91-7601-950-4
ISSN: 0346-6612
New series: 1993
Cover design: Anne-Laure Lécrivain
Electronic version available at: http://umu.diva-portal.org/
Printed by: Print & Media (Umeå University)
Umeå, Sweden 2018
To everyone who supported me
“C est ce que nous pensons déjà connaître
qui nous empêche souvent d apprendre”
Claude Bernard
(It is what we think we already know that often prevents us from learning)
i
Table of Contents
Table of Contents ........................................................................... i
Abstract ....................................................................................... iii
Abbreviations ............................................................................... v
Publications included in this thesis ............................................ vii
I- Introduction ............................................................................... 1
Post-transcriptional regulation of gene expression .................................................. 1
Ribonucleases .............................................................................................................3
RNase Y .................................................................................................................3
RNase III ............................................................................................................... 5
RNase J1 ............................................................................................................... 6
PNPase .................................................................................................................. 7
RNase R ................................................................................................................ 9
YhaM ................................................................................................................... 10
NanoRNases ....................................................................................................... 11
Interplay of RNases .................................................................................................. 11
RNA degradosome and other RNase complexes .............................................. 11
Models of RNA decay ......................................................................................... 13
RNA sequencing........................................................................................................ 14
General principle ................................................................................................ 14
RNA sequencing to identify ribonuclease targets ............................................ 14
Different methods to identify ribonuclease targets.......................................... 15
Streptococcus pyogenes ........................................................................................... 16
A human pathogen ............................................................................................. 16
Post-transcriptional regulation of gene expression in S. pyogenes ................ 18
II- Aims of the thesis .................................................................... 19
III- Results and discussion ......................................................... 20
Paper I ...................................................................................................................... 20
Methodology ...................................................................................................... 20
Characteristics of S. pyogenes RNase III .......................................................... 21
RNase III role in S. pyogenes ............................................................................ 22
ii
Paper II ..................................................................................................................... 24
Methodology ...................................................................................................... 24
YhaM has a global nibbling activity on transcript 3′ ends ............................ 25
Role and peculiarities of S. pyogenes YhaM .................................................... 26
PNPase role in RNA decay ................................................................................ 28
Limited RNase R activity .................................................................................. 29
Possible redundancy between RNase R and PNPase ...................................... 30
Paper III .................................................................................................................... 31
RNase Y cleaves after a guanosine.................................................................... 31
RNase Y and PNPase act in concert to degrade transcripts .......................... 33
Is RNase Y the major RNase initiating RNA decay?........................................35
The fate of transcript 5′ ends produced by RNase Y ....................................... 36
IV- Main findings of the thesis .................................................... 38
V- Acknowledgements ................................................................ 39
References ................................................................................... 41
iii
Abstract
Ribonucleases (RNases) are proteins that adjust cellular RNA levels by
processing RNA transcripts, leading to their stabilization or degradation.
RNases are grouped based on their ability to cleave the transcript internally
(endoRNases) or degrade the transcript starting from the ends (exoRNases).
Specificities of RNA degradation vary among bacterial species, attributable to
different sets of endo- and exoRNases. Most of the current knowledge gathered
about the roles of RNases and their targets relies on the study of a few model
bacteria, such as Escherichia coli and Bacillus subtilis. The aim of this thesis
was to understand how Streptococcus pyogenes, a strict human pathogen,
controls and adjusts gene expression by characterizing in vivo RNase activities.
The transcriptome of S. pyogenes was inspected to identify cleavages in vivo
performed by RNases of interest using RNA sequencing. For this purpose, we
developed a method to compare transcript 5′ and 3′ ends in RNase deletion
mutants with those in the parental strain. We first applied our method for the
study of endoRNase III, which cleaves ds RNA, and endoRNase Y, which is
specific for ss RNA. We accurately retrieved RNase III cleavage positions in
structured regions, characterized by 2 nucleotide (nt) 3′ overhangs, and we
showed RNase III nicking activity in vivo. We observed that RNase Y processed
transcripts after a guanosine. The upstream and downstream fragments
generated by a single cleavage event were never both identified, indicating that
RNase Y processing always led to the degradation of one of the two fragments.
To investigate further the degradation of the upstream fragment subsequent to
RNase Y processing, we characterized the 3′-to-5′ exoRNases R, YhaM, and
PNPase. RNase R did not have any detectable activity in standard laboratory
conditions. YhaM is an intriguing enzyme that removed on average 3 nt of the
majority of cellular transcripts. PNPase fully degraded fragments originating
from endoRNase processing and is the main 3′-to-5′ exoRNase involved in RNA
decay in S. pyogenes.
To conclude, in this work, we developed a novel method to analyze RNA
sequencing data. This method was successfully applied to the study of both
iv
endo- and exoRNases. Most importantly, we identified the targetomes of
RNases III, Y, R, YhaM, and PNPase and we highlighted the distinctive features
of these enzymes.
v
Abbreviations
5′ OH 5′ hydroxyl
5′ P 5′ monophosphate
5′ PPP 5′ triphosphate
cDNA complementary DNA
CRISPR clustered regularly interspaced short palindromic
repeats
crRNA CRISPR RNA
DNA deoxyribonucleic acid
DNase deoxyribonuclease
ds double-stranded
endoRNase endoribonuclease
exoRNase exoribonuclease
FDR false rate discovery
G guanosine
mRNA messenger ribonucleic acid
NAD 5′- nicotinamide-adenine dinucleotide
nt nucleotide
ORF open reading frame
PAP I poly(A) polymerase I
PFK phosphofructokinase
Pi inorganic phosphate
PNK polynucleotidekinase
poly(A) polyadenine
poly(N) polynucleotide
RBS ribosome binding site
RNA ribonucleic acid
RNase ribonuclease
rRNA ribosomal RNA
SAM S-adenosyl-L-methionine
sRNA small RNA
vi
ss single-stranded
tmRNA transfer messenger RNA
tracrRNA trans-activating CRISPR RNA
TRAP trp RNA-binding attenuation protein
tRNA transfer RNA
TSS transcriptional start site
UTR untranslated region
WT wild type
∆rnase RNase deletion mutant
vii
Publications included in this thesis
Paper I
Le Rhun A.*, Lécrivain A.-L.*, Reimegård J., Proux-Wera E., Broglia L., Della
Beffa C., and Charpentier E. (2017). Identification of endoribonuclease specific
cleavage positions reveals novel targets of RNase III in Streptococcus pyogenes.
Nucleic Acids Research. 45:(5), 2329-2340. *Contributed equally.
doi: 10.1093/nar/gkw1316
Paper II
Lécrivain A.-L.*, Le Rhun A*, Renault T. T., Ahmed-Begrich R., Hahnke K., and
Charpentier E. (2018). The in vivo 3′-to-5′ exoribonuclease targetomes of
Streptococcus pyogenes. PNAS. *Contributed equally.
doi: 10.1073/pnas.1809663115
Paper III
Lécrivain A.-L., Broglia L., Renault T. T., Hahnke K., Ahmed-Begrich R., Le
Rhun A. and Charpentier E. Interplay between 3′-to-5′ exoRNases and RNase Y
in Streptococcus pyogenes. Manuscript.
Published but not included in this thesis
Fonfara I.*, Le Rhun A.*, Chylinski K., Makarova K. S., Lécrivain A.-L., Bzdenga
J., Koonin E. V. and Charpentier E. (2014). Phylogeny of Cas9 determines
functional exchangeability of dual-RNA and Cas9 among orthologous type II
CRISPR-Cas systems. Nucleic Acids Research. 42:4, 2577-2590. *Contributed
equally. doi: 10.1093/nar/gkt1074
Broglia L., Materne S., Lécrivain A.-L., Hahnke K., Le Rhun A. and Charpentier
E. (2018). RNase Y-mediated regulation of the streptococcal pyrogenic exotoxin
B. RNA Biology. doi: 10.1080/15476286.2018.1532253.
1
I- Introduction
In order to survive, every organism needs to adapt to multiple changes in
their environment, protect itself from predators and fight various threats. At the
microscopic scale, this translates into adjusting gene expression spatially and
temporally to regulate the levels of RNAs and proteins in the cell. The central
dogma describes the transfer of genetic information, from deoxyribonucleic acid
(DNA) to protein (1): DNA that composes the gene is first transcribed into
messenger ribonucleic acid (mRNA), which is then translated into a protein.
However, RNA – compRising all classes of mRNA, transfer RNA (tRNA),
ribosomal RNA (rRNA) and small RNA (sRNA) − is not merely an intermediate
between DNA and proteins but can alter the information delivered by the DNA
and can have functions on its own (2). Indeed, RNA has the ability to catalyze
enzymatic reactions, to be reverse transcribed into DNA, to be modified and to
control gene expression (2). Therefore, bacteria have developed many ways to
regulate gene expression at the RNA level, also called post-transcriptional
regulation.
Post-transcriptional regulation of gene expression
Post-transcriptional regulation in prokaryotes occurs at the level of
transcriptional elongation, RNA stability and translational initiation (3). Among
the elements that control gene expression without affecting mRNA stability, the
riboswitches are regulatory elements located in the 5′ UTRs of the transcripts
that they regulate. They directly sense environmental changes by binding
ligands, such as uncharged tRNA or S-adenosyl-L-methionine (SAM), or by
sensing temperature changes (4–6). The binding of the ligand triggers the
formation of a terminator or anti terminator hairpin (transcriptional
elongation), or it sequesters or free the ribosome binding site (RBS)
(translation). For more information, refer to the recent review (7).
This section focuses on how a bacterium controls protein expression through
RNA maturation and degradation, affecting mRNA stability. Many factors are
2
involved but eventually the ribonucleases (RNases) act as the final agents to
clear unwanted RNA from the cells. RNA stability refers to the time a transcript
remains in the cell before it is being degraded (chemical stability) or inactivated
(functional stability) (8, 9). For instance, the functional stability of an mRNA is
defined by the time it undergoes translation and its chemical stability is the time
required for RNases to degrade it (8). Therefore, RNA stability depends on
intrinsic characteristics of the transcript, RNA-binding proteins, and sRNAs; all
further described below.
Each transcript has characteristics that will affect its half-life in the cell. The
5′ end of a primary transcript harbors a triphosphate nucleotide (5′ PPP) or a
5′- nicotinamide-adenine dinucleotide (NAD) cap that blocks endoribonucleases
(exoRNases) and 5′-to-3′ exoRNases (10–14). The sequence of a transcript
controls its targeting by proteins such as decapping enzymes or by sRNAs (15,
16). Structural elements at the transcript 5′ end (e.g. stem-loop or RNA duplex)
and at the 3′ end (e.g. transcriptional terminators) can efficiently prevent the
transcript degradation by exoRNases (17–21). However, these RNA duplexes
can also promote processing by double-stranded (ds) specific endoRNases (22).
