Bacillus subtilis RNase J1 endonuclease and 59 exonuclease
activities in the turnover of DermC mRNA
SHIYI YAO, JOSH S. SHARP,1and DAVID H. BECHHOFER
Department of Pharmacology and Systems Therapeutics, Mount Sinai School of Medicine of New York University, New York, New York 10029,
RNase J1, a ribonuclease with 59 exonuclease and endonuclease activities, is an important factor in Bacillus subtilis mRNA
decay. A model for RNase J1 endonuclease activity in mRNA turnover has RNase J1 binding to the 59 end and tracking to
a target site downstream, where it makes a decay-initiating cleavage. The upstream fragment from this cleavage is degraded by
39 exonucleases; the downstream fragment is degraded by RNase J1 59 exonuclease activity. Previously, DermC mRNA was used
to show 59-end dependence of mRNA turnover. Here we used DermC mRNA to probe RNase J1-dependent degradation, and the
results were consistent with aspects of the model. DermC mRNA showed increased stability in a mutant strain that contained
a reduced level of RNase J1. In agreement with the tracking concept, insertion of a strong stem–loop structure at +65 resulted in
increased stability. Weakening this stem–loop structure resulted in reversion to wild-type stability. RNA fragments containing
the 39 end were detected in a strain with reduced RNase J1 expression, but were undetectable in the wild type. The 59 ends of
these fragments mapped to the upstream side of predicted stem–loop structures, consistent with an impediment to RNase J1 59
exonuclease processivity. A DermC mRNA deletion analysis suggested that decay-initiating endonuclease cleavage could occur
at several sites near the 39 end. However, even in the absence of these sites, stability was further increased in a strain with
reduced RNase J1, suggesting alternate pathways for decay that could include exonucleolytic decay from the 59 end.
Keywords: Bacillus subtilis; mRNA decay; RNase J1; endonuclease cleavage; 59 exoribonuclease
For some time, we have been using the mRNA encoded by
the erythromycin-resistance gene, ermC, to study aspects of
mRNA metabolism in Bacillus subtilis. ermC encodes a
ribosomal RNA methylase that confers resistance to eryth-
romycin (Em) by altering the Em binding site on the ribo-
some. Addition of low levels of Em results in induction of
ermC gene expression, and this occurs at the translational
level (Dubnau 1984; Weisblum 1985). In the translational
attenuation mechanism, addition of Em results in ribosome
stalling, now known to occur after incorporation of amino
acid 9 of the ermC leader peptide coding sequence (CDS)
(Vazquez-Laslop et al. 2008; Yao et al. 2008). This, in turn,
causes the opening of an inhibitory RNA secondary
structure, allowing high-level methylase translation. Our
initial interest in the ermC message came from the
observation that induction of ermC mRNA translation
was accompanied by strong stabilization of ermC mRNA.
The stabilization of ermC mRNA in the presence of Em was
attributed to the protective effect of an Em-bound ribo-
some that is stalled in the 59-proximal leader peptide
coding sequence (Bechhofer and Dubnau 1987; Bechhofer
and Zen 1989; DiMari and Bechhofer 1993). More recently,
we have worked with a deleted version of ermC, called
DermC (Fig. 1A), which has the first 16 codons of the leader
peptide CDS fused in-frame to the carboxy-terminal
portion of the ermC methylase CDS, giving a z260-
nucleotide (nt) mRNA that includes a 62-amino acid
CDS (Drider et al. 2002). The small size, high level of
transcription, and induced stability of DermC mRNA make
it a useful model for studying mRNA decay (Sharp and
Bechhofer 2003, 2005).
We have observed that, upon Em-induced ribosome
stalling, DermC mRNA undergoes a processing event that
results in the accumulation of a 215-nt RNA fragment
whose 59 end maps to +45 (Fig. 1A), which is the beginning
1Present address: Children’s Hospital Boston, Department of Infectious
Diseases, 300 Longwood Avenue, Enders Building, Room 750.7, Boston,
MA 02115, USA.
Reprint requests to: David H. Bechhofer, Department of Pharmacology
and Systems Therapeutics, Mount Sinai School of Medicine of New York
University, New York, NY 10029, USA; e-mail: firstname.lastname@example.org;
fax: (212) 996-7214.
Article published online ahead of print. Article and publication date are at
RNA (2009), 15:2331–2339. Published by Cold Spring Harbor Laboratory Press. Copyright ? 2009 RNA Society.
of codon 5 of the leader peptide CDS (Drider et al. 2002).
We showed recently that this processing event, whose
product is only detectable when Em is added and ribosome
stalling occurs, was dependent on RNase J1 (Yao et al.
2008). Several experiments provided strong evidence that
processing of DermC mRNA at the +45 site was the result
of RNase J1 endonucleolytic cleavage, rather than exo-
nuclease decay from the 59 end up to the stalled ribosome.
It is not known if cleavage at this site only occurs when
a ribosome stalls or if cleavage occurs constitutively, but the
downstream cleavage product is too unstable to be detected
unless a stalled ribosome is present to protect its 59 end.
RNase J1 was recently identified as a B. subtilis ribonu-
clease that has both endonuclease and 59-to-39 exonuclease
activities (Even et al. 2005; Britton et al. 2007; Mathy et al.
2007; Deikus et al. 2008; Li de la Sierra-Gallay et al. 2008).
RNase J1 is an essential enzyme and is believed to be
a major player in mRNA decay in B. subtilis. A similar
ribonuclease, RNase J2, is not essential. A strain in which
RNase J1 is under control of an IPTG-inducible promoter
contains a reduced level of RNase J1 when grown in the
presence of IPTG. This strain shows a small but significant
increase in overall mRNA half-life (Even et al. 2005). A
recent microarray study showed that the level of many
RNAs is affected in a strain that is deleted for RNase J2 and
that has reduced expression of RNase J1 (Mader et al.
