Mycobacterium smegmatis RNase J is a 5′-3′ exo-/endoribonuclease and both RNase J and RNase E are involved in ribosomal RNA maturation


  • Valerio Taverniti,

    1. Department of Biomolecular Sciences and Biotechnology, University of Milano, Via Celoria 26, 20133 Milano, Italy
    2. CNRS UPR 9073 (affiliated with Univ Paris Diderot, Sorbonne Paris Cité), Institut de Biologie Physico-Chimique, 13 rue Pierre et Marie Curie, 75005 Paris, France
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  • Francesca Forti,

    1. Department of Biomolecular Sciences and Biotechnology, University of Milano, Via Celoria 26, 20133 Milano, Italy
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  • Daniela Ghisotti,

    1. Department of Biomolecular Sciences and Biotechnology, University of Milano, Via Celoria 26, 20133 Milano, Italy
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  • Harald Putzer

    Corresponding author
    1. CNRS UPR 9073 (affiliated with Univ Paris Diderot, Sorbonne Paris Cité), Institut de Biologie Physico-Chimique, 13 rue Pierre et Marie Curie, 75005 Paris, France
      E-mail; Tel. (+33) 1 58 41 51 27; Fax (+33) 1 58 41 50 20.
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E-mail; Tel. (+33) 1 58 41 51 27; Fax (+33) 1 58 41 50 20.


The presence of very different sets of enzymes, and in particular the presence of RNase E and RNase J, has been used to explain significant differences in RNA metabolism between the two model organisms Escherichia coli and Bacillus subtilis. However, these studies might have somewhat polarized our view of RNA metabolism. Here, we identified a RNase J in Mycobacterium smegmatis that has both 5′-3′ exo- and endonucleolytic activity. This enzyme coexists with RNase E in this organism, a configuration that enabled us to study how these two key nucleases collaborate. We demonstrate that RNase E is responsible for the processing of the furA-katG transcript in M. smegmatis and that both RNase E and RNase J are involved in the 5′ end processing of all ribosomal RNAs. In contrast to B. subtilis, the activity of RNase J, although required in vivo for 23S rRNA maturation, is not essential in M. smegmatis. We show that the pathways for ribosomal RNA maturation in M. smegmatis are quite different from those observed in E. coli and in B. subtilis. Studying organisms containing different combinations of key ribonucleases can thus significantly broaden our view of the possible strategies that exist to direct RNA metabolism.


Ribonucleases are important enzymes that intervene in the degradation, processing and quality control of all RNA species in the cell. These vital processes play an essential role in the control of gene expression and in the adaptation to environmental changes. Current knowledge suggests that the RNases E, J and Y are key players in eubacterial RNA metabolism. All known eubacteria contain at least one of these three ribonucleases and several of them have all three (Laalami and Putzer, 2011).

mRNA degradation/maturation has been extensively studied in Escherichia coli and Bacillus subtilis. Important differences in the mechanisms underlying mRNA metabolism have been demonstrated between these two model organisms and reflect the fact that they contain quite different sets of RNases. In the Gram-negative bacterium E. coli, RNase E is the key enzyme initiating mRNA decay. It generally catalyses a rate limiting endonucleolytic cleavage within an AU rich single-stranded region, causing the rapid 3′-5′ exonucleolytic degradation of the fragment upstream of the cleavage site (Carpousis et al., 2009). The strong preference of RNase E for 5′ monophosphorylated RNA substrates (Mackie, 1998) stimulates the subsequent endonucleolytic cleavage of the 3′ cleavage products. In some cases, this preference allows an alternative decay pathway in which internal cleavage by RNase E is triggered by prior conversion of the 5′ terminal triphosphate to a monophosphate by the pyrophosphohydrolase RppH (Deana et al., 2008). Inactivation of RNase E increases global mRNA half-life, and the non-catalytic C-terminal half of RNase E serves as the scaffold for the degradosome complex (Carpousis et al., 2009). RNase E also plays a major role in the maturation of ribosomal RNA (Deutscher, 2009). RNase G, a shorter paralogue of RNase E, which coexists with RNase E only in the β- and γ-subdivisions of the Proteobacteria (Condon and Putzer, 2002), plays a minor role in mRNA metabolism in E. coli (Arraiano et al., 2010). RNase E orthologues in other species are classified as RNase E/G enzymes because they usually share significant similarity with the catalytic N-terminal half of E. coli RNase E and RNase G (Condon and Putzer, 2002).

Despite its central role in mRNA stability in E. coli, RNase E is absent from many bacterial species including most Firmicutes, such as B. subtilis, and even some proteobacteria (Condon and Putzer, 2002; Laalami and Putzer, 2011). These bacteria instead contain the dual activity endo-/5′-3′ exoribonuclease RNase J (Even et al., 2005; Mathy et al., 2007) and/or the endonuclease RNase Y (Shahbabian et al., 2009). For example, in B. subtilis the essential RNase J1 (rnjA) and its paralogue RNase J2 (rnjB) have endonucleolytic cleavage specificity similar to that of RNase E (Even et al., 2005). RNase J1 and to a lesser extent RNase J2 also possess a 5′-3′ exonucleolytic activity that strongly prefers a substrate with a single phosphate at the 5′ end (Li de la Sierra-Gallay et al., 2008; Mathy et al., 2010). This property fitted well with the observation that, in B. subtilis, obstacles such as a stalled ribosome or secondary structure near the 5′ end can strongly stabilize downstream RNA for several kilobases indicative of a strong 5′-3′ directionality for mRNA degradation (Bechhofer and Dubnau, 1987; Agaisse and Lereclus, 1996; Hambraeus et al., 2002). Indeed, RNases J1 and J2, which have overlapping substrate specificities, together affect the expression levels of hundreds of genes (Mäder et al., 2008). The exonucleolytic activity of RNase J1 also participates in 16S rRNA maturation (Mathy et al., 2007). On the other hand, in a rnjA/rnjB double mutant (RNase J1 was depleted) global mRNA half-life was only slightly increased (Even et al., 2005). However, 5′ exonucleolytic degradation of a native transcript can be stimulated by a recently discovered RppH-like activity in B. subtilis (Richards et al., 2011). To what extent pyrophosphate removal from the 5′ end of original transcripts affects mRNA metabolism in B. subtilis remains to be analysed. The most significant effect on mRNA turnover in this organism can be attributed to RNase Y, a recently characterized novel endoribonuclease whose depletion increases bulk mRNA stability in B. subtilis to a similar extent as that of RNase E mutants in E. coli (Shahbabian et al., 2009). Transcriptome studies confirmed the important role of RNase Y in determining the stability of hundreds of mRNA species (Lehnik-Habrink et al., 2011) (our unpubl. results). In addition, the activity of RNase Y is significantly increased by the presence of a 5′-P group on the RNA substrate, another similarity shared with RNase E (Shahbabian et al., 2009). These observations raised the possibility that, after all, mRNA processing and degradation might be more similar between B. subtilis and E. coli than currently assumed, with an endonucleolytic cleavage being the key step in initiating mRNA decay.

In eubacteria the RNases E/G, J and Y probably represent the key enzymes initiating mRNA decay (Laalami and Putzer, 2011). In this context, mycobacteria are especially interesting to study because a genome sequence analysis shows that they contain both an RNase E/G and an RNase J but no RNase Y (Table 1). This combination has not been found in any of the organisms where RNA metabolism has been studied in some detail.

