Download NON-CANONICAL TRANSCRIPTION INITIATION: THE EXPANDING

NON-CANONICAL TRANSCRIPTION INITIATION: THE EXPANDING UNIVERSE
OF TRANSCRIPTION INITIATION SUBSTRATES
Ivan Barvík1, Dominik Rejman2, Natalya Panova3, Hana Šanderová3, and Libor Krásný3*
1
Division of Biomolecular Physics, Institute of Physics, Faculty of Mathematics and Physics,
Charles University in Prague, Ke Karlovu 5, 121 16 Prague 2, Czech Republic.
2
Institute of Organic Chemistry and Biochemistry, Czech Academy of Sciences v.v.i., Flemingovo
nám. 2, 166 10 Prague 6, Czech Republic.
Institute of Microbiology, Czech Academy of Sciences v.v.i., Vídeňská 1083, 142 20 Prague 4,
Czech Republic.
3
* [email protected]
Abstract
RNA polymerase (RNAP) is the central enzyme of transcription of the genetic information
from DNA into RNA. RNAP recognizes four main substrates: ATP, CTP, GTP, and UTP.
Experimental evidence from the past several years suggests that, besides these four NTPs,
other molecules can be used to initiate transcription: (i) ribo-oligonucleotides (nanoRNAs),
and (ii) coenzymes such as NAD, NADH, CoA, and FAD. The presence of these molecules
at the 5’ ends of RNAs affects the properties of the RNA. Here, we discuss the expanding
portfolio of molecules that can initiate transcription, their mechanism of incorporation,
effects on RNA and cellular processes, and we present an outlook towards other possible
initiation substrates.
Key Words: RNA polymerase, non-canonical transcription initiation, nicotinamide adenine
dinucleotide (NAD), coenzymes
1
Recently, coenzymes such as nicotinamide adenine dinucleotide (NAD) and coenzyme A (CoA)
were identified to be covalently attached to the 5’ ends of specific RNAs (Cahova et al., 2015;
Kowtoniuk et al., 2009). These coenzymes contain an adenosine linked to a second moiety that is
positioned at their 5’ end (Fig. 1). In the cell, NAD is involved in redox reactions as an electron
carrier (Lin and Guarente, 2003); CoA plays a role in the synthesis and oxidation of fatty acids
(DiRusso et al., 1999). This review focuses on current structural and biochemical findings of
how these coenzymes are incorporated into RNA by RNA polymerase (RNAP), and what their
biological function(s) might be in these molecules. Of these coenzymes, the main focus is on
NAD as it is presently the best studied representative of the 5’ end modifications. This is set into
the context of the transcription machinery, and the regulation of its activity by initiating
substrates, such as initiating nucleoside triphosphates (iNTPs) and nanoRNAs (2-4 nt long
oligoribonucleotides).
Figure 1. Compounds that can be present at 5’ RNA ends. 7-methylguanosine is specifically
eukaryotic. NAD and CoA were detected to form 5’ RNA ends in vivo, while for FAD and NADH
the evidence comes only from in vitro experiments.
RNAP
To survive, the cell has to sense environmental cues and deliver the information from the outside
to the transcription machinery inside to alter gene expression. The central enzyme of
transcription in bacteria is DNA-dependent RNA polymerase. It consists of an alpha dimer that
holds together the catalytic subunits  and ’, and the small subunit  that binds to ’. This core
enzyme (2’) is conserved throughout all bacteria (Murakami, 2015). Gram positive
2
Firmicutes contain additional subunits termed  and  (Keller et al., 2014; Rabatinova et al.,
2013). The RNAP core is capable of transcription elongation but not initiation. To initiate, it
must associate with a subunit that recognizes specific sequences in the DNA, promoters. The
number of sigma factor can vary from one to over 100 (Paget, 2015). In addition, a large number
of auxiliary factors associate with RNAP and modulate its activity (Helmann, 2009).
Transcription initiation
When RNAP recognizes the promoter DNA, it first forms the so called closed complex in which
the two DNA strands are still unwound. Subsequently, RNAP isomerizes and forms the open
complex in which the transcription bubble is formed (from -11 to +2) (Ruff et al., 2015), and
scrunching of DNA is involved in transcription start site (+1) selection (Winkelman et al., 2016).
After binding the initiating nucleoside triphosphate (iNTP) to the i site (base pairing with +1 in
the template strand) the second nucleotide binds to the i+1 site (e. i. base pairing with +2) and
the first covalent bond is formed. The NTP binding, catalysis, and subsequent translocation
depend on conformational changes of the trigger loop, an important mechanistic element of
RNAP. Upon catalysis by the SN2 mechanism (Steitz et al., 1994), and pyrophosphate release,
the trigger loop of RNAP and another element, the bridge helix, change the geometry of the
active site, and move the nascent RNA by one position. During initiation, short abortive RNA
products may be generated (Pupov et al., 2014), or reiterative initiation (slippage) may occur
where RNA does not translocate but slips upstream relative to the DNA template strand, the
initiating NTP is then added again (the number of additions may vary), and, hence, the 5’ end
sequence of the resulting RNA does not match the sequence of the DNA template from which it
was derived (Turnbough, Jr., 2011). Only when RNAP severs its contacts with the promoter and
sigma factor is released, mature elongation complex is formed (Winkelman et al., 2015). Nascent
RNA is threaded through the exit channel by which it leaves RNAP (Hein et al., 2014).