RNA-binding molecules can interfere with the stability of transcripts by
preventing or promoting the access of RNases to their binding. RNA helicases
unwind structured transcripts and thereby promote degradation (23).
Polyadenylation performed by the poly(A) polymerase I (PAP I) at the 3′ ends of
transcripts promotes exoribonucleolytic degradation (24–26). Conversely, the
host factor I protein Hfq protects transcripts by binding polyadenine (poly(A))
tails (27). In addition, Hfq promotes the interaction of sRNAs to their mRNA
targets, which can result either in protection or triggering of RNA decay (28).
Ribosomes protect mRNA during translation initiation by binding the RBS (29–
31) and during translation elongation (32, 33). For more detail about the
influence and mechanisms of RNA-binding proteins on RNA stability, refer to
(3, 34).
sRNAs bind mRNAs and can stabilize them by masking an RNase binding-
site or by promoting RNase processing (35–39). Alternatively, sRNAs
destabilize their targets by triggering degradation, either directly by RNase
3
recruitment or indirectly by blocking translation (40, 41). For more information
about the mechanisms involved, refer to (16, 42).
The final players are the RNases, which catalyze the cleavage of a
phosphodiester bond in a transcript. This leads to the degradation of the
substrate or, conversely, to the maturation of stable RNAs, such as rRNAs and
tRNAs (43, 44).
Ribonucleases
Ribonucleases are classified in endoRNases, which cleave in the body of a
transcript, and in exoRNases, which degrade a transcript starting from the 5′
end (5′-to-3′ exoRNase) or from the 3′ end (3′-to-5′ exoRNase). Some RNases
are phosphorolytic, i.e. they use inorganic phosphate (Pi) to break the
phosphodiester bond and release a nucleoside diphosphate, and others are
hydrolytic enzymes, i.e. they use a water molecule to break the bond and release
a nucleoside monophosphate.
This section focuses on the main RNases that regulate gene expression
and/or perform RNA decay and are present in S. pyogenes. Following is an
overview of the current general knowledge about RNase Y and RNase III
(endoRNases), RNase J1 (5′-to-3′ exoRNase), and YhaM, PNPase, RNase R, and
nanoRNase (3′-to-5′ exoRNases).
RNase Y
The single-stranded (ss) specific RNase Y is a dimeric enzyme that
preferentially targets transcripts with a 5′ monophosphate (5′ P) end (45, 46).
RNase Y is described as the main endoRNase initiating RNA decay in Gram-
positive bacteria such as B. subtilis, Clostridium perfringens and S. pyogenes
(39, 47–50), but has a smaller effect on global RNA stability and abundance in
Staphylococcus aureus (51, 52). This enzyme is involved in sRNA-mediated
RNA decay (39, 53) and in the degradation of 5′ UTR regulatory elements and
sRNAs (45, 47, 48, 51, 54, 55). Notably, RNase Y influences virulence gene
expression and the pathogenicity of S. aureus, S. pyogenes and C. perfringens
(39, 50, 56–60). In S. pyogenes, many virulence genes are controlled by
4
RNase Y in a glucose-restricted medium (57). Some of these genes are likely
affected by an indirect transcriptional regulation. For example, our laboratory
recently showed that the RNase Y-dependent regulation of the speB transcript
abundance was mainly indirect (60).
RNase Y is located at the bacterial membrane via a transmembrane domain
(45, 61–63); however this does not seem to restrict RNase Y target selection in
vivo (51, 64). RNase Y orthologues present variable RNA structure and sequence
specificities. In S. pyogenes, a guanosine (G) is required for the in vivo
processing of the speB transcript by RNase Y (60). The presence of a G was also
observed directly upstream of 58% of RNase Y processing sites identified in S.
aureus (51) but it was not necessary for the processing of the saePQRS
transcript (65). In this bacterium, RNase Y activity is instead controlled by a
structured element located 6 nucleotides (nt) downstream of the processing site
(65). RNA duplexes and riboswitches were also observed around RNase Y
processing sites in B. subtilis and C. perfringens (39, 40, 45). These structured
regions were not detected in B. subtilis and S. aureus genome-wide studies of
RNase Y processing sites (51, 66). It should be noted that intermolecular
pairings (e.g. sRNA-mRNA duplexes) and complex structured regions are very
difficult to predict on a global scale. Last but not least, RNase Y interacts with
other proteins (see „RNA degradosome and other RNase complexes“ in this
thesis). For instance, the Y-complex – composed of the proteins YlbF, YmcA,
and YaaT – interacts in vivo with RNase Y and is required for the processing of
several polycistronic transcripts in B. subtilis (66, 67). Interestingly, the Y-
complex is not necessary for RNase Y activity but rather is an accessory factor
that potentially directs RNase Y towards specific transcripts (66).
In B. subtilis, RNase Y expression is growth-phase dependent (49). RNase Y
regulates other RNases; in B. subtilis, RNase Y matures RNase P RNA (44) and
potentially regulates its own gene expression and that of other RNases in S.
aureus (51).
5
RNase III
The dimeric enzyme RNase III cleaves specifically in ds RNA, generating
products with a characteristic 2 nt 3′ end overhang (68). RNase III activity
mostly depends on structural motifs in the target RNA, as presented below.
Some reports have shown that this enzyme is able to cleave only one of the two
strands (i.e. nick) when an internal bulge or loop is present (69–72). The
biological effect of the nicking is illustrated by the study of the bacteriophage T7
transcript (73). The authors inserted a T7 stem-loop nicked by RNase III in a
human gene and showed that the in vivo mRNA stability was increased
compared to the wild type (WT) mRNA (73). Other specific motifs, such as a
bulge–helix–bulge on the dsRNA, allow RNase III to bind but prevent further
processing (70). In vivo, this leads to the stabilization of transcripts (74, 75) or
could activate translation initiation (70, 76, 77). These examples highlight the
various ways by which RNase III regulates gene expression by acting at the post-
transcriptional or translational level.
In addition to maturing the 16S and 23S rRNAs (78), RNase III participates
in the control of gene expression. Studies in E. coli and B. subtilis show that
RNase III affects approximatively 11-12% of the transcriptome (47, 79) and
many processing and binding sites have been identified genome-wide (80–82).
Due to its ability to process RNA duplexes, RNase III is involved in clearing
antisense transcripts generated from pervasive transcription in S. aureus (82,
83), in sRNA processing, and sRNA-mediated gene regulation in many bacteria
(82, 84–87). In B. subtilis, RNase III is essential due to the expression of
prophage-encoded toxins whose transcripts are degraded by the enzyme (40).
Moreover, the enzyme was shown to regulate directly translation initiation in
vivo (82, 88).
In many bacterial species, the intracellular level of RNase III is negatively
autoregulated, via RNase III processing its own transcript (rnc) leading to rnc
decay (80, 82, 89, 90). In addition, the enzymes activity is regulated at the post-
translational level. For instance, in E. coli, RNase III is bound by the protein
YmdB, which correlates with an increase in RNase III activity (91, 92).
RNase III activity is also hijacked by the T7 bacteriophage. The phage produces
6
a kinase that enhances RNase III activity to increase the maturation of T7
transcripts (93).
RNase J1
RNase J1 is the only bacterial 5′-to-3′ exoRNase described so far and is
mostly present in Gram-positive bacteria (94, 95). This relatively recent
discovery – the 5′-to-3′ ribonucleolytic activity was described in 2007– had a
huge impact in the field, as it opened the possibility of a degradation pathway
starting at the 5′ ends of transcripts, like in eukaryotes (96). An
endoribonucleolytic activity was also described in vitro for RNase J1 (95),
although the in vivo relevance of this activity is now questioned (97). Some
bacteria also encode a paralogue of RNase J1, named RNase J2 (95). In contrast
to RNase J1, RNase J2 mostly has an endoribonucleolytic activity in vitro (94).
RNases J1 and J2 form heteroduplexes (95, 98), which alter the in vitro
cleavage pattern in comparison to the one of each individual enzyme (98). In
vivo, global changes in RNA and protein abundances were observed between
the corresponding single and double deletion strains (99). RNase J1 degrades 5′
P and 5′ hydroxyl (5′ OH) transcripts (94, 95), hence it is likely to rely on the
activity of RppH or endoRNases that initiate the degradation of transcripts (11,
12, 29). In addition, the enzyme requires at least 9 unstructured nucleotides at
the 5′ end to access its substrate and to degrade it (94, 100).
The deletion of RNase J1 affected approximatively 30% of B. subtilis
transcriptome and more than 80% of the Gram-negative Helicobacter pylori
transcriptome (47, 101), highlighting the central role of this enzyme in RNA
degradation. RNase J1 decays transcripts at their 5′ end upon removal of the 5′
PPP by RppH (11) or participates in clearing the decay intermediate fragments
that are generated by endonuclease cleavages (47, 51, 102, 103). Particularly,
RNase J1 is involved in the degradation of decay intermediate fragments
originating from transcript 3′-proximal ends, which contain transcriptional
terminators (80, 102, 103). Interestingly, the 30S ribosomal subunits present on
the 5′ UTRs during translation initiation block RNase J1 5′-to-3′
exoribonucleolytic activity and stabilize the transcripts (38, 94, 100, 104, 105).
7
The role of RNase J2 is more obscure. In B. subtilis and S. aureus, it does not
seem to have any activity and is rather a structural support for RNase J1 (99,
106, 107). However, in Streptococcus mutans, RNase J2 processes the gbpC
transcript in vivo (37) and the deletion of Enterococcus faecalis RNase J2
influenced RNA abundance independently of RNase J1 (108).
While RNases J1 and J2 are dispensable in B. subtilis, S. aureus and S.
mutans (106, 107, 109), both proteins are essential in S. pyogenes (110).
PNPase
Polynucleotide phosphorylase, or PNPase, is a phosphorolytic enzyme that
has two functions. It uses Pi to degrade RNA in a 3′-to-5′ fashion, a reaction
that releases nucleoside diphosphates, and it also performs the reverse reaction,
i.e. the polymerization of nucleoside diphosphates without requiring a template
or a primer (111, 112). In vivo, however, PNPase behaves mostly as a degradative
enzyme and is thought to use its polymerase activity to add polynucleotide
(poly(N)) tails to transcript 3′ ends in E. coli (113–115). In B. subtilis, the
addition of poly(N) is not a major role of PNPase (116).
