A feedback loop regulates splicing of the spinal
muscular atrophy-modifying gene, SMN2
Francine M. Jodelka1, Allison D. Ebert3, Dominik M. Duelli2and Michelle L. Hastings1,∗
1Department of Cell Biology and Anatomy and2Department of Pathology, The Chicago Medical School, Rosalind
Franklin University of Medicine and Science, North Chicago, IL, USA and3Stem Cell and Regenerative Medicine
Center, Department of Neurology, University of Wisconsin, Madison,WI, USA
Received July 14, 2010; Revised September 10, 2010; Accepted September 24, 2010
Spinal muscular atrophy (SMA) is a neurological disorder characterized by motor neuron degeneration and
progressive muscle paralysis. The disease is caused by a reduction in survival of motor neuron (SMN) protein
resulting from homozygous deletion of the SMN1 gene. SMN protein is also encoded by SMN2. However, spli-
cing of SMN2 exon 7 is defective, and consequently, the majority of the transcripts produce a truncated,
unstable protein. SMN protein itself has a role in splicing. The protein is required for the biogenesis of spli-
ceosomal snRNPs, which are essential components of the splicing reaction. We now show that SMN protein
abundance affects the splicing of SMN2 exon 7, revealing a feedback loop inSMN expression. The reduced
SMN protein concentration observed in SMA samples and in cells depleted of SMN correlates with a decrease
in cellular snRNA levels and a decrease in SMN2 exon 7 splicing. Furthermore, altering the relative abun-
dance or activity of individual snRNPs has distinct effects on exon 7 splicing, demonstrating that core
spliceosomal snRNPs influence SMN2 alternative splicing. Our results identify a feedback loop in SMN
expression by which low SMN protein levels exacerbate SMN exon 7 skipping, leading to a further reduction
in SMN protein. These results imply that a modest increase in SMN protein abundance may cause a dispro-
portionately large increase in SMN expression, a finding that is important for assessing the therapeutic
potential of SMA treatments and understanding disease pathogenesis.
Proximal spinal muscular atrophy (SMA) is an autosomal
recessive disorder characterized by progressive muscle weak-
ness and paralysis, resulting from the specific degeneration of
lower motor neurons in the spinal cord. SMA affects approxi-
mately one in 6000 live births and is a leading genetic cause of
infant mortality (1,2).
SMA is caused by homozygous mutation or deletion of the
survival of motor neuron 1 (SMN1) gene that codes for SMN
protein (3). Two genes, SMN1 and SMN2, code for SMN
protein in humans, and the copy number of SMN2 is a deter-
minant of disease severity (4,5). SMN1 and SMN2 are nearly
identical, and both are ubiquitously expressed. However,
SMN2 produces less SMN protein than SMN1 due to a
C-to-T change in exon 7 of SMN2 (6,7) that compromises
exon 7 recognition by the splicing machinery (Fig. 1A). As
a result of this nucleotide difference, the majority of SMN2
mRNA transcripts lack exon 7 and code for truncated SMN
protein that is unstable and rapidly degraded (8). Thus,
SMN2 cannot fully compensate for the loss of SMN1 in
SMA because the amount of full-length mRNA and functional
SMN protein produced from SMN2 is considerably lower than
that from SMN1. For unknown reasons, the reduced abundance
of SMN protein results in the specific degeneration of motor
SMN protein is essential for the biogenesis of spliceosomal
snRNPs U1, U2, U4, U4atac, U5, U11 and U12 (10–12).
SMN, in a complex with other proteins (SMN complex),
assembles the Sm proteins B/B’, D1, D2, D3, E, F and G,
∗To whom correspondence should be addressed at: Department of Cell Biology and Anatomy, The Chicago Medical School, Rosalind Franklin
University of Medicine and Science, 3333 Green Bay Road, North Chicago, IL 60064, USA. Tel: +1 8475788517; Fax: +1 8475783253;
# The Author 2010. Published by Oxford University Press.
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License
(http://creativecommons.org/licenses/by-nc/2.5), which permits unrestricted non-commercial use, distribution, and reproduction
in any medium, provided the original work is properly cited.