We note that although bacterial RNAP is by definition a DNA dependent enzyme it can also
transcribe from an RNA template [from 6S RNA (Wehner et al., 2014)], and it tolerates a
number of DNA modifications, both natural and artificial (Raindlova et al., 2016).
Canonical substrates of RNAP and gene expression regulation by their concentration
NTPs are the canonical substrates of RNAP. Both the 5’ and 3’ ends of the iNTP are required for
its recognition by RNAP. The non-bridging oxygen of the alpha phosphate (5’ end) of the iNTP
interacts with Lys-838 and Lys-846 (Thermus thermophilus [Tth]; corresponding E. coli [Eco]
residues are Lys-1065 and Lys-1073). Discrimination between NTP and dNTP (3’ end) is
mediated by Asn737 (’, Tth) that hydrogen bonds with both O3’ and O2’ of the second NTP
and Arg704 (’; Eco Arg425) that forms hydrogen bonds with the O2’ atoms of both the iNTP
the second NTP (Basu et al., 2014; Vassylyev et al., 2007b; Vassylyev et al., 2007a) (Fig. S1).
3
The intracellular concentrations of NTPs play important regulatory roles for a number of
promoters. An example of a positive regulation by a high iNTP concentration are promoters with
relatively unstable open complexes where the time window for the entry of iNTPs is short. The
higher the concentration of the iNTP, the higher its chance to penetrate into the active site and
initiate transcription (Haugen et al., 2008). Among these promoters belong e. g. rRNA promoters
in Escherichia coli and Bacillus subtilis (Krasny and Gourse, 2004; Murray et al., 2003). In the
latter organism, the identity of the +1 position (A or G) is of pivotal importance during the
stringent response (amino acid starvation) when the concentrations of ATP and GTP change
reciprocally (Krasny et al., 2008; Tojo et al., 2010). An example of a negative regulation by a
high iNTP level is the reiterative transcription initiation of the pyrimidine biosynthetic operons
carAB and pyrBI of E. coli where high UTP causes promoter sequence-dependent slippage of the
nascent transcript and its premature release, and thus prevents unnecessary transcription of the
genes (Turnbough and Switzer, 2008). Regulation of transcription initiation by the concentration
of the iNTP was described also for eukaryotes (Kuehner and Brow, 2008).
Non-canonical substrates of RNAP (i): NanoRNAs
Besides the four NTPs, non-canonical substrates called nanoRNAs (2-4 nt long
oligoribonucleotides) can prime bacterial transcription in vitro and in vivo (Goldman et al.,
2011). NanoRNAs are generated during the degradation of cellular RNA, and their level in the
cell is controlled by oligoribonuclease Orn, an essential enzyme in E. coli (Ghosh, PNAS 1999).
Use of nanoRNAs as primers for transcription initiation is especially prevalent in stationary
phase (Vvedenskaya et al., 2012; Druzhinin et al., PLOS Genetics 2015). Furthermore, changes
in gene expression that occur as a direct consequence of nanoRNA-mediated priming of
transcription modulate biofilm formation in E. coli (Druzhinin et al., PLOS Genetics 2015).
NanoRNAs base-pair with the template strand whereby their 3’ end nucleotide aligns with the +1
position. In cells, the use of nanoRNAs as primers for transcription initiation introduces a 5’ end
hydroxyl to the resulting RNAs, which is likely to have consequences for their biological halflife. In particular, the 5’ end phosphorylation status is known to affect the stability of RNAs that
do not contain a palindromic sequence near this terminus. The triphosphate is considered as a
prokaryotic cap, stabilizing the RNA. An enzyme called RNA pyrophosphohydrolase (RppH)
removes the pyrophosphate, leaving a 5’ end monophosphate (Vasilyev and Serganov, 2015),
and thus decreases the stability of some RNAs (Mathy et al., 2007; Piton et al., 2013). To the
contrary, a 5’ hydroxyl has a stabilizing effect, rendering the RNA less susceptible to
degradation by RNases (Nickels, 2012; Celesnik, Mol cell 2007).
Non-canonical substrates of RNAP (ii): Coenzymes
In 2003 Huang demonstrated that phage T7 RNAP can use adenosine-containing coenzymes
NAD, CoA, and FAD to initiate transcription in vitro (Huang, 2003) (Fig. 1). Several years later,
it was shown in E. coli and Streptomyces venezuelae that NAD and CoA (including 3’dephospho-CoA and its succinyl-, acetyl-, and methylmalonyl-thioester) are part of cellular
4
RNAs, especially those smaller than 200 nt, covalently attached to their 5’ ends (Chen et al.,
2009; Kowtoniuk et al., 2009). The identity of the modified RNAs was unknown.
