The group IIA Ll.LtrB intron interrupts the ltrB relaxase gene, which resides on the conjugative plasmid pRS01 and is required for pRS01 conjugative transmission (Belhocine et al., 2004; Belhocine et al., 2005; Mills et al., 1996). Both intron-containing (Int⁺) and intron-less (Int^-) ltrB genes, under control of a nisin-inducible promoter on a pCY20 plasmid, were expressed in the native host L. lactis strain IL1403 (Figure 1A). Northern blotting analysis of RNA (Figure 1B) indicated that the relaxase mRNA abundance was much lower in the presence of the intron than in its absence (21% vs. 100%). Consistent with the mRNA difference, the LtrB relaxase determined by Western blotting also accumulated to lower levels in the Int⁺ cell (Figure 1C, 36% vs. 100%). Notably, a previous study showed an even larger (20-fold) difference in protein levels in Int^- versus Int⁺ cells (Chen et al., 2005). The dramatic decrease in the ltrB relaxase mRNA level in the presence of the intron was confirmed by performing quantitative reverse transcription PCR (qRT-PCR). The difference in mRNA levels between the Int⁺ and the Int^- cells was 23-fold (Figure 1D, Figure 1—source data 1, Figure 1—figure supplement 1, 4.3% vs. 100%). The greater difference by qRT-PCR and Northern blotting, may be accounted for by the different reference RNAs used (copA mRNA for qRT-PCR vs. 16S rRNA for Northern blot) or by the inherent difference between the two techniques in determining RNA levels.

Figure 1 with 4 supplements see all

Download asset Open asset

Figure 1—source data 1

RT-qPCR data for mRNA.

mRNA quantitation cycle (C_q) values for Intron^- and Intron⁺ samples. Contents are the experimental or unknown sample (Unkn), no RT enzyme control (NRT) and no template control (water, NTC).

https://doi.org/10.7554/eLife.34268.008

Download elife-34268-fig1-data1-v1.xlsx

By performing both reverse transcription and qRT-PCR analyses we determined that the decrease in mRNA was not simply due to group II intron-promoted reduction in transcription rate (Figure 1—figure supplement 1). We analyzed nascent ltrB primary transcripts in Int^-/Int⁺ cells, utilizing a DNA primer that bound 50 nucleotides downstream from the 5′-end of the transcript (Figure 1—figure supplement 1A–B). Because of the proximity of the primer to the 5′-end, we presumed that the yield of the cDNA or RT-PCR products generated was a reflection of transcription initiation, and indeed there was only a 20% difference in the amount of cDNAs between Int^- and Int⁺ (Figure 1—figure supplements 1A, 100% vs. 79%; 1B, 100% vs. 74%). In addition, comparison of the relative mRNA expression level in the Int^- cell with that of total intron RNAs (pre-mRNA + Intron) in the Int⁺ cell, also indicated small transcription differences between Int^-/Int⁺ cells (Figure 1—figure supplement 1C, primers, 1D, relative levels, 20.04 ± 2.00 vs. 25.65 ± 6.47).

In regard to splicing, we used Northern blotting, primer extension and qRT-PCR to determine the relative abundance of the spliced intron. The data showed that the sum of intron-containing RNAs (pre-mRNA + Intron) was relatively abundant, in contrast to the spliced mRNA (Figure 1B, Figure 1—figure supplement 1D). Notably, a previous analysis of Ll.LtrB group II intron expression in the natural pRS01 context also showed that the ltrB mRNA level was much lower than that of the total intron RNA (Chen et al., 2005). When the intron splicing efficiency was defined as the ratio of spliced intron against the sum of the intron-containing RNAs (pre-mRNA + Intron), the numbers ranged from 63% to 91% (Figure 1B, 91%; Figure 1—figure supplement 1D, 73.8%; Figure 1—figure supplement 2B, 63%). Thus, it does not appear that a splicing deficiency alone resulted in the mRNA reduction.

Furthermore, we compared the decay rate of mRNAs expressed in Int^-/Int⁺ cells. After a 30 min induction of ltrB gene expression, the cells were treated with rifampicin to stop transcription, and mRNA level was monitored over time and analyzed by Northern blotting (Figure 1—figure supplement 3A–B). Surprisingly, the decay rate of spliced mRNA for the Int⁺ cells was ¼ to ½ of the rate for the mRNA of the Int^- cells. Because the rates apply to a small residual fraction of the mRNA, these results must be interpreted with caution. Nevertheless, they suggest that the reduction in mRNA is not due to increased degradation.