Human Molecular Genetics, 2010, Vol. 19, No. 24
Advance Access published on September 30, 2010
onto the snRNA in the first steps of snRNP biogenesis. The
decrease in SMN protein resulting from the loss of SMN1
protein alters the repertoire of snRNAs in the cell and leads
to deficits of fully assembled snRNPs (13–15). In fact, a
decrease in SMN protein has been reported to have an effect
on a number of splicing events (13–15), although it is
unclear whether splicing changes are a cause or consequence
of SMA disease pathology (16). The effect that SMN protein
abundance has on splicing could explain the motor neuron
degeneration in SMA if splicing of a transcript that encodes
a protein with critical motor neuron-specific function is
altered by the decrease in SMN protein (17). Motor neurons
may also have a higher demand for spliceosomal snRNPs,
and thus changes in the abundance of snRNPs may result in
more dramatic changes in splicing in motor neurons than in
other cell types.
One question that has not yet been addressed is whether
splicing of SMN2 exon 7 itself is sensitive to changes in
snRNP levels. Splicing of this exon is under the control of a
number of splicing factors. The C-to-T change in SMN2 that
results in an increase in exon 7 skipping compared with
SMN1 disrupts an exonic splicing enhancer (ESE) motif recog-
nized by the SR protein SF2/ASF (18). The loss of this ESE
weakens exon 7 recognition, making its splicing more sensi-
tive to control by a number of splicing factors. For example,
in the absence of this ESE, inhibitory interactions between
splicing silencer elements and hnRNP A1 predominate and
result in exon skipping (19,20). Additional proteins and
sequence elements have also been identified that can influence
SMN2 exon 7 splicing (21–32). For example, the Tra2 family
of SR-like proteins (33) and the splicing factor hnRNP Q/R
(25) influence exon 7 inclusion. RNA secondary structure is
another determinant of exon 7 splicing (34,35).
One way that cis-acting sequences influence exon 7 splicing
is by binding splicing factors that help to recruit spliceosomal
snRNPs to exon 7 (22). The importance of efficient
Figure 1. Low SMN protein abundance correlates with low SMN2 exon 7 splicing in SMA. (A) SMN1 and SMN2 gene structures. Boxes indicate exons and
horizontal lines are introns. Dominant splicing pattern is shown with solid diagonal lines, and minor alternative splicing is indicated with hatched lines.
Shaded boxes indicate the coding region of the most abundant mRNA. (B) Immunoblot analysis of SMN protein in lysates from tissues of normal
(Smn+/+) or SMA (Smn2/2) hSMN2 transgenic mice. Blotting of b-actin protein was used as a loading control. (C) Semi-quantitative radiolabeled RT–
PCR analysis of human SMN2 mRNA from tissues of human SMN2 transgenic mice that are either homozygous (+/+) or (2/2) for murine Smn. Products
were separated on a 6% native polyacrylamide gel. Transcripts that include or skip exon 7 are indicated, and results are quantitated and shown as % exon 7
inclusion [included/(included + skipped)×100]. (D) Immunoblot analysis of SMN, hnRNP Q/R, Tra2b1 (SFRS10) and b-actin (loading control) in cell
lysates from human-induced pluripotent stem (iPS) cells derived from an SMA patient or an unaffected family member (wt) (42). The SMN/b-actin ratio is
shown beneath the blot. (E) RT–PCR analysis of RNA from iPS cells. Reaction products were incubated with DdeI (D) to digest SMN2 RNA and identities
of each transcript are indicated. The percent of SMN2 transcripts that include exon 7 is shown beneath the gel and reflect the average of three RT–PCR reactions
from independent RNA isolations. (F) Real-time PCR analysis of snRNAs from iPS-WT and iPS-SMA cells. The percent change in snRNA quantity in SMA
samples compared with WT samples was normalized to Rpl39 and graphed as the percentage of WT. Results are the average of three independent tests of the
WT-iPS and SMA-iPS samples, which were repeated in triplicate for each reaction. Error bars represent the standard error of the means (SEM).
Human Molecular Genetics, 2010, Vol. 19, No. 244907
recruitment of snRNPs to the exon suggests that splicing of the
exon may be sensitive to alterations in the snRNP abundance
caused by a decrease in SMN protein levels in the cell. In
addition, the 5’ splice site of exon 7 is suboptimal, suggesting
that U1 snRNP recruitment to the exon may be a limiting
factor in the efficiency of exon 7 splicing (36,37). The impor-
tance of snRNP recruitment to exon 7 splice sites suggests that
a change in the snRNP abundance, as is seen when SMN
protein is reduced in SMA, may influence exon 7 splicing.