Attempts were made at establishing whether it is bacterial RNAP that could use NAD or CoA as
initiation substrates but they failed (Chen et al., 2009; Kowtoniuk et al., 2009). Therefore, it was
proposed recently for NAD that it may be the NAD synthetic machinery that attaches
nicotinamide to the RNAs posttranscriptionally (Luciano and Belasco, 2015).
Very recently, using elegant chemistry that substituted nicotinamide with 4-pentyn-1-ol,
subsequent azide-alkyne cycloaddition (azide was biotinylated) followed by next generation
sequencing, Cahova and coworkers (2015) identified specific RNAs that are modified with NAD
in E. coli. The most frequently modified RNA was RNAI (13% in exponentially growing cells in
rich media). RNAI is a plasmid-encoded small RNA (sRNA), and it is essential for plasmid
replication control (Helmercitterich et al., 1988). Other identified NAD-modified RNAs included
those involved in stress response (sRNA: DsrA, ChiX, GadY, GcvB, MgrR, and 5’ gene regions:
bsmaA, hdeD, uspE), amino acid synthesis (hisL, ilvL), and other functions, such as affecting the
5’ end phosphorylation status (rppH, see above). Importantly, other sRNAs (e. g. tRNA, 6S
RNA) were not found to be among the NAD-enriched fraction (Cahova et al., 2015), suggesting
a discriminatory mechanism for NAD incorporation.
In the cell, the nicotinamide ribonucleoside moiety can be removed by a Nudix enzyme. Nudix
enzymes are pyrophosphohydrolases that catalyze the breakage of pyrophosphate bonds in a
range of substrates (McLennan, 2013). Consistently, deletion of nudC resulted in about a twofold increase in the abundance of the modified RNAs (Cahova et al., 2015).
This recent identification of specific NAD-modified RNAs sparked new interest into the
mechanism of NAD incorporation into RNA (Jaschke et al., 2016; Marbaniang and Vogel, 2016)
and the possibility that, despite the previous unsuccessful attempts, it still might be RNAP that
could use NAD as a non-canonical transcription substrate. In favor of this hypothesis was the
ability of T7 RNAP to utilize NAD and other cofactors (Huang, 2003), the ability of RNAP to
use nanoRNAs as primers (the presence of additional nucleotides at the 5’ end does not hinder
transcription initiation) (Goldman et al., 2011), and spatio-structural considerations that argued
that NAD/CoA could be accommodated in RNAP during transcription initiation. In a recent
paper, Bird and coworkers (2016) designed new experiments and reinvestigated the possibility
that RNAP may use NAD, NADH, desphospho-CoA as substrates.
The in vitro experiments conclusively demonstrated that E. coli RNAP (in complex with the
primary sigma factor, 70) and also yeast RNA pol II can use the above mentioned coenzymes as
transcription initiation substrates, provided the +1 position encodes ATP. Nevertheless, NAD
appeared to be a less favorable substrate than ATP. The relative efficiency of NAD utilization
from the RNAI promoter that drives transcription of RNAI was 0.15 when that of ATP was set as
1. The use of iATP vs initiating NAD (iNAD) in vivo likely also depends on the relative
5
intracellular concentrations of these compounds. Both compounds are present in the cell in the
low mM range and their concentrations can change, depending on the nutritional status of the
cell. Importantly, the intracellular ratio of free NAD to NADH is ~ 100-600:1 (Lin and Guarente,
2003), suggesting that the incorporation of NADH (Fig. 1) is a rare event. CoA is present in the
cell at 20 to 600 M (Takamura and Nomura, 1988), and probably it is used less frequently than
NAD. FAD is present at an even lower concentration (~10 M) (Louie et al., 2003), making it an
even less likely substrate for RNAP in vivo. Hence, [ATP]~[NAD]>>[CoA]>[NADH]~[FAD].
So far, NAD-capping has been demonstrated with RNAP in complex with the main sigma factor.
It is possible that NAD may be an initiating substrate for RNAP associated with alternative
sigma factors. A candidate in E. coli is e. g. 54 as the promoter of the GlmY sRNA, one of the
sRNAs modified with NAD identified by Cahova and colleagues (2015), contains recognition
sequences for both 70 and 54 (Gopel et al., 2011; Papenfort and Vanderpool, 2015;
Reichenbach et al., 2009). Further research will be required to elucidate whether alternative
sigma factors allow initiation with coenzymes.
Structural consideration of NAD-capping
Bird and colleagues (2016) presented structural insights into transcription initiation with NAD
and CoA. The T. thermophilus RNAP crystal structures of the initial product complex obtained
upon initiation with NAD/CoA and CTP showed that the pC residue and the adenosine part of
NAD/CoA base pair with the DNA template strand in the RNAP active-center in positions +1
and -1, respectively, making the same interactions as RNAP initiating with ATP+CTP.