PNPase is present in bacteria, chloroplasts, and in mitochondria, but not in
archaea and yeast (117). PNPase is a trimeric enzyme that forms a ring-like
structure, with the catalytic site buried in the channel, and three RNA-binding
domains, which are presented in front of the channel entry (118, 119). The
distance between the RNA-binding domains and the catalytic sites determines
PNPase binding to ss tails of RNA of at least 7 nt (118, 120, 121). PNPase is
generally blocked by secondary structures such as intrinsic transcriptional
terminators (18, 122, 123). To overcome this obstacle, PNPase can use (i)
repeated rounds of tail polymerization at transcript 3′ ends by PAP I, or
allegedly by PNPase itself (124–127), (ii) the help of RNA-unwinding partners,
such as RNA helicases (126, 128–130), or (iii) the help of an endoRNase that
removes the structure (103, 122).
In B. subtilis and S. aureus, RNA decay is the major function of PNPase (34,
114, 131). In E. coli, PNPase is involved in RNA decay together with another 3′-
to-5′ exoRNase, RNase II (113, 132, 133). Despite this important role, the
8
deletion of PNPase only mildly affects bulk RNA stability (133, 134), which
indicates that there is redundancy between PNPase and the other exoRNases to
degrade transcripts. PNPase principally targets decay intermediate fragments
and degrades them very efficiently, since these fragments are not detectable in
vivo (131, 132, 135, 136). A small fraction of PNPase is bound to the E. coli RNA
degradosome (137, 138), a multiprotein complex organized around the
endoRNase E, where it participates in mRNA and sRNA degradation (see „RNA
degradosome and other RNase complexes“ in this thesis).
In parallel, PNPase acts as a regulatory protein in vivo. In E. coli, the
exoRNase has an important protective role for sRNAs that are dependent on
Hfq (139–141). Conversely, sRNAs independent of Hfq are degraded by PNPase
(139, 140, 142). PNPase matures RliB, a transcript originating from a peculiar
Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) array
that protects Listeria monocytogenes against invading plasmids (143). In B.
subtilis, PNPase matures the SR5 antisense RNA that is part of the toxin-
antitoxin system bsrE/SR5 (144) and specifically degrades the trp (tryptophan
operon) leader when it is bound to TRAP (trp RNA-binding attenuation
protein), resulting in the recycling of TRAP (145, 146).
As expected, the deletion of PNPase is associated with various phenotypes
(e.g. reduced survival at cold temperatures, changes in bacterial morphology
and biofilm formation) and affects bacterial competence or DNA damage repair,
among many cellular processes (147).
The expression and the activity of PNPase are tightly controlled. The
mechanisms vary from one bacterium to another and, to keep this section
simple, only a global overview is presented here. At the post-transcriptional
level, RNase III and PNPase act in concert to keep PNPase expression at a basal
level by processing the pnpA (coding for PNPase) 5′ UTR (148, 149). The master
carbon storage regulator (CsrA) prevents the processed pnpA transcript from
being translated (150). pnpA stability and translation is repressed by the
antisense RNA SraG (151). Additionally, PNPase binds its transcript in an
RNase III-independent manner and prevents its translation (152). PNPase
responds to cellular metabolites, such as ATP, magnesium-chelated citrate, or
ppGpp that activates or represses PNPase depending on the bacteria studied
9
(153–155). During adaptation to cold, PNPase levels in the cell increases (156–
158).
In S. pyogenes, a study reported that PNPase destabilizes a few transcripts
coding for virulence factors (159).
RNase R
RNase R is a hydrolytic protein that belongs to the RNR family, compRising
unspecific 3′-to-5′ exoribonucleolytic enzymes (160). RNase R is a processive
enzyme, meaning that it stays bound to the substrate during the complete
degradative process (121, 161).
A particularity of RNase R is its ATP-independent helicase activity that
allows the efficient degradation of structured fragments (162–165). The
biological importance of the helicase activity is highlighted by the fact that
RNase R rescues the lethal deletion of the helicase CsdA in E. coli independently
of its RNase activity (162).
Like PNPase, RNase R needs a ss tail at the substrate 3′ end to bind and
degrade it (166–168). While 7 nt is a minimum, E. coli RNase R has the best in
vitro degradation rate with 10 nt (163, 168). The composition of the tail is
important, as RNase R prefers poly(A) tails to poly(U) tails downstream of a
structured RNA and does not efficiently bind poly(G) RNA or generally
substrates with a high G content (163, 168).
RNase R is mostly known for participating in quality control and maturation
of rRNAs (169–171) and transfer messenger RNA (tmRNA or SsrA RNA) (172).
tmRNA is involved in the trans-translation system that rescues ribosomes
stalled on mRNA lacking a stop codon and RNase R clears the cells from these
defective transcripts (172, 173). RNase R plays a role in mRNA decay by
degrading structured RNA fragments (136, 167) and transcripts (174, 175).
However, the deletion of RNase R has little impact on RNA abundance in H.
pylori, Pseudomonas putida, and in E. coli (176–178). In addition, the deletion
of RNase R together with other 3′-to-5′ exoRNases does not increase bulk RNA
stability in B. subtilis (179), suggesting a minor role of the protein in RNA decay.
The importance of RNase R becomes obvious during bacterial adaptation to cold
10
shock, where global mRNA stabilization was observed in absence of RNase R
(180).
In E. coli, RNase R expression and activity are tightly controlled at multiple
levels. Protein abundance and/or activity are increased at elevated temperature
(175), upon cold shock, at the entry in the stationary phase of growth and under
starvation (172, 181). RNase R is normally unstable and the regulation of
expression described above occurs at the post-translational level (182). Indeed,
in absence of acetylation, RNase R interacts less tightly with tmRNA/SmpB
(trans-translation system actors). The weak interaction with tmRNA prevents
RNase R from being targeted by proteases (183–186). In addition, during the
exponential phase of growth, RNase R is mostly bound to the ribosomes, and
free RNase R is degraded to prevent unwanted global RNA degradation (187).
YhaM
YhaM has only been identified so far in Gram-positive bacteria (179). B.
subtilis YhaM efficiently degrades nanoRNA (5-mer) and long ss RNA regions in
vitro (179, 188), but is blocked by secondary structures (179). The S. aureus
orthologue of YhaM, Cbf1, is also able to degrade RNA in vitro, although less
efficiently than B. subtilis YhaM (179). The ribonucleolytic activity of YhaM was
detected in vivo as well (136, 189).
The physiological role of YhaM is unclear. B. subtilis YhaM restored the
viability of E. coli lacking the oligoRNase Orn, which is responsible for
degrading nanoRNAs produced by exoRNases (see “NanoRNases” below) (188).
Bulk RNA stability was not significantly affected when YhaM was removed from
B. subtilis (179) and the study of specific transcripts showed a very limited
participation in RNA decay (136). Interestingly, this enzyme is linked to DNA
replication and repair. Indeed, Cbf1 binds the replication enhancer of the
plasmid pT181 (190) and interacts with DnaC, a helicase involved in DNA
replication (191). B. subtilis YhaM presents a 3'-to-5' exoDNase activity in vitro
(179) and the expression of the protein is induced by UV or mitomycin C (DNA
damaging agent) in a RecA-dependent manner (192).
11
Little is known about the regulation of YhaM. The enzyme is degraded when
B. subtilis enters the stationary phase of growth (193). In B. subtilis, the YhaM
promoter is bound by the carbon catabolite transcriptional repressor CcpN
(194) and harbors a binding site for the transcriptional repressor LexA (192).
NanoRNases
The nanoRNases NrnA, NrnB in Gram-positive bacteria and Orn in E. coli
are enzymes that degrade nanoRNAs into mononucleotides, an essential process
to replenish the pool of nucleotides available (188, 195–197). These nanoRNAs
(2 - 5 nt long) are usually produced by the exoRNases, such as PNPase and
RNase R, at the end of the RNA degradation process (196).
Interplay of RNases
How do organisms ensure the efficient and coordinated action of different
actors in crowed cellular environments? The compartmentalization of processes
is very well known in eukaryotic cells and is allowed by the presence of
organelles that concentrate the different proteins required for a same process
(e.g. the DNA replication or transcription machineries located in the nucleus).
Some prokaryotic organelles have been described, usually for very specific
processes (198). However, absence of organelles does not mean absence of
compartmentalization. In all organisms, proteins assemble in complex
machineries that gather distinct enzymatic activities in one location and
bacteria are not an exception. For instance in E. coli, the RNA degradosome is a
complex composed of an endoRNase, a 3′-to-5′ exoRNase, an RNA helicase and
a glycolytic enzyme. The characteristics of the RNA degradosome, as well as
other complexes described in Gram-positive bacteria, and its role in mRNA
decay is described below.
RNA degradosome and other RNase complexes
The RNA degradosome in E. coli and other Gram-negative bacteria is very
well studied. It is an assembly of multiple proteins that is involved in bulk
mRNA decay (137, 199). The central protein, RNase E, is generally anchored at
12
the bacterial membrane through an amphipathic helix (200, 201). This enzyme
is essential (202) and it is presumably due to its importance for several major
processes, such as stable RNA maturation and the initiation of mRNA decay
(203–205). In E. coli, the C-terminal domain of RNase E interacts with the
other proteins from the degradosome, which are PNPase, the RNA helicase
RhlB and the glycolytic enzyme enolase (206). The composition of the
degradosome is dynamic and, under specific growth conditions, it involves other
partners, for instance Hfq or PAP I (207). The degradosome is not necessary for
bulk mRNA decay in E. coli, however this association between an endoRNase, a
3′-to-5′ exoRNase and an RNA helicase facilitates the degradation of structured
or otherwise inaccessible transcripts (206).
In Gram-positive bacteria, the existence of a stable RNA degradosome is
subject to debate (reviewed in (208, 209)). Several RNases have been shown to
interact with other RNases, RNA helicases, and glycolytic enzymes (see below).
Early studies proposed that B. subtilis RNase Y acted as the scaffold protein and
the central endoRNase in this complex (46, 210, 211). Multiple interactions
between RNase Y, PNPase, RNases J1/J2, the RNase RnpA, the RNA helicase
CshA, and the glycolytic enzymes enolase and phosphofructokinase (PFK) were
reported in B. subtilis and S. aureus (210–212). Some were contradicted by
other reports, such as the in vivo interaction between RNase Y and
RNases J1/J2 (94) and other were confirmed by in vitro techniques (213). In
addition, a recent in vivo localization study showed that PNPase, RNases J1/J2,
enolase, PFK, and CshA do not localize to the B. subtilis membrane, where
RNase Y is anchored (62). Many investigations were done on the binary
interactions between these enzymes. For instance, in B. subtilis, the glycolytic
enzyme glyceraldehyde 3-phosphate dehydrogenase (GapA) binds to RNase J1
and to RNase Y (214). In H. pylori and S. aureus, RNase J1 forms a complex
with an RNA helicase (215, 216). In S. pyogenes, RNase Y interacts with enolase
(57). In S. epidermidis, PNPase binds RNase J1 (217). In addition, RNase Y
interacts with the Y-complex in B. subtilis (see “RNase Y” in this thesis). In
conclusion, these studies suggest that the interactions between RNases,
helicases, and glycolytic enzymes are transient and unlikely to happen
altogether in vivo.