A correlation between SMN2 exon 7 splicing and SMN
protein abundance is apparent in transgenic mice (25,38) as
well as in human cells (8,39,40). However, this correlation
has not been previously attributed to a feedback mechanism,
and the direct effect of reduced SMN protein levels on exon
7 splicing has not been investigated. We now demonstrate
that a decrease in SMN protein results in a decrease in
SMN2 exon 7 splicing. We also show that splicing of SMN2
exon 7 is sensitive to changes in the relative abundance of spli-
ceosomal snRNPs. Our results indicate that SMN expression is
controlled, in part, by a feedback mechanism that allows
homeostatic control of exon 7 splicing as a means to poten-
tially regulate snRNP biogenesis and other functions of
SMN. Feedback regulation of SMN expression has impli-
cations for understanding cellular defects in SMA. This mech-
anism also suggests that a small increase in SMN2 exon 7
splicing would result in a disproportionately high increase in
SMN protein levels, a concept that will be important when
assessing the potential therapeutic effect of compounds and
other small molecules in SMA.
Low SMN protein levels correlate with decreased exon 7
splicing in SMA
Based on the premise that a decrease in SMN protein levels in
cells results in changes in the relative snRNP abundance,
which can affect alternative splicing in general (13–15), we
analyzed RNA from an SMA patient and a mouse model of
SMA to determine whether splicing of SMN2 exon 7 is
affected by a decrease in the SMN protein abundance in
vivo. We first compared SMN protein abundance in mice
transgenic for human SMN2 either with both copies of the
mouse Smn gene (hSMN2; Smn+/+) or with a homozygous
deletion of the mouse Smn gene (hSMN2; Smn 2/2) at post-
natal day 1. Transgenic mice with homozygous deletion of the
mouse SMN gene are referred to as SMA mice as they exhibit
an SMA phenotype (41). SMN protein was not detectable in
the SMA mouse tissues analyzed, indicating that very little
SMN protein is produced from the hSMN2 gene (Fig. 1B).
A protein with lower mobility than full-length SMN was
detected in the spinal cord of SMA mice (Fig. 1B). This
product may be truncated SMN protein arising from SMN2
mRNA lacking exon 7. The analysis of exon 7 splicing in
the human SMN2 gene RNA transcript revealed that ,10%
of the hSMN2 transcripts include exon 7 (Fig. 1C). It has
been shown previously that the protein product encoded by
the SMN mRNA transcripts lacking exon 7 is unstable and
undetectable by immunoblot in mice (8), which is consistent
with the lack of SMN protein detected in the SMA tissues
(Fig. 1B). In contrast, hSMN2 transgenic mice that have an
intact mouse SMN gene (hSMN2, mSMN+/+) produce
SMN protein from the endogenous gene and exhibit a
4–8-fold increase in SMN2 transcripts that include exon 7
compared with the mSMN 2/2 samples (Fig. 1C). These
results demonstrate a correlation between SMN2 exon 7 spli-
cing and SMN protein levels.
To further test whether low SMN protein abundance corre-
lates with a decrease in SMN2 exon 7 splicing, we analyzed
SMN protein and exon 7 splicing in induced pluripotent
stem (iPS) cells generated from an SMA patient (iPS-SMA)
or from an unaffected family member (iPS-WT) (42).
SMN1 and SMN2 spliced RNA products were analyzed by
reverse transcription followed by polymerase chain reaction
(PCR) amplification. SMN1 and SMN2 products can be distin-
guished from one another by digestion with the restriction
enzyme DdeI, which has a unique site in exon 8 of SMN2
that is not present in SMN1. Following digestion, SMN2
spliced products are smaller than SMN1 as visualized by
gel electrophoresis. We found that in the iPS-SMA cells,
SMN protein levels are reduced by 45% compared with
WT (Fig. 1D), reflecting the loss of the SMN1 gene in
these cells. The low SMN protein levels in the iPS-SMA
cells corresponded with a 48% reduction in the percent of
SMN2 transcripts that include exon 7 splicing in iPS-SMA
cells compared with WT cells (Fig. 1E). This result further
demonstrates the correlation between SMN protein abun-
dance and exon 7 splicing, suggesting a possible auto-
regulatory feedback mechanism.
Feedback regulation of exon 7 splicing could reflect either a
direct or an indirect effect of SMN protein on SMN2 exon 7
splicing. We also analyzed by immunoblot whether depletion
of SMN protein in cells causes a change in known regulators
of SMN2 exon 7 splicing. We found that neither hnRNP Q/R
nor Tra2b1 abundance changed between the WT and SMA
iPS cells, suggesting that the reduction in SMN protein
levels does not cause exon 7 skipping indirectly due to the
reduction of these factors, which have been previously
shown to affect exon 7 splicing (25,33) (Fig. 1D).
