Accumulation of noncoding RNA due to an RNase P
defect in Saccharomyces cerevisiae
MICHAEL C. MARVIN,1SANDRA CLAUDER-MU¨NSTER,2SCOTT C. WALKER,1ALI SARKESHIK,3
JOHN R. YATES III,3LARS M. STEINMETZ,2and DAVID R. ENGELKE1,4
1Department of Biological Chemistry, University of Michigan, Ann Arbor, Michigan 48109-0606, USA
2European Molecular Biology Laboratory, 69117 Heidelberg, Germany
3Department of Chemical Physiology, The Scripps Research Institute, La Jolla, California 92037, USA
Ribonuclease P (RNase P) is an essential endoribonuclease that catalyzes the cleavage of the 59 leader of pre-tRNAs. In addition,
a growing number of non-tRNA substrates have been identified in various organisms. RNase P varies in composition, as bacterial
RNase P contains a catalytic RNA core and one protein subunit, while eukaryotic nuclear RNase P retains the catalytic RNA but
has at least nine protein subunits. The additional eukaryotic protein subunits most likely provide additional functionality to
RNase P, with one possibility being additional RNA recognition capabilities. To investigate the possible range of additional
RNase P substrates in vivo, a strand-specific, high-density microarray was used to analyze what RNA accumulates with
a mutation in the catalytic RNA subunit of nuclear RNase P in Saccharomyces cerevisiae. A wide variety of noncoding RNAs
were shown to accumulate, suggesting that nuclear RNase P participates in the turnover of normally unstable nuclear RNAs. In
some cases, the accumulated noncoding RNAs were shown to be antisense to transcripts that commensurately decreased in
abundance. Pre-mRNAs containing introns also accumulated broadly, consistent with either compromised splicing or failure to
efficiently turn over pre-mRNAs that do not enter the splicing pathway. Taken together with the high complexity of the nuclear
RNase P holoenzyme and its relatively nonspecific capacity to bind and cleave mixed sequence RNAs, these data suggest that
nuclear RNase P facilitates turnover of nuclear RNAs in addition to its role in pre-tRNA biogenesis.
Keywords: Ribonuclease P; lncRNA; RNA turnover
Nuclear ribonuclease P (RNase P) in Saccharomyces cer-
evisiae plays an essential role in maturing the 59 end of pre-
tRNA via an endonucleolytic cleavage (for review, see Frank
and Pace 1998; Walker and Engelke 2006). In bacteria the
holoenzyme contains a large, catalytic RNA subunit and
a small protein subunit that stabilizes the folding of the
catalytic RNA and helps to bind pre-tRNA substrate leader
sequences (for review, see Smith et al. 2007; Koutmou et al.
2009). In addition, a recent crystal structure of the bacterial
holoenzyme complex with mature tRNA reinforced earlier
biochemical results by showing that the protein subunit
acts to primarily distinguish between tRNA product and
pre-tRNA substrate via interaction with the 59 leader
(Reiter et al. 2010). The simple bacterial enzyme primarily
recognizes substrate structure, and several additional non-
tRNA substrates have been previously identified (Bothwell
et al. 1976; Peck-Miller and Altman 1991; Giege ´ et al. 1993;
Alifano et al. 1994; Komine et al. 1994; Liu and Altman
1994; Hartmann et al. 1995; Jung and Lee 1995; Gimple and
Scho ¨n 2001; Hansen et al. 2001; Li and Altman 2003;
Altman et al. 2005; Wilusz et al. 2008). The archaeal and
eukaryotic nuclear enzymes are far more complex, and
although they retain the catalytic RNA subunit, they have
multiple proteins with largely unknown functions (for review,
see Walker and Engelke 2006). The yeast nuclear enzyme has
nine essential protein subunits in addition to the RNA sub-
unit, Rpr1r (Lee et al. 1991; Chamberlain et al. 1998). Like its
bacterial counterpart, in addition to pre-tRNA, other possible
substrates have been identified in eukaryotes (Chamberlain
et al. 1996; Yang and Altman 2007; Coughlin et al. 2008;
Wilusz et al. 2008). In addition, purified nuclear RNase P’s
affinity for single-stranded RNA and its ability to cleave at
multiple sites in vitro without obvious sequence specificity
(see Marvin et al. 2011) indicate a broad capacity to serve as
a relatively nonspecific RNA endonuclease.
In an earlier study showing RNase P bound to many RNAs
in vivo, we used double-stranded probes to entire open
reading frames (ORFs) that did not discriminate between
Article published online ahead of print. Article and publication date are
RNA (2011), 17:1441–1450. Published by Cold Spring Harbor Laboratory Press.
sense and antisense transcripts (Coughlin et al. 2008). To
obtain a more nuanced view of what transcripts are sensitive
to a well-characterized mutation in the catalytic RNA subunit
of nuclear RNase P, we have examined the cellular RNA
population at high resolution in a strand-specific, high-
density microarray. The results show surprisingly broad
RNA accumulations, primarily of normally unstable non-
coding RNAs and short-lived pre-mRNAs containing in-
trons. The results are discussed in the context of a possible
endonucleolytic role for nuclear RNase P in the turnover of
RNA that does not enter defined pathways for ribonuclear
protein (RNP) assembly or mRNA utilization.
High-density, strand-specific identification
of RNAs that accumulate in an RNase P
We have previously identified several RNase P temperature-
sensitive (ts) mutations that result in the accumulation of
pre-tRNA (Paga ´n-Ramos et al. 1996; Xiao et al. 2005, 2006).