Table 1.  Occurrence of ribonucleases.
 E. coliB. subtilisM. smegmatisM. tuberculosis
  1. The sequences of the E. coli and B. subtilis ribonucleases were downloaded from the Comprehensive Microbial Resource database (CMR: The homology analysis was carried out (using BLAST program: by matching the known sequences with the whole mycobacterial genomes.

 RNase E+++
 RNase G+
 RNase III++++
 Mini III+
 RNase M5+
 RNase P++++
 RNase Z++++
 RNase I+
 RNase Y+
 RNase J+++
 RNase J+++
 RNase T+
 RNase R++
 RNase II+
 RNase PH+++

Mycobacteria can be divided into two classes: the slow-growers that include pathogens like Mycobacterium tuberculosis and Mycobacterium leprae and the fast-growers like Mycobacterium smegmatis. It is noteworthy that the putative RNases present in M. tuberculosis and M. smegmatis are the same (Table 1). This implies that RNA metabolism is likely to be very similar and studies in M. smegmatis, which is nonpathogenic and simpler to handle than M. tuberculosis, should be pertinent for both species.

Mycobacterium smegmatis has functional orthologues for both RNase E (Zeller et al., 2007) and, as we show here, RNase J. We demonstrate that the RNase J orthologue is a dual activity 5′ exo- and endonuclease very similar to RNase J1 of B. subtilis. To analyse in more detail the relative roles of RNase J and RNase E in vivo in M. smegmatis, we created mutant strains and studied their effect on the maturation of the ribosomal RNA and processing of the furA-katG operon mRNA. Our results show that rRNA maturation in M. smegmatis is different from that in both E. coli and B. subtilis.


The M. smegmatis genome encodes a protein with high similarity to B. subtilis RNases J1 and J2

RNases J1 and J2 were discovered in B. subtilis but orthologous enzymes are present in about half of the sequenced bacterial and archeal genomes (Even et al., 2005). A blast search with the B. subtilis RNase J1 and J2 sequences against protein sequences encoded by the M. smegmatis mc2155 chromosome (Fig. S1) identified a single gene product (MSMEG_2685) with a high similarity over its entire length with both RNases J1 and J2. The 558 amino acids long mycobacterial protein shares 35% identical residues with RNase J1 (55% similarity) and 33% identical residues with RNase J2 (59% similarity). Analysis of the amino acid sequence revealed the presence of all known motifs important for the ribonuclease activity (Li de la Sierra-Gallay et al., 2008) and clearly categorizes this mycobacterial protein as a member of the β-CASP subfamily of zinc-dependent metallo-β-lactamases (Callebaut et al., 2002). Based on the recently resolved 3D structure of the Thermus thermophilus RNase J (Fig. S2A (Li de la Sierra-Gallay et al., 2008) we deduced the 3D structures of B. subtilis RNase J1 (Fig. S2B) and the putative M. smegmatis RNase J (Fig. S2C). The three structures are very similar and can practically be superposed. Minor differences observed for the mycobacterial protein with respect to the other two RNases include a somewhat less structured linker (shorter α-helix) and C-terminal domain where only the straight part of the kinked helix found in RNase J1 is present (Fig. S2).

The MSMEG_2685 protein has 5′-3′ exoribonucleolytic activity

The 5′ exonucleolytic activity of B. subtilis RNase J1 requires a 5′ monophosphorylated RNA; the γ-phosphate of a 5′-PPP terminal nucleotide in the phosphate binding pocket would place the scissile bond out of phase (Li de la Sierra-Gallay et al., 2008). We used the well characterized B. subtilis thrS leader RNA to assay the M. smegmatis enzyme activity in vitro. A 280 nt long RNA, which comprises thrS leader sequence and extends to include the intrinsic terminator structure, was synthesized in vitro, labelled at its 5′ end using kinase and γ-32P-ATP, and incubated with the purified MSMEG_2685 protein for up to 30 min (see Experimental procedures). Radioactive mononucleotides accumulated in a time-dependent manner with the bulk of the mononucleotides liberated after about 10 min (Fig. 1 lanes 5–8). A very similar activity pattern was obtained with purified B. subtilis RNase J1 (Fig. 1, compare lanes 5–8 with lanes 9–10). In order to confirm that the exoribonucleolytic activity was carried by the mycobacterial protein MSMEG_2685, we also tested a mutant form of the protein, which carries two substitutions, Asp85Lys and His86Ala (see Experimental procedures for the construction). Both amino acids are conserved in the B. subtilis and the T. thermophilus RNase J proteins and are important for enzyme function. They are directly implicated in the co-ordination of one of the two Zn ions in the catalytic centre (Li de la Sierra-Gallay et al., 2008). The mutant MSMEG_2685 protein was severely impaired in its 5′ exonucleolytic activity liberating only very low levels of mononucleotides even after 30 min (Fig. 1, lanes 20–22). We thus considered the MSMEG_2685 protein to be an RNase J orthologue.

Figure 1.

M. smegmatis RNase J endo- and exo-ribonucleolytic activity in vitro. The B. subtilis thrS leader transcript was used as substrate for the activity assays. Both M. smegmatis wild type RNase J (J) and an active site mutant (D85K and H86A) protein (J*) were used. B. subtilis RNase J1 (J1) was used as a control. Each reaction was stopped at the times indicated on top of the lanes (in min) and the reaction products were separated on a 20% (w/v) denaturing polyacrylamide gel. The 5′ monophosphorylated thrS transcript (5′-P), labelled at its 5′ end (see Experimental procedures) was used to monitor the exonucleolytic activity: J = M. smegmatis wild-type RNase J (lanes 5–8), J1 = B. subtilis RNase J1 (lanes 9 and 10) and J* = mutant RNase J (lanes 20 −22). The transcript was incubated for 30 min with the reaction buffer as control (lane 4). The 5′ triphosphorylated thrS transcript (5′-PPP), labelled at its 5′ end with γ-32P GTP, was used to monitor the endonucleolytic activity of M. smegmatis wild-type RNase J (lanes 11–14), B. subtilis RNase J1 (lanes 15 and 16) and the mutant RNase J* (lanes 17–19). As a control the purified transcript was migrated before and after the 30 min incubation with the reaction buffer (lanes 2 and 3). M: the 5′ monophosphorylated thrS transcript was incubated with an alkaline buffer to obtain a nucleotide-by-nucleotide ladder (lane 1). The positions corresponding to the nucleotides GTP and GMP, respectively, are indicated.

M. smegmatis RNase J has endoribonucleolytic activity

The B. subtilis RNases J1 and J2 were initially identified as endoribonucleases that can cleave the thrS and thrZ leader RNAs (Even et al., 2005). We therefore chose the same thrS leader RNA and reaction conditions that were used for the exonuclease assay, to detect a potential endoribonucleolytic activity of the mycobacterial RNase J in vitro. In order to avoid exonucleolytic digestion of the initial transcript, the thrS leader RNA was 5′ triphosphorylated by end-labelling the in vitro transcript with γ-32P-GTP.