Because a major function of SMN protein involves assem-
bly of Sm proteins on snRNAs (12), which stabilizes the
snRNAs (14), we next analyzed snRNA levels in the iPS
cells to determine whether reduced SMN protein abundance
in iPS-SMA cells correlates with a reduction in snRNAs. A
reduction in snRNAs could account for alterations in SMN2
exon 7 splicing that is observed when SMN protein abundance
is low. Exon 7 is spliced via the major spliceosomal pathway
involving U1, U2, U4, U5 and U6 snRNPs. We focused on U1,
U2, U4 and U5, because SMN assembles Sm proteins onto
these, whereas the U6 snRNA assembly involves a different
pathway (10). We measured snRNA levels by reverse tran-
scription followed by real-time PCR. We observed a 23,
41and 34% reduction in U1, U2and U4 snRNA abundance,
respectively, in iPS-SMA relative to iPS-WT when normalized
to a control transcript (Fig. 1F), whereas U5 snRNA levels
were slightly higher in iPS-SMA compared with iPS-WT
cells. Our results suggest that the decrease in exon 7 splicing
triggered by low SMN protein levels may be a result of
lowered snRNA abundance or a change in the relative abun-
dance of the snRNAs (13,14). Overall, our results reveal that
4908 Human Molecular Genetics, 2010, Vol. 19, No. 24
iPS cells derived from SMA patients have a deficit in the
abundance of some U snRNA species as well as lower
SMN2 exon 7 splicing.
Cellular depletion of SMN leads to an increase in SMN2
exon 7 skipping
To test directly whether SMN protein abundance affects SMN2
exon 7 splicing, we assayed SMN2 splicing in HEK-293T cells
depleted of SMN using an siRNA that targets both SMN1 and
SMN2. RNAi against SMN resulted in a 64% reduction in
SMN protein (Fig. 2A). To verify that any effect of the RNAi-
mediated knockdown of SMN is a direct result of lowered SMN
protein levels, we introduced an RNAi rescue plasmid expres-
sing full-length SMN2 cDNA with silent point mutations that
render its mRNA insensitive to RNAi. Expression of the
rescue SMN plasmid replenishes SMN protein in the siRNA-
treated cells in a dose-dependent manner (Fig. 2A).
We also analyzed by immunoblot analysis the effect of
SMN depletion on the expression of a number of known reg-
ulators of exon 7 splicing, including hnRNP Q/R and A1,
Tra2b1, SF2/ASF and SRp55 (Fig. 2B) (25,43,44). There
was no change in the abundance of these proteins in
SMN-depleted cells compared with control cells.
Analysis of snRNA levels in SMN-depleted cells confirmed
that the snRNA levels are affected by a decrease in SMN
protein. Real-time PCR indicated a reduction in U1, U2 and
U4, but not in U5 snRNA levels in cells with reduced SMN
protein (Fig. 2C). Expression of SMN protein in siSMN-treated
cells partially restored U1 snRNA, but not U2 or U4 snRNA
dance in cells with reduced SMN protein, with some snRNAs
more affected than others. These changes result in an overall
alteration in the relative abundance of individual snRNPs,
which could influence the dynamics of splice site recognition.
The relatively subtle effect that restoration of SMN protein
expression had on snRNA levels led us to test the degree to
which the abundance of mature snRNPs, which require SMN
for assembly, was affected by SMN protein depletion and
rescue in this system. We focused on U1 snRNP for the
purpose of comparing the two snRNA assays. Mature snRNPs
were immunoprecipitated from lysates of treated cells using an
snRNAs were subsequently linkered at the 3’ end and reverse-
transcribed using a linker-specific primer. PCR was carried out
with an snRNA-specific primer and a linker-specific primer.
The amount of U1 snRNA in the starting cell lysate was
measured as an input control. SnoRNA48 was also measured to
normalize input RNA concentration and to demonstrate the
specificity of the immunoprecipitation for Sm-bound snRNAs.
of SMN and restored in cells expressing the SMN rescue protein
(Fig.2D). Theseresultssuggestthat althoughtotal snRNA abun-
on mature snRNPs is more dramatic and that U1 snRNP abun-
dance is restored by expression of the SMN rescue protein in
We next analyzed the effect of SMN depletion on exon 7
splicing. Depletion of endogenous SMN1 and 2 from the
cells resulted in a reduction in mRNA and protein levels,
precluding analysis of endogenous SMN2 exon 7 splicing in
these experiments because it is eliminated in the siSMN-
treated cells. Instead, a minigene plasmid expressing the 3’
comprised exons 6, 7 and 8 and the intervening introns and
recapitulates SMN2 exon 7 alternative splicing in a manner
similar to the endogenous transcript (20). Reverse transcrip-
tion (RT)–PCR analysis of SMN2 exon 7 splicing from the
minigene showed a nearly 48% reduction in SMN2 mRNA
containing exon 7 in the cells depleted of SMN (Fig. 2E).