2008). RNase J1 59-to-39 exonuclease activity requires
a monophosphate or hydroxyl 59 end and is inhibited by
FIGURE 1. Diagrams of DermC mRNA wild type and mutants. (A) Wild-type DermC mRNA showing ribosome binding site (hatched box near
59 end), site of RNase J1 cleavage upon Em-induced ribosome stalling (downward arrow), amino acids required for ribosome stalling (boxed),
codons in the ribosome P site and A site when ribosome stalling occurs, transcription terminator structure (39TT), and extent of 215-nt RNA
observed upon ribosome stalling. (Note that this latter processing product was formerly called the ‘‘209-nt RNA’’ [Drider et al. 2002; Yao et al.
2008], since the size of full-length ermC mRNA was taken as 254 nt. We now estimate the size of full-length DermC mRNA to be 260 nt; hence the
processed product is a 215-nt RNA.) At right are Northern blot analyses of DermC mRNA decay in wild-type and RNase J1 conditional mutant
strains, grown in the presence of IPTG. Time indicated above each lane is minutes after rifampicin addition. The half-life value 6 standard
deviation, which was the average of at least three determinations, is shown below the blots. As a control for the amount of RNA loaded, blots were
stripped and reprobed with a 5S rRNA probe. (B) Mutation of AU-rich DermC codons to GC-rich codons. (C) DermC mRNA with stop after
codon 2 and strong stem–loop inserted after codon 11. (D) Nucleotide sequence and predicted structure of strong and moderate stem–loops
inserted after codon 11. Arrowhead points to single nucleotide change that substantially weakens the structure. Free energies (kcal mol?1) of the
predicted structures are at the bottom. (E) Northern blot analysis of decay of constructs with strong or moderate stem–loop insert and in the
presence and absence of translation. (F) Plot of half-life data (average of three experiments) for DermC in wild-type strain (open circles), DermC
in RNase J1 mutant strain (closed squares), and DermC SSL>11 (closed triangles).
Yao et al.
RNA, Vol. 15, No. 12
a triphosphate 59 end (Mathy et al. 2007; Deikus et al. 2008;
Li de la Sierra-Gallay et al. 2008). On the other hand, the
endonuclease activity of RNase J1 is insensitive to the
phosphorylation state of the 59 end (Deikus et al. 2008; Li
de la Sierra-Gallay et al. 2008).
Current models of mRNA decay in B. subtilis (Bechhofer
2009) propose that RNase J1 can initiate mRNA decay in
one of two ways: RNase J1 may bind to the 59 end and de-
grade mRNA processively in the 59-to-39 direction. Pre-
sumably this activity is preceded by conversion of the 59
triphosphate nucleoside of the transcription product into
a 59 monophosphate nucleoside, as has been proposed for
the initiation of RNase E activity in E. coli (Celesnik et al.
2007; Deana et al. 2008). Alternatively, RNase J1 binds to
the 59 end and tracks to an endonucleolytic target site
(Condon 2007). Cleavage at this site results in an upstream
fragment that has an unprotected 39 end and that is de-
graded processively by 39-to-59 exonucleases, and a down-
stream fragment that is the target of further RNase J1
endonuclease cleavages and/or RNase J1 59-to-39 exonu-
clease activity. In this study, we examined involvement of
RNase J1 in DermC mRNA decay.
DermC mRNA decay is RNase J1 dependent
To determine the dependence of DermC mRNA decay on
RNase J1, a strain was constructed that carried a DermC
gene on a high-copy plasmid and that conditionally
expressed RNase J1. In this strain, RNase J1 transcription
is under the control of an IPTG-inducible promoter, and it
has been shown that addition of IPTG results in a level of
RNase J1 that is approximately fivefold lower than in the
wild-type strain (Daou-Chabo et al. 2009). DermC mRNA
half-life measurements were performed in the presence of
IPTG to avoid conditions of severe growth retardation that
occur in the absence of IPTG, where the essential RNase J1
is present at severely reduced levels and where one might
expect an effect on mRNA half-life due to slow growth rate.
Northern blot analysis of DermC mRNA decay in a wild-
type versus RNase J1 conditional strain showed a 2.5-fold
increase in half-life under conditions of reduced RNase J1
(Fig. 1A, right, Fig. 1F). This result suggested that decay of
DermC mRNA is largely dependent on RNase J1 activity, as
even a reduction (and not elimination) of this function
substantially increased mRNA half-life.
We next determined whether cleavage at the +45 site,
which we detect only in the presence of Em-induced
ribosome stalling (Drider et al. 2002; Yao et al. 2008),
was a factor in DermC mRNA half-life. A DermC mRNA
derivative was made in which the AU-rich nucleotide
sequence at the RNase J1 cleavage site was mutated to
a GC-rich sequence (Fig. 1B). The RNase J1 endonuclease
cleavage sites known to date all occur in AU-rich sequences
(Bechhofer 2009). We reasoned that, if RNase J1 cleavage at
this site was constitutive and was involved in initiating
decay, then altering the site such that RNase J1 would no
longer recognize it might result in a longer DermC mRNA
half-life. A similar sequence change at an RNase J1 target
site in trp leader RNA resulted in a fourfold increase in
RNA stability (Deikus and Bechhofer 2007). The effect of
the GC-rich sequence on RNase J1 recognition was con-
firmed by in vitro analysis of RNase J1 cleavage of 59-end-
labeled DermC mRNA (data not shown). A band represent-
ing cleavage in the AU-rich region around nucleotide 45
was observed for wild-type DermC mRNA, but was absent
for GC-rich mutant mRNA. Northern blot analysis (data
not shown) of the decay of RNA encoded by the construct
with the GC-rich sequence in codons 3–6 showed a half-life
of 6.8 6 0.4 min, which was barely significantly different
from the wild-type half life of 7.5 6 0.1 min (P-value =
0.044). Thus, RNase J1 cleavage at the +45 site is not
a factor in DermC mRNA decay, and there are likely target
sites further downstream on the DermC sequence.