A major cleavage of the thrS leader RNA in the presence of M. smegmatis RNase J was observed at position +245, immediately upstream of the terminator structure (Fig. 2A, lane 3). In a control experiment B. subtilis RNase J1 was shown to cleave at the same site (Fig. 2A, lane 5), which was previously shown to be the major cleavage position in the thrS leader RNA for B. subtilis RNases J1 and J2 (Even et al., 2005). We conclude that M. smegmatis RNase J is a dual activity 5′ exo- and endoribonuclease with a cleavage specificity identical or very similar to that of B. subtilis RNase J1.

Figure 2.

Characterization of RNase J in vitro endoribonucleolytic activity. The B. subtilis thrS leader transcript (thrS native) and its modified derivatives thrS+3, thrS+5, thrS+10, carrying polyA extensions (Fig. S5) were synthesized as described in Experimental procedures, 5′ end-labelled and incubated with M. smegmatis RNase J (J), Bsubtilis RNase J1 (J1) or with the reaction buffer as control. The reactions were stopped at the times indicated on top of the lanes (in min). A. Resolution of reaction products on a 5% denaturing polyacrylamide gel together with a DNA molecular weight marker (M); the size of the ladder bands is indicated on the left of the gel. The size of the +10A thrS transcript (290) and its processed form (255) together with a scheme of the relevant leader structures are shown on the right side of the gel. B. The reactions products were resolved on a 20% denaturing polyacrylamide gel together with an RNA alkaline ladder (A). The positions of the 5′ proximal residues is indicated.

We also observed significant cleavage very close to the RNA 5′ end, notably within the first four 5′ terminal nucleotides (Fig. 1, lanes 12–14). Again, the activity of the M. smegmatis and B. subtilis proteins were very similar (Fig. 1, compare lanes 12–14 and 15–16). Interestingly, the Asp85Lys/His86Ala RNase J double mutant, which has almost no 5′ exonuclease activity (Fig. 1, lanes 21–22), retained considerable endonuclease activity on the 5′ proximal nucleotides compared with the wild-type protein (Fig. 1, compare lanes 12–14 and 18–19). In order to evaluate if cleavage close to the 5′ end is dependent on a specific sequence context we modified the thrS transcript by inserting 3, 5 and 10 A residues, respectively, downstream of the second nucleotide of the original transcript (Fig. S3). Incubation of these RNAs with M. smegmatis RNase J and B. subtilis RNase J1 produced the major fragment resulting from cleavage upstream of the terminator. As expected, the cleaved RNA increased in size corresponding to the number of residues inserted at the 5′ end (Fig. 2A, lanes 7–10, 12–15 and 17–20). In contrast, the endonucleolytic cleavage pattern of the 5′ proximal RNA region remained unchanged (Fig. 2B, lanes 2–5, 7–10, 12–15 and 17–20) suggesting that cleavage close to the 5′ end might not depend on a specific sequence. However, the fact that RNase J can endonucleolytically cleave a 5′ triphosphoylated RNA at or very close to the 5′ end could have important repercussions on the initiation of degradation of a given mRNA (see Discussion).

M. smegmatis RNase J is not essential

Both RNase J1 in B. subtilis and RNase E in E. coli are essential enzymes that significantly affect RNA processing and degradation in their respective organisms. Because Mycobacteria have both an RNase E/G type enzyme and RNase J this raises the intriguing question of how these two key nucleases cooperate within the same organism.

We first analysed whether either or both enzymes are essential in M. smegmatis by creating two conditional mutants in the rne and rnj genes respectively. Expression of the genes was put under the control of the ptr promoter that can be induced by the addition of pristinamycin I, which removes the Pip repressor from the operator region of ptr (Forti et al., 2009). The mutants were created by constructing plasmids in which the 5′ proximal region of the rne or rnj genes was cloned downstream of the ptr promoter (pMYS820 for rne and pMYS823 for rnj). The recombinant plasmids that also carry the pip repressor gene were integrated in the M. smegmatis strain mc2155 chromosome by a single cross-over event (see Experimental procedures). The growth of both mutant strains was analysed in the presence and absence of inducer: tenfold serial dilutions of log phase cultures were plated on solid media. In the presence of the inducer, both mutants grew as the wild-type strain (Fig. 3A). In the absence of inducer, the rne mutant was not viable, but the rnj mutant grew as well as the wild-type strain (Fig. 3A). In liquid media we obtained similar results. The rne mutant stopped growing about seven generations after removal of the inducer while the rnj conditional mutant grew equally well in the presence or absence of the inducer (Fig. 3B). We conclude that in M. smegmatis, RNase E is an essential protein while RNase J is not.

Figure 3.

Growth of M. smegmatis rnj, rne and rne/rnj mutants. A. Growth on solid medium. Tenfold serial dilutions of log phase cultures of the wild type, the rne conditional mutant (Pptr-rne), the rnj conditional mutant (Pptr-rnj) and the Pptr-rnernj double mutant strains were replicated on LB-agar in the presence (+Pr) or absence (−Pr) of Pristinamycin I. B and C. Growth curves in liquid medium. Precultures of the different strains were grown in LD medium in the presence of Pristamycin I to an OD600 = 0.5, washed twice and diluted 1:200 (B) and 1:20 (C) in fresh LD medium, with or without Pristinamycin I (Pr); The zero time point corresponds to the dilution of the culture in fresh medium.

In order to definitely confirm this conclusion we constructed an RNase J null mutant where the rnj gene was partially deleted following a double cross-over event with plasmid pMYS824 (see Experimental procedures). In this mutant 418 out of a total 558 amino acids (positions 140 to 558) were replaced by the hygromycin resistance cassette. The rnj deletion strain had no growth defects (data not shown).

We also created an rne/rnj double mutant strain carrying the rnj deletion and the rne gene under control of the inducible ptr promoter. As expected, in the absence of inducer, the double mutant strain was not viable on either solid or in liquid media (Fig. 3A and C).

RNase E but not RNase J is involved in the processing of the furA-katG mRNA

Processing/degradation of mycobacterial mRNA has not been studied in any detail. The only known example where mRNA processing plays a role in gene expression is the M. smegmatis furA-katG operon (Sala et al., 2008) (Fig. 4A) encoding a Fur-like protein and catalase-peroxidase. The original bicistronic furA-katG mRNA is processed at position +443 (with respect to the transcription start point), 17 nts upstream of the furA stop codon (Fig. 4A and D). Processing at this site generates two cleavage products of very different stabilities. The upstream furA mRNA is rapidly degraded while the downstream katG mRNA is stabilized (Sala et al., 2008).

Figure 4.