We also investigated if gene expression was affected at the translation level. Comparison of mRNA polysome profiling showed that there was only a marginal difference in mRNAs enriched in the polysomal fractions between Int^-/Int⁺ cells (Figure 1—figure supplement 4, 100% vs. 85%), thereby rendering unlikely the possibility that the spliced mRNA experienced translational repression.

Additionally, although ltrB RNA levels can be condition-dependent (Chen et al., 2005), the differential expression levels between Int⁺ and Int^- genes were not influenced by stressors and environmental changes, such as temperature, pH, salt and redox (data not shown). Taken together, we concluded that the group II intron-promoted inhibition in gene expression is likely to be mainly at the mRNA level.

We also investigated if this substantial group II-intron mediated decrease in mRNA was independent of plasmids and promoters from which the ltrB relaxase gene was expressed (Figure 2A). When the Int^-/Int⁺ ltrB genes were expressed from different plasmids, including pDL278 and pAMJ328, where gene expression was driven by either constitutive or pH-inducible promoters, respectively, we observed similar levels of mRNA reduction in the presence of the intron (pDL: 100% vs. 28%; pAMJ: 100% vs. 45%) (Figure 2A). This demonstrates that group II intron-promoted inhibition was independent of the plasmid and promoter from which the ltrB relaxase gene was expressed.

Figure 2

Download asset Open asset

In addition, we extended this comparative study to other types of bacterial group II introns, including a typical IIB intron, EcI5 (Dai and Zimmerly, 2002b) and a IIC intron, BhI1 (Candales et al., 2012). The Int^-/Int⁺ constructs of these introns were fused to an enhanced GFP reporter at the N-terminus. When expressed in Escherichia coli, levels of the mRNA dropped substantially for both EcI5 and BhI1 in the presence of the intron (Figure 2B,C). Also the GFP reporter was greatly reduced for EcI5 with the intron present (Figure 2B bottom). Therefore, it appears that inhibiting gene expression in bacteria is a common property of group II introns.

Because ltrB relaxase is required for initiation of plasmid conjugation (Belhocine et al., 2004; Belhocine et al., 2005; Mills et al., 1996), and because ltrB gene expression is reduced in the presence of the intron, we wished to determine if the intron may inhibit conjugal transfer of pRS01. To test this hypothesis, we measured conjugation frequency where the donor strain, IL1403, co-hosted erythromycin resistant (erm^R) conjugative plasmid pRS01 ltrB^- (ΔLtrB::tet) and the pCY20 Int^- or Int⁺ plasmid. After mating with fusidic acid-resistant (FA^R) ILI403 the frequency of the Erm^RFA^R exconjugants was measured (Figure 3). Consistent with the LtrB relaxase protein levels (Figure 1C), the conjugation frequency of the Int⁺ donor was 3- to 4-fold lower that of the Int^-, at 2.0 ± 0.1 × 10⁻⁴ versus 7.3 ± 10⁻⁴ exconjugants per donor (n = 2, p<0.05) (Figure 3 and data not shown). This result ties into a previous study (Novikova et al., 2014) suggesting that these two disparate mobile elements, the mobile intron and the conjugative pRS01 plasmid, have a functional interplay.

Figure 3

Download asset Open asset

The first crystal structure of a group II intron lariat has revealed that ligated exons (mRNA) remain bound to the spliced intron RNA (Robart et al., 2014). The recently determined cryo-electron microscopy (cryo-EM) structure of the Ll.LtrB group II intron-IEP complex, that was isolated from its native host, also shows that after splicing the mRNA is retained within the ribonucleoprotein (RNP) (Qu et al., 2016). To confirm the presence of this intron-mRNA interaction in the bacterial cell, we performed intron-RNA pull-down assays (Qu et al., 2014). Briefly, a streptavidin-specific RNA aptamer was introduced into the Ll.LtrB intron in order to purify it over streptavidin resin (Figure 4A). After purification, the intron and any associated RNAs, were isolated and analyzed by reverse transcription. With the aptamer-containing intron, we observed that while the full-length pre-mRNA and spliced intron were pulled down as expected, the small mRNA (smRNA), generated from splicing of the aptamer-containing pre-mRNA with small exons (S-smRNA), was co-isolated. However, the smRNA produced either from the Int^- cell (C-smRNA) or from the aptamer-free Int⁺ cell (S-smRNA) were not co-isolated (Figure 4B). These results demonstrate that the intron is interacting with the spliced mRNA in the bacterial cell.