13
Models of RNA decay
A general model of mRNA decay was proposed based on studies using E. coli,
which expresses endoRNases and 3′-to-5′ exoRNases (218). This model has
been updated with the numerous discoveries made over the years and accounts
for the different sets of RNases expressed in Gram-negative and Gram-positive
bacteria (219). The two main degradation pathways in E. coli (representative of
Gram-negative bacteria) and in B. subtilis (representative of Gram-positive
bacteria) are presented below.
In E. coli, the major endoRNase responsible for mRNA decay is RNase E
(206) and two main pathways have been described: the 5′ P-dependent pathway
and the direct entry pathway, each of them initiated by a different enzyme. In
the 5′ P-dependent pathway, the 5′ PPP present on primary transcripts is
removed sequentially by a yet unidentified enzyme and the RNA
pyrophosphohydrolase RppH (10, 220–222). This event triggers RNase E
cleavage in the transcript (10, 220). RNase E processing generates an upstream
decay intermediate fragment, which is not protected by a 3′ end structure and is
therefore degraded by 3′-to-5′ exoRNases; and a downstream decay
intermediate fragment with a 5′ P end. Because RNase E has a strong preference
for 5′ P transcripts, the downstream fragment becomes in turn a substrate for
RNase E that repeats the processing several times on each downstream decay
intermediate fragment generated (223, 224). The 5′ P-independent pathway,
also called “direct entry” pathway, relies on the ability of RNase E to bypass the
5′ P requirement and process a transcript with a 5′ PPP end (225–227). The
final steps of transcript degradation are performed by the two major 3′-to-5′
exoRNases: the hydrolase RNase II and PNPase (113, 132, 133).
In B. subtilis, which expresses the 5’-to-3’ exoRNase J1, two different
pathways are known and they differ at the initiation step. The first pathway
relies on an endoRNase that initiate the degradation of a primary transcript,
generating an upstream and a downstream decay intermediate fragment. The
initiator is mainly RNase Y, and in some cases RNase III (16, 228). The
upstream fragment is degraded by 3’-to-5’ exoRNases, mainly PNPase and
RNase R, and the downstream fragment is degraded by RNases J1/J2. The
14
second pathway is initiated by RppH. The enzyme converts the transcript 5′ PPP
end in a 5′ P end in two steps, thereby making the transcript available for
RNase J1 exoribonucleolytic activity (11, 15).
For both E. coli and B. subtilis, mRNA degradation is an “all-or-none decay”
process (131, 229). Indeed, the decay intermediate fragments are only visible in
the absence of exoRNases, as shown in the global study of Liu et al. (131).
RNA sequencing
General principle
RNA sequencing relies on the reverse transcription of RNA into
complementary DNAs (cDNAs), which will be then sequenced (230). Many RNA
sequencing methods have been developed to answer specific scientific
questions, recently reviewed in (231) and (232). This section presents the global
workflow used in this thesis, which is based on the RNA sequencing method of
Dotsch et al. (233).
RNA sequencing to identify ribonuclease targets
First, the RNA samples are prepared for the ligation of the adapters. Total
RNA is extracted, DNase-treated and depleted from rRNAs. At this stage,
different treatments allow the selection of different transcripts. For instance, the
treatment with Tobacco Acid Pyrophosphatase (TAP) that converts 5′ PPP to 5′
P ensures the ligation of both primary and processed transcripts. Conversely, in
the absence of TAP treatment, only processed transcripts with a 5′ P are ligated.
The transcripts are mechanically fragmented before the treatment with T4
polynucleotidekinase (PNK), which converts 5′ OH to 5′ P.
The second step consists of preparing the cDNA library. The RNA fragments
are ligated to 5′ adapters (ligated at fragment 3′ ends) and to 3′ adapters
(ligated at fragment 5′ ends). The use of two adapters allows the identification
of the DNA strand of origin (sense or antisense). A primer that binds the 5′
adapter is used for the reverse transcription. Finally, the cDNA libraries are
sequenced.
15
The last step is the analysis. The sequencing outputs are called “reads”,
which include the transcript sequences. The reads undergo quality control check
and adaptor sequence removal before being mapped to the corresponding
genome. The mapping is visualized as “coverage” of reads over the genome. The
“total coverage” represents the number of reads that cover each nucleotide of
the genome. In addition, depending on the conditions of library preparation and
the RNA sequencing parameters, it is possible to calculate the “read end
coverage”. This corresponds to the number of reads that start (5′ end) or stop
(3′ end) at each nucleotide on the genome, which define the transcript
boundaries.
To compare the gene expression between wild type (WT) and RNase deletion
mutant (∆rnase), the differential expression analysis is done with the
normalized total coverages using edgeR (234) or DEseq2 (235). To determine
the position of a processing, the differential expression analysis is done with the
end coverage between WT and ∆rnase.
Different methods to identify ribonuclease targets
To have a glance at the global impact of RNases in vivo, the simplest method
is to compare global RNA steady-state levels in WT and ∆rnase − i.e. differential
expression analysis. However, the distinction between transcriptional regulation
(i.e. indirect effect) and post-transcriptional regulation (i.e. direct effect) is not
possible with this analysis.
Another method is to arrest the cellular transcription and follow the stability
of transcripts over time and genome-wide. The stability of each mRNA as well as
the global mRNA stability can be calculated. For instance, the median mRNA
half-lives in Bacillus cereus and S. pyogenes were estimated at 2.4-2.6 min and
0.89 min, respectively, and the average mRNA half-life in E. coli was estimated
at 2.5 min (50, 236, 237). The deletion of RNase Y in S. pyogenes increased the
median mRNA half-life to 1.81 min, due to the stabilization of the majority of
the transcripts (50).
During the last years, different techniques have been developed to identify
more accurately the mRNA bound or processed by the RNase of interest. Some
16
techniques were based on the selection of particular transcripts (structure or
nature of the transcript end) and some examples are described below. A
catalytically inactive variant of RNase III was used to isolate the dsRNA bound
to the enzyme in vivo (82). Alternatively, RNA duplexes were
immunoprecipitated with J2 antibody from WT and RNase III deletion strains,
which revealed a major role of RNase III on degrading dsRNA in E. coli (84).
The ability of metal-ion independent RNases to produce fragments with a 2′, 3′-
cyclic phosphate 3′ end was exploited to determine the transcripts processed by
RNase L (238).
In addition to identify the transcripts targeted by an RNase of interest, the
mapping of transcript ends allows the determination of the exact cleavage
positions (i.e. targetome). This is done by comparing the ends present in WT
and absent in ∆rnase strains. For instance, the mapping of transcript 5′ ends
pinpointed the exact cleavage sites of RNase J1 (5′-to-3′ exoribonucleolytic
activity) (106), RNase Y (51, 66), RNase III (80, 239), RNase E (225, 240) and of
the toxin MazF (241).
The mapping of the transcript 3′ ends has been principally used to study
operon architectures and specificities of transcriptional termination (242–246).
Streptococcus pyogenes
Streptococcus pyogenes (also called GAS, for Group A Streptococcus) is a
non-motile, non-spore forming bacteria that grows in chains. It belongs to the
Firmicutes phylum, the Bacilli class − like B. subtilis and S. aureus, two
bacterial species often mentioned in this thesis − and the Lactobacillales order.
The strains are classified into serotypes according to the sequence of the emm
gene, which encodes a virulence factor called the M protein (247).
A human pathogen
S. pyogenes is a strict human pathogen that is responsible for one of the
widest range of human diseases (248). S. pyogenes is transmitted via direct
17
contact with a contaminated person or, very rarely, via contaminated food
(249).
Commonly, the bacteria infect the skin and the throat, leading to superficial
conditions (e.g. impetigo or pharyngitis) or to more severe diseases (e.g. scarlet
fever). S. pyogenes can progress deeper in the tissues and cause erysipelas,
cellulitis and necrotizing fasciitis. This last disease gave S. pyogenes the
nickname of “flesh-eating bacteria”. The bacteria are able to spread into the
bloodstream, leading to the streptococcal toxic shock syndrome (STSS), a life-
threatening condition. Due to cross-reactivity of some antibodies developed
against GAS during the infection, several complications can occur after the
bacteria have been efficiently eliminated (250). The most common ones are the
acute post-streptococcal glomerulonephritis (inflammation of the kidneys), the
acute rheumatic fever (mainly inflammation of the joints and heart) and the
rheumatic heart disease (inflammation of the heart).
S. pyogenes is responsible for more than 700 million cases of impetigo and
pharyngeal infections (251) and more than half a million deaths per year
worldwide (252). The treatment of choice is ampicillin, as no resistance to the
antibiotic has ever been developed by S. pyogenes (253). For patients who are
allergic to ampicillin, or in case of deep infections, macrolides are used but the
resistance rates to this molecules are increasing (254). The development of a
vaccine has proven to be challenging, due to the lack of an invariant and safe
antigen, but not impossible, as some vaccines are being evaluated in human
clinical trials (255).
S. pyogenes expresses many virulence factors to successfully establish an
infection (250). For instance, adhesins allow the bacterium to adhere to host
cells (256). SpeB, an unspecific protease that degrades both human and
bacterial proteins, promotes S. pyogenes dissemination (257, 258). To protect
itself against the immune response of the host, the bacterium produces
peptidases that degrade antibodies or a hyaluronic acid capsule that prevents
phagocytosis (259, 260). In addition, S. pyogenes produces toxins, such as
Streptolysin S, which create pores in the host cells (261).
18
Post-transcriptional regulation of gene expression in S.
pyogenes
The bacterial strain studied in this thesis is S. pyogenes SF370 serotype M1
(ATCC 700294), which was originally isolated from a wound infection (262).
The S. pyogenes chromosome has a size of 1.8 Mb and a low GC content of
38.6% (262, 263).
The study of the post-transcriptional regulation of gene expression in S.
pyogenes focused mainly on the sRNA-mediated regulation and the discovery of
new sRNAs (85, 264, 265). The role of RNases was investigated, in particular of
RNase Y, RNases J1/J2, and PNPase (refer to the corresponding sections in this
thesis). Based on the studies of RNases J1/J2 and PNPase, a model of mRNA
decay dependent on the phase of growth was proposed (110, 159, 266).