In vivo analysis of M. smegmatis furA-katG mRNA processing. A. Scheme of furA-katG region. The positions of the oligos are indicated by horizontal arrows. The figure is not to scale. The 5′ ends of the furA and katG genes are indicated by vertical arrows. SD: Shine–Dalgarno sequence. B. Primer extension analysis of katG transcript processing. Oligonucleotide V244 was hybridized and extended on total RNA extracted from the wild-type M. smegmatis mc2155 strain, the rne conditional mutant (Pptr-rne) grown in the presence (+) or absence (−) of pristinamycin I (Pr) and the rnj deletion mutant (Δrnj). Reaction samples were migrated on a 5% denaturing polyacrylamide gel. The sequence ladder was generated with the same oligonucleotide (V244). For each strain, the analysis was performed on RNA extracted from cultures in early and late exponential growth phase (the respective OD600 is indicated on top of each lane). For the RNase E conditional mutant grown in the absence of pristinamycin I, the first sample (OD600 = 0.17, lane 5) corresponds to the time at which the strain stopped growing (about 7 generations after Pr depletion), the following points (lanes 6–8) were taken at 2 h intervals. The positions of the various signals are indicated with respect to the furA transcription start (5′furA). C. Identification of the furA transcription start. The primer extensions reactions were carried out using oligonucleotide V26 on total RNA extracted from the M. smegmatis wild type strain mc2155 (wt) and the RNase E mutant strain (Pptr/rne) grown in the presence (+) or absence (−) of pristinamycin I (Pr) and harvested at mid-exponential growth (OD600 = 0.4). The experiment was performed as described in (B), and the sequence ladder was generated with oligonucleotide V26. D. Secondary structure model of the furA-katG processing region. The structure was obtained using the M-fold algorythm. All co-ordinates refer to the furA transcription start point. The 5′ end of the katG mRNA (443) identified by primer extension and the stop codon of the furA gene are indicated.

In order to investigate the possible involvement of either RNase E or RNase J in this mRNA processing we performed a primer extension analysis of total RNA extracted from the Δrnj and the inducible ptr-rne mutant, grown in the presence or absence of pristinamycin I (see Experimental procedures). In a wild-type strain, extension of primer V244 complementary to 5′ proximal sequences within the katG mRNA (Fig. 4A) produced a signal corresponding to the previously identified 5′ end of the processed katG mRNA (Fig. 4B, lanes 1–2). In the RNase E depletion strain, this signal was strongly attenuated when the strain was grown in the absence of inducer (Fig. 4B, compare lanes 3–4 and 5–8), strongly suggesting that cleavage of the furA-katG mRNA is mediated by RNase E. In accordance, RNase E depletion caused an increase of the signal corresponding to the full-length transcript initiating at the furA promoter (Fig. 4B, lanes 5–8 and 4C).

In contrast, in the rnj deletion mutant, processing at position +443 was not impaired, and only minor differences were observed in the primer extension pattern compared with the wild-type strain (Fig. 4B, compare lanes 1–2 and 9–10). The most evident is the disappearance of a signal at 330 nts and the appearance of a 312 nts band. This suggests that RNase J has at best only a minor role in the decay of the furA-katG mRNA.

The role of RNases E and J in ribosomal RNA maturation

In E. coli RNase E is involved in 16S and 5S rRNA maturation (Li et al., 1999), whereas in B. subtilis RNase J1 participates in 16S rRNA maturation (Britton et al., 2007). Nothing is known about how ribosomal RNA processing occurs in mycobacteria.

Mycobacterium smegmatis has two ribosomal RNA operons, rrnA and rrnB. The two operons differ only in the 5′ proximal part of the leader region (shown in light grey in Fig. 5A) (Ji et al., 1994a). The rest of the two operons are identical. Transcription of the rrnA operon has been shown to initiate predominately at one of three potential promoters (P2), the rrnB operon is transcribed from a single promoter (P) (Gonzalez-y-Merchand et al., 1996) (Fig. 5A).

Figure 5.

Analysis of the 5′ end maturation of the rRNA precursors. Primer extension experiments were performed as indicated in Experimental procedures on M. smegmatis strain mc2155 (WT), the rnj null mutant (Δrnj), the Pptr-rne mutant (rne-) and the double Pptr-rnernjrnj rne-) in the absence of the inducer (Pr), as indicated on top of the lanes, using specific primers (V30, V32, and V37, in B, C, and D, respectively). For the rne depletion mutant the RNA was isolated as soon as a growth defect became apparent (generally 7 generations after removal of pristinamycin I). The extension products were separated on a 5% (w/v) denaturing polyacrylamide gel. The first four lanes of each gel show the DNA sequences obtained using the same primers on the rrnB operon. On the left, the DNA sequences surrounding the positions of the signals found by primer extension (circled nucleotides) are reported. The numbering on the right indicates the position of the signals with respect to the +1 of the rrnB operon. A. Schemes of the two rRNA operons (rrnA and rrnB) in M. smegmatis. The two operons have identical sequences except for the 5′ proximal half of the leader region that is shown in light grey. The mature rRNA sequences are represented by hatched boxes, and the sequences removed by processing are indicated (leader, spacers and the 3′ terminal region). P1/2/3 and P indicate the promoters that express the rrnA and rrnB operons respectively (Gonzalez-y-Merchand et al., 1996). The figure is not to scale. The sizes of the leader and spacer regions are indicated. The bar below the rrnA operon leader indicates the distance of P2 from the 5′ end of 16S rRNA (275 nt). In rrnB the distance of P to the 5′ end of 16S is 301 nt. The positions of the oligonucleotides V30/V32/V37 used for sequencing and primer extension experiments are shown by arrows. A scale indicating the co-ordinates of the 5′ and 3′ ends of each rRNA encoded by the rrnB operon is shown at the bottom of the figure. B. Analysis of the leader region. Primer V30 was extended to detect 16S 5′ end precursors. C. Analysis of the spacer 1 region. Primer V32 was extended to detect 23S intermediate rRNA precursors. D. Analysis of the spacer 2 region. Primer V37 was extended to detect 5S intermediate rRNA precursors.

Here we used rne and rnj single and double mutant strains in order to gain a first insight into whether RNases E and J are involved in rRNA maturation in M. smegmatis.

We performed a primer extension analysis on total RNA isolated from the wild type and mutant strains (Fig. 5B, C and D) using oligonucleotides complementary to regions close to the 5′ end of the mature rRNA transcripts (labelled V30, V32 and V37 in Fig. 5A). The primer extension probes thus hybridize to both the rrnA and rrnB operon transcripts. When not specified, positions of primer extension arrests are given with respect to the single promoter P of the rrnB operon.

Pre-16S rRNA maturation

Extension of primer V30, located downstream of the 16S 5′ end (position +301, Fig. 5A), showed that in a wild-type strain more than 90% of the 16S rRNA exists in its fully matured form (Fig. 5B, lane 1), although traces of full-length transcripts originating at the rrnA P2 and the rrnB promoter are present. A significant amount of a precursor 16S rRNA with a 5′ end at position +148 could also be detected. This processing site lies within a region forming a double-stranded stem made up of sequences upstream of the 16S rRNA and sequences of the spacer 1 region (between the 16S and 23S rRNA, Figs 5A and 6A). The presence of a concomitant cleavage observed on the opposite strand within a bulge (position +1977/79, Fig. 5C, see below) strongly suggested a maturation by the double-stranded specific RNase III (Fig. 6A). Indeed, this configuration is reminiscent of RNase III cleavages observed in similar positions in the long processing stalks of 16S rRNA precursors of E. coli (Fig. 7A) (Young and Steitz, 1978) and B. subtilis (Loughney et al., 1983; Herskowitz and Bechhofer, 2000).

Figure 6.