Figure 4

Download asset Open asset

Next, we investigated the possibility that the intron-mRNA interaction might account for the group II intron-promoted mRNA reduction. Similar to a previous study, we developed an in trans, two-plasmid expression system (Qu et al., 2014). We therefore co-expressed the intron-less ltrB relaxase from the pCY20 plasmid with either the Ll.LtrB intron-containing (with flanking small exons), or intron-less (small exons only) second plasmid, pLNRK (Figure 5A, left). Northern blotting analysis again showed that the level of relaxase mRNA dramatically dropped in the presence of the intron (21% vs. 100%). Additionally, an unexpected band smaller than the mRNA was detected (we call this band RNA 3, see below) (Figure 5A, right). Also, Western blotting indicated less accumulation of LtrB relaxase protein (24% vs. 100%).

Figure 5 with 2 supplements see all

Download asset Open asset

It was necessary to elucidate if mRNA loss in this case is due to retrohoming or RNA-RNA interaction alone. To this end, mutants of the intron’s IEP, LtrA, in which retrohoming was stopped due to inactivation of either the reverse transcriptase (RT) or DNA endonuclease (EN), were created (Figure 5—figure supplement 1A). RNA analysis showed that strong mRNA inhibition still persisted in these mutants (Figure 5—figure supplement 1B), thus indicating that the inhibition of gene expression was due to the proposed RNA-RNA interaction rather than retrohoming.

Using the trans-expression system, we validated that the intron-mRNA interaction requires EBS-IBS base-pairing, as shown previously in yeast (Qu et al., 2014). Mutants with nucleotide substitutions either in the IBS of the relaxase mRNA, or in the EBS in the intron expressed in trans, were compared to the respective wild-type counterparts (Figure 5B,C). The results showed that the relaxase mRNA level was recovered in the EBS mutant strain (Figure 5C, 21% to 92%), and increased ~2-fold in the IBS mutant (Figure 5D, 24% to 45%). Similar results were obtained with a different set of IBS-EBS mutants (m*), with recovery of mRNA levels in the EBS and IBS mutants from 23% to 116% and 23% to 51% respectively (Figure 5—figure supplement 2). Importantly, relaxase mRNA level was again reduced when the IBS mutation in the mRNA complemented the mutated EBS from the intron in the cell (Figure 5—figure supplement 2, down to 18%).

The observation that splicing ability of the intron is required for mRNA inhibition (data not shown) raised the possibility that the intron interaction with mRNA enables biochemical reactivity. We were also curious about the unexpected bands (RNAs 3 and 4) observed in Figure 5C and D respectively. Biochemical reactivity was validated and the origin of the extraneous bands was examined with the in trans system through a series of Northern blotting assays with multiple probes and confirmed by Rapid Amplification of cDNA Ends (RACE), which generates a cDNA copy for sequencing of the desired RNA. Four RNA molecules (RNA 1-RNA 4) with identity clearly distinct from the RNAs that were predicted to be expressed, were revealed (Figure 6A–C, Figure 6—figure supplement 1A–D, Figure 6—figure supplement 2).

Figure 6 with 2 supplements see all

Download asset Open asset

Among the four RNA molecules, two of them appeared in the intron-specific probing (Figure 6A; Figure 6—figure supplement 1A, lane 7). The larger of the two, termed RNA 1, was also revealed with both relaxase mRNA 5′- and 3′-exon probes (Figure 6B–C; Figure 6—figure supplement 1C–D, lane 7), and was identified as the intron-containing relaxase pre-mRNA. This was confirmed by probing a splicing-inactive catalytic triad mutant (T), which produced only this pre-mRNA band, RNA 1, and not RNAs 2–4 (Figure 6—figure supplement 1A,C,D, lane 3), and also by 5′-RACE followed by PCR amplification and sequencing of the cDNA (Figure 6—figure supplement 2A). This result suggests that the intron expressed in trans, is reverse splicing into the relaxase mRNA (Figure 6A). The smaller of the two intron-specific RNAs, termed RNA 2, was also revealed with the probe for 3′-exon but was absent in the probing for the 5′-exon (Figure 6B–C; Figure 6—figure supplement 1C–D, lane 7). With selective Northern blotting, along with 5′-RACE followed by cDNA sequencing, this RNA was identified as the unspliced intron expressed in trans, with its small 3′-exon replaced by the full-length 3′-exon of the relaxase mRNA (Figure 6A–B, Figure 6—figure supplement 1A,D, Figure 6—figure supplement 2A). This result suggests that there was shuffling of exons or RNA recombination, possibly between two reverse splicing reactions, that resulted in the formation of a chimeric precursor (Figure 6B).