This effect was specific to the reduction in SMN protein
levels as evidenced by the restoration of exon 7 splicing
when the SMN2 rescue plasmid (WT) is expressed to restore
SMN protein levels (Fig. 2E).
We also rescued SMN knockdown cells with an SMN gene
that is insensitive to RNAi and codes for the SMN mutant pro-
teins A111G, which has a moderately decreased affinity for
Sm proteins, and Y272C, which has more severe defects in
the Sm core assembly (Fig. 2A) (2,45–47). The SMN/
A111G protein partially rescued the splicing defect, whereas
the SMN/Y272C protein was completely inactive (Fig. 2E).
These results are consistent with the documented activity of
these mutant proteins and support the idea that SMN2 exon
7 splicing is affected by SMN protein abundance and activity
related to snRNP assembly and abundance. Overall, our results
strongly support a role for SMN protein in SMN exon 7 spli-
cing by altering the abundance of a subset of snRNPs.
To test the dynamics of the feedback loop, we performed a
time-course experiment in which cells were treated with
siSMN or the control siRNA, and exon 7 splicing of SMN2
minigene transcripts was measured at different time points fol-
lowing SMN depletion. A correlation between SMN protein
levels and SMN2 exon 7 splicing was observed with
maximum exon 7 skipping occurring when SMN protein
levels are lowest at 72 h post-transfection (Fig. 2F). These
results demonstrate that SMN protein abundance correlates
with the degree of exon 7 splicing.
snRNP abundance influences exon 7 splicing
The differences in snRNA levels in SMA compared with WT
iPS cells and in SMN knockdown cells compared with control
cells suggest that the abundance of some snRNA species may
be more sensitive to a decrease in SMN protein abundance
than others. This differential change in snRNA species could
explain the effect of SMN protein abundance on SMN2 exon
7 splicing if splicing is regulated by changes in the relative
abundance of snRNPs. To directly test the idea that changes
in individual snRNPs influence exon 7 splicing, we used a
cell-free in vitro splicing assay. In this assay, in vitro tran-
scribed SMN1 or SMN2 pre-mRNAs comprised exons 6 and
7 and a portion of exon 8, and the intervening introns (18)
were spliced in the HeLa cell nuclear extract. U1, U2, U4
and U5 snRNAs were targeted by oligonucleotide-directed
RNase H cleavage, a method commonly enlisted to digest
snRNAs in order to test their requirement in splicing (48–
52). Oligonucleotides were tested at different concentrations
that were optimized for each snRNA in order to assess a dose-
dependent effect of snRNA depletion on splicing.
RNase H cleavage of each species of snRNA resulted in
strikingly different effects on exon 7 splicing. At the lowest
Human Molecular Genetics, 2010, Vol. 19, No. 244909
Figure 2. SMN protein reduction causes an increase in SMN2 exon 7 skipping. (A) Immunoblot analysis of SMN protein from HEK-293T cells transfected with a
minigene expressing SMN2 and either a scrambled control (siC) siRNA or an siRNA targeted to SMN1/2 (siSMN). Rescue refers to samples from cells trans-
fected with different quantities of a plasmid expressing SMN2 cDNA with silent point mutations in the siRNA target sequence. Blots were probed with SMN or
b-actin (loading control)-specific primary antibodies followed by a fluorescent and HRP-conjugated secondary antibody. Bands were quantitated by phosphor-
image analysis. Graph shows results as the ratio of SMN to b-actin. Error bars represent SEM. The number of independent experimental measurements (n) is
shown above the graph. For all data points, P , 0.0005 relative to siSMN as determined by the Student’s t-test. (B) Immunoblots probed with antibodies against
hnRNP R/Q, hnRNP A1, SFRS10 (Tra2b1), SRp55, SF2/ASF and b-actin. (C) Quantitation of snRNAs by real-time PCR. The snRNA levels of the SMN RNAi
cells are normalized to Rpl39 and plotted as percentage of control (siC). Error bars represent SEM, n ¼ 4 independent experiments for the siSMN/siC comparison
and n ¼ 3 for siSMN + rescue/siC. (D) PCR analysis of 3’ end-linkered U1 snRNAs immunoprecipitated from cell lysates using an anti-Sm antibody. U1 snRNA
levels are normalized to snRNA in the input lysate. SnoRNA48 was used as a control. (E) RT–PCR analysis of SMN2 minigene transcript splicing in cells treated
as in (A). Reaction products were digested with DdeI as in Figure 1. Results are represented graphically as percent exon 7 inclusion. Error bars represent SEM; n
is shown above the graph.∗P , 0.05;∗∗P , 0.005. (F) Immunoblot (top) and RT–PCR (bottom) analysis of SMN protein expression and exon 7 splicing as a
function of time following siRNA-mediated depletion of SMN protein from cells. Quantitation was performed as in (A) and (C) above.