According to this model, the transcripts are separated in two classes: the Class I,
which compRises transcripts detectable in exponential phase of growth, and the
Class II, which compRises transcripts mostly detected in stationary phase of
growth (159). Class I transcripts are short-lived, whereas Class II transcripts are
very stable and present a biphasic degradation pattern (159). RNases J1/J2 are
responsible for initiating the decay of transcripts from the two classes, with a
preference for Class I (110). Therefore, in stationary phase of growth,
RNases J1/J2 start to degrade the Class II transcripts after the cells were
depleted in Class I transcripts (110). The transcripts belonging to both classes
are further degraded by PNPase (266).
In conclusion, the post-transcriptional regulation of gene expression is still
poorly characterized in S. pyogenes and further investigations are required,
particularly about the roles of RNases and their specificities.
19
II- Aims of the thesis
In this thesis, the roles of various RNases on the regulation of gene
expression in the human pathogen S. pyogenes were investigated. The approach
used was to establish the targetomes of these RNases, i.e. the exact processing
sites on a genome-wide scale. The specific objectives were:
1- To develop a novel analysis tool that uses RNA sequencing datasets to
identify the targetome of a given RNase.
2- To characterize the roles and specificities of the endoRNases Y and III,
and the 3′-to-5′ exoRNases R, YhaM, and PNPase.
3- To investigate the interplay between RNase Y and the 3′-to-5′ exoRNases.
20
III- Results and discussion
Paper I
Identification of endoribonuclease specific cleavage positions
reveals novel targets of RNase III in Streptococcus pyogenes.
RNA sequencing is widely used to identify in vivo genome-wide RNase
cleavage positions (i.e. targetome). However, when our project to establish
RNase III targetome started, very few studies were available and relied only on
the sequencing of transcript 5′ ends (106, 225, 241). The disadvantage of this
method is that the RNase processing events leading to subsequent degradation
of transcript 5′ ends are not identified. We sequenced both transcript 5′ and 3′
ends to increase the detection of processing events. To determine the global
processing positions of an endoRNase of interest, we compared the abundances
of transcript 5′ and 3′ ends in WT and ∆rnase strains. The ds specific RNase III
was a perfect model to validate this method of analysis, as it generates
theoretically two new 5′ ends and two new 3′ ends for each cleavage event.
Methodology
Two different kinds of libraries (p and Pp) were prepared using total RNA of
WT, ∆rnc (lacking RNase III), ∆rny (lacking RNase Y), and double mutant
∆rny_∆rnc strains (Paper I, Supplementary Figure S1). The Pp library
contained the processed and primary transcripts and the p library was enriched
in processed transcripts. The total coverage as well as the coverages of the 5′ end
and the 3′ end (i.e. the ends of the read) were calculated for each position on the
genome. To insure that we compared positions with enough coverage in all
strains, we filtered out the positions with less than 10 reads in the reference
(expressing RNase III, i.e. WT and ∆rny) and mutant strains (not expressing
RNase III, i.e. ∆rnc and ∆rny_∆rnc). Then, we defined several parameters to
isolate the specific cleavage positions from the background noise. These
parameters were less stringent for the p libraries, where the processed
21
transcript ends were enriched, than for the Pp libraries. The following settings
were used for the p libraries (Paper I, Figure 1): (i) the position of interest
cumulated a minimal read end coverage of 10 in the two strains that were
compared; (ii) the reads that ended at the position of interest represented at
least 16% of all the read ends in a 20 nt-window centered at this position; (iii)
the percentage of reads ending at the position of interest compared to the total
coverage at the position of interest was at least 5%; and (iv) the ratio of the
percentages obtained in (iii) for the reference strain and for the mutant strain
was at least 3.
Characteristics of S. pyogenes RNase III
We detected 92 cleavage positions that were dispatched on 25 transcripts
and on the six pre-rRNA transcripts. For more than half of the 25 transcripts,
we retrieved both RNase III cleavage positions in a stem-loop and in two cases,
in antisense transcripts (Paper I, Figure 3A). The 2 nt 3′ overhang characteristic
of RNase III processing was detected, which validated the accuracy of our
method. We identified a single cleavage site (referred to as stand-alone
positions, or SA) for the rest of the transcripts (Paper I, Figure 3A). Although a
relaxed consensus sequence was described for E. coli RNase III around the
cleavage site (72, 239, 267), we did not observe any conserved sequence or motif
for S. pyogenes RNase III.
When we visually inspected the individual reads mapped at the RNase III
double cleavage positions, we discovered that, for a given target, a portion of the
transcripts were cleaved only at one position (i.e. cleavage in one side of the
RNA stem) (Paper I, Figure 3C). This nicking activity was visible on each strand
for a given target, which means that RNase III did not favor one strand in
particular. It was previously observed that E. coli RNase III had a nicking
activity in vitro and in vivo (70, 71, 267). Our primer extension analyses
confirmed the presence of two alternative 5′ ends (nicking at one or the other
side of the RNA stem) for four transcripts (Paper I, Supplementary Figure S2).
The transcripts where we could observe RNase III nicks harbored at least one
bulge or a loop located near one of the two cleavage positions (Paper I, Figure
22
3B-C, Supplementary Figure S4). These structural features are known to
destabilize RNase III binding to the RNA after the first cleavage (69, 72). The
accessibility of the upstream fragment, generated by RNase III nicking, to 3′-to-
5′ exoRNases was investigated in Paper II. The study of RNase III nicking
activity on pnpA 5′ UTR showed that PNPase was able to degrade the two
alternative 3′ ends (Paper II, SI appendix, Fig. S7). Thus, for this specific target,
the nicks did not alter the degradation of the upstream fragment. Up to this
date, there is only the T7 transcript that is nicked and stabilized by RNase III,
which illustrates the biological impact of RNase III nicking activity (73). We
showed that RNase III nicking activity is a general phenomenon in S. pyogenes
but the impact on gene regulation remains unknown.
SA processing positions were detected in ss regions of eight transcripts. We
first reasoned that these transcripts base-paired with an unknown transcript
and that the second processing position was in the complementary mRNA or
sRNA. Unfortunately, we could not identify base-pairing partners for these
transcripts. Other explanations were considered: (i) RNase III cleaved in an
unstable ds region (predicted as ss) and the fragment generated by the other
processing was degraded and therefore not detected; (ii) RNase III nicked in an
unstable ds region (predicted as ss); (iii) RNase III cleaved in a RNA stem-loop
that was subsequently trimmed by the 5′-to-3′ exoRNase J1 for the 5′ end
positions, or by the 3′-to-5′ exoRNases for the 3′ end positions, hence the
information about the original cleavage positions of RNase III was not retrieved.
Two SA positions were associated with a change in abundance of neighborhood
transcript (Paper I, Supplementary Table S3), therefore the study of these SA
targets could reveal novel mechanisms of gene regulation by RNase III.
RNase III role in S. pyogenes
As described in other bacteria, we found that S. pyogenes RNase III
participated in rRNA maturation (78). The enzyme processed the pre-rRNA
transcripts in the stem-loops delimiting the 16S and 23S rRNAs, which were
further matured by other RNases (Paper I, Supplementary Figure S3). RNase III
was not essential to this process, as RNase III deletion did not influence the
23
maturation of the 16S and 23S rRNAs. In this context, we detected additional
processing positions (called substitute processing events) on the pre-rRNAs that
were not present in the WT (Paper I, Supplementary Table S4). This indicated
that the rRNA maturation was performed by other RNases to ensure that
ribosomes were still functional in absence of RNase III.
S. pyogenes RNase III targeted mostly UTRs, as for S. aureus RNase III (82),
but in contrast to what has been described in E. coli and B. subtilis (80, 239).
This suggests that the cleavages would affect gene expression. However, out of
the 25 transcripts cleaved by RNase III (excluding rRNAs), the abundance of
only seven was affected. It is possible that RNase III regulates transcript
stability or translation without affecting the transcript abundance. For instance,
in S. pyogenes, the global transcript stabilization caused by RNase Y deletion
did not reflect on the transcript abundance (50). Therefore, alternative methods
should be used to investigate RNase III regulatory effects.
We identified several mRNAs that were known targets of RNase III in other
bacteria, such as pnpA. In E. coli, RNase III cleaves a stem-loop in the 5′ UTR of
pnpA, which leads to pnpA destabilization (147). We observed a similar cleavage
by S. pyogenes RNase III in pnpA 5′ UTR and pnpA abundance was increased
in absence of RNase III (Paper I, Figure 3B and Supplementary Table S6).
Another target, the secY-adk transcript (coding for Sec translocase and
adenylate kinase), was cleaved by RNase III in S. aureus (82). More specifically,
RNase III cleaves the transcript in the secY ORF and the absence of cleavage
leads to an increased secY abundance and stability (82). We showed that a
different regulatory mechanism occurs in S. pyogenes. RNase III cleaved in the
UTR between the secY and adk, which uncoupled the expression of the two
genes as only adk was more abundant in ∆rnc (Paper I, Figure 3C and
Supplementary Table S6). These examples highlight that, while some regulatory
mechanisms are conserved, bacteria also develop alternative mechanisms to
regulate the expression of similar transcripts using orthologue RNases.
RNase III is involved in sRNA-mediated gene regulation (84, 85, 87, 268). In
this study, few sRNAs were cleaved by RNase III or differentially abundant
(Paper I, Supplementary Table S7). For instance, we retrieved RNase III
processing in the putative 23S methyl regulatory element, in the antisense RNA
24
SPy_sRNA477741 (85), in CRISPR RNAs (crRNAs), and trans-activating
CRISPR RNA (tracrRNA) (87). sRNAs were lost during our library preparation,
which could explain the limited number of RNase III targets.
In conclusion, we determined that RNase III participates in conserved
regulatory and maturation pathways and has a limited specific role in S.
pyogenes. Further work is required to evaluate the impact of RNase III on
sRNAs, which may be broader than what was described in this study.
Paper II
The in vivo 3′-to-5′ exoribonuclease targetomes of Streptococcus
pyogenes.
In Paper I, we identified and validated RNase processing positions by
sequencing both the 5′ and the 3′ ends of transcripts. This opened the
possibility to study the targetomes of 3′-to-5′ exoRNases. PNPase and RNase R
have been intensively studied in various organisms, yet the global identification
of their direct targets was missing. We applied our method to the study of
YhaM, PNPase, and RNase R, the three 3′-to-5′ exoRNases present in S.
pyogenes.