One of the ts mutations used in this study of Rpr1r, the
catalytic RNA subunit of RNase P, is implicated in magne-
sium coordination and primarily affects the kcatfor pre-
tRNA substrates (Paga ´n-Ramos et al. 1996). Although the
mutation is lethal at 37°C, there are also significant defects
under conditions that allow growth. As seen in Supplemental
Figure S1, pre-tRNALeuaccumulates strongly at both 37°C
and 30°C in the ts strain relative to an otherwise isogenic
wild-type (wt) strain, confirming previous observations of
severely defective RNase P activity.
Using high-density, strand-specific microarrays, we deter-
mined how RNA levels change with the RNase P mutation
relative to wild type at 30°C and 37°C. Table 1 shows a
summary of the most highly enriched RNAs organized by type
of encoded gene product, taken from the larger list provided
with numerical values in Supplemental Table S1. Two classes
of RNA that were the most affected by the RNase P mutation
were a large variety of noncoding RNAs from across the
genome and ribosomal protein mRNA containing introns,
both large and small subunits. The great majority of the most
TABLE 1. Top nuclear-encoded RNAs that enrich with RNase P mutation
Ribosomal small subunit mRNAs
RPS10A (3.64), RPS29A (2.93), RPS10B (2.73), RPS25A (2.63), RPS18B (2.42), RPS4A
(2.40), RPS30A (1.96), RPS6B (1.92), RPS19A (1.91), RPS26B (1.73), RPS19B (1.70),
RPS21B (1.67), RPS16A (1.67), RPS23B (1.54), RPS24B (1.42), RPS30B (1.35),
RPS8A (1.35), RPS29B (1.32), RPS24A (1.22), RPS26A (1.18), RPS27B (1.17)
RPL27B (3.84), RPL39 (3.77), RPL37A (3.73), RPL34B (3.72), RPL26B (3.46),
RPL13B (3.21), RPL34A (3.01), RPL37B (2.97), RPL19B (2.82), RPL31A (2.58),
RPL23B (2.35), RPL36A (2.17), RPL14A (2.09), RPL40B (2.07), RPL27A (2.04),
RPL40A (2.02), RPL43A (1.90), RPL29 (1.86), RPL33A (1.80), RPL33B (1.77),
RPL35B (1.73), RPL21A (1.56), RPL7A (1.52), RPL26A (1.46), RPL6A (1.26),
CUT324 (2.50), CUT526 (2.43), CUT680 (2.22), CUT843 (2.07), CUT008 (1.82),
CUT073 (1.66), CUT128 (1.62), CUT846 (1.58), CUT149 (1.58), CUT732 (1.53),
CUT523 (1.52), CUT347 (1.51), CUT009 (1.49), CUT249 (1.47), CUT461 (1.39),
CUT734 (1.39), CUT168 (1.36), CUT055 (1.35), CUT595 (1.32), CUT190 (1.31),
CUT894 (1.30), CUT791 (1.29), CUT085 (1.28), CUT046 (1.27), CUT572 (1.25),
CUT376 (1.24), CUT689 (1.22), CUT456 (1.22), CUT125 (1.22), CUT325 (1.21),
CUT432 (1.21), CUT440 (1.20), CUT030 (1.20), CUT676 (1.18), CUT837 (1.17),
SUT582 (2.03), SUT677 (1.96), SUT741 (1.96), SUT116 (1.81), SUT625 (1.76),
SUT139 (1.68), SUT699 (1.67), SUT248 (1.65), SUT631 (1.62), SUT279 (1.61),
SUT205 (1.61), SUT517 (1.61), SUT249 (1.58), SUT074 (1.57), SUT542 (1.57),
SUT343 (1.56), SUT287 (1.55), SUT771 (1.53), SUT101 (1.49), SUT045 (1.46),
SUT442 (1.43), SUT129 (1.42), SUT008 (1.42), SUT346 (1.42), SUT814 (1.39),
SUT035 (1.39), SUT827 (1.37), SUT200 (1.36), SUT636 (1.35), SUT313 (1.34),
SUT700 (1.34), SUT553 (1.31), SUT844 (1.31), SUT535 (1.30), SUT756 (1.29),
SUT617 (1.29), SUT278 (1.28), SUT691 (1.26), SUT519 (1.25), SUT808 (1.25),
SUT347 (1.25), SUT001 (1.20), SUT411 (1.20), SUT404 (1.19), SUT114 (1.19)
YNR073C (2.01), YOR053W (1.83), YIL127C (1.58), COS12 (1.46), YGR121W-A (1.45),
YGR169C-A (1.43), YJL144W (1.39), YNL162W-A (1.26), YOL014W (1.21),
AIF1 (2.01), SOM1 (1.79), ATG8 (1.66), SPG4 (1.66), YCL058W-A (1.48), QCR9 (1.40),
MAG1 (1.31), SNR9 (1.30), JID1 (1.17)
Ribosomal large subunit mRNAs
Cryptic unstable transcripts (CUTs) (36/925)
Stable unannotated transcripts (SUTs) (45/847)
Core Sm transcripts
Other transcripts containing introns
Uncharacterized or unknown function
Accumulation values are indicated in parenthesis for 37°C only with an arbitrary cutoff of 1.17. Full listing of the most enriched RNAs can be
found in the Supplemental Data. Underlined RNAs have one or more intron(s).
Marvin et al.
RNA, Vol. 17, No. 8
accumulated mRNAs were from intron-containing genes,
although a smaller number of other mRNA and uncharac-
terized ORF transcripts are also represented in Table 1.