Evidence for RNase J1 tracking from the 59 end
Since we had demonstrated earlier that DermC mRNA is
stabilized by the presence of secondary structure at the 59
end (Sharp and Bechhofer 2005), it was reasonable to
assume that RNase J1 binds at the 59 end and then tracks to
a target site downstream, as has been suggested for
initiation of B. subtilis mRNA decay generally (Condon
2003). We tested whether the presence of a strong second-
ary structure in the DermC coding sequence affected mRNA
half-life, as this could slow down RNase J1 tracking. A
construct was made that had a strong stem–loop inserted
after codon 11 (Fig. 1C). This construct was named SSL>11
(strong stem–loop after codon 11). The stem–loop se-
quence was the 59-terminal stem–loop structure of a mutant
mdr (or bmr3) RNA (Fig. 1D), which was shown to increase
mdr mRNA stability at least fourfold when located at the 59
end (Ohki and Tateno 2004). The SSL>11 construct was in
the context of a stop codon after codon 2, to insure that the
strong stem–loop was not unwound by ribosome transit. In
a Northern blot analysis, SSL>11 RNA was found to be
significantly more stable than wild-type DermC mRNA,
with a half-life of 10.9 min (Fig. 1E,F). Since we had found
earlier that the stability of DermC mRNA was unaffected by
the presence of a stop after codon 2 (Sharp and Bechhofer
2003), the increased stability of SSL>11 RNA was attributed
to the stem–loop structure insert, despite it being located
relatively far from the 59 end. This result suggested that the
presence of the strong stem–loop structure hindered RNase
J1 tracking, causing a delay of downstream target site
binding and decay-initiating cleavage. Two predictions of
this model were tested.
First, the absence of a stop after codon 2 might result in
a wild-type half-life, as the translating ribosome could melt
RNase J1-dependent decay of DermC mRNA
the strong structure. An SSL>11 construct was made that
did not contain the stop after codon 2. As shown in Figure
1E, the half-life of this construct was 8.1 min, similar to
that of wild-type DermC mRNA (P-value = 0.12).
Second, replacement of the strong stem–loop structure
after codon 11 with a moderately strong structure should
decrease half-life, as there would be less inhibition of
tracking from the 59 end. A construct was made that was
almost identical to SSL>11, but that had a single nucleotide
change in the stem sequence that lowered the stability of the
predicted structure substantially (Fig. 1D). This construct
was designated MSL>11 (moderate stem–loop after codon
11). The RNA encoded by this construct gave a half-life that
was 7.1 min, similar to that of wild-type DermC mRNA
(Fig. 1E). Furthermore, the stability of MSL>11 RNA was
only marginally sensitive to the stop codon after 2. When
the stop codon was changed to the wild-type isoleucine
codon, the half-life was 6.3 min (data not shown).
Detection of 39-end-containing decay intermediates
in the RNase J1 mutant strain
Our working hypothesis was that initiation of DermC
mRNA decay is dependent on cleavage at an RNase J1
target downstream from the site SSL>11 insertion, which
generates unstable upstream and downstream RNA frag-
ments. The upstream product of such cleavage would be
expected to be rapidly degraded by 39-to-59 exonuclease
activity. The downstream product of such cleavage would
be expected to be rapidly degraded by RNase J1.
We attempted to detect downstream products in the
RNase J1 conditional mutant strain. RNA isolated from the
wild-type strain and from an RNase J1 conditional mutant
strain grown in the absence of IPTG (i.e., severely reduced
RNase J1 levels) was probed with a labeled oligonucleotide
that was complementary to nucleotides 205–240 near the 39
end of DermC mRNA. The results in Figure 2A (DermC
blot) show that this probe did not detect any decay
intermediates in the wild-type strain. In the RNase J1
mutant strain, however, two prominent sets of bands,
designated ‘‘z140’’ and ‘‘z85,’’ in addition to several less
prominent bands, were detected (see the schematic of the
prominent bands in Fig. 2C). We postulated that these
bands were the result of endonuclease cleavage followed by
limited 59 exonuclease processing. In the wild-type strain,
which had a much higher level of RNase J1, these fragments
were rapidly degraded by the 59-to-39 exonuclease activity
of RNase J1, as we have shown previously that 39-terminal,
structured fragments can be degraded by RNase J1 (Deikus
et al. 2008). In the RNase J1 mutant strain, with a sharply
reduced level of RNase J1, these fragments were detectable
since they were not degraded as rapidly. We assumed that
the 39 end of these fragments was protected from 39-to-59
exonuclease decay by the extremely stable DermC transcrip-
tion terminator stem–loop, which has a DG0of ?16.2 kcal
mol?1. In data not shown, the prominent bands at z140
and z85 nt were also detectable in the RNase J1 conditional
strain grown in the presence of IPTG (i.e., only a fivefold
reduction in the RNase J1 level), but not nearly as strongly
as in the same strain grown in the absence of IPTG.
Additional Northern blotting was performed using
a high-resolution gel to resolve the multiple RNA frag-
ments that constituted the two major bands detected on the
low-resolution gel in Figure 2A. This analysis (Fig. 2B, lane
De) showed that the two sets of bands detected by the
DermC 39-terminal probe ran between nucleotides 128 and
143 and between nucleotides 80 and 95. These corre-
sponded to RNAs with 59 ends in the regions of nucleotides
117–132 and 165–180 (Fig. 2C).