4910 Human Molecular Genetics, 2010, Vol. 19, No. 24
concentration of U1 snRNA-targeting oligonucleotides, there
was a greater than 50% reduction in spliced products that
include exon 7, whereas transcripts lacking exon 7 did not
change dramatically relative to reactions with untreated
extracts (Fig. 3A). This change in alternative splicing
dynamics is reflected in the dramatic decrease in the percent
exon 7 inclusion (Fig. 3A). In contrast, targeting U2 snRNA
resulted in a nearly 50% reduction in exon 7 skipping and
an increase in exon 7 inclusion at the lowest concentration
of oligonucleotide (Fig. 3B). This results in an overall increase
in exon 7 inclusion, suggesting that when U2 snRNA is limit-
ing, exon 7 splicing out-competes exon 7 skipping (Fig. 3B). A
similar effect was observed when U4- and U5-targeted oligo-
nucleotides were tested (Fig. 3C and D).
Splicing of both SMN1 and SMN2 exon 7 was similarly
responsive to the RNase H depletion of individual snRNAs,
suggesting that the effect that low snRNA levels have on spli-
cing is not dependent on the C-to-T difference in exon 7
between SMN1 and SMN2. To demonstrate that the RNase H-
depleted extracts were effectively targeting snRNAs, we tested
splicing of a constitutively spliced b-globin transcript. The
RNase H depletion caused a dose-dependent inhibition of
b-globin splicing (Fig. 3E), demonstrating a progressive
block to constitutive splicing after degradation of snRNAs.
Depletion of snRNP proteins reduces SMN2 exon 7 splicing
To further test whether altering snRNP levels can affect exon 7
splicing, we depleted snRNPs from cells using siRNAs
directed against several protein components of the mature
snRNP-associated protein, B" (SNRPB2) by 51%, or the U1
snRNP-associated protein, U170K (SNRNP70) by 63%, led
to a 37% and 44% decrease in SMN2 exon 7 splicing, respect-
ively (Fig. 4A and B). Similarly, a 15% knockdown of the Sm
protein D3 (SNRPD3), a direct target of SMN in the assembly
onto U1, U2, U4 and U5 snRNAs (53), resulted in a 52%
reduction in exon 7 splicing. Cellular depletion of D3 also
resulted in a 30% reduction in another Sm protein, B/B’
(SNRPB). This effect likely reflects the instability of Sm pro-
teins when not complexed together (54). Together, these
results demonstrate that SMN2 exon 7 splicing is sensitive to
changes in the abundance of snRNP components.
Sequestering U1 snRNP in cells reduces exon 7 splicing
Our results indicate that U1 snRNA abundance goes down in
cells with low SMN protein levels and that a decrease in U1
snRNPs dramatically reduces exon 7 splicing. We further
explored the role of U1 snRNP abundance in exon 7 splicing
through the use of RNA decoys. The expression of RNA
decoys with sequence similarity to a 5’ splice site has been
shown to sequester U1 snRNP and to alter the splicing of
exon 7 from SMN2 minigene-expressed pre-mRNAs (37)
(Fig. 5A). This system offers a means to independently test
the effect of lowered U1 snRNP levels on the splicing of
endogenous SMN2 exon 7 to determine whether the modu-
lation of U1 snRNP activity alters splicing in a manner
Figure 3. snRNAs differentially affect exon 7 splicing in vitro. In vitro spli-
cing analysis of SMN1 and SMN2 transcripts in HeLa cell nuclear extracts.