CUT and SUT noncoding RNA were identified in recent
studies that investigated the prevalence of transcription in
the yeast genome, in which as much as 85% of the genome
was found to be transcribed (David et al. 2006; Neil et al.
2009; Xu et al. 2009; for review, see Jacquier 2009). These
two classes of RNA were differentiated by relative stability,
with CUTs, which are normally rapidly degraded, stabilized
enough for detection by a deletion mutant of an exonuclease
component of the nuclear exosome, RRP6 (for review, see
Vanacova and Stefl 2007). Both of these noncoding RNA
classes are transcribed by RNA polymerase II, with many of
them transcribed in the opposite direction of associated
protein-coding gene promoters (Neil et al. 2009; Xu et al.
2009). CUTs are typically 200–600 nt, 59-capped, and have
heterologous 39 ends (for review, see Jacquier 2009). SUTs
are usually longer then CUTs with an average length of 761
nt (Xu et al. 2009). The two classes are not rigidly dif-
ferentiated since independent methods were used to identify
them in multiple studies (Neil et al. 2009; Xu et al. 2009).
The accumulation of CUT and SUT noncoding RNA
shown in Table 1 is also presented visually with expression
maps shown in Figure 1. Figure 1A–D shows a representa-
tive sample of regions corresponding to CUTs and SUTs
that are enriched in the temperature-sensitive mutant at the
Accumulated protein-coding mRNAs were largely in-
tron-containing as represented by the underlined RNAs in
Table 1 and illustrated in Figure 2. This is a highly specific
group of pre-mRNAs, since in S. cerevisiae introns are only
present in z5% of genes (Spingola et al. 1999; Juneau et al.
2006; Roy and Gilbert 2006). Of this small percentage of
genes that have introns, ribosomal protein genes dominate
with 71% containing one or more introns (Planta and
Mager 1998). Table 1 indicates that the majority of most
accumulated intron-containing mRNAs encode for ribo-
somal proteins, with one exception (DYN2). Importantly,
all of the top enriched mRNA that encodes for ribosomal
protein contained introns (Supplemental Table S1). Thus,
the accumulation of a high number of mRNAs coding for
ribosomal protein appears to correlate with the presence of
introns and not simply being a ribosomal protein mRNA.
To determine if other introns were accumulating that we
might not have observed in Table 1, because the intron is
only a small part of the overall gene signal, we used visual
inspection of the transcript map of the entire genome
identify the changes in mRNA intron abundance compared
to corresponding exon signals (for examples, see Fig. 2). It
was clear that the introns of ribosomal protein pre-mRNAs
FIGURE 1. Noncoding RNA accumulates in an RNase P mutant strain. RNA expression data are shown along various positions of the indicated
chromosomes for the Watson (W, top) and the Crick (C, bottom) strands. Abundance data across the whole genome are found in a searchable
online database (see Materials and Methods). Normalized signal intensities are shown for indicated samples. Biological replicates for either wild-
type (wt) or RNase P mutant (ts) strains were grown at either 30°C or 37°C in synthetic media. (Vertical red lines) Inferred positions of
transcription boundaries. Genome annotations are shown in the center with annotated open reading frames (ORFs) (blue) and untranslated
regions (UTRs) (gray lines). (Orange boxes) Previously annotated (Xu et al. 2009) SUTs; (purple boxes) CUTs; (light blue box) an
uncharacterized ORF; (arrows) direction of transcription. Coordinates are indicated in base pairs in the center. (A–D) Examples of SUTs (A–C)
and CUTs (B,D) accumulating in the RNase P ts strain at either 30°C or 37°C. (B,C) Reciprocal examples in which ORF RNAs decline in abundance
with the RNase P mutation, while antisense RNAs accumulate. As shown in A, sometimes the inferred transcription boundaries (Xu et al. 2009) do
not precisely align with the apparent expression in these experiments. This could reflect larger RNAs being present in the RNase P mutants.
Expanded cellular roles for RNase P
and other intron-containing pre-mRNAs were selectively
enriched. A typical ribosomal protein mRNA that does not
contain an intron is shown in Figure 2D for comparison.
Intron-containing messages where the intron accumulates
are listed in Table 2. These include many that are not rep-
resented in Table 1, since the majority of the signal strength
that defines accumulation of genes in Table 1 derives from
the much stronger exon signals.
Sense/antisense RNA balance affected by defective
A number of noncoding RNAs, 36 out of 925 total anno-
tated CUTs and 45 out of 847 total annotated SUTs, were
among the most enriched transcripts in the RNase P mu-
tant strain. Upon inspection of the transcript map results
interesting changes were evident (Fig. 1A–D). In addition to
cases of CUT and SUT RNA accumulation, sometimes the
sense and antisense changes were inverse in the RNase P
mutant (Fig. 1B,C; Supplemental Fig. S2A). In addition,
some cases were observed where an unannotated RNA
accumulated while an overlapping ORF showed de-enrich-
ment (Supplemental Fig. S2B). For two cases, enrichment
of SUT RNAs that are antisense to de-enriched ORF RNAs
was confirmed by Northern blots (Fig. 3A,B). These results
also indicated that the SUT RNA was accumulating in
multiple larger forms, as large as 6000 nt, rather than only
the smaller forms observed previously (Fig. 3). The anno-
tations of the boundaries of CUTs and SUTs were mapped
differentially using, in the case of CUTs, an automated
segmentation algorithm in an Rrp6 deletion strain, or in
the case of SUTs, automated segmentation followed by
manual curation in wild-type cells (Xu et al. 2009). The
appearance of these various sizes coincides with a reciprocal
loss of overlapping mRNA signal at these loci. The exact
nature of these larger SUT RNAs, whether they are pre-
cursors of the smaller RNAs, and how they might be related
to loss of transcripts from the opposing strand remains to be
investigated. These multiple sizes, added to the fact that the
RNase P mutant accumulates overlapping but nonidentical
noncoding RNAs, suggest that the RNase P mutant is not
affecting precisely the same turnover events as seen with the
Rrp6 deletion mutant in previous work (Xu et al. 2009).