The third RNA, RNA 3, previously seen in Figure 5, was observed in both the exon splice junction (Figure 6—figure supplement 1B, lane 7) and the relaxase mRNA 3′-exon-specific probings (Figure 6B; Figure 6—figure supplement 1D, lane 7), but it was absent from the probing for 5′-exon of the relaxase (Figure 6C; Figure 6—figure supplement 1C, lane 7). This RNA was further defined by 5′-RACE followed by cDNA sequencing as a ligation product of the small 5′-exon expressed in trans and the 3′-exon of the ltrB relaxase mRNA (Figure 6—figure supplement 2A). Identification of RNA 3 suggested that RNA 2, the chimeric intron precursor resulting from reverse splicing and RNA recombination described above, could be subject to forward splicing in the cell (Figure 6B). This result is in accord with splicing being favored over reverse splicing in vivo. Notably, RNA 3 appeared much more abundant than the relaxase mRNA (Figure 6B; Figure 6—figure supplement 1B,D).

Besides reverse splicing and its related RNA recombination, these experiments also suggested the occurrence of a spliced exons reopening (SER) reaction (Jarrell et al., 1988) (Figure 6C). This was evidenced by identification of the fourth RNA product, RNA 4, a free 5′-exon of the relaxase mRNA. RNA 4 was observed in the probing specific for the 5′-exon of the relaxase mRNA (Figure 6C; Figure 6—figure supplement 1C, lane 7) but was absent with all other probes (Figure 6A–B; Figure 6—figure supplement 1A,B,D, lane 7). Additionally, its presence was validated by 3′-RACE analysis followed by PCR amplification and sequencing of the cDNA products (Figure 6—figure supplement 2B). However, free 3′-exon of the relaxase mRNA, which should be produced simultaneously with the free 5′-exon from the SER reaction, was not detected either by Northern blotting or 3′-RACE (Figure 6B; Figure 6—figure supplement 1D, lane 7; Figure 6—figure supplement 2B). This could be attributed to bacterial mRNAs usually having a triphosphate at the 5′ end that is more resistant to mRNA decay than monophosphate (Celesnik et al., 2007; Schoenberg, 2007). Notably, this free 5′-exon RNA was also revealed when the intron was expressed in cis (Figure 6C; Figure 6—figure supplement 1C, lane 2; Figure 6—figure supplement 2B). Thus, these results suggest the SER, followed by RNA decay of the opened 3′-exon, to be a mechanism resulting in mRNA loss in the cell.

To further elucidate how these retargeting reactions are responsible for mRNA disappearance, we sought to examine mutants that were already created for this study. Initially we chose to identify the four RNA molecules in the intron_EBSm* and mRNA_IBSm* mutants (Figure 5—figure supplement 2). Using the in trans system, Northern blotting analyses showed that none of the four RNA products 1–4, that appeared with the wild-type strain existed in the mutants (Figure 7, lanes 3,5 versus lane 2). However, they reappeared when the EBS from the intron was mutated to complement the IBS within the relaxase mRNA in this mutant (Figure 7, lanes 6). This result confirmed that all of the identified mRNA retargeting reactions rely on the intron-mRNA interaction that is based on EBS-IBS base pairing.

Figure 7

Download asset Open asset

Inspired by the recently determined group II intron RNP structure and by the distribution features of group II introns in bacterial genomes, we performed a comparative expression study of intron-containing and intron-less variants of otherwise identical genes, to investigate the impact of group II introns on host gene expression. We discovered that group II introns reduce gene expression by robustly decreasing the spliced mRNA level in bacteria (Figure 1). This group II intron-induced mRNA disappearance appears independent of plasmids and promoters from which the genes are expressed (Figure 2), and is independent of stresses and changes of environmental conditions (data not shown). While this intron-induced reduction of mRNA levels was revealed mainly for the well-defined group IIA intron Ll.LtrB and its host ltrB relaxase gene in its native bacterial host Lactococcus lactis, it was also shown for two other types of bacterial group II introns, the group IIB EcI5 and group IIC BhI1 introns (Figure 2). Using the Ll.LtrB intron as the model, we also demonstrated that this mRNA reduction results from interaction between the intron with its spliced mRNA, via EBS-IBS base pairing (Figures 4, 5 and 7). Since ltrB is part of an operon containing also ltrC, ltrD, ltrE and ltrF (Chen et al., 2005), it will be interesting to determine how ltrB targeting by the group II intron affects levels of the entire transcript.

Interestingly, group II introns can also cause gene silencing in eukaryotic cells (Chalamcharla et al., 2010; Zerbato et al., 2013). Here, EBS-IBS-based intron-mRNA interactions shut down nuclear gene expression (Qu et al., 2014). Taken together, we conclude that group II introns are inherent and general inhibitors of gene expression.