To understand why these fragments were accumulating
in the strain with a reduced level of RNase J1, we looked for
secondary structure near the 59 ends of these fragments. The
DermC sequence was analyzed by the Zuker mfold version
3.3 program, using 50-nt increments 10 nt apart, i.e., 91–
140, 101–150, 111–160, etc. We used an arbitrary cutoff of
DG0 < ?3.0 kcal mol?1for consideration as a stable
structure. Only three predicted structures with significantly
low free energy were predicted. The first two of these, from
nucleotides 125–150 and 143–184, contained overlapping
sequences, so we chose the sequence from 125–150, which
was considerably more stable with a DG0 = ?6.4 kcal
mol?1. The other sequence was from 167–202, with a DG0=
?5.1 kcal mol?1. The sequence of these predicted secondary
structures, which were designated eSL1 (ermC stem–loop 1)
and eSL2, are shown in Figure 2C. We surmised that
endonuclease cleavage at sites upstream of these structures
(Fig. 2C, downward open arrows) would, under conditions
of low RNase J1 activity, result in the accumulation of decay
intermediates with 59 ends that map to the upstream side of
these structures. To test this, the 39 stem sequence of eSL2
was changed to a NotI sequence (Fig. 2C), which eliminated
eSL2 secondary structure. Northern blot analysis of this
RNA on a low-resolution gel (Fig. 2A, NotI blot) showed
that this change had no effect on the z140-nt band, but
resulted in loss of the z85-nt band and appearance of
a z70-nt band. Results from a high-resolution gel (Fig. 2B,
NotI lane) showed that the upstream fragments with 59 ends
in the region from 117–132 were unchanged by this
mutation whereas the downstream fragments with 59 ends
from 165–180 were no longer present. Instead, fragments
that had 59 ends mapping from 187–200 were observed (see
Discussion). In a separate construct, we made a change in the
59 stem sequence of eSL1 to give a ClaI site (Fig. 2C), which
eliminated eSL1 secondary structure. The low-resolution gel
in Figure 2A (ClaI blot) showed loss of the z140-nt band
and retention of the z85-nt band. These results were con-
sistent with RNase J1 endonucleolytic cleavage upstream of
eSL1 and eSL2, followed by RNase J1 59-to-39 exonucleo-
lytic degradation being hindered by the eSL1 and eSL2
Yao et al.
RNA, Vol. 15, No. 12
Pattern of 39-proximal decay
intermediates in the absence
of ribosome transit
We tested whether detection of the decay
intermediates in the strain with reduced
RNase J1 levels was dependent on ribo-
some transit. Decay intermediates from
2 were analyzed using the 39-proximal
RNase J1 mutant background was identi-
mRNA. Thus, the putative endonuclease
cleavages in the body of the message were
not dependent on ribosome flow.
Differential accumulation of
According to our hypothesis, the de-
tection of the 39-end-containing frag-
ments in the RNase J1 conditional mu-
tant strain was possible because of slower
decay of these fragments due to a re-
duction in the level of RNase J1 59-to-39
exonuclease activity. However, long ex-
posure times were required to observe
these decay intermediates, suggesting
that they do not accumulate. These de-
cay intermediates differed from the 215-
nt RNA seen upon RNase J1 cleavage at
+45 (Fig. 1A), which accumulates with
time after rifampicin addition and is easy
to detect (Drider et al. 2002; Yao et al.
2008). We analyzed the decay of pro-
cessing intermediates in the RNase J1
conditional mutant strain (grown in the
presence of IPTG), after addition of Em and subsequent
addition of rifampicin. As can be seen in the shorter ex-
posure in Figure 3B, the amount of the 215-nt fragment in-
creased with time, as the full-length RNA was processed by
RNase J1 endonuclease cleavage at +45. (The 215-nt RNA ac-
cumulates to higher levels in a wild-type strain that has a
higher level of RNase J1.) By contrast, the amount of the two
sets of bands that represent 39-end-containing fragments
generated by downstream processing decreased with time
(Fig. 3B, long exposure). Most likely, the 215-nt fragment is
protected from further 59-to-39 degradation due to the
stalled ribosome at its 59 end, whereas the downstream
fragments are subject to ongoing 59-to-39 decay.
If endonuclease cleavage at RNase J1 target sites in the body
of DermC mRNA was responsible for its decay, then
deletion of these sites should result in a more stable DermC
mRNA. Thus, we analyzed the stability of RNAs encoded by
a number of DermC deletion mutants. The half-life of wild-
type DermC mRNA in these experiments was about 7 min
(Fig. 4A). Deletion of small regions starting from near the
39 end (Fig. 4B–D) had no significant effect on DermC
mRNA half-life. Even deletion of a 95-nt segment that
included both regions in which endonuclease cleavages
were thought to occur (Fig. 4E) did not significantly affect
DermC mRNA half-life. These results suggested that elim-
ination of one or more endonuclease target sites was
insufficient to confer increased stability, and perhaps
additional target sites were still present. Interestingly, de-
letion of a 65-nt segment that was closer to the 39 end (Fig.
4F) resulted in a significant, 40% increase in half-life,
despite the fact that the upstream target site should still be
present in this construct.
FIGURE 2. Northern blot analysis of 39-end-containing decay intermediates. (A) Steady-state
pattern of 39-end-containing decay intermediates in the wild-type (wt) and RNase J1 conditional
mutant strain (J1), grown in the absence of IPTG. Blots of wild-type DermC RNA, NotI mutant
RNA, and ClaI mutant RNA are shown, as indicated. On the left are the sizes (nucleotides) of
59-end-labeled pSE420 TaqI DNA fragments (Brosius 1992) run in a parallel lane. On the right,
the migration of full-length (FL) RNA and major decay intermediate bands are indicated. (B)
High-resolution Northern blot analysis of DermC wild-type (De) and NotI construct RNA.
Locations of 59 ends of detected decay intermediates are indicated on the left. The sequencing
ladder was generated from M13mp18 single-stranded DNA. Sizes of the sequencing ladder bands
are indicated on the right. (C) Partial nucleotide sequence of the downstream portion of DermC
mRNA, showing DermC stem–loops 1 and 2 and their predicted free energies, as well as location
of NotI and ClaI mutations. Approximate 59 ends of the two groups of decay intermediates are
shown schematically below, as well as the location of the complementary 39 probe.