Secondary structure models of the M. smegmatis rRNAs precursors. The structures are based on an M-fold analysis with some manual adjustments. All co-ordinates refer to the transcription start from the rrnB promoter (+1). The scissors symbol indicates the location of signals observed by primer extension. Where known, the name of the RNase responsible for the cleavage is indicated. A. Pre-16S rRNA structure. The boxed region corresponds to the mature 16S rRNA. The region upstream of the mature 5′ end corresponds to the leader region. The downstream region (170 nt from the 3′ end of the 16S rRNA) corresponds to the 5′ half of the spacer 1 region. B. Pre-23S-5S rRNA secondary structure. The boxed regions correspond to the mature 23S and 5S rRNAs. The region upstream of the 5′ end of the 23S rRNA (215 nucleotides) corresponds to the 3′ half of the spacer 1 region. The sequences downstream of the mature 23S 3′ end represent the spacer 2 and the 5S rRNA. The 12 nucleotides shown in grey are likely degraded by the 5′-3′ exoribonucleolytic activity of RNase J.

Figure 7.

Models of the 5′ end maturation of the 16S, 23S and 5S rRNA in M. smegmatis, E. coli and B. subtilis. Simplified models of the precursors for the three ribosomal subunits are presented. The grey boxes represent the mature subunits. The positions of the different cleavages are marked by arrows with the name of the enzyme involved or a scissor if unknown. The 5′-3′ exoribonucleolytic activity is shown by Pacman symbol. See text for details.

Inactivation of the rnj gene had no major effect on 16S rRNA maturation (Fig. 5B, lane 2). By contrast, depletion of RNase E caused a twofold increase in the putative RNase III processing product (position +148) but also a significant accumulation of a +36 nt precursor (position +265, Fig. 5B, lane 3). In the rnj/rne double mutant we observed, in addition to the +36 nt species, a +6 and a +11 nt precursor (positions 290 and 295 Fig. 5B, lane 4). Their absence in the rne single mutant suggests that RNase J is responsible for their degradation.

Pre-23S rRNA maturation

The 23S rRNA structural gene lies downstream of the 16S rRNA gene separated by a 368 nt region (spacer 1, Fig. 5A) that does not encode any known gene. Extension of primer V32, hybridizing to the 5′ proximal region of the mature 23S rRNA, revealed that in the wild-type strain about 90% of the 23S rRNA is fully processed (position +2196, Fig. 5C, lane 1). Two minor upstream signals were observed at positions +1977/79 and +2040. The signal at position +1977/79 (217/219 nt upstream of the mature 23S 5′ end) most likely corresponds to a precursor species generated by RNase III cleavage in the double-stranded region formed by sequences 5′ to the 16S rRNA and the spacer 1 region that separates the 16S and 23S rRNAs on the original transcript (Fig. 6A); cleavage of the opposite strand occurs at position +148, as described above. The exact positions of the high molecular weight bands in Fig. 5B and C were deduced by long runs on separate gels (Fig. S4).

The 23S precursor RNA with a 5′ end at position +2040 was present at lower levels in a strain depleted for RNase E (Fig. 5C, lane 3) suggesting that this transcript may be generated by an RNase E cleavage. It is located in the single-stranded region between two extensive secondary structures of the 23S precursor (Fig. 6B). In accordance, the upstream precursor likely generated by RNase III (position +1977/79) is present at higher levels when RNase E expression is reduced (Fig. 5C, lane 3).

The absence of RNase J has a major effect on the 23S rRNA maturation. Under these conditions the cell contains no mature 23S rRNA at all (Fig. 5C, lane 2). More than 80% of the total 23S rRNA accumulates as a +14 nt precursor (position +2182, Fig. 5C, lane 2 and Fig. 6B). A similar result was observed in the rnj/rne double mutant, where a few additional minor precursor species were observed (Fig. 5C, lane 4). This indicates that RNase E cannot directly generate a mature 23S rRNA, but probably cleaves 14 nts further upstream.

The rnj null mutation also caused a small but significant accumulation of two closely spaced precursors at positions +2068/2069. Together with a signal observed at position +5366 within spacer 2 (see below, Fig. 5D, lanes 3 and 4) these bands probably correspond to the remnants of a precursor generated by RNase III processing within the 23S processing stalk aimed at separating the 23S and 5S rRNAs (Fig. 6B).

Pre-5S subunit maturation

The 5S rRNA structural gene is separated from the upstream 23S rRNA gene by a 101 nt spacer. Primer extensions with the V37 primer in the wild-type strain showed that more than 95% of the total RNA corresponded to that of the mature 5S rRNA (+5457, Fig. 5D, lane 1). A precursor rRNA 29 nt longer than the mature 5S rRNA (position +5428) was present in minor quantities. Disruption of the rnj gene had no significant impact on the processing of the 5S rRNA (Fig. 5D, lane 2). Depleting RNase E also did not substantially reduce the extent of fully processed 5S rRNA (Fig. 5D, lane 3). However, under these conditions two new precursor 5S rRNA species appeared. The first (position +5366) likely corresponds to the RNase III generated precursor described above, together with a cleavage on the opposite strand at position +2067/2068 (Fig. 5C, lane 2). The most prominent precursor detected during RNase E depletion contained 37 additional nucleotides (position +5420). In the rnj/rne double mutant, a similar pattern was observed, although in this case a clear reduction of the amount of mature 5S rRNA was observed, suggesting that both RNases are involved in its maturation.


Mycobacterial RNA metabolism has not been well studied but these high GC Gram-positive organisms contain an RNase E/G type enzyme and a putative RNase J orthologue. In addition, a compilation of potential RNase orthologues (Table 1) suggests that mycobacteria constitute an interesting class of bacteria that combine key enzymes present in E. coli and B. subtilis. Sequence and 3D structure comparisons of the M. smegmatis protein 2685 with known RNases J of T. thermophilus and B. subtilis (Figs S1 and S2) showed good overall similarities (55% and 59%, respectively, with B. subtilis RNases J1 and J2) and the conservation of all important motifs, notably the five classical β-lactamase motifs, the three β-CASP motifs and the amino acids constituting the mononucleotide binding pocket (Li de la Sierra-Gallay et al., 2008). However, this comparison provided no clues as to the major enzymatic activity. It was important to verify that the single putative M. smegmatis RNase J orthologue (MSMEG_2685) actually is an RNase and whether it has endo- and/or exoribonucleolytic activities. In fact, in B. subtilis both RNase J1 and RNase J2 show a similar endoribonucleolytic activity (Even et al., 2005), but RNase J2 contrary to RNase J1 has a very weak 5′-3′ exoribonucleolytic activity (Mathy et al., 2010). In contrast, archeal RNase J orthologues appear to have 5′ exonuclease but no endonuclease activity (Clouet-d'Orval et al., 2010; Hasenohrl et al., 2011). In vitro assays with the purified mycobacterial protein showed that it actually has both activities, 5′-3′ exo- and endonucleolytic. The exonucleolytic activity is comparable with that of B. subtilis J1 when assayed on the same 5′ monophosphorylated B. subtilis thrS leader substrate (Li de la Sierra-Gallay et al., 2008). The Asp85Lys and His86Ala mutant protein has almost completely lost its 5′ exonuclease activity, confirming that the activity attributed to M. smegmatis RNase J is genuine and not due to contamination during protein purification.