Methodology
We improved the previous method presented in Paper I by adding a step of
normalization between the samples and including statistical power using false
discovery rate (FDR). Total RNA was extracted in triplicates from WT, ∆yhaM
(lacking YhaM), ∆pnpA (lacking PNPase), and ∆rnr (lacking RNase R). EdgeR
was used to analyse the differential abundance of transcript 5′ and 3′ ends
between the reference strain and the mutant strain (234). The positions with an
absolute fold change of at least 2 and an FDR below 0.05 were used for further
analysis. We applied the parameters from Paper I on these positions: the
percentage of reads ending at the nucleotide of interest compared to all the
reads that mapped at the nucleotide of interest was at least 2%; and the ratio of
25
the percentage obtained for the reference and the mutant strains was at least 3.
We used the Pp libraries for all the strains.
The positions with a negative fold change (i.e. more abundant in the WT)
corresponded to exoRNase trimming stop positions and those with a positive
fold change (i.e. more abundant in the ∆exornase) corresponded to exoRNase
trimming start positions. The distance between them represented the
processivity of the enzyme. Approximatively 90% of all the positions more
abundant in the WT were transcript 3′ ends, which confirmed the in vivo 3′-to-
5′ exoribonucleolytic activity of these enzymes.
YhaM has a global nibbling activity on transcript 3′ ends
We identified 602 trimming stop positions for YhaM. The conservation of a
secondary structure and a stretch of uridines signaled the presence of intrinsic
transcriptional terminators upstream half of these positions (Paper II, Fig. 2
and SI appendix, Fig. S2A-B). The second half of the positions presented no
particular conservation of sequence or structure (Paper II, SI appendix, Fig.
S2C-D). These positions were mostly found in ORFs, which suggested that these
transcript 3′ ends originated from endoRNase processing. We estimated that
the enzyme trimmed approximatively half of the terminated transcripts ends, in
addition of the transcript 3′ ends produced by endoRNases. Therefore, YhaM
has a broad targetome in S. pyogenes. It would be of interest to understand why
some transcripts are not targeted by YhaM. Perhaps factors such as low
transcript abundance, too short or absent ss tail, or the localization of the
transcripts in the cell can restrict YhaM activity.
The 602 trimming stop positions were associated with the closest 171
trimming start positions (maximum distance of 10 nt) to estimate YhaM
processivity. We obtained 112 pairs located within 10 nt from transcriptional
terminators and the 42 pairs located at putative processing sites. The two
groups of pairs revealed that YhaM trimmed 3 nt on average (Paper II, Dataset
S2), which implied that the terminator structure did not limit YhaM
processivity. YhaM nibbling activity was additionally validated by Northern
blotting analyses (Paper II, SI appendix, Fig. S4). Almost all the trimming start
26
positions were paired with the stop positions, supporting that the short
processivity was not a bias of the distance of pairing. Oppositely, the paired
trimming stop positions represented only a small fraction of the total number of
trimming stop positions. This can be explained by the fact that many YhaM
trimming start positions were not identified (for more details about the
limitation of our method, refer to Paper II, SI appendix, Fig. S3A-B). Indeed, by
visual screening, we could observed these trimming start positions 1-3 nt
downstream of the trimming stop positions (examples presented in Paper II, SI
appendix, Fig. S3C). We cannot exclude that, among the unpaired trimming
start positions, some may be indicative of a processivity for YhaM higher than 3
nt. However, this would be a minor activity compared to YhaM nibbling activity.
Role and peculiarities of S. pyogenes YhaM
It has been suggested that YhaM acted more as a DNase than as an RNase in
vivo (269). Our study shows that S. pyogenes YhaM is an unspecific RNase that
broadly targets transcript 3′ ends originating from termination or processing by
other RNases. The removal of 3 nt on average at transcript ends raised several
questions.
First, what prevents YhaM from trimming further the transcript ends? (i) It
could be due to the intrinsic biochemical properties of the enzyme. However, B.
subtilis YhaM was able to degrade a 110 nt substrate in vitro (179). (ii) It could
be transcript secondary structures; yet, we failed to detect any structure that
could have blocked YhaM at the transcript 3′ ends generated by endoRNases.
(iii) It could be a balance between the cellular concentration of YhaM and of the
transcripts, yet we did not observe a correlation between transcript abundance
and trimming. In E. coli, a similar observation was recently made for the 3′-to-
5′ exoRNase II, which nibbles 1-3 nt at transcript intrinsic terminators (243).
The authors did not comment on whether RNase II nibbled only at the
transcript intrinsic terminators or at unstructured transcript 3′ ends as well. As
RNase II is blocked approximately 9 nt downstream of secondary structures
(270), this physical constraint could be an explanation. The mechanisms behind
the short processivity of YhaM and RNase II are likely to be different.
27
Interestingly, the same study described a narrow YhaM nibbling activity at
intrinsic transcriptional terminators in B. subtilis (15% of the intrinsic
terminators) (243). It is possible that YhaM has a broader targetome in S.
pyogenes than in other bacteria. The comparison of the targetomes of YhaM
orthologues belonging to the Bacillales order (e.g. B. subtilis and S. aureus) and
to the Lactobacillales order (e.g. S. pyogenes) will help answering this question.
Secondly, what is the biological purpose of this nibbling? RNase II is
suggested to have a protective role on transcripts by shortening the tail required
for PNPase and RNase R binding and degradation (133, 271, 272). We observed
that the ss tail downstream of terminators was reduced from 9 to 6 nt on
average in the WT compared to ∆yhaM (Paper II, SI appendix, Fig. S1). For E.
coli PNPase, these tails would be too short and the transcripts would be
protected (120). To gain some insight on the YhaM possible protective role, we
looked at the differential gene expression in ∆yhaM compared to the WT. Forty
transcripts were differentially expressed and nineteen were downregulated in
∆yhaM (Paper II, Dataset 3). This small number of transcripts did not correlate
with the global nibbling performed by YhaM. Additionally, the transcript 3′
ends generated by endoRNases did not have a secondary structure that would
protect them from PNPase or RNase R (Paper II, SI appendix, Fig. S2D). Thus,
removing 3 nt from the ss tail of these transcripts seems unnecessary.
We observed that S. pyogenes growth at cold temperature was greatly
impacted by the deletion of YhaM (Paper II, SI appendix, Fig. S9B).
Interestingly, the deletion of PNPase or RNase R, which are known to be
essential or important for optimal growth at cold temperatures in various
bacteria (147, 172), did not have such an effect. It was previously shown in B.
subtilis that the deletion of YhaM in the double mutant ∆pnpA_∆rnr slowed
bacterial growth at cold temperatures (179). We observed a synergic effect when
YhaM was deleted in ∆rnr but not in ∆pnpA (Paper II, SI appendix, Fig. S9B). It
is tempting to speculate that YhaM activity becomes important when S.
pyogenes encounters specific growth conditions. In these conditions, it is not
known whether the slower growth phenotype is due to the absence of global
nibbling on the transcript 3′ ends, to an indirect effect of the enzyme deletion on
specific targets, or to its putative DNA-binding activity.
28
PNPase role in RNA decay
The deletion of PNPase revealed 1255 trimming start positions and 183
trimming stop positions. The small amount of trimming stop positions implied
that PNPase fully degraded its targets (i.e. until the 5′ end). Therefore, we
assumed that transcript 5′ ends should be more abundant in absence of PNPase.
It should be noted that the fragments whose 5′ end corresponds to
transcriptional start sites (TSS) were not identified with our method (for a
detailed explanation, refer to Paper II, SI Appendix, Fig. S3). Indeed, we noticed
336 transcript 5′ ends that accumulated in ∆pnpA (24 and 8 times more than in
∆yhaM and ∆rnr, respectively; Paper II, Fig. 1A). By pairing the transcript 5′
ends with the closest trimming start positions, we showed that 185 fragments
with a median size of 115 nt were present in ∆pnpA. As 85% of these fragments
were located in ORFs, we assumed that they originated principally from the
mRNA decay pathway initiated by endoribonucleolytic activities. These
fragments were not detected in the WT, therefore they were immediately
degraded by PNPase. In B. subtilis, PNPase degrades principally decay
intermediate fragments originating from the 5′ proximal part of transcripts
(131). We could not observe a similar pattern, due to the impossibility to detect
fragments accumulating at the TSS in ∆pnpA (for a detailed explanation, refer
to Paper II, SI Appendix, Fig. S3).
Less than 20% of PNPase total trimming start positions were located in
UTRs. Some of them were downstream of regulatory 5′ UTRs, for instance T-
boxes that controlled aminoacyl-tRNA synthetase expression by binding tRNAs.
Northern blotting analyses of 10 regulatory 5′ UTRs revealed that decay
intermediate fragments accumulated in ∆pnpA for 9 of them (Paper II, Figs. 3B
and 4, and SI Appendix, Fig. S8). For example, when the pyrR leader was
probed, the decay intermediate fragments were longer than the full length
leader, indicating that the initial processing events were in the pyrR coding
sequence instead of the 5′ UTR. Indeed, several PNPase trimming start
positions were detected downstream of the leader (Paper II, SI Appendix, Fig.
S8). With the notable exception of FMN, the stability of the full-length leaders
was unaffected by the deletion of PNPase, hence PNPase is not initiating the
29
decay. Additionally, we observed that the dpr leader was matured by PNPase
(Paper II, Fig. 3C and SI Appendix, Fig. S8). Interestingly, this transcript codes
for a peroxide resistance protein (273), thus the function of this stable sRNA in
oxidative stress should be investigated. In view of these results, our study shows
that PNPase is also responsible for degrading 5′ UTRs in S. pyogenes. Many
regulatory 5′ UTRs adopt different structural conformations based on external
stimuli such as temperature or binding of a ligand (274). For instance, B.
subtilis PNPase preferentially degrades the regulatory 5′ UTR of trp when it is
bound by TRAP (145, 146). It would be of interest to study whether this
conformational preference is observed for S. pyogenes PNPase.
As PNPase relies on endoRNases to degrade the decay intermediate
fragments, we compared PNPase trimming stop positions with RNase III
cleavage positions (Paper I) (275). Indeed, PNPase trimmed fragments
produced by RNase III, for example the decay intermediate fragments generated
during rRNA maturation and the processing of pnpA 5′ UTR (Paper II, SI
appendix, Figs. S6 and S7). PNPase also trimmed approximatively 25 nt of rplQ
3′ UTR upstream of RNase III cleavage (Paper II, SI appendix, Fig. S7).
In conclusion, PNPase is the major 3′-to-5′ exoRNase involved in S.
pyogenes RNA decay and the processing of the dpr leader hints toward a
possible regulatory role for PNPase in S. pyogenes, as it is the case in many
bacteria.