RNase J1-dependent decay of DermC mRNA
When a large, 129-nt deletion was made that included
nucleotides 87–215 (Fig. 4G), the half-life of DermC mRNA
jumped twofold to about 15 min. The increase in stability of
this RNA could have been due to its small size, which was
only 131 nt. Two controls for RNA size were constructed, in
which deletions of the same size were made elsewhere in the
RNA (Fig. 4H,I). In both cases, the DermC mRNA half-life
was similar to that of wild type. Taken together with the
results obtained with 39-end-containing decay intermediates
(Fig. 2), the deletion analysis indicated that there were at
least three target sites for initiation of decay by endonucle-
ase cleavage (Figure 4A, arrows), and that elimination of
these three targets had a profound effect on mRNA half-life.
A prediction of this conclusion was that insertion of an
endonuclease target site into the stable construct shown in
Figure 4G should result in destabilization. This was the case:
insertion of nucleotides 151–215 into an upstream region of
the stable deletion mutant (Fig. 4J) resulted in a half-life
that was actually somewhat shorter than wild type.
Finally, the stability of the 129-nt
deletion construct (Fig. 4G) was tested
in the RNase J1 conditional mutant.
If, in fact, the deletion eliminated all
RNase J1 recognition sites, and decay was
proceeding by an RNase J1-independent
pathway, we expected that, unlike the
case with wild-type DermC mRNA (Fig.
1A), the reduction in RNase J1 levels
should not affect stability. Surprisingly,
we found that the stability of the RNA
with the large deletion was even greater
in the RNase J1 mutant (Fig. 4G, right),
with a half-life of more than 40 min. This suggested that
RNase J1 is involved even in decay of the RNA encoded by
the 129-nt deletion construct, perhaps by a 59-to-39
exonucleolytic pathway or by cleavage at sites in the
upstream half of the RNA.
Although the DermC gene is an artificial construct, it has all
the elements of a small mRNA, and, we believe, can be used
as a model to study mRNA decay. It should be noted that
the DermC mRNA sequence has a low GC content (27%),
even lower than that of B. subtilis (43%), which is char-
acterized as a ‘‘low GC’’ organism. It is therefore possible
that DermC mRNA contains a higher than average density
of target sites for RNase J1, which appears to cleave pref-
erentially at AU-rich sequences (Bechhofer 2009).
Decay intermediates are normally difficult to detect.
Once the initial, rate-determining step in mRNA decay
FIGURE 3. Northern blot analysis of (A) 39-end-containing decay intermediates detected with
(De) and without (stop > 2) translation, and (B) processed RNA and decay intermediates in
the presence of Em-induced ribosome stalling.
FIGURE 4. Deletion analysis of DermC mRNA. Schematic diagram of (A) wild-type (wt) DermC mRNA is shown at the top, with downward
arrows indicating putative sites of RNase J1 endonucleolytic cleavage. Half-lives (average of at least three determinations) are shown at right,
together with P-value relative to wild-type half-life. For the deletion constructs B–J, the extent of the deletion in nucleotides is indicated at the left
and represented by parentheses on the schematic diagrams. Construct J contained an insertion of nucleotides 151–215 into the 59-proximal region
of construct G. Northern blot analysis of construct G half-life, in wild-type and RNase J1 conditional mutant strains (grown in the presence of
IPTG), is shown at the right of the construct G schematic.
Yao et al.
RNA, Vol. 15, No. 12
occurs, RNA fragments that are generated in the course of
complete turnover are rapidly degraded. The use of DermC
mRNA enabled the analysis of intermediates in the process
of mRNA decay. With relatively long exposure times, we
were able to detect 39-end-containing DermC mRNA decay
intermediates in the RNase J1 conditional mutant strain.
The combination of DermC on a high-copy plasmid, tran-
scription from a strong promoter, secondary structure to
slow rapid 59-to-39 decay, and a strain with reduced RNase
J1 levels made this detection possible.
We found that DermC mRNA half-life doubled even
under conditions where RNase J1 was still present, but at
reduced levels (Fig. 1A), which indicated strongly that
DermC mRNA decay is dependent on RNase J1 activity. An
important goal of this study was to determine which of
the two ribonuclease activities of RNase J1, 59 exonuclease
or endonuclease (or both) is (are) involved in decay of a
particular mRNA. Several findings argue against DermC
mRNA being degraded solely exonucleolytically from the 59
end (which would initiate after 59 pyrophosphate removal).
First, we found that the presence of a strong stem–loop
structure at nucleotide +65, which presumably is too far
downstream to affect the 59 end as a substrate for RNase
J1 59 exonuclease activity, resulted in increased stability
(Fig. 1D). Even if 59-to-39 exonuclease processivity would
be slowed by the inserted stem–loop, this would not affect
a half-life measurement that is dependent on the amount of
full-length RNA remaining. Second, if DermC mRNA were
degraded only by an exonucleolytic pathway, there should
be no effect of deletions situated well downstream from the
59 end. We found, however, that a 129-nt deletion start-
ing at nucleotide 87 caused a substantial increase in half-
life (Fig. 4G). Furthermore, insertion of a 65-nt segment
from the downstream half of DermC mRNA, in which we
mapped possible endonuclease cleavage sites, resulted in
destabilization of the 129-nt deletion construct (Fig. 4J),
which is also inconsistent with attack solely from the 59
end. We conclude, therefore, that DermC mRNA decay is,
at least to some extent, dependent on endonucleolytic
cleavage. The stability conferred by 59-terminal stem–loop
structure (Sharp and Bechhofer 2005) is presumably due to
interference with RNase J1 binding at the 59 end, which is
required not only for exonuclease activity, but also for
initiation of tracking to downstream target sites.