Confirmation of intron-containing
We used Northern blots and semi-quantitative RT-PCR
analyses of total RNA to confirm microarray results for
FIGURE 2. Introns accumulate in an RNase P mutant strain. Examples are shown of intron accumulation, with representations as in Figure 1.
(Gray lines) Introns and untranslated 59 or 39 segments of transcripts outside the ORFs are shown connecting ORF exons (blue boxes). (Blue
arrows) Transcript starts; (black arched lines) known splicing events. Panel A does not show a blue box for the 59 exon because the intron is part
of the 59 UTR. (A–C) Examples of introns accumulating in mRNA-coding regions with RNase P mutation, either ribosomal protein mRNA (A,B)
or nonribosomal protein mRNA (C), compared to a ribosomal mRNA not containing an intron (D). An overlapping CUT in C does not show
significant change in abundance at this locus.
Marvin et al.
RNA, Vol. 17, No. 8
selected mRNA intron regions and to determine the nature
of the RNAs that were accumulating in RNase P mutant
samples. Figure 4 shows Northern blots of RNA from both
wt and ts strains that were grown for 2 h at 30°C or 37°C.
Figure 4 shows total RNA probed for ribosomal protein
mRNA and nonribosomal protein mRNA that contain
introns with probes that annealed to the coding sequences
(CDS). Accumulation of more slowly migrating bands than
the mRNAs was observed for all intron-containing mRNAs
tested, having sizes consistent with failure to remove the
intron. Since these Northern blots were carried out with
probes complementary to 39 exons, RT-PCR from the introns
to the 59 exons or 59 UTR was used to confirm the accu-
mulation of unspliced pre-mRNAs rather than intermediates
lacking 59 exons (data not shown). The accumulation in pre-
mRNAs is accompanied by a slight but consistent drop in
abundance of the mature-sized mRNAs relative to an Scr1
small cytoplasmic RNA loading control. Thus, it is possible
that the pre-mRNA accumulates either because of a mild
defect in splicing efficiency or a failure to efficiently destroy
pre-mRNAs that have not entered the splicing pathway
(Bousquet-Antonelli et al. 2000; Sayani et al. 2008).
Shortening of RNAs
An additional effect of the RNase P mutation was discov-
ered when the sizes of small RNAs were examined by
Northern blot. In probing the U6 snRNA signal as a loading
control on high-resolution denaturing gels, we noticed that
there was a small size difference in U6 in the ts strain
relative to the wt strain. This analysis was subsequently
carried out repeatedly for all of the nuclear spliceosome
snRNAs, as well as two RNAs found primarily in the cy-
toplasm, the Scr1 RNA (signal recognition particle RNA)
and the 5S ribosomal RNA. There was a 2–4-nt shortening
of the RNAs observed for RNAs that were small enough for
this difference to be resolved (Fig. 5A). Shortening of the
small RNAs is consistent with 39–59 exonuclease trimming
until a substantial block is reached due to RNA or RNP
structure (Brow 2002), and we used primer extension to
confirm that the 59 ends of U4 and U6 RNAs were
unaffected (Fig. 5B). Levels of the small RNAs do not
change significantly so this slight trimming does not result
in the complete degradation of these RNAs (Fig. 5C).
It is not obvious why an RNase P defect would affect all
of these RNAs. They are primarily found in different
cellular compartments, involved in different processes,
bound by different proteins, and synthesized by different
RNA polymerases (RNA polymerase III for U6, 5S, and
Scr1; RNA polymerase II for the other RNAs). The only
clear commonality is that they are made in the nucleus,
suggesting that the shortening might occur there. It is
interesting to note that the one short RNA tested that
appears unaffected is tRNA, along with its precursors
(Supplemental Fig. S1). A reason could be that pre-tRNAs
and tRNAs are routinely exposed to 39 exonuclease attack
in wild-type cells, but are rebuilt by CCA addition and
aminoacylation at the 39 termini, which might repair any
damage and seal them against further attack.