Limited RNase R activity
A total of 82 trimming positions were retrieved for RNase R. Only two
transcripts harboring a trimming start position and a trimming stop position
and six decay intermediate fragments were identified. One trimming start
position was located in the glyQ leader, a T-box riboswitch, which was also
degraded by PNPase (Paper II, SI Appendix, Fig. S8). Northern blotting
analyses confirmed that glyQ leader was degraded by PNPase and RNase R
(Paper II, Fig. 4). However, the decay intermediate fragment was more stable in
absence of RNase R than of PNPase. We hypothesized that (i) RNase R could be
more efficient in degrading this decay intermediate fragment than PNPase
30
(redundant activity), or (ii) there could be two populations of intermediate
fragments (tRNA-bound and free) and the population accumulating in ∆pnpA is
different from the one accumulating in ∆rnr, as the trimming start positions of
RNase R and PNPase were few nt apart (specific activity) (Paper II, SI
Appendix, Fig. S8). In the second case, each 3′-to-5′ exoRNase would then be
responsible for degrading one population of glyQ leader. In absence of RNase R,
PNPase would degrade both glyQ leader populations and would need more time
to degrade the RNase R-specific population. Reciprocally, in absence of PNPase,
RNase R would be able to degrade both glyQ leader populations, and would
need more time to degrade the PNPase-specific population.
Possible redundancy between RNase R and PNPase
Several reasons could explain the very limited targetome observed for
RNase R. First, it could indicate a low expression or a reduced activity of the
enzyme under standard laboratory conditions of growth (mid-logarithmic phase
of growth, rich medium, optimal temperature). Indeed, RNase R in Gram-
negative bacteria is more stable and more expressed under stress conditions
(276). Secondly, RNase R could degrade transcripts from transcriptional
terminator to TSS, which were not detected by our method (Paper II, SI
appendix, Fig. S3E). The third explanation is that PNPase would compensate
when RNase R is absent, thereby preventing the identification of RNase R
targets. In favor of this hypothesis, RNase R activity is mostly observed in
absence of PNPase in E. coli and B. subtilis (136, 167). In addition, the double
mutant ∆pnpA_∆rnr could not be obtained in the conditions tested in S.
pyogenes. In E. coli, ∆pnpA_∆rnr is also lethal, probably due to the
accumulation of abnormal rRNA precursors (169, 277). Interestingly, in B.
subtilis, all the 3′-to-5′ exoRNases (PNPase, RNase R, YhaM, and RNase PH)
can be deleted together (136), which suggests a different pathway of rRNA
quality control than in E. coli. If the lethality of ∆pnpA_∆rnr in S. pyogenes is
confirmed, it would be of interest to investigate the biological reason behind.
The decay intermediate fragments originating from the regulatory 5′ UTRs
that accumulated in ∆pnpA were eventually degraded by another exoRNase
31
(Paper II, SI Appendix Fig. S8). Therefore another RNase, presumably RNase R
or RNase J1, can compensate the loss of PNPase in RNA decay pathway. In this
case, the redundancy is only partial, as we observed many specific PNPase
trimming positions in our screening (Paper II, Dataset S1). B. subtilis PNPase is
also the major 3′-to-5′ exoRNase (131, 136) and in its absence, RNase R degrade
some decay intermediate fragments (136). However, the combined deletion of
PNPase, RNase R, RNase PH, and YhaM is not lethal in B. subtilis (136),
suggesting either that the 3′-to-5′ degradation pathway is not necessary or that
an additional 3′-to-5′ exoRNase remains to be identified.
Both B. subtilis and S. pyogenes express RNases J1 and J2, however these
enzymes are only essential in S. pyogenes (107, 110). The essentiality of the 3′-
to-5′ degradation pathway and of the 5′-to-3′ degradation pathway can imply
that they have distinct functions in S. pyogenes.
Paper III
Interplay between 3′-to-5′ exoRNases and RNase Y in
Streptococcus pyogenes.
As a continuation of Paper II, we decided to investigate the implication of
RNase Y in the generation of transcript 3′ ends that are targeted and degraded
by the 3′-to-5′ exoRNases YhaM, PNPase, and RNase R. First, we applied the
analysis method used in Paper II (see Paper II, ”Methodology” in this thesis) to
determine the targetome of RNase Y. Total RNA was extracted from WT, ∆rny
(lacking RNase Y), and ∆rny::rny (complemented strain) that was used as a
second reference strain. Secondly, we compared the targetome of RNase Y with
the targetomes of the 3′-to-5′ exoRNases.
RNase Y cleaves after a guanosine
We identified 320 processing positions, 60% were detected as 5′ ends (190)
and 40% were detected as 3′ ends (130). The majority of transcript 5′ ends
(87.4%) generated by RNase Y were located immediately after a guanosine (G)
(Paper III, Figure 1D). A recent study from our laboratory showed that RNase Y
32
requires a G to process the speB transcript in vivo (60). The conserved presence
of a G in our screening suggests that this requirement is a general feature of S.
pyogenes RNase Y.
It is difficult to envision how the G alone can restrict S. pyogenes RNase Y
activity to the limited number of processing positions we identified. A G residue
was previously identified upstream of 58% of the 99 transcript 5′ ends produced
by RNase Y in S. aureus (51), which is similar to what we observed. In
comparison, Salmonella enterica RNase E prefers a U located 2 nt downstream
of the processing position and 22000 processing sites were reported in vivo
(240). Thus, the specificity towards 1 nt is unlikely to restrict RNase Y activity
enough to explain the small targetomes detected in S. pyogenes and S. aureus.
The specificity of S. aureus RNase Y was further investigated on the saePQRS
transcript, which was processed just downstream of a U (65). Instead of a
sequence motif, S. aureus RNase Y requires a downstream structured element
to cleave the saePQRS transcript (65). Global studies done in B. subtilis, in S.
aureus as well as our study in S. pyogenes did not reveal structured regions near
RNase Y processing positions (51, 66). The identification of complex structured
regions among the targets is difficult; therefore, it is possible that undetected
structures are important to direct RNase Y processing of some transcripts.
Another possibility is that S. pyogenes RNase Y specificity would be altered
when RNase Y associates with other proteins, such as the Y-complex described
in B. subtilis (66). The localization to the membrane could restrict RNase Y
access to the transcripts. However, this was not the case in S. aureus and S.
pyogenes (51, 64). It is possible that RNase Y has a broader targetome than
what we detected, which is undetectable because the exoRNases degraded the
transcripts produced by RNase Y. It is not clear whether RNase Y orthologues
have different specificities or whether RNase Y responds to multiple signals that
have not been fully characterized. The co-crystal structure of this protein with
its target is crucially missing to improve our understanding on RNase Y
specificity.
33
RNase Y and PNPase act in concert to degrade transcripts
The RNase Y-dependent 130 transcript 3′ ends did not present the conserved
G. We assumed that these transcripts were initially processed by RNase Y and
were then further trimmed by 3′-to-5′ exoRNases. The transcript 3′ ends
detected in our screening would therefore correspond to the trimming stop
positions of the exoRNases. To prove this hypothesis, we compared the
targetome of RNase Y with the trimming start and stop positions of YhaM,
PNPase, and RNase R previously published in Paper II (Paper III, Figure 2).
The positions of six and forty-six RNase Y-dependent transcript 3′ ends
corresponded to PNPase trimming start and stop positions, respectively (Paper
III, Figure 2A, panels 1-2 and Tables S2-S3). The six PNPase trimming start
positions, which would correspond to the nucleotide upstream of RNase Y
original processing positions, were located at a G (Paper III, Table S3). These
results showed that most of the transcript 3′ ends produced by RNase Y and
trimmed by PNPase were immediately trimmed (i.e. mostly trimming stop
positions detected). It was shown that RNase Y and PNPase interact in B.
subtilis, which could explain the immediate degradation of transcripts produced
by RNase Y (278). However, the disruption of the complex did not affect the
decay of the transcripts studied (278). In addition, we retrieved 27 transcript 3′
ends nibbled by YhaM and one transcript 3′ end trimmed by RNase R, in
accordance with YhaM broad nibbling activity and RNase R limited detectable
activity described in Paper II. The rest of the RNase Y-dependent transcript 3′
ends did not present a conservation of G. They could originate from PNPase and
RNase R redundant trimming (which we cannot detect using our method), from
another unidentified 3′-to-5′ exoRNase expressed in S. pyogenes, or from
another RNase whose activity is dependent on RNase Y.
We did not retrieve both the transcript 3′ end (i.e. upstream fragment) and
5′ end (i.e. downstream fragment) produced by a single RNase Y processing
event. Because our results showed that 3′-to-5′ exoRNases trimmed transcripts
produced by RNase Y, we searched for the 3′-to-5′ exoRNase trimming start
positions located near the transcript 5′ ends produced by RNase Y (Paper III,
Figure 2A, panel 3). Both transcript ends were observed for 23 RNase Y
34
processing events and PNPase trimmed all the transcript 3′ ends identified
(Paper III, Table S4). Many RNase Y-dependent transcript 5′ ends (167) were
not paired with the corresponding upstream fragments. This implies that other
3′ ends are missing, probably because they were fully degraded or because they
were trimmed by RNase R in absence of PNPase.
PNPase generally cannot degrade Rho independent terminators and depends
on the activity of endoRNases to remove terminators in E. coli and B. subtilis
(103, 122, 123, 203). S. pyogenes does not have a Rho orthologue (279),
therefore all the transcriptional terminators are in theory intrinsic terminators.
Some RNase Y processing positions were located upstream of transcriptional
terminators and attenuators. For instance, RNase Y removed the transcriptional
attenuator from the glyQ leader (Paper III, Table S2), which was further
processed by PNPase and RNase R (Paper II, SI Appendix, Fig. S8). An RNA
decay mechanism such as the one initiated by RNase Y on the glyQ leader must
be general. We do not know whether RNase Y is the global initiator of decay by
removing intrinsic terminator structures, because few positions close to the
transcriptional terminators were observed. In some cases, the small size of the
downstream fragments containing the terminators prevented the detection of
RNase Y processing positions at the terminators. Their identification could be
prevented as well by RNase J1, shown to degrade the fragments containing the
transcriptional terminators in B. subtilis (80, 102, 103).
Because PNPase degraded the transcript 3′ ends generated by RNase Y, we
had a closer look at the transcript ends that accumulated in ∆pnpA (Paper II, SI
appendix, Fig. S2E). A conserved G was visible directly upstream of transcript 5′
ends, which pinpointed towards RNase Y activity. We repeated the sequence
analysis on the decay intermediate fragments (Paper III, Fig. S5) and showed
that RNase Y was responsible for the generation of some decay intermediate
fragment 5′ ends but not for the 3′ ends (Paper III, Figure 2C). Therefore,
PNPase degrades the decay intermediate fragment 3′ end generated by an
unidentified endoRNase until it reaches the fragment 5′ end produced by
RNase Y. This highlights that the interplay between PNPase and RNase Y is not
exclusive and that PNPase processes transcript 3′ ends that are generated by
other endoRNases.