Although we have not formally proved that the endo-
nuclease activity involved in DermC mRNA decay is that of
RNase J1, this is a reasonable interpretation of our results.
The demonstrated 59-end-dependence of DermC mRNA
stability (Sharp and Bechhofer 2005) suggests involvement
of RNase J1, whose 59 exonuclease activity is known to be
59-end dependent and perhaps its endonuclease activity
requires a 59 end as well. An earlier review of mRNA decay
in B. subtilis (Condon 2003) compared the 59-end-
dependence of decay in this organism versus 59-end-
dependence of RNase E in E. coli. From many studies,
a model was suggested in which E. coli RNase E binds to the
59 end of a message, but then loops around potential
roadblocks (ribosomes, bound protein, RNA secondary
structure) to its endonuclease target sites, whereas the sim-
ilarly functioning 59-dependent endonuclease in B. subtilis
(not known to be RNase J1 at the time) depends on
tracking on the mRNA in the 59-to-39 direction to reach its
target cleavage site. The observation of mRNA stabilization
by an internal strong stem–loop structure (the SSL>11
RNA) (Fig. 1C) is consistent with this model. When this
structure was disrupted, either by ribosome transit or by
introduction of a destabilizing base change, the half-life
reverted to wild type (Fig. 1E). Thus, we hypothesize that
RNase J1 binds to the 59 end of DermC (or to a site close to
the 59 end) and then tracks downstream until it reaches
a target site. Insertion of the SSL after codon 11 hinders
RNase J1 tracking to downstream target sites, which is
necessary for initiation of decay. From the fact that the
stabilizing effect of SSL>11 was abolished by translation of
the message (Fig. 1E), we surmise that translated mRNAs
have naturally few roadblocks to RNase J1 tracking to down-
stream endonuclease target sites. More likely, the regulation
of mRNA stability by interference with RNase J1 activity
occurs at the binding step at the 59 end, upstream of ribo-
some flow. We have found that, when the same stem–loop
structure that is present in the SSL>11 construct was placed
at the 59 end of an RNA that has a 59-proximal RNase J1
target site, cleavage by RNase J1 at this site was virtually
abolished (Yao and Bechhofer 2009).
Our earlier finding of RNase J1 cleavage at the +45 site
(Drider et al. 2002; Yao et al. 2008) was shown here not to be
a factor in constitutive mRNA decay (based on results with
the construct shown in Fig. 1B). Rather, RNase J1 cleavage at
+45 is likely a consequence of Em-induced ribosome stalling.
RNases J1 and J2 were originally isolated from a high salt
ribosomal wash fraction (Even et al. 2005), and biochemical
studies have shown that RNase J1 colocalizes with ribosomes
(Hunt et al. 2006), suggesting a strong interaction. Thus, the
observed association of RNase J1 with ribosomes fits well
with the notion of RNase J1-mediated cleavage at the +45
site in response to ribosome stalling. However, it is clear that
RNase J1 endonuclease cleavage is not completely ribosome
dependent, since such cleavage occurs in trp leader RNA
(Deikus and Bechhofer 2007; Deikus et al. 2008) and in small
cytoplasmic RNA (Yao et al. 2007), which are untranslated
RNAs. Here, too, we showed that the pattern of DermC
mRNA decay intermediates, which we suggest arises follow-
ing RNase J1 endonuclease cleavage, was the same in the
presence and absence of ribosome flow (Fig. 3A). Thus,
ribosome-dependent cleavage by RNase J1 at the +45 site is
likely a special case of endonuclease cleavage in response to
an urgent situation (ribosome stalling), but endonuclease
activity of RNase J1 is generally not ribosome dependent.
We reported earlier that fragments containing the
39-terminal sequence are detectable in an RNase J1
RNase J1-dependent decay of DermC mRNA
conditional mutant (Deikus et al. 2008). These were
fragments that contained the transcription terminator
sequence, and, as such, we could not study the effect of
sequence changes in these structures, since changes would
also affect transcription termination. Here, we obtained
evidence that even weaker structures, such as eSL1 and
eSL2, are capable of affecting RNase J1 59-to-39 processing.
The fact that the pattern of 39-end-containing intermedi-
ates was the same, whether or not DermC mRNA was
translated (Fig. 3A), was somewhat surprising, since trans-
lation made a significant difference in the effect of SSL>11
(Fig. 1E). Perhaps the process of tracking in the 59-to-39
direction and the process of degradation in the 59-to-39
direction relate differently to the flow of ribosomes. More
experiments are needed to address this issue.
The 39-end-containing fragments were visible in the
mutant strain with reduced RNase J1 levels, but not in
the wild-type strain (Fig. 2A). We hypothesize that the
wild-type level of RNase J1 is enough to cause rapid
degradation of such fragments. When the level of RNase
J1 is reduced, the endonucleolytic cleavages, which are
required only once per molecule to initiate decay, still
occur at a wild-type or near wild-type rate, but the impeded
processivity of RNase J1 in the 59-to-39 direction by
secondary structure may cause RNase J1 to release the
substrate, and only when high levels of RNase J1 are present
does the continual binding and 59-to-39 degradation ensure
rapid removal of the 39-end-containing fragments.
The ‘‘bands’’ detected on Northern blots from low-
resolution gels were shown to be a large group of bands
on Northern blotting from a high-resolution gel (Fig. 2B).
Since the intensity of these bands was much reduced for
DermC mutants with sequence changes that disrupted eSL1
or eSL2, it is likely that these structures are somewhat
resistant to RNase J1 59 exonuclease processivity. However,
our mapping of these bands is not entirely consistent with
a fall-off of RNase J1 upstream of the structure. Rather, the
bands map part way up the 59 side of the structure itself.