Using the B. subtilis thrS leader region in its 5′-triphosphorylated form we showed that M. smegmatis RNase J also has endonucleolytic activity. The major cleavage site was found at position +245 immediately upstream of the terminator structure, which is also the principal site of cleavage by B. subtilis RNase J1/J2, as described previously (Even et al., 2005). Interestingly, significant cleavage by the M. smegmatis as well as the B. subtilis enzyme was also observed within the first four to five 5′ proximal nucleotides that are part of a region previously shown to be single-stranded (Luo et al., 1998). RNase J1 has been shown to cleave single-stranded sequences non-specifically under high enzyme concentrations (Daou-Chabo and Condon, 2009). However, under the conditions used here RNase J cleaved the thrS leader with a clear bias towards two distinct sites, i.e. upstream of the leader terminator and at sites near the 5′ end. Insertion of three to ten A residues after position +2 of the thrS RNA also did not create new cleavage sites. On the contrary, even in these modified thrS leader substrates, with extended 5′ single-stranded regions, cleavage by RNase J in vitro occurred predominantly within the first four 5′ proximal nucleotides (Fig. S3).

Interestingly, the Asp85Lys and His86Ala mutant protein that is almost devoid of 5′ exonucleolytic activity retained considerable endonucleolytic activity. This was evident not only for the cleavage of the 5′ proximal residues but also for the cleavage upstream of the terminator structure (data not shown). Because the double mutation concerns two residues that co-ordinate to one of the catalytic Zn+2 ions, it is unlikely that the mutant protein binds both Zn+2 ions. This configuration is reminiscent of that encountered in RNase J2. This enzyme has three active site residues that are not conserved with respect to the paralogous RNase J1, all of which are involved in the co-ordination of the equivalent Zn+2 ion. Similar to the mutated mycobacterial protein, B. subtilis RNase J2 has endo- but almost no exonucleolytic activity.

The capacity of RNase J to endonucleolytically remove the 5′ terminal triphosphorylated nucleotide of a native transcript could play an important role in initiating mRNA decay by creating an entry site for the 5′ exonucleolytic activity of RNase J. In the most straightforward model, RNase J would simply switch from endo- to exonucleolytic mode without releasing the RNA (Li de la Sierra-Gallay et al., 2008). Even though pyrophosphate removal by an RppH-like activity has also been observed in B. subtilis (Richards et al., 2011), RNase J cleavage close to the 5′ end might well constitute an alternative pathway to initiate 5′ exonucleolytic decay of native transcripts.

The mode of action by which RNase J releases the first nucleotide(s) must be endonucleolytic, because co-ordination of the γ-phosphate of the terminal nucleotide in the phosphate binding pocket would place the scissile bond out of phase (Li de la Sierra-Gallay et al., 2008). The recent discovery that the RNA entry channel of RNase J actually extends past the active site (Dorléans et al., 2011; Newman et al., 2011) created a rationale for how the catalytic cleft could directly accommodate an RNA in endonucleolytic mode (Fig. S5). Direct access to the site of cleavage is thus likely to be the principal mode of action to cleave an RNA far from the 5′ end.

In what concerns cleavage very close to the 5′ end we would like to suggest an alternative model in which RNase J would function as a ‘sliding endonuclease’. In this mechanism, the native triphosphorylated RNA enters the RNA entry channel of the dimer and is threaded towards the active site in the same way as 5′ monophosphorylated RNA. Because the 5′ PPP moiety cannot interact productively with the phosphate binding pocket, we propose that the RNA can slide past the catalytic centre and be cleaved endonucleolytically at the downstream nucleotides. This sliding mode would be expected to be very sensitive to secondary structure and probably not go beyond a few nucleotides. Indeed, the cleavable phosphodiester fits into the monophosphate binding pocket much in the same way as the 5′ terminal monophosphate (Dorléans et al., 2011). It is therefore likely that once the 5′ PPP group has slided past the active site one the following phosphodiester groups approaching the catalytic centre should readily snap into the phosphate binding pocket. This would explain why the observed endonucleolytic cleavage is essentially observed for the 5′ proximal nucleotides.

RNase E and RNase J1 are essential for viability in E. coli and B. subtilis respectively. These ribonucleases are clearly very important for RNA metabolism but the precise reason for their essentiality has not been established, neither for RNase E nor for RNase J1. In M.  smegmatis RNase E is essential while RNase J is not, implying that the 5′-3′ exonucleolytic activity of RNase J is dispensable. A similar configuration exists in the α-proteobacterium Sinorhizobium where the RNase J orthologue is dispensable. However, it is not known whether RNase J in this organism has 5′ exo-, endo- or both nucleolytic activities (Madhugiri and Evguenieva-Hackenberg, 2009).

Processing of mRNA precursors has also been observed in Mycobacteria but available data are extremely scarce. In fact, the sole known case is the furA-katG operon encoding a Fur-like transcriptional repressor (furA) and a catalase peroxidase (katG) involved in the activation of isoniazide, an antitubercular drug (Milano et al., 2001; Sala et al., 2003). The major katG transcript in M. smegmatis is produced by processing of the native bicistronic furA-katG mRNA close to the end of the furA ORF (Sala et al., 2008). Here we showed that the ribonuclease responsable for this cleavage is RNase E. Processing of the bicistronic furA-katG mRNA is also observed in M. tuberculosis and, similarly, prevented by a depletion of M. tuberculosis RNase E (data not shown). Processing occurs 17 nucleotides upstream of the furA stop codon 3′ to two A residues in a single-stranded region between two potential secondary structures. This is similar to the sequence and structural context identified for ectopic cleavage sites for E. coli RNase E (Mackie, 1992; Baker and Mackie, 2003). Maturation of the native transcript thus generates a relatively stable katG mRNA and an unstable furA transcript. Because FurA negatively autoregulates its own transcription from pfurA, this allows the cell to produce a sufficient amount of KatG peroxidase and at the same time keep FurA repressor biosynthesis low.

Processing of ribosomal RNAs

In Mycobacteria, rRNA processing has not been studied in much detail (Ji et al., 1994a,b) and the nucleases involved were unknown. The M. tuberculosis single rRNA operon (rrnA) and the two M. smegmatis operons (rrnA and rrnB) contain the genes for the 16S, 23S and 5S rRNA, co-transcribed as a single precursor molecule (Ji et al., 1994a; Gonzalez-y-Merchand et al., 1996). Co- or post-transcriptional folding of the precursor rRNAs is predicted to involve extensive base pairing between the leader and spacer-1 region (16S), spacer-1 and spacer-2 (23S and 5S) as suggested by the secondary structures predicted by the M-fold algorithm (Fig. 6). The M. smegmatis rrnA and rrnB operons have identical sequences downstream of position −165 with respect to the 16S rRNA structural gene. Thus the folded precursor structures that are the substrates for a series of maturation reactions to produce the mature rRNAs should be the same for both operons. The large processing stalks are thus a common feature shared by most if not all eubacteria. They are generally cleaved by the double-stranded specific RNase III or a related nuclease that physically separate the individual rRNA precursors. We have not formally shown that RNase III is responsible for the initial cleavage of these processing stalks in M. smegmatis. However, we have identified 5′ ends in the different rRNA precursors that when the stalks are folded, are in proximity on the opposite strand and near a bulge. Such a configuration is very characteristic of an RNase III cleavage site. Thus, initial separation of the 16S, 23S and 5S rRNA most likely occurs through two RNase III cleavages (Fig. 6).