During the early phase of transcription, when the nascent RNA moves through the exit channel,
the nicotinamide ribonucleoside moiety of NAD interacts with RNAP differently than the 5’ end
triphosphate. At the stage when cytidine monophosphate is already covalently attached to NAD,
the nicotinamide ribonucleoside interacts with RNAP  Asp396, Tyr998 and His999 (Eco
Asp516, Asn1236 and His1237) (Bird et al., 2016) (Fig. 2), and it is directed toward the pocket
of RNAP that can be occupied by rifampicin-like inhibitors, which prevent the nascent RNA to
be extend beyond -3 to -6 nt (Molodtsov et al., 2013). The effect of these additional interactions
of NAD on transcription is not known, yet.
A
B
6
Figure. 2. POST-translocated NAD-CMP in the exit channel. A. C, cytosine of CMP; A, adenine
of NAD; N, nicotinamide of NAD. Carbon atoms are in yellow, nitrogen atoms in blue, oxygen
atoms in red. In green is the nascent RNA as it normally threads through the RNA exit channel.
In gray is the surface of the subunit of RNAP that forms the RNA exit channel. B. A detail of
NAD-CMP and the relevant amino acid residues that contact the nicotinamide ribonucleoside.
Another important step in transcription initiation where the 5’ end of RNA interacts with RNAP
is the moment when it contacts the 3.2 region of sigma subunits of RNAP. The 3.2 loop of sigma
70 (Tth residues 321 to 327) physically occupies the path of the elongating RNA and must be
displaced to form a 5bp POST-translocated RNA:DNA hybrid (Bae et al., 2015).The 5’triphosphate of the 5nt-long PRE-translocated RNA appears to contact the acidic sigma 3.2 loop
as shown by the well-defined electron density bridging these two elements (Zuo and Steitz,
2015). In contrast, the complex with a 6 nt-long RNA reveals that the 5’-end of the RNA
transcript contacts sigma region 3.2 causing the tip of this region (residues 321-327) to become
disordered. This is likely due to the charge repulsion between the acidic cluster of the sigma
region 3.2 and the 5’-triphosphate of RNA as it pushes away the sigma factor (Basu et al., 2014)
(Fig. S2). Possibly, an RNA with a positively charged 5’ end, due to the presence of
nicotinamide, may form a more stable interaction with the negatively charged 3.2 loop tip and
alter the kinetics of sigma displacement.
The sequence/kinetic determinants of coenzyme incorporation into RNA are mostly unknown.
Alignment of promoter regions of genes determined to contain NAD at the 5’ ends of their RNAs
revealed that they contained no obvious sequence motif that distinguished them from other
promoters (Cahova et al., 2015). Nevertheless, Bird and colleagues (2016) showed that mutating
the -1 position affects the affinity of RNAP for NAD and suggested that the nicotine amide
moiety forms “pseudo-base pair” with the -1 base of the template strand. Further experimental
work is needed to fully understand the structural requirements for NAD accommodation.
Biological function(s) of the presence of coenzymes at the 5’ RNA ends
Previously, besides its electron carrier function, NAD was also reported to play a role in
transcription regulation as a cofactor of eukaryotic sirtuins that deacetylate histones and thereby
inhibit transcription (Chen et al., 2015). To the contrary, the biological role of the presence of the
coenzymes at the 5’ end of RNAs is only now beginning to be unraveled. It is possible that in
some cases, where the percentage of NAD capping is low, it is merely a byproduct of the
promiscuous ability of RNAP to utilize these compounds as initiating substrates, and it may lack
a physiological role. However, in vitro and in vivo experiments demonstrated a markedly
7
improved stability of NAD-modified RNAs (Bird et al., 2016; Cahova et al., 2015). RNases that
depend on the 5’ end RNA status are RNase E and RNase J1. E. coli RNase E cleaves RNA
internally but for optimal activity it requires the 5’ terminus of the substrate to be
monophosphorylated (Callaghan et al., 2005). Likewise, B. subtilis RNase J1 (5’-3’ exo) requires
a 5’end monophosphate (Mathy et al., 2007). In this respect NAD reminds of 7-methylguanosine
(Fig. 1) of eukaryotic mRNAs that (among other functions) increases mRNA stability
(Topisirovic et al., 2011). The percentage of NAD capping increases in stationary phase (Bird et
al., 2016) when the gene expression pattern is being reprogrammed and many RNAs are
degraded. Thus, NAD capping of RNA may contribute to survival of the cell under adverse
conditions.
The eukaryotic cap is critical for translation. In bacteria, such a possibility is more difficult to
envision. Nevertheless, here we outline a potential scenario that involves the S1 ribosomal
protein. E. coli S1 acts at the interface of transcription, translation and mRNA decay, and copurifies with RNA polymerase in stoichiometric amounts (Hajnsdorf and Boni, 2012). In E. coli
and other gram-negative bacteria, S1 is composed of six homologous repeats (72-75 amino acid
residues each), called the S1 motifs or S1 domains. RNase E, mentioned above, contains an S1
domain that binds RNA and helps orient its 5’ end for recognition by the enzyme. During
translation the ribosomal S1 protein binds mRNA and is important for translation of mRNAs
with weak ribosome binding sites (Delvillani et al., 2011; Malys and McCarthy, 2011).