Perhaps the slowing of RNase J1 processivity caused by
these structures increases the likelihood of substrate release,
and such release can occur several nucleotides downstream
from where processivity begins to be affected. Future
studies will look at the effects of a number of secondary
structures predicted to form in vivo, as well as RNase J1
activity on eSL1 and eSL2 in vitro.
The increased intensity of bands 60-73 nt long in the
NotI mutant construct, representing 39 decay intermediates
with 59 ends between 187 and 200 nt, was of interest. The
inserted GC-rich sequence, which constitutes the NotI site
in the DNA, starts at nucleotide 189. We have found in
studies on trp leader RNA that the GC-rich sequence of a
NotI site is inhibitory to PNPase processivity in the 39-to-59
direction, both in vivo and in vitro (Deikus and Bechhofer
2009). We speculate that this particular sequence is also
inhibitory to RNase J1 processivity in the 59-to-39 di-
rection. Therefore, disrupting the eSL2 structure eliminates
decay intermediates with 59 ends between 165 and 180,
which we hypothesize are due to partial blockage to RNase
J1 processivity, but results in intensification of decay
intermediates with 59 ends between 187 and 200 (Fig.
2B). Again, further in vitro studies with purified RNase J1
are required to test this.
The deletion construct analysis (Fig. 4) suggested that
there are at least three preferred RNase J1 target sites
located in the downstream half of DermC mRNA. These are
located upstream of eSL1, between eSL1 and eSL2, and
between eSL2 and the 39 transcription terminator. Deletion
of all these three target sites resulted in increased mRNA
half-life (z15 min) (Fig. 4G). RNase J1 cleavage sites that
have been mapped in other RNAs occur between or near
regions of structured RNA, so the presence of eSL1 and
eSL2 may render the AU-rich sequences near these struc-
tures preferred target sites. Nevertheless, the stability of the
RNA encoded by this deletion construct was further
enhanced in an RNase J1 conditional mutant, where the
half-life was >40 min (Fig. 4G). It is possible that increased
stability conferred by the 129-nt deletion is due to
elimination of only one element in the RNase J1 decay
pathway. Other elements could include secondary RNase J1
targets in the upstream half of the mRNA, as well as
exonucleolytic decay from the 59 end. It will be interesting
to determine half-lives of DermC mRNA constructs that
combine 59 stabilizer sequences at the 59 end (Sharp and
Bechhofer 2005) and the 129-nt deletion, to see whether
even such constructs are further stabilized in an RNase J1
MATERIALS AND METHODS
The B. subtilis host was BG1, which is trpC2 thr-5. Chromosomal
DNA from the RNase J1 conditional mutant strain (Britton et al.
2007) was used to transform BG1 to Em resistance. The RNase J1
conditional strain also contained plasmid pMAP65, which carries
extra copies of the lacI gene (Petit et al. 1998). The preparation
and transformation of B. subtilis competent cell cultures were as
described previously (Dubnau and Davidoff-Abelson 1971). E. coli
strain DH5a (Grant et al. 1990) was the host for plasmid
The wild-type DermC gene was carried on plasmid pYH250, an
E. coli/B. subtilis shuttle plasmid that contains a chloramphenicol
resistance marker (Sharp and Bechhofer 2005). In pYH250,
a HindIII site is present at the beginning of the DermC transcrip-
tional unit and an EcoRI site is located downstream from the
DermC transcription terminator. The HindIII site was incorpo-
rated into mutagenic PCR primers that included nucleotide
sequence changes in the DermC CDS. These 59 primers were used
in conjunction with a 39 primer containing the downstream EcoRI
Yao et al.
RNA, Vol. 15, No. 12
site to amplify DermC, giving an amplicon that was cloned as
a HindIII-EcoRI fragment into the pYH250 vector. For some
constructs, a mutagenic oligonucleotide that contained the HpaI
site consisting of codons 17 and 18 was used in conjunction with
a primer that contained the XbaI site located about 600 base pairs
upstream of DermC. For construction of the NotI and ClaI
mutants (Fig. 2C), complementary mutagenic oligonucleotides
containing the mutated sequence were used in separate PCR
reactions with an oligonucleotide containing either the upstream
XbaI sequence or the downstream EcoRI sequence. The two PCR
amplicons were annealed to each other and were amplified in
a second round with the upstream and downstream primers. The
mutated fragment was digested with XbaI and EcoRI and used to
replace the small XbaI-EcoRI fragment of pYH250.
Supplemented minimal medium for RNA isolation, Northern blot
analysis, and 5S rRNA probing as a quantitation control were as
described (Oussenko et al. 2005). For expression of RNase J1 in the
conditional mutant strain, IPTG was added to 1 mM. For the
analysis of steady-state RNA shown in Figure 2A,B, strains were
length DermC mRNA and RNA processing products was done with
a Storm 860 PhosphorImager (Molecular Dynamics) or a Typhoon
TRIO variable mode imager (GE Healthcare).
This work was supported by Public Health Service grant GM-
48804 from the National Institutes of Health.
Received May 22, 2009; accepted August 24, 2009.
Bechhofer DH. 2009. Chapter 6 messenger RNA decay and matura-
tion in Bacillus subtilis. Prog Nucleic Acid Res Mol Biol 85: 231–273.
Bechhofer DH, Dubnau D. 1987. Induced mRNA stability in Bacillus
subtilis. Proc Natl Acad Sci 84: 498–502.
Bechhofer DH, Zen KH. 1989. Mechanism of erythromycin-induced
ermCmRNAstability inBacillussubtilis.J Bacteriol171:5803–5811.
Britton RA, Wen T, Schaefer L, Pellegrini O, Uicker WC, Mathy N,
Tobin C, Daou R, Szyk J, Condon C. 2007. Maturation of the 59
end of Bacillus subtilis 16S rRNA by the essential ribonuclease
YkqC/RNase J1. Mol Microbiol 63: 127–138.