By using the in trans system, we revealed that through RNA-RNA interactions, the spliced intron can retarget the mRNA in the cell (Figures 6–7, Figure 6—figure supplement 1 and 2). The intron’s ribozyme activities include reverse splicing into the spliced mRNA, and the SER reaction. Because forward splicing is usually favored in the cell, based on the high splicing efficiency shown in Figure 1B, Figure 1—figure supplements 1–2, we speculate that reverse splicing accounts for a relatively small portion of mRNA loss. Additionally, a robust SER, which was also revealed in the in cis system (Figure 6C), splits the 5′-exon from the mRNA, and the free 3′-exon appears to be degraded (Figure 8A).

Figure 8

Download asset Open asset

Although group II introns commonly invade DNA sequences, targeting RNA in vivo has not been previously described. However, RNP invasion of DNA is a relatively unfavorable event (Aizawa et al., 2003), which makes it tempting to speculate that the RNA SER and subsequent RNA degradation may favor DNA invasion and retromobility.

The RNA decay study with rifampicin indicated slower mRNA degradation in the Int⁺ cell (Figure 1—figure supplement 3). However, this result may be misleading as the residual RNA that escapes intron targeting may be a more stable fraction. Alternatively, the intron may act as ribosomes do on bacterial mRNA, providing a ‘protective barrier’ by binding the mRNA that escapes degradative attack by the group II intron (Deana and Belasco, 2005). Additionally, enhanced stability may be a reflection of the reciprocal relationship between the stability and concentration of bacterial mRNAs (Nouaille et al., 2017), such that lower mRNA levels in the Int⁺ cell, would lead to higher stability of the mRNA. Whichever mechanism underlies the phenomenon, the consequence is that a fraction of the mRNA is maintained for translation.

The discovery that group II introns inhibit gene expression likely explains why they are located mostly outside of genes or in non-essential genes, and why they are maintained at low frequency in bacterial genomes (Dai and Zimmerly, 2002a; Zimmerly and Semper, 2015). Group II intron-promoted reduction of gene expression could be the major driver that selected for group II introns residing in intergenic or non-essential regions. It is therefore intriguing that there are rare cases of essential genes containing group II introns in bacterial genomes (Candales et al., 2012; Dai et al., 2003; Dai and Zimmerly, 2002a; Zimmerly and Semper, 2015). We speculate that, in these cases, either robust gene expression is not needed, that reduction in gene expression is modest or that the intron plays a regulatory role (Belfort, 2017). The effect of organellar group II introns that are commonly present in essential genes is largely unknown (Dai and Zimmerly, 2002a; Zimmerly and Semper, 2015). Perhaps many of these introns have lost the ability to target mRNA and cause its loss, or do so very inefficiently. Alternatively, a lower expression level might be required for proper functions of these host genes. Notably, favoring the latter notion, a recent study showed that the presence of self-splicing introns including group II introns in the mitochondrion of Saccharomyces cerevisiae was required for inefficient expression of host genes that was essential for maintaining proper mitochondrial function (Rudan et al., 2018).

Group II introns are also frequently found in mobile DNAs (Candales et al., 2012; Dai et al., 2003; Dai and Zimmerly, 2002a; Zimmerly and Semper, 2015). One of the examples is the Ll.LtrB group II intron that naturally resides in the conjugative plasmid pRS01, interrupting the ltrB relaxase gene whose expression is required for the horizontal gene transfer (HGT) of the plasmid. A recent study indicated that pRS01 promotes retromobility of the intron by providing LtrB relaxase that stimulates both the frequency and diversity of retrotransposition (RTP) events with its off-target DNA nicking activity (Novikova et al., 2014). Here our study showed that the Ll.LtrB group II intron reduces the conjugal transfer of pRS01 via inhibition of its host gene expression (Figure 3). Thus, these findings together suggest that the two mobile genetic elements (MGEs) functionally interact by exploiting ltrB relaxase gene expression. Whereas relaxase expression stimulates intron RTP and HGT of the conjugative element, inhibition of relaxase expression by the group II intron opposes the promotion of RTP and HGT (Figure 8B), effects that are likely in equilibrium. In general, expression of conjugative transfer genes is tightly controlled to minimize the burden on the host (Zatyka and Thomas, 1998). Indeed conjugative plasmid gene expression is kept in a default ‘off’ state and is switched on only under conditions that are optimal for transfer of the conjugative element (Singh and Meijer, 2014). Silencing of the relaxase gene by its resident group II intron therefore represents a novel way in which a conjugative element is down-regulated.