Extracts were untreated (2) or treated with an increasing concentration of
antisense oligonucleotides (oligo) targeting (A) U1 (0.3125, 0.625, 1.25 mM),
(B) U2 (1.25, 2.5, 5 mM), (C) U4 (0.156, 0.3125, 0.625 mM) and (D) U5
(1.25, 2.5, 5 mM) snRNAs for RNase H-mediated cleavage. Unspliced
pre-mRNA and spliced products are indicated. Open arrowheads represent
debranched intron lariats. Quantitation of in vitro splicing, shown as the
percentexon7 inclusion [included/(included + pre-mRNA)×100],
graphed to the right in each panel. Error bars represent + SEM with n indi-
cated above the bars,
Bars with no error represent the average of two independent experiments.
(E) In vitro splicing of b-globin transcripts. Splicing reactions were carried
out as in (A).
∗P , 0.05,
∗∗P , 0.005 relative to untreated (2).
Human Molecular Genetics, 2010, Vol. 19, No. 244911
similar to that seen with the SMN or snRNP component
Expression of the 5’ss decoy in HEK-293T cells resulted in
a 50% reduction in endogenous SMN2 exon 7 splicing
(Fig. 5B). SMN1 exon 7 splicing was also reduced. Expression
of decoys with point mutations in the 5’ splice site sequence
did not affect exon 7 splicing (Fig. 5B). The 5’ splice site
decoys caused a nearly 50% overall reduction in full-length
exon 7, including transcripts from both SMN1 and SMN2.
This reduction in full-length transcripts leads to a 42%
reduction in SMN protein levels (Fig. 5C). These results
demonstrate the specific sensitivity of exon 7 splicing to
alterations in the U1 snRNP abundance and also show that a
reduction in full-length SMN1/2 transcripts leads to a
reduction in SMN protein. Our findings further link a modu-
lation of snRNP abundance to the control of SMN2 exon 7
splicing and SMN protein expression.
A major goal in SMA research has been to identify approaches
to improve expression of SMN protein from SMN2 for the
treatment of the disease. Increasing SMN2 exon 7 splicing
has been studied intensely as a means to elevate full-length
SMN protein levels in SMA. One key question is how much
of an increase in SMN2 exon 7 splicing is required for thera-
peutic value. Because SMN protein itself functions in the
pre-mRNA splicing pathway, it is important to understand
how the protein may influence splicing of its own pre-mRNA.
We now demonstrate that the abundance of SMN protein deter-
mines, in part, the outcome of SMN2 alternative splicing. The
discovery of a feedback loop in SMN protein expression has
important implications for SMA by suggesting that treatment
strategies that lead to modest increases in SMN protein may
have significant therapeutic value for the disease.
The SMN feedback loop likely involves the key function of
SMN in the snRNP assembly. We (Figs 1 and 2) and others
(13,14) have found that some spliceosomal snRNA species
decrease, following a reduction in the SMN protein abun-
dance. This change in snRNAs consequently lowers snRNP
abundance and correlates with changes in alternative splicing
of a number of gene transcripts, presumably due to a change in
the relative concentration of snRNPs (14). It is not clear how
changes in the relative abundance of snRNPs lead to changes
in alternative splicing. Using an in vitro splicing assay, we
tested directly the possibility that subtle changes in the relative
Figure 4. Reduction of snRNP proteins causes SMN2 exon 7 skipping. (A) Immunoblot analysis of HEK-293T cells transfected with either a scrambled control
(C) siRNA or an siRNA targeted to U2-B", Sm-D3 or U1-70K. Blots were probed with primary antibodies to indicated proteins. Fluorescent and HRP-conjugated
secondary antibodies were used for detection, bands were quantitated by phosphorimage analysis and results are graphed as the ratio of indicated protein to
b-actin signal normalized to control samples. (B) RT–PCR analysis of SMN1 and SMN2 endogenous transcripts in cells treated as in (A). Reaction products
were digested with DdeI. Results are represented graphically as the percent exon 7 inclusion. For both (A) and (B), error bars represent the SEM, n ¼ 3,
∗∗P , 0.005.
4912 Human Molecular Genetics, 2010, Vol. 19, No. 24
abundance of the snRNPs can have effects on alternative spli-
cing of SMN2 exon 7 (Fig. 3). We found that altering the abun-
dance of individual snRNPs in vitro had dramatically differing
effects on exon 7 splicing, demonstrating that shifts in the
balance of snRNPs can regulate alternative splicing (Fig. 3).
Our results suggest that a change in the relative amount of
individual snRNPs alters SMN2 exon 7 splicing, which, in
turn, affects SMN protein levels which feeds back to further
disrupt snRNP abundance.