Hypothetically, during translation initiation the RNA 5’ end (tri-, monophosphate, NAD) may be
recognized by one of the S1 domains and affect binding of the particular mRNA to the ribosome,
and the status of the 5’ end may play a role in mRNA recognition. The possible binding of NAD
to the S1 motif is supported by the recent finding that pyrazinoic acid (POA), the active form of
pyrazinamide (PZA, a first-line drug for tuberculosis treatment), binds to Mycobacterium
tubercolosis S1 (RpsA) (Yang et al., 2015). Interestingly, there is apparent structural similarity
of PZA/POA with the nicotinamide/nicotinate moiety of NAD (Fig. 3).
8
Figure 3. Structures of pyrazinamide,
pyrazinoic acid, nicotinamide and
nicotinate.
Finally, the ability of yeast RNA pol II to use CoA and NAD as transcription initiation substrates
in vitro strongly suggests that RNAs modified with these coenzymes may be present in
eukaryotic cells. The identity of these transcripts is unknown and it is of high interest to
determine whether such modified RNAs are present in eukaryotic cells including mitochondria,
or whether they may be present also in viral/phage derived RNAs and possibly play a role in
pathogenesis.
Other possible substrates for RNAP
In theory any molecule with attached nucleotide containing free 3’-hydroxyl group may serve as
a transcription initiation substrate provided its 5’ moiety is not too bulky to fit into RNAP. One
such a family of molecules are dinucleoside polyphosphates (ApxN) (Ismail et al., 2003). In
ApxN two nucleoside moieties are linked 5’-5’ via a polyphosphate chain containing two to
seven phosphoryl groups (Fig. 4). The dinucleoside polyphosphates (ApxN) are ubiquitous noncanonical nucleotides found in all types of cells in micromolar to submicromolar concentrations.
The low concentration of ApxN (similar to those of NADH and FAD) makes them less likely
substrates for RNAP in vivo. However, it was reported that their concentrations increased during
oxidative stress in Helicobacter pylori (Lundin et al., 2003), and it may be relevant also for other
species.
9
Figure 4. Structure of dinucleoside polyphosphates (ApxN). B (A, G, U, C) indicates any of the
four bases.
CONCLUDING COMMENTS
RNAP can use, besides its canonical substrates and nanoRNAs, NAD and desphospho-CoA as
transcription initiation substrates in vivo. In addition, there is a number of other molecules that
could be used by RNAP in a similar manner. These include FAD, NADH, and ApxN. Thus,
RNAP has a relatively wide repertoire of molecules to select from to initiate transcription
(Fig.5). The competition between these compounds is likely affected both by their relative
intracellular concentrations and the not-yet-identified promoter sequence/kinetic properties of the
resulting complexes with RNAP.
A relatively small fraction of a given cellular RNA bears these modifications in exponentially
growing cells. However, this fraction increases in stationary phase and it will be of interest to
determine how the frequency of coenzyme incorporation into RNA depends on various types of
stress and/or environmental changes. The NAD capping stabilizes RNA in vivo and it still
remains to be demonstrated what the physiological consequences are. Nevertheless, it is possible
to envision the use of synthetic promoters that would favor initiation with NAD in
biotechnological applications to boost the stability of the mRNA and subsequently increase the
production of the protein of choice.
Finally, it is desirable to identify whether such (or similar) modifications of RNA are present
also in other domains of life (Archea, Eukarya). Hence, in the near future, we anticipate an
explosion of research activity into this new phenomenon of 5’ end modified RNAs.
NAD
10
ATP
RNAP

DNA
CoA
+1A
NADH
5’ NAD
5’ PPP
5’ P
RNA
ApxN
stability
FAD
Figure 5. Known (ATP, NAD, CoA) and hypothetical substrates (NADH, FAD, ApxN) of RNAP
in vivo at promoters where +1 encodes adenine. The size of the circles represents the relative
pools of the inscribed molecules in the cell (not drawn to scale). In the inset, three types of 5’
RNA ends are depicted in brown; the triangle on the right hand side shows their relative
stability.
11
12
Figure S1. Interactions between RNAP, iNTP and the second NTP in i (+1) and i+1 (+2)
positions. The depicted lysines from  coordinate the a-phosphates. The discrimination for
ribonucleosides is mediated by R704 and N737 from ’. The blue spheres are the two Mg+II ions.
13
a)
b)
c)
Figure S2.
a) The 5’ end of RNA contacts the 3.2 region of sigma subunits of RNAP.
b) The 3.2 loop of sigma 70 physically occupies the path of the elongating RNA and must be
displaced to form a 5bp POST-translocated RNA:DNA hybrid.
c) The complex with a 6 nt-long RNA reveals that the 5’-end of the RNA transcript contacts
sigma region 3.2 causing the tip of this region to become disordered.