The demonstration that intron RNAs are not only able to transpose and cleave exogenous RNAs, but can also recombine RNAs from different sources, suggests that progenitors of group II introns could have processed RNAs in primitive genomes. These actions could have contributed to the generation of an evolved genome and a cell that has novel functions and evolutionary advantages.

In regard to the ancestral relationship between group II and spliceosomal introns, expression of group II introns in nuclei causes gene silencing in eukaryotic cells (Chalamcharla et al., 2010; Zerbato et al., 2013). In this study, we demonstrated that group II introns also hinder gene expression in bacteria. Notably, a common mechanism underlying the group II intron-promoted inhibition of gene expression in bacteria and eukaryotes is based on interactions between intron and mRNA via EBS-IBS base pairing (Qu et al., 2014). In contrast to group II introns, many spliceosomal introns are shown to enhance gene expression in plants and other organisms (Le Hir et al., 2003; Rose, 2008). Interactions between the intron and mRNA no longer exist in the splicing of spliceosomal introns, where such pairings have been functionally substituted with interactions between U5 snRNA and mRNA (Hang et al., 2015; Qu et al., 2016; Yan et al., 2015). We speculate that, through these changes, introns can promote mRNA levels and gene expression, or at very least not be inhibitory, and therefore provided a force that could have stimulated evolution of group II introns into spliceosomal introns in primitive nuclear genomes.

Lactococcus lactis IL1403 was used for ltrB gene expression and RNA and protein analysis. IL1403 pRS01 ltrB^- (ΔLtrB::tet) and IL1403 (FA^R) were used as donor and recipient strains for conjugation, respectively. Cultures were grown in M17 media supplemented with 0.5% glucose (GM17) in tightly-capped tubes or bottles at 30°C without shaking. For gene expression, cultures were grown to OD₆₀₀ ~0.6 and nisin was added to a final concentration of 0.4 μg/ml, for 2–3 hr. Cultures were spun at 5000 x g and pellets were stored at −80°C. Where appropriate, the media contained spectinomycin (Spec) at 300 μg/ml, chloramphenicol (Cam) at 10 μg/ml, erythromycin (Erm) at 10 μg/ml, or fusidic acid (FA) at 25 μg/ml.

Escherichia coli MC1061(DE3) was used for over-expression of EcI5 and BhI1 plasmids. Cultures were grown in LB media with 100 μg/ml ampicillin at 37°C with aeration. At OD₆₀₀ ~0.3, cultures were induced with 0.1 mM IPTG for 3 hr. Cultures were spun at 5000 x g and pellets were stored at −80°C.

All plasmids created or used are listed in Supplementary file 1, and DNA oligonucleotides used in this study are listed in Supplementary file 1. Plasmids pCY20LtrB (Int^-) and pCY20LtrB::Ll.LtrB (Int⁺) were the vector pair used in the intron in cis system for comparative analysis of ltrB relaxase gene expression in the absence and presence of the Ll.LtrB intron. With the in trans system, pCY20LtrB was used for ltrB relaxase gene expression while plasmid pLNRK smEx::Ll.LtrB was used to express the intron. For an intron-less control in this system, pLNRK smEx::Ll.LtrB was replaced with plasmid pLNRK smEx, that contained only the small flanking exons. The ltrB IBS mutants, mRNA_IBSm and mRNA_IBSm*, and Ll.LtrB intron mutants including EBSm, EBSm* and Triad, and the IEP EN and RT mutants were created with the above pCY20 and pLNRK plasmids by site-directed mutagenesis (SDM) using QuickChange Lightning kit (Agilent, Santa Clara, CA). More specifically, to create EBSm*, the IBS region of the small exons of EBSm were mutated to make the intron splicing competent. This was done with 2 rounds of SDM PCR, using the plasmid product from PCR1 as the template for PCR2. All plasmids were confirmed by sequencing (see oligonucleotides used in Supplementary file 1). Construction of other plasmids used in this study are detailed below.

For RNA analysis, 7.5 μg of total RNA was separated on a 1.2% agarose/formaldehyde gel and transferred to hybond XL membrane (GE Healthcare, Pittsburgh, PA), which was then UV cross-linked and probed with ³²P-labeled oligonucleotides using Rapid-hyb buffer (GE Healthcare). For the Ll.LtrB intron, the membranes were hybridized for detection of mRNA exon splice junction (IDT4685), 5′-Exon (IDT5374), 3′-Exon (IDT5012), and Ll.LtrB intron (IDT1073). For the EcI5 intron, membranes were hybridized for the detection of mRNA (IDT4972) and intron (IDT4970), and for BhI1 intron, membranes were hybridized for mRNA (IDT4975) and intron (IDT4973). All membranes were probed for 16S rRNA (IDT861) as a loading control. Images were exposed on a phosphor screen, scanned on a Typhoon Trio, and quantified using ImageQuant.