In this study, we identified a broad and diverse set of
noncoding RNAs that accumulate with a mutation in the
TABLE 2. Top mRNA introns affected by RNase P mutation
Enriched RPL27B, RPL39, RPL37A, RPL34B, RPS10A, RPL26B, RPL13B, RPL34A, RPL37B, RPS29A, RPL19B, RPS10B,
RPS25A, RPL31A, RPS18B, RPS4A, RPL23B, RPL36A, RPL14A, RPL40B, RPL27A, RPL40A, RPS30A, RPS6B,
RPS19A, RPL43A, RPL29, RPL33A, RPL33B, RPL35B, RPS26B, RPS19B, RPS21B, RPS16A, RPL21A, RPS23B,
RPL7A, DYN2, RPL26A, RPS24B, RPS30B, RPS8A, RPS29B, RPL6A, RPL14B, RPS24A, RPS26A, RPS27B,
YNL050C, RPL24A, RPS16B, RPS11B, RPL42A, RPS18A, RPL23A, RPS17B, RPS17A, RPL42B, RPS6A, RPS8B,
RPL35A, RPL24B, RPS25B, YDL012C, RPL21B, RPL31B, RPL43B, NHP6B, RPS4B, RPL20B, RPL19A, YBL059W,
MUD1, RPL2B, RPS27A, LSM7, RAD14, IWR1, APS3, PCC1, MOB2, RPS9A, RUB1, YJL041W, MRPL44, VMA10,
UBC13, BET1, RPS7B, YSC84, RPL32, KEI1, RPS9B, LSM2, PRE3, RPS7A, TMA20, TAD3, RPL25, RPL17A, RPS0A,
RPL17B, COX4, RPS21A, VPS29, RPL30, RPL6B, YOP1, SEC14, OST5, RPL16A, RPS14A, RPL16B, STO1, ERD2,
RPP1B, RPS0B, COF1, VPS75, RPL13A, RPL2A, RPL36B, RPS23A
MND1, HOP2, PCH2, SAE3, REC107, MTR2, RPL18A, MATa1, NCE101, MMS2, SNC1, AIM11, SFT1, SMD2,
YDR381C-A, HFM1, RPL7B, NOG2, TAF14, SPO1, HNT2, DCN1, NYV1, CNB1, UBC5, MEI4, UBC8, DMC1,
YLR211C, MAF1, OM14, LSB3, RPO26, POP8, RIM1, BIG1, RPS14B, RRT8, AMA1, DID4, COX5B, MRK1, CMC2,
UBC9, YPR063C, MOB1, UBC12, GCR1, IST1, RPL22B, DTD1, SCS22, ERV41, KIN28, SEC17, TDA5, RPL18B,
ERV1, HNT1, MPT5, GLC7, BOS1, YRA1, RPS13, QCR10, ARP2, SPT14, ECM33, TEF4, APE2, ASC1, SAR1, SAC6,
CIN2, YBR220C, SRB2, PMI40, TUB1, DBP2, IMD4, CPT1, YIP3, YPR098C, EPT1, RPS22B
YOS1, GIM5, RPL28, RPS11A, ARP9, YHR097C, TAN1, YML6, UBC4, PHO85, NCB2, TUB3, QCR9, EFB1, AML1,
NMD2, RBS1, GOT1, ACT1, TFC3, RFA2, PFY1, RPL22A, ARF2, SEC27
YOL047C, REC114, YBR225C-A, SRC1
Relative intron enrichment with RNase P mutation at 37°C is indicated via visual inspection of transcript map data. Quantitative analysis of
microarray data, which is a composite of probes in exons and introns for each RNA, can be found in the Supplemental Data. Information on
intron content is drawn from the Ares Lab Yeast Intron Database: http://metarray.ucsc.edu/yeast_intron_db/.
Expanded cellular roles for RNase P
catalytic subunit of RNase P. There is increasing evidence
that long, noncoding RNAs (lncRNAs) serve regulatory
functions in eukaryotes (Mercer et al. 2009; Ponting et al.
2009; Lee 2010; Ponting and Belgard 2010; van Leeuwen
and Mikkers 2010), although the mechanisms by which the
lncRNA transcripts are turned over are only beginning to
be elucidated. Originally, the widespread occurrence of
CUTs and SUTs in S. cerevisiae was difficult to interpret
in light of the lack of a small interfering RNA (siRNA)
pathway for RNA or the equivalent of the Schizosacchar-
omyces pombe chromatin modification pathway, although
recent observations suggest that alternative chromatin
modification pathways might be engaged (Camblong
et al. 2007; Berretta et al. 2008; Houseley et al. 2008).
Confirming that these RNAs are RNase P substrates in
vivo is difficult given that in vitro studies (Chamberlain
et al. 1996; Coughlin et al. 2008; Marvin et al. 2011) have
shown that nuclear RNase P can bind and cleave single-
stranded RNA at multiple positions without clear sequence
or structural requirements. Our in vitro study of cleavage
preferences deliberately chose both sense and antisense
RNAs across a region where the balance was affected by
RNase P mutations, showing that RNase P could bind all
segments across the region and cleave RNAs from both
strands at multiple places (Camblong et al. 2007; Coughlin
et al. 2008; Marvin et al. 2011). Thus, we are confident that
RNase P is potentially capable of cleaving all of the RNAs
that accumulate in the ts mutant strain, if it is allowed
access to the deproteinized RNA. Although it is formally
possible that the RNAs are accumulating due to some
indirect effect of the RNase P mutation elevating pro-
miscuous transcription from many sites in the genome, the
most likely hypothesis is loss of normal participation by
RNase P in RNA turnover.
The question of whether RNase P normally does cut
these unstable RNAs in vivo to speed their turnover, or
whether their accumulation in the RNase P mutant is more
of an indirect effect, is difficult to approach. We explored
the possibility that RNase P might be part of a complex that
includes other enzymes known to participate in nuclear
RNA turnover (e.g., exosome or TRAMP components) (for
review, see Houseley and Tollervey 2009). Previous work
has shown that the exosome uses both exonuclease and
endonuclease activity for degradation of highly structured
RNAs (Lebreton et al. 2008), and one could envision the
RNase P active site having been adapted to serve this
function. However, rapid, low-stringency affinity isolation
of RNase P complexes and sensitive proteomic analysis
showed a large number of co-isolating proteins, but these
did not include significant amounts of any obvious protein
partners that are known to participate in RNA turnover
(Supplemental Table S2). Thus, if RNase P is acting in
concert with other nucleolytic activities, it does not appear
to result in stable complexes with a high percentage of the
In previous studies, the removal of the nuclear component
of the exosome, Rrp6, resulted in a significant accumulation
of a large number of ncRNAs (Jacquier 2009; Xu et al. 2009;
Lardenois et al. 2011). The accumulation of noncoding
RNAs that we observe with our RNase P mutant is not as
widespread as in this earlier work, and contains RNAs from
some regions not previously annotated. In addition, RNAs
from some loci in the RNase P mutant strain are clearly
larger and more heterogeneous than previously annotated.