14
Reference List
Bae,B., Feklistov,A., Lass-Napiorkowska,A., Landick,R., and Darst,S.A. (2015). Structure of a
bacterial RNA polymerase holoenzyme open promoter complex. Elife in press.
Basu,R.S., Warner,B.A., Molodtsov,V., Pupov,D., Esyunina,D., Fernandez-Tornero,C.,
Kulbachinskiy,A., and Murakami,K.S. (2014). Structural Basis of Transcription Initiation by
Bacterial RNA Polymerase Holoenzyme. Journal of Biological Chemistry 289, 24549-24559.
Bird,J.G., Zhang,Y., Tian,Y., Panova,N., Barvík,I., Greene,L., Liu,M., Buckley,B., Krásný,L.,
Lee,J.K., Kaplan,C.D., Ebright,R.H., and Nickels,B.E. (2016). The mechanism of RNA 5'
capping with NAD+, NADH, and desphospho-CoA. Nature.
Cahova,H., Winz,M.L., Hofer,K., Nubel,G., and Jaschke,A. (2015). NAD captureSeq indicates
NAD as a bacterial cap for a subset of regulatory RNAs. Nature 519, 374-377.
Callaghan,A.J., Marcaida,M.J., Stead,J.A., McDowall,K.J., Scott,W.G., and Luisi,B.F. (2005).
Structure of Escherichia coli RNase E catalytic domain and implications for RNA turnover.
Nature 437, 1187-1191.
Chen,B., Zang,W.W., Wang,J., Huang,Y.J., He,Y.H., Yan,L.L., Liu,J.J., and Zheng,W.P. (2015).
The chemical biology of sirtuins. Chemical Society Reviews 44, 5246-5264.
Chen,Y.G., Kowtoniuk,W.E., Agarwal,I., Shen,Y.H., and Liu,D.R. (2009). LC/MS analysis of
cellular RNA reveals NAD-linked RNA. Nature Chemical Biology 5, 879-881.
Delvillani,F., Papiani,G., Deho,G., and Briani,F. (2011). S1 ribosomal protein and the interplay
between translation and mRNA decay. Nucleic Acids Res. 39, 7702-7715.
DiRusso,C.C., Black,P.N., and Weimar,J.D. (1999). Molecular inroads into the regulation and
metabolism of fatty acids, lessons from bacteria. Progress in Lipid Research 38, 129-197.
Goldman,S.R., Sharp,J.S., Vvedenskaya,I.O., Livny,J., Dove,S.L., and Nickels,B.E. (2011).
NanoRNAs Prime Transcription Initiation In Vivo. Molecular Cell 42, 817-825.
Gopel,Y., Luttmann,D., Heroven,A.K., Reichenbach,B., Dersch,P., and Gorke,B. (2011).
Common and divergent features in transcriptional control of the homologous small RNAs GlmY
and GlmZ in Enterobacteriaceae. Nucleic Acids Res. 39, 1294-1309.
Hajnsdorf,E. and Boni,I.V. (2012). Multiple activities of RNA-binding proteins S1 and Hfq.
Biochimie 94, 1544-1553.
Haugen,S.P., Ross,W., and Gourse,R.L. (2008). Advances in bacterial promoter recognition and
its control by factors that do not bind DNA. Nature Reviews Microbiology 6, 507-519.
15
Hein,P.P., Kolb,K.E., Windgassen,T., Bellecourt,M.J., Darst,S.A., Mooney,R.A., and Landick,R.
(2014). RNA polymerase pausing and nascent-RNA structure formation are linked through
clamp-domain movement. Nature Structural & Molecular Biology 21, 794-802.
Helmann,J.D. (2009). RNA polymerase: A nexus of gene regulation Introduction. Methods 47,
1-5.
Helmercitterich,M., Anceschi,M.M., Banner,D.W., and Cesareni,G. (1988). Control of Cole1
Replication - Low Affinity Specific Binding of Rop (Rom) to Rnai and Rnaii. Embo Journal 7,
557-566.
Huang,F.Q. (2003). Efficient incorporation of CoA, NAD and FAD into RNA by in vitro
transcription. Nucleic Acids Research 31.
Ismail,T.M., Hart,C.A., and McLennan,A.G. (2003). Regulation of dinucleoside polyphosphate
pools by the YgdP and ApaH hydrolases is essential for the ability of Salmonella enterica
serovar Typhimurium to invade cultured mammalian cells. Journal of Biological Chemistry 278,
32602-32607.
Jaschke,A., Hofer,K., Nubel,G., and Frindert,J. (2016). Cap-like structures in bacterial RNA and
epitranscriptomic modification. Curr. Opin. Microbiol. 30, 44-49.