35
Is RNase Y the major RNase initiating RNA decay?
S. pyogenes RNase Y is involved in RNA decay, as illustrated by the interplay
with PNPase to degrade RNA fragments. Several facts support that RNase Y has
a minor role in this process and are developed below: the small targetome of
RNase Y, the absence of strong phenotype for ∆rny, and the presence of
RNase Y-independent but PNPase-dependent degradative pathway.
First, with the addition of putative processing positions of RNase Y found in
∆pnpA (336 transcript 5′ ends with a small conservation of the G, Paper II), the
total number of RNase Y processing positions that we detected in these
conditions would not be above 500. The S. pyogenes strain SF370 used in our
studies has a genome of 1.85 Mb and 1801 genes (262, 263). Thus, the RNase Y
targetome represents a small fraction of processed transcripts, as described for
B. subtilis and S. aureus (51, 66). This is opposite to the 22000 processing
positions published for RNase E, the major initiator of mRNA decay, in S.
enterica (240). Secondly, the deletion of RNase Y mildly affected the bacterial
growth (Paper III, Figure S3). We observed an absence of hemolysis on agar
blood plates and of protease activity on silk-milk plates (it has been shown to be
due to SpeB (60)), but no susceptibility to several antibiotics, no susceptibility
to osmotic stress, no differences in metabolic reactions with Api 20 Strep
(Biomérieux) were detected. This is opposite to the severe phenotypes observed
in B. subtilis (107) and to the essentiality of RNase E in Gram-negative bacteria
(202). Thirdly, the size of the fragments degraded by PNPase (120 nt on
average) indicates that transcripts are processed multiple times. The
conservation of sequence around the fragments degraded by PNPase (Paper III,
Figure S5) suggested that other RNases were involved. Thus, PNPase degrades
transcripts in an RNase Y-independent manner. This was supported by the
growth of the double deletion mutant ∆rny_∆pnpA, which is slower than the
single deletion mutants ∆rny or ∆pnpA (Paper II, SI appendix, Fig. S9; to
compare with Paper III, Figure S3).
Taken together, our data supports the existence of a degradation pathway
initiated by RNase Y and involving PNPase for the final degradation steps.
However, this pathway does not seem central in mRNA decay when bacteria are
36
grown under standard laboratory conditions. The study of a different S.
pyogenes strain (NZ131) grown in C-medium, which is a peptide-rich and
carbon-poor medium that mimics host environment during deep tissue invasion
(280, 281), revealed that the bulk RNA half-life was increased by 2 fold in
absence of RNase Y (50). It is possible that RNase Y has a more prominent role
under other conditions.
The fate of transcript 5′ ends produced by RNase Y
The main finding of our study is the different fate of the transcript 3′ ends
and 5′ ends produced by RNase Y. On one side, we showed that most of the
transcript 3′ ends are further trimmed by 3′-to-5′ exoRNases and on the other
side, we observed that 87.4% of the transcript 5′ ends corresponded to the
original RNase Y processing positions. Therefore, these transcript 5′ ends are
not subsequently partially trimmed by RNase J1, which would give observable
RNase J1 trimming stop positions. However, since 107 transcript 3′ ends were
not paired with their corresponding downstream transcript 5′ ends, a number of
transcript 5′ ends were not detected in our study. A part of these transcript 5′
ends were degraded by PNPase, which started at the downstream 3′ end until it
reached the 5′ end produced by RNase Y (Paper III, Figure 2D). We assume that
RNase J1 is also involved in degrading these transcript 5′ ends. The interplay
between RNase Y and RNase J1 was indeed observed in vivo in B. subtilis and S.
aureus (40, 51). An interesting observation was that either the transcript 5′ ends
were original RNase Y processing positions or they were not detected, i.e.
degraded to completion. The signals deciding the different fates of these
transcript 5′ ends should be further investigated.
The interplay between RNase Y and RNase J1 does not seem as broad as the
interplay between RNase Y and PNPase. S. pyogenes RNases J1 and J2 are
essential (110) and the fact that the deletion of RNase Y is not lethal suggests
that RNase J1 and/or RNase J2 perform RNA decay mainly in an RNase Y-
independent manner. To this date, we have been unable to obtain the single
deletions of RNase J1, RNase J2 and the double deletion of PNPase and RNase
R in S. pyogenes, suggesting that these deletions are lethal. This highlights that
37
both 3′-to-5′ degradation and 5′-to-3′ degradation pathways are vital for S.
pyogenes. Whether it is due to a global role in one particular cellular process
(such as RNA decay) or to the maturation or degradation of a specific target is
currently unknown.
38
IV- Main findings of the thesis
The aim of this thesis was to investigate the poorly characterized post-
transcriptional regulation mediated by RNases in S. pyogenes. A method was
developed to identify the specific in vivo targetomes of endoRNases and
exoRNases. It was applied to study the roles of RNase Y, RNase III, YhaM,
PNPase, and RNase R in S. pyogenes, as well as the interplay between these
RNases. The three papers included in this thesis led to the conclusions
presented below.
We reported a general in vivo nicking activity of RNase III and a broad
nibbling activity of YhaM, whose impact on the regulation of gene expression in
S. pyogenes is currently unknown.
PNPase is the major 3′-to-5′ exoRNase responsible for RNA decay in S.
pyogenes. Our results additionally suggest a redundant role between PNPase
and RNase R in this process. However, while PNPase compensated the loss of
RNase R, the reverse was only partial.
S. pyogenes RNase Y processes transcripts downstream of a G and initiates
the subsequent degradation of the upstream fragment by 3′-to-5′ exoRNases.
These fragments can be fully or partially degraded. The downstream fragments
are either stable or fully degraded, as we did not observe partial degradation.
RNase Y is not a major player in S. pyogenes mRNA decay in standard
laboratory conditions. Instead, S. pyogenes, like other Gram-positive bacteria,
seems to rely on several endoRNases to initiate mRNA decay, opposite to E. coli
that relies principally on RNase E.
In conclusion, the method presented in this thesis is particularly suitable for
the study of sequential processing events performed by multiple RNases. Given
the importance of S. pyogenes as a human pathogen, it will be relevant to
extend the use of this method to study the role of these RNases in conditions
mimicking human infection.
39
V- Acknowledgements
Many persons made this journey possible and unforgettable. It started in
Umeå seven years ago and I am glad to come back there for the final step (even
if it is in December).
I am grateful to my supervisor Emmanuelle Charpentier for the opportunity
she gave me to work in her lab, to develop my projects independently, to attend
international conferences, and for making me discover Berlin.
I am grateful to my co-supervisors and my PhD committee members Bernt
Eric Uhlin and Tracy Nissan, to my PhD committee member Jan Larsson, to
Sven Bergström for being my examiner and to the members of the BEU journal
club and of the RNA group for the fruitful conversations. In particular, Tracy,
thank you for including me in your group during my “lonely” years in Umeå, for
the lab meetings and the journal clubs “à l’américaine”, for the time you
invested in me, for your critics and your support and for your knowledge (I am
sure that your brain is more complete that my Endnote libraries will ever be). I
learnt so much from you during this time, you are a truly inspiring scientist!
Thanks to my third co-supervisor, AnaÏs, for your support, dedication, constant
diverging opinions and for improving my writing .
For their precious help to organize the defense from Berlin and for their fast
answers to any question, thank you Eva-Maria Diehl and Ulrich von Pawel-
Rammingen! Thank you Jörgen Johansson for your help when you were
responsible for the PhD students, and for the numerous “C’est la vie” at the
coffee machine. I am also very grateful to the persons working at the
administration of Mol Biol, MPIIB and MPUSP.
From my time in Umeå: the original lab members : Sophie, for constantly
mocking my French accent, which improved it (a bit); Geet, for the best
cheesecakes ever; Khan, for being the kindest person in the lab; and of course,
where is Shi Pey? Already gone to Berlin… The beer corner times will stay
forever in my memory, as much for the people, the discussions and the music as
for the affordable beers… Susanne, well, you know… thank you for all of that. I
remember the night when half of the EC lab left, I was really down. Then, truly
40
amazing persons appeared in my life: Rosh, you are the best kind of crazy
person; Akbar: keep moving those hips! And keep the discussions about
everything and nothing; Ala: ma petite, I will always fight to keep the last bit,
even though I know I don’t stand a chance; Nabil: Exaaaaactlyyyyyy…; I love
you guys, you made Umeå the best place in the world. The lovely members of
the Spanish group and Vassili group, Ági, Mridula, Radha, Chinmay, Hélène,
Sveta, Sandrine, thank you for all the parties, outings, the curling, ultimate
Frisbee, etc… Thanks to the TAs during the fun/stressful teaching times.
From my time in Berlin: Laura, my PE buddy, my double loading gel partner,
it was so fun and motivating to work with you in the lab! Andres, there is
something I still want to say: “Yes, I did it with the radioactive substrate and it
did not work!!”; Karin, you are the best technician and such a sweet person;
Thibaud, for your constant flow of ideas, discussions and ideas (and
discussions) (and ideas..); Stefan, you are one of a kind, I am glad that I met
you!; Rina, thanks for introducing me to bouldering; Hagen, James, Majda,
Lina, Eric, Thomas, Frank, Katja, Sandra, Dior, Marlene, Vanessa, thank you for
the fun, the outings, Metallica concert, Croatian dinner, etc... I want to thank as
well the students who helped me: Franziska, Juan, Ann, and Quentin. And of
course all the member of the EC/RIIB/MPUSP lab! Lya and Lisa, thank you for
your kindness; Teresa, Paul, Sara, thank you for the strand bar, French-German
conversations and diners! Ines (there you are ), thank you for being my friend,
for being such a lovable person, thank you for being there when I needed it and
thank you for just being yourself… Andrey and your incessant flow of fun facts,
historical facts, and fabulous facts, thank you for being part of my life and a
great friend. Laurianne, thank you for all your caring, support and friendship!
Du fond de mon coeur, merci à ma famille pour son soutien: Maman, tu es l’une
des femmes les plus solides que je connaisse; Papa, tu me manques…; mes
frères Emmanuel et Christophe, pour être là, tout simplement; Odile, Jean-
Marc, Véro et le reste de la famille. Loïc, tu étais un grand soleil dans ma vie…
Mes amies de toujours: Anso, Laura, Titia, Aude, Titi, merci d’être là, même de
loin :)
Finally, thanks to all the musicians who put their souls in their music and
release it out there…
41
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3. Van Assche E, Van Puyvelde S, Vanderleyden J, Steenackers HP (2015) RNA-binding proteins involved in post-transcriptional regulation in bacteria. Front Microbiol 6:141.
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