This lack of congruity between the RNase P mutant and the
RRP6 deletion accumulations suggests that RNase P might
intersect the function of the exosome, but operate indepen-
dently in turning over lncRNA. Given that RNase P can
recognize both highly structured RNA and preferred sites in
less structured sequences, the subset of CUTs/SUTs that
accumulate could have nonobvious features or protein
partners in vivo that mark them for cleavage. Thus, our
de-enrichment with an RNase P mutation. Total RNA isolated from
replicate cultures of wild-type (wt) or RNase P mutant (ts) samples
grown at either 30°C or 37°C are shown separated on 1.4% de-
naturing agarose gels with subsequent Northern blot analysis. Sizes (in
nucleotides, nt) were estimated from known markers (Materials and
Methods), and W and C strands are as in Figure 1. (A) Northern blots
were probed for Sut428 or Opt2 RNAs, reprobing for Scr1 RNA as
a loading control. (B) Northern blots were probed for Sut116 or
Hnm1 and reprobed for Scr1 as a loading control. In both cases
shown, the SUT loci accumulated RNA that differs from previous
annotations, which were used for this data set (Xu et al. 2009).
Numbers are shown indicating different RNA species. Importantly,
the RNA species most consistent with established annotation is
indicated at the transcription site on the loci diagram below the
Northern blots: Sut428 (1) and Sut116 (2). Also, the ‘‘sense’’ mRNA is
shown significantly de-enriched in both panels (see also Fig. 1).
Antisense RNA accumulation and overlapping mRNA
Marvin et al.
RNA, Vol. 17, No. 8
results are consistent with RNase P possibly playing a role in
the degradation of a select subgroup of CUT and SUT RNAs,
although further work is needed to determine how RNase P
carries out this role.
The other major type of RNA that was shown to accu-
mulate with the RNase P mutation was intron-containing
pre-mRNA. These pre-mRNA species are similar to the
CUTs/SUTs only in that both species are relatively transient
under wild-type conditions. The observed accumulation of
pre-mRNA introns was not completely unprecedented, as a
previous study showed accumulation of precursors to in-
tron-encoded box C/D snoRNAs, consistent with an RNase
P–mediated pathway for excising the intronic snoRNAs that
does not proceed through a spliced intermediate (Villa et al.
2000; Coughlin et al. 2008). The much broader accumulation
of intron-containing mRNAs that we were able to detect
using higher-density microarrays could have resulted from
one of at least two effects. First, the RNase P mutation could
be indirectly and mildly compromising the splicing appara-
tus (for review, see Wahl et al. 2009), which would be con-
sistent with the shortening of the snRNA components of the
spliceosome, but would not explain shortening of the other
RNAs or the accumulation of lncRNAs. Alternatively, a small
population of unspliced pre-mRNAs might normally fail to
enter the splicing pathway and require nuclear turnover
involving RNase P. The observed low-level association of
RNase P with a wide variety of transcripts from spliced genes
(Coughlin et al. 2008) and the concomitant accumulation of
many CUTs and SUTs would be consistent with the second
There is not a clear explanation for how RNase P muta-
tions might indirectly cause most of the effects we see here.
The best understood effect of this RNase P mutation is the
accumulation of pre-tRNA, long known to be the evolu-
tionarily conserved RNase P substrate from bacteria to
vertebrates. The accumulation of pre-tRNAs is one possible
indirect mechanism for the accumula-
tion of other RNAs and/or the slight
shortening of RNAs that we observe in
our study. Accumulated pre-tRNAs might
compete for RNA-binding proteins that
would otherwise protect many small
RNAs in the nucleus and possibly con-
tribute to rapid turnover of CUTs,
SUTs, and ‘‘escaped’’ intron-containing
pre-mRNAs. One obvious candidate for
such a protein was the ubiquitous La
protein, Lhp1 in S. cerevisiae, which
binds to the 39 ends of RNA polymerase
III transcripts and other poly(U) se-
quences (Pannone et al. 1998, 2001;
Mayes et al. 1999). Although overex-
pression of Lhp1 did not suppress the
observed shortening of small RNAs
(data not shown), it remains possible
that it or another RNA-binding complex (e.g., Lsm) is
competed for by the accumulated pre-tRNAs (Mayes et al.
1999; Beggs 2005). An alternative interpretation might be
that the accumulation of RNAs that are normally turned
over with the help of RNase P (CUTs, SUTs, and pre-
mRNAs) frees up exonuclease capacity or competes for
RNA-binding proteins, either of these resulting in the
increased exonuclease attack on normally resistant RNAs.
RNase P has evolved to recognize the structure of pre-
tRNA and cleave at a precise position relative to the tertiary
structure (Frank and Pace 1998; Walker and Engelke 2006).
However, it appears that nuclear RNase P has gained
additional RNA-binding and cleavage capability due to
increased protein content of the RNA core (Marvin et al.