Keller,A.N., Yang,X., Wiedermannova,J., Delumeau,O., Krasny,L., and Lewis,P.J. (2014).
epsilon, a New Subunit of RNA Polymerase Found in Gram-Positive Bacteria. Journal of
Bacteriology 196, 3622-3632.
Kowtoniuk,W.E., Shen,Y.H., Heemstra,J.M., Agarwal,I., and Liu,D.R. (2009). A chemical
screen for biological small molecule-RNA conjugates reveals CoA-linked RNA. Proceedings of
the National Academy of Sciences of the United States of America 106, 7768-7773.
Krasny,L. and Gourse,R.L. (2004). An alternative strategy for bacterial ribosome synthesis:
Bacillus subtilis rRNA transcription regulation. Embo Journal 23, 4473-4483.
Krasny,L., Tiserova,H., Jonak,J., Rejman,D., and Sanderova,H. (2008). The identity of the
transcription+1 position is crucial for changes in gene expression in response to amino acid
starvation in Bacillus subtilis. Molecular Microbiology 69, 42-54.
Kuehner,J.N. and Brow,D.A. (2008). Regulation of a eukaryotic gene by GTP-dependent start
site selection and transcription attenuation. Molecular Cell 31, 201-211.
Lin,S.J. and Guarente,L. (2003). Nicotinamide adenine dinucleotide, a metabolic regulator of
transcription, longevity and disease. Current Opinion in Cell Biology 15, 241-246.
Louie,T.M., Xie,X.S., and Xun,L.Y. (2003). Coordinated production and utilization of FADH(2)
by NAD(P)H-flavin oxidoreductase and 4-hydroxyphenylacetate 3-monooxygenase.
Biochemistry 42, 7509-7517.
16
Luciano,D.J. and Belasco,J.G. (2015). NAD in RNA: unconventional headgear. Trends in
Biochemical Sciences 40, 245-247.
Lundin,A., Nilsson,C., Gerhard,M., Andersson,D.I., Krabbe,M., and Engstrand,L. (2003). The
NudA protein in the gastric pathogen Helicobacter pylori is an ubiquitous and constitutively
expressed dinucleoside polyphosphate hydrolase. Journal of Biological Chemistry 278, 1257412578.
Malys,N. and McCarthy,J.E. (2011). Translation initiation: variations in the mechanism can be
anticipated. Cell Mol Life Sci. 68, 991-1003.
Marbaniang,C.N. and Vogel,J. (2016). Emerging roles of RNA modifications in bacteria. Curr.
Opin. Microbiol. 30, 50-57.
Mathy,N., Benard,L., Pellegrini,O., Daou,R., Wen,T.Y., and Condon,C. (2007). 5 '-to-3 '
exoribonuclease activity in Bacteria: Role of RNase J1 in rRNA maturation and 5 ' stability of
mRNA. Cell 129, 681-692.
McLennan,A.G. (2013). Substrate ambiguity among the nudix hydrolases: biologically
significant, evolutionary remnant, or both? Cellular and Molecular Life Sciences 70, 373-385.
Molodtsov,V., Nawarathne,I.N., Scharf,N.T., Kirchhoff,P.D., Showalter,H.D.H., Garcia,G.A.,
and Murakami,K.S. (2013). X-ray Crystal Structures of the Escherichia coli RNA Polymerase in
Complex with Benzoxazinorifamycins. Journal of Medicinal Chemistry 56, 4758-4763.
Murakami,K.S. (2015). Structural biology of bacterial RNA polymerase. Biomolecules 5, 848862.
Murray,H.D., Schneider,D.A., and Gourse,R.L. (2003). Control of rRNA expression by small
molecules is dynamic and nonredundant. Molecular Cell 12, 125-134.
Nickels,B.E. (2012). A new way to start: NanoRNA-mediated priming of transcription initiation.
Transcription 3, 300-304.
Paget,M.S. (2015). Bacterial Sigma Factors and Anti-Sigma Factors: Structure, Function and
Distribution. Biomolecules 5, 1245-1265.
Papenfort,K. and Vanderpool,C.K. (2015). Target activation by regulatory RNAs in bacteria.
FEMS Microbiol. Rev. 39, 362-378.
Piton,J., Larue,V., Thillier,Y., Dorleans,A., Pellegrini,O., Sierra-Gallay,I.L., Vasseur,J.J.,
Debart,F., Tisne,C., and Condon,C. (2013). Bacillus subtilis RNA deprotection enzyme RppH
recognizes guanosine in the second position of its substrates. Proceedings of the National
Academy of Sciences of the United States of America 110, 8858-8863.
Pupov,D., Kuzin,I., Bass,I., and Kulbachinskiy,A. (2014). Distinct functions of the RNA
polymerase Sigma subunit region 3.2 in RNA priming and promoter escape. Nucleic Acids
Research 42, 4494-4504.
17
Rabatinova,A., Sanderova,H., Matejckova,J.J., Korelusova,J., Sojka,L., Barvik,I.,
Papouskova,V., Sklenar,V., Zidek,L., and Krasny,L. (2013). The delta Subunit of RNA
Polymerase Is Required for Rapid Changes in Gene Expression and Competitive Fitness of the
Cell. Journal of Bacteriology 195, 2603-2611.