Brosius J. 1992. Compilation of superlinker vectors. Methods Enzymol
Celesnik H, Deana A, Belasco JG. 2007. Initiation of RNA decay in
Escherichia coli by 59 pyrophosphate removal. Mol Cell 27: 79–90.
Condon C. 2003. RNA processing and degradation in Bacillus subtilis.
Microbiol Mol Biol Rev 67: 157–174.
Condon C. 2007. Maturation and degradation of RNA in bacteria.
Curr Opin Microbiol 10: 271–278.
Daou-Chabo R, Mathy N, Benard L, Condon C. 2009. Ribosomes
initiating translation of the hbs mRNA protect it from 59-to-39
exoribonucleolytic degradation by RNase J1. Mol Microbiol 71:
Deana A, Celesnik H, Belasco JG. 2008. The bacterial enzyme RppH
triggers messenger RNA degradation by 59 pyrophosphate re-
moval. Nature 451: 355–358.
Deikus G, Bechhofer DH. 2007. Initiation of decay of Bacillus subtilis
trp leader RNA. J Biol Chem 282: 20238–20244.
Deikus G, Bechhofer DH. 2009. Bacillus subtilis trp leader RNA: RNase
J1 endonuclease cleavage specificity and PNPase processing. J Biol
Chem 284: 26394–26401.
Deikus G, Condon C, Bechhofer DH. 2008. Role of Bacillus subtilis
RNase J1 endonuclease and 59-exonuclease activities in trp leader
RNA turnover. J Biol Chem 283: 17158–17167.
DiMari JF, Bechhofer DH. 1993. Initiation of mRNA decay in Bacillus
subtilis. Mol Microbiol 7: 705–717.
Drider D, DiChiara JM, Wei J, Sharp JS, Bechhofer DH. 2002.
Endonuclease cleavage of messenger RNA in Bacillus subtilis.
Mol Microbiol 43: 1319–1329.
Dubnau D. 1984. Translational attenuation: The regulation of
bacterial resistance to the macrolide–lincosamide–streptogramin
B antibiotics. CRC Crit Rev Biochem 16: 103–132.
Dubnau D, Davidoff-Abelson R. 1971. Fate of transforming DNA
following uptake by competent Bacillus subtilis. I. Formation
and properties of the donor–recipient complex. J Mol Biol 56:
Even S, Pellegrini O, Zig L, Labas V, Vinh J, Brechemmier-Baey D,
Putzer H. 2005. Ribonucleases J1 and J2: Two novel endoribonu-
cleases in B. subtilis with functional homology to E. coli RNase E.
Nucleic Acids Res 33: 2141–2152.
Grant SG, Jessee J, Bloom FR, Hanahan D. 1990. Differential plasmid
rescue from transgenic mouse DNAs into Escherichia coli methylation-
restriction mutants. Proc Natl Acad Sci 87: 4645–4649.
Hunt A, Rawlins JP, Thomaides HB, Errington J. 2006. Functional
analysis of 11 putative essential genes in Bacillus subtilis. Micro-
biology 152: 2895–2907.
Li de la Sierra-Gallay I, Zig L, Jamalli A, Putzer H. 2008. Structural
insights into the dual activity of RNase J. Nat Struct Mol Biol 15:
Mader U, Zig L, Kretschmer J, Homuth G, Putzer H. 2008. mRNA
processing by RNases J1 and J2 affects Bacillus subtilis gene
expression on a global scale. Mol Microbiol 70: 183–196.
Mathy N, Benard L, Pellegrini O, Daou R, Wen T, Condon C. 2007.
59-To-39 exoribonuclease activity in bacteria: Role of RNase J1 in
rRNA maturation and 59 stability of mRNA. Cell 129: 681–692.
Ohki R, Tateno K. 2004. Increased stability of bmr3 mRNA results in
a multidrug-resistant phenotype in Bacillus subtilis. J Bacteriol 186:
Oussenko IA, Abe T, Ujiie H, Muto A, Bechhofer DH. 2005.
Participation of 39-to-59 exoribonucleases in the turnover of
Bacillus subtilis mRNA. J Bacteriol 187: 2758–2767.
Petit MA, Dervyn E, Rose M, Entian KD, McGovern S, Ehrlich SD,
Bruand C. 1998. PcrA is an essential DNA helicase of Bacillus
subtilis fulfilling functions both in repair and rolling-circle
replication. Mol Microbiol 29: 261–273.
Sharp JS, Bechhofer DH. 2003. Effect of translational signals on
mRNA decay in Bacillus subtilis. J Bacteriol 185: 5372–5379.
Sharp JS, Bechhofer DH. 2005. Effect of 59-proximal elements on
decay of a model mRNA in Bacillus subtilis. Mol Microbiol 57: 484–
Vazquez-Laslop N, Thum C, Mankin AS. 2008. Molecular mechanism
of drug-dependent ribosome stalling. Mol Cell 30: 190–202.
Weisblum B. 1985. Inducible resistance to macrolides, lincosamides,
and streptogramin type B antibiotics: The resistance phenotype,
its biological diversity, and structural elements that regulate
expression—a review. J Antimicrob Chemother 16 Suppl A: 63–90.
Yao S, Bechhofer DH. 2009. Processing and stability of inducibly
expressed rpsO mRNA derivatives in Bacillus subtilis. J Bacteriol
Yao S, Blaustein JB, Bechhofer DH. 2007. Processing of Bacillus
subtilis small cytoplasmic RNA: Evidence for an additional
endonuclease cleavage site. Nucleic Acids Res 35: 4464–4473.
Yao S, Blaustein JB, Bechhofer DH. 2008. Erythromycin-induced
ribosome stalling and RNase J1-mediated mRNA processing in
Bacillus subtilis. Mol Microbiol 69: 1439–1449.
RNase J1-dependent decay of DermC mRNA