2011). The simplest hypothesis for why noncoding RNAs,
and possibly unspliced pre-mRNAs, accumulate is that they
are normally cleaved by RNase P for rapid turnover, but
this raises an interesting question of how they become
exposed to the enzyme. Localization of RNase P RNA by
fluorescent in situ hybridization shows that the enzyme is
present primarily in the nucleolus in S. cerevisiae (Bertrand
et al. 1998), where the majority of the tRNA genes and pre-
tRNAs are also found (Thompson 2003). It is possible that
smaller quantities of active enzyme are also more dispersed
in the nucleoplasm, or that the noncoding RNAs must
traffic to the nucleolus to encounter RNase P. The related
complex that processes pre-rRNAs, RNase MRP, is also
primarily in the nucleolus, but also has been detected in the
cytoplasm, where it appears to be involved in turnover of
a set of mRNAs that are entirely distinct from the ones that
accumulate in our hands with the RNase P mutant (Gill
et al. 2006). No RNase P has ever been detected by our
group in the cytoplasm through fluorescent tags on either
the RNA subunit or the unique protein subunit, Rpr2p, but
we are unable to definitively exclude lesser, non-nucleolar
pools (Bertrand et al. 1998).
FIGURE 4. Confirmation of pre-mRNA accumulation in an RNase P mutant strain. Total RNA
was separated using 1.4% denaturing agarose gels and subjected to Northern blot analysis.
Indicated mRNAs were probed with oligos specific to 39-exon regions. Total RNA is shown for
wild-type (wt) and RNase P mutant (ts) strains grown at either 30°C or 37°C. RNA size markers
are indicated next to each blot (Materials and Methods). The predicted positions of pre-mRNA and
mature mRNA are shown next to the bands on the blots. For Rps29A/B the pre-mRNA cartoon
indicates the 59 UTR intron. Mature mRNA levels are not significantly enriched in the RNase P
mutant strain when normalized to internal Scr1 RNA control levels (values range from 0.6-fold to
1.0-fold), but new bands appear at positions predicted to be the indicated pre-mRNA. The presence
of 59 exons, as well as introns and 39 exons, was confirmed by RT-PCR (data not shown).
Expanded cellular roles for RNase P
It was recently shown that RNase MRP has a limited
sequence preference for a C at position +4 relative to the site
of cleavage (Esakova et al. 2011). This preference does not
seem to occur with RNase P, which appears to recognize
RNA with little regard to primary sequence (Marvin et al.
2011). Why might RNase P and RNase MRP recognize
different types of RNA substrate? Although the two enzymes
share eight of the same protein subunits, there are one or
two proteins that are distinctive to each enzyme and the
RNAs are different, although evolutionarily related (for
review, see Walker and Engelke 2006). It is certainly possible
that the RNA subunit differences could play a role in
contributing to the observed differences in RNA recognition
between the two complexes. However, the structure of the
bacterial enzyme shows that, as expected, the RNA subunit
recognizes primarily the structured portion of pre-tRNAs,
while single-stranded extensions are in contact with the
single small protein (Reiter et al. 2010). Consistent with this,
the in vitro activity of the nuclear RNase P suggests that two
or more of the nine protein subunits are involved in binding
non-tRNA substrates, but not necessarily pre-tRNA sub-
strates. Interestingly, one of these proteins is Rpr2p, the sole
protein that is found in RNase P, but not
RNase MRP (see Marvin et al. 2011).
Thus, while multiple proteins and the
RNA subunit in the holoenzyme might
be involved in contacting unstructured
RNA substrates, Rpr2p is a prime can-
didate for providing contacts by which
RNase P recognizes features in non-
MATERIALS AND METHODS
S. cerevisiae strain JLY1 (MATa ade2-1 his3-
11,15 leu2-3,112 trp1-1 ura3-1 can1-100
RPR1THIS3), with a background of W3031A,
was the parent strain with RPR1 on a LEU2-
marked plasmid. We investigated the effect of
a temperature-sensitive mutation in Rpr1r at
position G207G211(Paga ´n-Ramos et al. 1996).
A secondary, neutral sequence alteration was
present in the P3 region of RPR1 (position
69–75: AUCAGAU to CAGGACG) as a unique
hybridization marker in both the wild-type
and the ts RPR1 RNA sequences.
The synthetic dropout media (SD-His) was
used for growth of JLY1 strains. For analysis
of RPR1 mutant effects (G207G211), previous
studies have identified the optimal growth
conditions to observe RNase P defects
(Coughlin et al. 2008). Both wild-type and
temperature-sensitive yeast were grown into mid-log phase then
diluted and shifted to 37°C in pre-warmed SD-His media for 2 h.
Hot acid phenol was used to isolate total RNA from yeast grown
at either 30°C or 37°C (Ko ¨hrer and Domdey 1991). RNA was
treated with Turbo DNase I per the manufacturer’s protocol
(Ambion). DNase was then inactivated using inactivation reagent
(Ambion). UV absorbance at 260/280 nm was used to measure
concentration of samples.
For first-strand cDNA synthesis, 20 mg of total RNA was mixed
with 1.72 mg of random hexamers and 0.034 mg of oligo(dT)
primer and incubated for 10 min at 70°C followed by 10 min at
25°C, then transferred to ice. The synthesis included 2000 units of
SuperScript II Reverse Transcriptase, 50 mM Tris-HCl, 75 mM
KCl, 3 mM MgCl2, 0.01 M DTT, dNTP + dUTP mix (0.5 mM for
dCTP, dATP, and dGTP; 0.4 mM for dTTP; and 0.1 mM for
dUTP; [Invitrogen]), and 20 mg/mL actinomycin D in a total
volume of 105 mL. Actinomycin D was added during reverse
transcription (Perocchi et al. 2007). The reaction was carried out
sequentially for 10 min at 25°C, 30 min at 37°C, 30 min at 42°C,
and 10 min at 70°C for heat inactivation. Samples were then
subjected to RNase treatment of 20 min at 37°C (30 units of
FIGURE 5. Small RNAs in RNase P mutant strain. (A) Total RNA was separated on either 10%
denaturing polyacrylamide gels (U1, U5, U4, U6, and 5S for loading control) or 1.4% denaturing
agarose gels (U2 and Scr1) and then subjected to Northern blot analysis. RNA was probed from
biological replicates of either wild-type (wt) or RNase P temperature-sensitive (ts) samples grown
at either 30°C or 37°C. Fold enrichment (ts/wt) for snRNA relative to loading controls showed
only modest increases in multiple experiments (1.2-fold to 2.3-fold enrichment). However, these
RNAs consistently appear 2–3 nt shorter in the RNase P mutant strain in repeated experiments.