RNase E was found to be involved in 16S rRNA maturation. Its partial depletion caused a twofold increase in the putative RNase III processing product (position +148) but also a significant accumulation of the +36 nt precursor. This is consistent with RNase E being able to create the mature 5′ end of 16S rRNA. The putative cleavage site is composed of a U rich single-stranded region (Fig. 6A) compatible with the known requirements for RNase E cleavage in E. coli (Ehretsmann et al., 1992; McDowall et al., 1994) and M. tuberculosis (Zeller et al., 2007). In contrast, the complete inactivation of RNase J had no effect on 16S rRNA except when RNase E was depleted at the same time. The stabilization of two precursors with the 5′ end close to the mature 5′ end, suggested that the RNase J collaborates with RNase E to complete the maturation of the 16S rRNA.

We propose the following model for 16S rRNA maturation (Fig. 7A): (i) ribonucleolytic cleavages at +148 and +1977/79, likely performed by RNase III, release the 16S rRNA precursor from the 30S original transcript; (ii) an unidentified nuclease cleaves at position +265 leaving 36 nt upstream of the mature 16S rRNA 5′ end; and (iii) this shortened precursor is then processed by RNase E and RNase J to create the mature 16S rRNA 5′ end.

When we compare M. smegmatis 16S rRNA maturation with that of E. coli and B. subtilis (Fig. 7A) we note that in all cases, maturation appears to occur in a two-step mechanism: the initial RNase III precursors are processed by RNase E (E. coli, +66 nt) or unknown endoribonucleases (B. subtilis, +38 nt and M. smegmatis, +36 nt) apparently to create a new 5′ end free from the processing stalk that could increase the efficiency of the final maturation steps at both the 5′ and 3′ extremities.

RNase J plays a major role in 23S rRNA maturation: in the Δrnj mutant strain no mature 23S rRNA was present in the cell. Because the strain is viable, the absence of a fully mature 23S rRNA does not compromise ribosome activity. This indicates that the precursor molecule, although 14 nucleotides longer at its 5′ end than the mature 23S rRNA, is able to efficiently assemble into a functional ribosome. It is likely that the +14 nt precursor is then trimmed by the RNase J 5′ exonuclease activity, but we cannot rule out an endonucleolytic cleavage by RNase J to create the mature 23S rRNA in a single step.

Ribonuclease E is also involved in 23S rRNA maturation, although its role is less relevant: notably the RNase III generated precursor 23S rRNA (from cleavages at positions +148 and +1977/79) is increased when RNase E is depleted and the precursor pattern of the rnj/rne double mutant shows an additional band corresponding to +26 nt precursor (Fig. 5C, lane 4).

Maturation of the 23S rRNA in M. smegmatis is thus quite different when compared with both E. coli and B. subtilis (Fig. 7B). In fact, RNase J, the nuclease involved in the final steps of 23S rRNA maturation in M. smegmatis, does not participate in the processing of 23S rRNA in a B. subtilis wild-type strain (Redko et al., 2008).

In the absence of RNase J, maturation of 5S rRNA is normal, whereas in the RNase E mutant several precursors (91 nt and 37 nt from the mature 5′ end) accumulate. However, the fact that in the rnj/rne double mutant the amount of mature 5S rRNA is clearly reduced suggests that both ribonucleases are involved in its maturation. The data are compatible with a direct maturation of the 5S rRNA 5′ end by RNase E using the +37 nt precursor and possibly also longer precursors. However, it is likely that another nuclease besides RNase E is involved in the final processing steps. This unknown nuclease might also account for the +37 nt precursor. Again, the maturation pathway in M. smegmatis differs from 5S rRNA processing in E. coli and B. subtilis (Fig. 7C).

More data are needed to fully understand rRNA processing in Mycobacteria particularly at the 3′ end. Nevertheless, our experiments clearly show that the mechanisms and enzymes involved in the 5′ end processing of all three rRNAs are significantly different compared with what we know from the two model organisms E. coli and B. subtilis. From an evolutionary and mechanistic point of view it will be interesting to better understand how combinations of RNases present in a given organism influence and direct RNA metabolism.

Experimental procedures

Bacterial strains and growth conditions

The prototrophic M. smegmatis mc2155 (Snapper et al., 1990) was used for the construction of the mutants. In the strain called rnj101, the rnj (MSMEG_2685) gene was put under the control of the ptr promoter by Campbell-type integration of plasmid pMYS823. For strain rnj102, the rnj (MSMEG_2685) gene was partially deleted by double cross-over recombination with plasmid pMYS824. In the derivative strain rne101, the rne (MSMEG_4626) gene was put under control of the ptr promoter by Campbell-type integration of plasmid pMYS820. For the double mutant strain, the rne (MSMEG_4626) gene was put under control of the ptr promoter by Campbell-type integration of plasmid pMYS843 on the chromosome of the rnj102 strain.

The following E. coli strains were used: DH10B (Grant et al., 1990) the host for plasmids construction, strain XL1Blue (Stratagene) for protein mutagenesis and strain BL21-CodonPlus (DE3) (Stratagene) carrying pMYS825 and pMYS841, for overexpression of His-tagged proteins.

Mycobacterium smegmatis mc2155 was grown in LD medium (Sabbatini et al., 1995) containing 0.2% (vol/vol) glycerol and 0.05% (vol/vol) Tween 80 and supplemented when necessary with hygromycin (50 µg ml−1), kanamycin (25 µg ml−1) and pristinamycin I (10 µg ml−1). E. coli was grown in LB medium supplemented when necessary with neomycin (25 µg ml−1), chloramphenicol (20 µg ml−1) and IPTG (20 mM ml−1) for the overexpression of the heterologous protein.

Oligonucleotides and plasmids

The oligonucleotides used are listed in Table S1.

pMYS825: the coding sequence of the MSMEG_2685 gene amplified with the oligos FG2360/V7, containing an additional N-terminal His-tag, was cloned in plasmid pKYB1 (New England Biolabs) between the sites NdeI and MfeI.

pMYS841: derivative of pMYS825, but the MSMEG_2685 coding sequence carries mutations that cause two amino acid changes in the active site (D85K & H86A), obtained with the oligos V34-V35.

pMYS820 and pMYS823: the 850 bp fragment of rne gene starting at position +2 with respect to the initiation codon was PCR amplified with the oligos V12/V13 and the 846 bp fragment of MSMEG_2685 gene starting at position +4 with respect to the initiation codon was PCR amplified with the oligos V14/V15, respectively, and the fragments were ligated as NcoI-SphI fragments into the respective sites of the integrative plasmid pAZI9479 (Forti et al., 2009) downstream of the ptr promoter. Campbell-type integration of pMYS820 or pMYS823 on the M. smegmatis chromosome renders rne and MSMEG_2685 expression pristinamycin I dependent.

pMYS824: A 1063 bp PCR fragment of the MSMEG_2685 5′ region (oligos FG2361/FG2362) beginning at the position −644 with respect to the initiation codon of the gene (with an additional stop codon at the end) was ligated as a SpeI-HindIII fragment into the plasmid pjsc284 [derivative of pYUB854 (Bardarov et al., 2002)], downstream of the hygromycin resistance cassette. On the same plasmid, a 1023 bp PCR fragment of the MSMEG_2685 3′ region (oligos FG2363/FG2364) beginning respectively at +1323 with respect to the initiation codon of MSMEG_2685 was ligated as a XbaI-KpnI fragment upstream of the hygromycin resistance cassette. The restriction fragment SpeI-KpnI from pMYS824 was utilized to transform M. smegmatis and by double cross-over, the central region of MSMEG_2685 on the chromosome (903 bp) was substituted by a hygromycin resistance cassette.

pMYS843: derivative of pMYS820 in which the hygromycin resistance cassette was deleted by PdiI digestion and substituted with the kanamycin resistance cassette, PCR amplified with the oligos FG2564/FG2564 from pMV261 (Stover et al., 1991).