For LtrB relaxase analysis, total cell lysate was separated on a 12% SDS-polyacrylamide gel and transferred to 0.2 μM Immuno-blot PVDF membrane (Bio-Rad) at 25V for 30 min. The membrane was blocked with 5% dry milk in TBS-T (20 mM Tris, 140 mM NaCl, 2% Tween), incubated with a 1/1500 dilution of primary anti-relaxase antibody for 1 hr, washed with TBS-T twice for 15 min, and incubated with a 1/10,000 dilution of secondary HRP-labeled anti-rabbit antibody (Advansta, Menlo Park, CA) for 1 hr. Chemiluminescent HRP substrate (Advansta WesternBright ECL) was used for detection. For lane normalization, total protein was visualized from a coomassie stained 12% SDS-polyacrylamide gel. All images were scanned using a Bio-Rad ChemiDoc MP. Relaxase bands and total protein were quantified using Bio-Rad Image Lab software.

cDNA synthesis was performed in a 20 μl reaction with 10 ng of DNase treated (Promega RQ1 DNase) RNA template using iScript (Bio-Rad, Hercules, CA), as per manufacturer’s protocol. Minus RT (RT^-) controls were also performed. Total RNA quality and primer specificity were analyzed by gel electrophoresis prior to qPCR. qPCR reactions were done in a total volume of 10 μl, using iTaq Universal SYBR Green Supermix (Bio-Rad) and contained 2 μl of the cDNA reaction as template and 5 pmol of each primer. Reactions were run in technical triplicates and a no-template control was included in every run. Reactions were amplified with the following conditions: 95°C for 30 s, (95°C for 5 s, 60°C for 10 s) x 40 cycles, melt curve 65°C to 95°C (0.5°C every 2 s), on a Bio-Rad CFX384 Touch Real-Time PCR Detection System. Three biological replicates were run for each sample. Amplification efficiencies (E) of all primers were calculated using a 10-fold dilution and standard curve. Efficiencies, C_t values and charts were obtained using Bio-Rad CFX Manager Software. Primers used, their percent amplification efficiencies and amplicon lengths are listed in Supplementary file 1. Relative gene expression of each target (tar) was normalized to a reference gene (ref), CopA, whose expression was demonstrated to be constant in a previous study (Magnani et al., 2008), and calculated using the following equation, with subtraction of RT^- background:

To identify splicing products, primer extension was performed using SuperScript III Reverse Transcriptase (Thermo Fisher Scientific, Waltham, MA), as per manufacturer’s protocol, with 4 μg of DNase-treated RNA and 0.4 pmol of ³²P-labeled oligonucleotides. Oligonucleotide IDT4836 was used with the addition of ddTTP for detection of intron precursor and spliced intron. Products were separated on an 8% Urea-polyacrylamide sequencing gel. To measure 5′-end transcription levels of the mRNA and intron precursor, IDT4916 was used, and products were separated on a 10% Urea-polyacrylamide gel. To analyze RNAs that were pulled down with streptavidin resin, IDT5078, IDT1073 and IDT5127 were used to probe the smRNAs, intron RNAs, and 6S non-coding RNA, respectively. To measure splicing efficiency of the intron_EBSm* mutant, oligonucleotide IDT1073 was used to detect the presence of intron precursor and spliced intron, and products were separated on a 10% Urea-polyacrylamide gel. Images were exposed on a phosphor screen, scanned on a Typhoon Trio, and quantified using ImageQuant.

L. lactis IL1403 was grown to OD₆₀₀ ~0.6 and nisin was added to a final concentration of 0.4 μg/ml for 0.5 hr. Rifampicin was then added to a final concentration of 0.2 mg/ml, and 10 ml of culture was removed for each aliquot and immediately spun down at 4°C. Total RNA was prepared and analyzed for Northern blots.