This shortening is inferred to be at the 39 end as shown by primer extension (B). (B) Primer
extension analysis of U4 and U6 RNA is shown at single-nucleotide resolution with total RNA
isolated from wt and ts samples grown at either 30°C or 37°C. The RNA 59 ends of U4 and U6
appear unchanged. (C) Quantitation of U4 and U6 RNA levels relative to 5S RNA loading
control. Fold enrichment (ts/wt) for U4 is 1.4 and U6 is 1.1, which indicates that even though
trimming of these RNAs is occurring, they are still stably expressed. RNA was loaded twice from
biological triplicate samples on the same 8% denaturing polyacrylamide gel, followed by
quantitation of bands after Northern blot with error bars indicating SEM.
Marvin et al.
RNA, Vol. 17, No. 8
RNase H [Epicentre], 60 units of RNase Cocktail [Ambion]).
First-strand cDNA was purified using the MinElute PCR purifi-
cation kit (QIAGEN), and 4 mg was fragmented and labeled using
the GeneChip WT Terminal labeling kit (Affymetrix) according to
the manufacturer’s protocol. The labeled cDNA samples were
denatured in a volume of 300 mL containing 50 pM control
oligonucleotide B2 (Affymetrix) and Hybridization mix (Gene-
Chip Hybridization, Wash and Stain kit; Affymetrix) of which 250
mL was hybridized per array (S. cerevisiae yeast tiling array;
Affymetrix, PN 520055). Hybridizations were carried out for 16 h
at 45°C with 60 rpm rotation. The staining was carried out using
the GeneChip Hybridization, Wash and Stain kit with fluidics
protocol FS450_0001 in an Affymetrix Fluidics station. The cDNA
hybridizations are available at ArrayExpress (http://www.ebi.ac.
uk/arrayexpress/) under the accession number E-MEXP-3140 and
the array design under A-AFFY-116.
Genomic DNA (gDNA) was prepared from the same back-
ground strain (W3031A) that was used for total RNA preparation
above to normalize signal from the RNA (Huber et al. 2006). Only
the probes matching exactly and uniquely to the W303 genome
were considered. Transcript boundaries were taken from the
Supplemental Table 3 of Xu et al. (2009). For each transcript,
expression level was estimated by the midpoint of the ‘‘shorth’’
(shortest interval that covers half the values) of the normalized
probe intensities lying within the transcript. A searchable
database of expression results from the microarray experiment
can be found at http://steinmetzlab.embl.de/engelkeArray/index.
Northern blots of RNA
Northern blots were carried out using both denaturing polyacryl-
amide and denaturing agarose gel systems. Denaturing polyacryl-
amide Northern blots were carried out as performed previously
(Coughlin et al. 2008). Agarose–formaldehyde gels (Sambrook et al.
2001) were used to fractionate total RNA followed by transfer to
Nytran Supercharge membranes using a TurboBlotter apparatus
(Schleicher & Schuell Biosciences). An RNA ladder (Lonza) was
loaded in the same way as total RNA and run for size estimation.
Specific oligonucleotide probes (Supplemental Table S3) were
labeled with [g-32P]ATP using polynucleotide kinase (NEB) per
the manufacturer’s protocol, hybridized to membranes, and sub-
sequently washed per the manufacturer’s protocol for Nytran
SuperCharge membranes (Schleicher & Schuell Biosciences).
Signal was detected using a Typhoon Trio+ imager and quanti-
tated using ImageJ software (http://rsbweb.nih.gov/ij/).
Analysis of the sizes of total cellular poly(A) tracts after
digestion of total RNA with a titration of RNase A followed by
Northern blot analysis using radiolabeled oligo(dT) (15 nt) as
a probe, also suggested that mRNA might be slightly shortened by
approximately five or more adenosine residues (data not shown),
but the inherent length heterogeneity of the poly(A) tails made
this difficult to interpret with confidence.
Total RNA that was isolated from wt and ts yeast strains was
reverse-transcribed using Omniscript reverse transcriptase per the
manufacturer’s protocol (QIAGEN) with radiolabeled U4 and U6
oligos (Supplemental Table S3) to determine if these RNAs are
being trimmed at the 59 ends. After separation on denaturing gels,
samples were exposed to a phosphor screen with visualization
using a Typhoon Trio+ imager.
Supplemental material is available for this article.
We thank Zhenyu Xu for help with microarray analysis. In addition,
we thank Francoise Stutz for discussions concerning antisense Pho84
RNA. This work was supported by grant GM034869 (to D.R.E.), and
an UM Cellular Biotechnology Training Grant T32-GM08353 and
a fellowship from the Horace H. Rackham Graduate School (both to
M.C.M.). Funding was also provided by grants from the NIH and
Deutsche Forschungsgemeinschaft (to L.M.S.) and NIH Grant P41
RR011823 (to J.R.Y.). This is TSRI Manuscript #21257.
Received March 19, 2011; accepted April 28, 2011.
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