Total RNA isolation

Twenty-five millilitres of M. smegmatis cultures (OD600 of 0.7–1) were pelleted, washed with 1 ml Tween 80 0.5%, 1 ml TSE buffer (10 mM Tris HCl pH 8, 100 mM NaCl, 1 mM EDTA) and resuspended in 200 µl of STET buffer (50 mM Tris HCl pH 8, 8% sucrose, 0.5% Triton X-100, 10 mM EDTA, 7 mg ml−1 lysozyme) then incubated at 0°C for 20 min. The cell suspension was added to a tube containing an equal volume of glass beads (0.2 mm diameter) and 200 µl phenol/H2O, incubated at 100°C for 3 min and vortexed 5 min at 4°C. The phases were separated by centrifugation (5 min 13000 r.p.m. at room temperature).

The aqueous phase was re-extracted twice with 200 µl phenol/H2O pH: 8 and 200 µl phenol/CHCl3. The nucleic acids were precipitated with 0.1 vol LiCl 5 M, 3 vol EtOH 100% and dissolved in RNase-free H2O.

Primer extension

The maturation of the different ribosomal RNAs of the rrn operons was studied by primer extension on the total RNA isolated from the wild type and various mutant strains. Primer extension was performed by adapting a previously described protocol (Sambrook et al., 1989). Briefly, an annealing mix (10 µl), containing 10 µg of total RNA, 1 U µl−1 RNasin (Promega), 0.5 pmole of 5′ labelled primer in 1× hybridization buffer (300 mM NaCl, 10 mM Tris HCl pH 7.5, 2 mM EDTA), was denatured 4 min at 80°C and incubated 2 h at 50°C for the annealing. Then, to the annealing mix were added 40 µl of 1.25× RT-buffer (1.25 mM of each dNTP, 12.5 mM DTT, 12.5 mM Tris HCl pH 8, 7.5 mM MgCl2), 5 U RNasin (Promega) and 10 U AMV Reverse Transcriptase (Finnzymes) and incubated 30 min at 50°C for the extension. Samples were precipitated with 1/10 volume NaAc 3 M, 2.5 Volumes EtOH, dissolved in 12 µl stop mix (95% formamide, 20 mM EDTA, 0.05% bromophenol blue, 0.05% xylene cyanol FF). Reaction products were separated on a 5% denaturing polyacrylamide gel.

Oligonucleotides used for rrn operons analysis were V30 (complementary to region +347/+367 internal to 16S), V32 (complementary to region +2246/+2268 internal to 23S) and V37 (complementary to region +5491/+5512 internal to 5S). All co-ordinates are relative to the transcription start (+1) of the rrnB operon. Oligonucleotides used for furA-katG operon analysis: V26 (complementary to region +66/+88 internal to furA gene), V244 (complementary to region +533/+552 internal to katG gene). All the co-ordinates are relative to the transcription start (+1) of the furA promoter.

The oligonucleotides were radiolabelled with γ-(32P) ATP using T4 Polynucleotide Kinase (New England Biolabs).

Site-directed mutagenesis

The mutations D85K and H86A in the RNase J protein of M. smegmatis were introduced on the pMYS825 plasmid by the Quikchange strategy (Stratagene) with the oligonucleotides V34-V35 and KOD DNA polymerase (Novagen).

In vitro transcription

In vitro transcription with T7 RNA polymerase was performed as described by the manufacturer (Promega) using a PCR fragment as template. For thrS RNA synthesis, the PCR template was prepared using oligonucleotides HP857 and HP1165. It carries the T7 promoter, followed by thrS leader sequence extending to a terminator element. Oligonucleotides HP1587, HP1588 and HP1589 were used for the synthesis of the thrS RNA carrying respectively 3,5, and 10 additional nucleotides at its 5′ end (see Fig. S3).

The 5′ triphosphorylated transcripts were 5′ end-labelled by addition of γ-(32P) GTP in the transcription reaction mix as described previously (Shahbabian et al., 2009), the 5′ monophosphorylated transcripts were 5′ end-labelled incubating the 5′ dephosphorylated RNA with γ-(32P) ATP using T4 Polynucleotide Kinase (New England Biolabs). Dephosphorylation of the RNA was carried out using calf intestinal phosphatase (New England Biolabs).

Transcripts were purified by gel filtration on Sephadex G-25 columns (GE Healthcare). Alternatively, the RNA substrate was purified from unwanted products by elution from a 5% polyacrylamide gel.

RNase J overexpression and purification

The M. smegmatis rnj open reading frame, in its N-terminal His6-tagged version, was cloned into the pKYB1 vector (New England Biolabs), between the NdeI-MfeI sites to obtain plasmid pMYS825. This plasmid was used introduce the double mutation Asp85Lys and His86Ala via the Quikchange strategy (Stratagene) using oligonucleotides V34 and V35. The RNase J wild type and mutant proteins were overexpressed from the T7 promoter in E. coli strain BL21(DE3) CodonPlus (Stratagene).

The proteins were isolated by affinity chromatography on a Ni-NTA Agarose resin using an imidazol elution as described by the supplier (Qiagen). The eluated protein was quantified by the Bradford assay (Bradford, 1976) and analysed on a 10% SDS-acrylamide gel.

RNase J processing assays

The assay mixture (10 µl) containing 20 mM HEPES-KOH (pH 8.0), 8 mM MgCl2, 100 mM NaCl, 0.24 U ml−1 RNasin (Promega), 35 µM of 5′ triphosphorylated or monophosphorylated labelled RNA substrate and 4 µM of purified proteins, was incubated at 37°C. For time-course experiments, samples of scaled-up reactions (60 µl) were taken at the indicated times. Reactions were stopped by the addition of 5 µl of 3× gel loading buffer (87.5% formamide, 0.05% xylene cyanol, 0.05% bromophenol blue, 5 mM EDTA). The control reaction was performed by incubating substrate with the reaction buffer under the same conditions. The samples were directly loaded on a 20% or a 5% denaturing polyacrylamide gel and the reaction products were visualized on a Phosphorimager.


We are grateful to Jackie Plumbridge for critical reading of the manuscript. This work was supported by funds from the CNRS (UPR 9073), Univ Paris Diderot, Sorbonne Paris Cité and a grant from the EC-VI Framework Contract No. LSHP-CT-2005–018923. V.T. was awarded a FEMS short-term fellowship.