Raindlova,V., Janouskova,M., Slavickova,M., Perlikova,P., Bohacova,S., Milisavljevic,N.,
Sanderova,H., Benda,M., Barvik,I., Krasny,L., and Hocek,M. (2016). Influence of major-groove
chemical modifications of DNA on transcription by bacterial RNA polymerases. Nucleic Acids
Res. 44, 3000-3012.
Reichenbach,B., Gopel,Y., and Gorke,B. (2009). Dual control by perfectly overlapping sigma
54- and sigma 70- promoters adjusts small RNA GlmY expression to different environmental
signals. Mol Microbiol. 74, 1054-1070.
Ruff,E.F., Record,M.T.Jr., and Artsimovitch,I. (2015). Initial Events in Bacterial Transcription
Initiation. Biomolecules 5, 1035-1062.
Steitz,T.A., Smerdon,S.J., Jager,J., and Joyce,C.M. (1994). A Unified Polymerase Mechanism
for Nonhomologous Dna and Rna-Polymerases. Science 266, 2022-2025.
Takamura,Y. and Nomura,G. (1988). Changes in the Intracellular Concentration of Acetyl-Coa
and Malonyl-Coa in Relation to the Carbon and Energy-Metabolism of Escherichia-Coli-K12.
Journal of General Microbiology 134, 2249-2253.
Tojo,S., Kumamoto,K., Hirooka,K., and Fujita,Y. (2010). Heavy Involvement of Stringent
Transcription Control Depending on the Adenine or Guanine Species of the Transcription
Initiation Site in Glucose and Pyruvate Metabolism in Bacillus subtilis. Journal of Bacteriology
192, 1573-1585.
Topisirovic,I., Svitkin,Y.V., Sonenberg,N., and Shatkin,A.J. (2011). Cap and cap-binding
proteins in the control of gene expression. Wiley Interdisciplinary Reviews-Rna 2, 277-298.
Turnbough,C.L., Jr. (2011). Regulation of gene expression by reiterative transcription. Curr.
Opin. Microbiol. 14, 142-147.
Turnbough,C.L. and Switzer,R.L. (2008). Regulation of pyrimidine biosynthetic gene expression
in bacteria: Repression without repressors. Microbiology and Molecular Biology Reviews 72,
266-300.
Vasilyev,N. and Serganov,A. (2015). Structures of RNA Complexes with the Escherichia coil
RNA Pyrophosphohydrolase RppH Unveil the Basis for Specific 5 '-End-dependent mRNA
Decay. Journal of Biological Chemistry 290, 9487-9499.
Vassylyev,D.G., Vassylyeva,M.N., Perederina,A., Tahirov,T.H., and Artsimovitch,I. (2007a).
Structural basis for transcription elongation by bacterial RNA polymerase. Nature 448, 157-1U3.
18
Vassylyev,D.G., Vassylyeva,M.N., Zhang,J.W., Palangat,M., Artsimovitch,I., and Landick,R.
(2007b). Structural basis for substrate loading in bacterial RNA polymerase. Nature 448, 1631U4.
Vvedenskaya,I.O., Sharp,J.S., Goldman,S.R., Kanabar,P.N., Livny,J., Dove,S.L., and
Nickels,B.E. (2012). Growth phase-dependent control of transcription start site selection and
gene expression by nanoRNAs. Genes & Development 26, 1498-1507.
Wehner,S., Damm,K., Hartmann,R.K., and Marz,M. (2014). Dissemination of 6S RNA among
Bacteria. Rna Biology 11, 1468-1479.
Winkelman,J.T., Vvedenskaya,I.O., Zhang,Y., Zhang,Y., Bird,J.G., Taylor,D.M., Gourse,R.L.,
Ebright,R.H., and Nickels,B.E. (2016). Multiplexed protein-DNA cross-linking: Scrunching in
transcription start site selection. Science 351, 1090-1093.
Winkelman,J.T., Winkelman,B.T., Boyce,J., Maloney,M.F., Chen,A.Y., Ross,W., and
Gourse,R.L. (2015). Crosslink Mapping at Amino Acid-Base Resolution Reveals the Path of
Scrunched DNA in Initial Transcribing Complexes. Mol Cell 59.
Yang,Y., Darbari,V.C., Zhang,N., Lu,D., Glyde,R., Wang,Y.P., Winkelman,J.T., Gourse,R.L.,
Murakami,K.S., Buck,M., and Zhang,X. (2015). TRANSCRIPTION. Structures of the RNA
polymerase-sigma54 reveal new and conserved regulatory strategies. Science 349, 882-885.
Zuo,Y.H. and Steitz,T.A. (2015). Crystal Structures of the E. coli Transcription Initiation
Complexes with a Complete Bubble. Molecular Cell 58, 534-540.
19