To perform polysome profiling, 200 ml of L. lactis IL1403 Int⁺/Int^- cells (1:20 dilution of saturated overnight culture) were grown and induced for gene expression with nisin for 2–3 hr. 100 mg/ml chloramphenicol was added and the cultures were chilled on ice for ~30 min with intermittent whirling and then collected by centrifugation at 5,000 rpm for 10 min at 4°C. Cell pellets were resuspended in 500 μl of ice-cold lysis buffer (20 mM Tris-HCl, pH 8.0; 140 mM KCl; 40 mM MgCl₂; 0.5 mM DTT; 100 μg/ml chloramphenicol; 1 mg/ml heparin; 20 mM EGTA; 1% Triton X-100) and washed twice with the same buffer. The cell pellets were resuspended again in 500 μl of lysis buffer and snap-frozen in liquid nitrogen. Then the cells were disrupted at 4°C in 15 ml falcon tubes with 500 μl of 0.1 mm ice-cold glass beads by rigorous vortexing (30 s for 20 times, with 1 min interval) and then briefly spun down at 4,000 rpm for 5 min. Crude cell lysate was then gently mixed, transferred into 1.5 ml tubes on ice, and cleared by 14,000 rpm for 25 min at 4°C. 400 μl of the cleared lysates were then loaded onto prepared 10–50% sucrose gradients in lysis buffer without 0.5 mg/ml heparin and centrifuged at 36,000 rpm for 153 min at 4°C (SW41 rotor). 30 fractions of ~400 μl each were collected for each gradient from top to bottom. RNAs were extracted from each fraction by using phenol/chloroform and analyzed by running gels and performing Northern blotting.

To measure conjugation efficiencies, L. lactis IL1403 (FA^R) was used as the recipient strain and was grown to an OD₆₀₀ ~0.6 and then for an additional 3 hr. L. lactis IL1403 ΔLtrB::tet (erm^R) was used as the donor strain, and contained either the Int^- plasmid pCY20LtrB, or the Int⁺ plasmid pCY20LtrB::Ll.LtrB. Cultures were grown to OD₆₀₀ ~0.6 and nisin was added to a final concentration of 0.4 μg/ml for 3 hr. Equal volume of donor and recipient were mixed, spotted on pre-incubated filters (0.45 μm pore size, Millipore) on GM17 plates, and incubated ~18 hr at 30°C. The filters were removed from the plates, washed with 5 ml GM17 in a 50 ml conical tube by vortexing, and the wash was spotted or plated on GM17 plus 25 μg/ml fusidic acid and 10 μg/ml erythromycin for selection of transconjugants. Plates were incubated ~18 hr at 30°C. To calculate input donor, the donor strain was diluted to 10⁻⁶ and spotted or plated on GM17 plus 10 μg/ml erythromycin. Conjugation frequency was expressed as exconjugants per donor (Erm^RFA^R/Erm^R).

Bacterial cells (100 ml) were collected after 2–3 hr of nisin induction of RNA expression and disrupted in 800 μl of CB₅₀₀ solution (20 mM Tris–HCl, pH 8.0, 500 mM NaCl, 0.1 mM EDTA, 1 mM PMSF) by vortexing (30 s, rest for 1 min, 24 cycles) using 500 μl of 0.1 mm ice-cold glass beads (Sigma). Lysates were cleared by centrifugation at 14,000 rpm for 25 min. To pull down RNAs, ~500 μl of cleared lysate was incubated with 100 μl of streptavidin resin (Thermo Scientific) that was equilibrated with CB₅₀₀. The resin was washed eight times with 1 ml CB₅₀₀ buffer. RNAs bound to resin were eluted with 400 μl of 5 mM biotin for 1 hr and were extracted from the eluates. RNA identities were determined by using reverse transcription primer extensions.

To further identify mRNA targeting products 1–4, 5' and 3' RACE (Rapid Amplification of cDNA Ends) experiments were performed. Briefly, Int⁺ and Int^- strains from cis and trans systems were grown and induced as previously described, and total RNA was prepared. RNAs were polyA-tailed using E. coli Poly(A) Polymerase (New England BioLabs, Ipswich, MA). 5' and 3' RACE experiments were then done using Invitrogen 5' or 3' RACE kits (Cat# 18374058 or 18373019), or the Takara SMARTer 5'/3' kit (Cat# 634859), following the kit protocols. Reverse transcriptase was used to synthesize cDNAs, which were then used as template for PCR amplification using GSP (gene specific primers IDT6074 and IDT6070 to ltrB 5' or 3' exon, respectively) along with a kit universal primer annealing to the synthesized cDNA. 5' and 3' RACE PCR products were run out on a 1.2% agarose gel, and the bands were excised, gel purified, and sequenced directly (Eton Bioscience, San Diego, CA) or cloned into pGEM-T vector (Promega), and then sequenced, to confirm their identity.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Alan Lambowitz for providing DNA for cloning the EcI5 group II intron. This work was supported by National Institutes of Health Grants GM39422 and GM44844 (to MB).

1. Received: December 12, 2017
2. Accepted: May 18, 2018
3. Version of Record published: June 15, 2018 (version 1)

© 2018, Qu et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.