SMN2 exon 7 splicing is particularly sensitive to U1 snRNP
levels. We present data showing that U1 snRNA is reduced
when SMN protein abundance is low (Figs 1 and 2) and that
exon 7 splicing decreases following a decrease in U1 snRNP
activity or abundance (Figs 3–5). U1 snRNP recognizes the
5’splicesite through direct
between the pre-mRNA and the snRNA (55). U1 snRNP
binding is an early determinant of splice site selection
(56,57). Thus, the 5’ splice site sequence, and likely also the
abundance of U1 snRNP, plays an important role in the
initial recognition of the site. 5’ splice sites with weak base-
pairing potential to U1 snRNA are not recognized as effi-
ciently as those with high base-pairing potential (58). The effi-
ciency of 5’ splice site selection is important when 5’ splice
sites are in competition with each other in alternatively
spliced exons. For example, in the case of SMN2, the 5’
splice site of exon 7 is an important determinant of exon
inclusion (36,37). Exon skipping will occur if exon 6 splicing
to exon 8 occurs before exon 7 can splice to exon 8. Improve-
ment of the base-pairing potential of its 5’ splice site to U1
improves exon 7 splicing, likely due, in part, to its improved
ability to compete with the 5’ splice site of exon 6 for splicing
to exon 8. We observed that a decrease in U1 snRNP abun-
dance in vitro results in a decrease in exon 7 inclusion relative
to exon skipping (Fig. 3). This may be due to the weak exon 7
5’ splice site, which is further weakened under conditions of
low U1 snRNP. This site may also not be strong enough to
promote exon definition across exon 7 for enhancement of
the 3’ splice site of exon 7, which has also been demonstrated
to be a relatively weak splice site (22,59). Limiting U1 snRNP
may further weaken exon 7 definition, thereby lowering the
occurrence of splicing from the 5’ splice site of exon 6 to
the 3’ splice site of exon 7. In this case, splicing from exon
6 to the 3’ splice site of exon 8 may be selected over the
exon 7 3’ splice site, resulting in exon 7 skipping.
Interestingly, targeted reduction of individual snRNAs had
differential effects on exon 7 splicing. Unlike U1 snRNA
depletion, exon 7 inclusion was much more resistant to the
depletion of U2, U4 and U5 snRNA species in vitro compared
with skipping (Fig. 3). These results suggest that exon 7 skip-
ping and inclusion are differentially affected by alterations in
the relative abundance of individual snRNPs. This phenom-
enon, in which the relative abundance of individual snRNPs
dictates exon 7 splicing, could result in differences in the rela-
tive amount of SMN2 exon 7 inclusion in different cell types
and at different stages of development. In the case of SMA,
in which SMN1 is lost, this feedback regulation of SMN2 spli-
cing could be crucial for cell survival. It is possible that
SMN2 exon 7 splicing in motor neurons is especially sensitive
tributing tomotorneuron degenerationinSMA. Indeed, snRNP
levels and the abundance and activity of SMN protein fluctuate
in the spinal cord throughout early development (60).
Modulation of individual snRNAs did not always have the
expected outcome on SMN2 exon 7 splicing. The relative
resistance of exon 7 inclusion to U2 snRNA depletion in
vitro (Fig. 3), for example, was surprising. U2 snRNP binds
to the 3’ss and is a determinant of 3’ss selection. U2 snRNP
binding to the 3’ splice site of exon 7 is impaired in SMN2
compared with SMN1 (22,59). Based on these results, it
might be predicted that the depletion of U2 snRNP would
impair SMN2 exon 7 3’ splice site recognition and lead to
an increase in skipping. Our results, however, are consistent
with our previous finding that depletion of U2AF65 or
PUF60, which recruit U2 snRNP to the 3’ splice site, results
in an increase in exon 7 inclusion (28). Modulation of 3’
splice site recognition may weaken the use of the distal
Figure 5. Overexpression of U1 snRNP binding sites causes SMN2 exon 7
skipping. (A) Diagram of U1 snRNA with 5’ sequence base-pairing to the
target sequence of the U1 5’ splice site (5’ss) decoy or U1 mutant decoys
(mut1, mut2) with one nucleotide change in the U1-recognition sequence
(bold and underlined). (B) RT–PCR analysis of endogenous SMN1 and
SMN2 transcripts from HEK-293T cells transfected with plasmids expressing
5’ss decoys (5’ss) or mutated decoys. Reactions were digested with DdeI and
separated on a 6% native polyacrylamide gel. Bands were quantitated and
graphed as the percent exon 7 inclusion [included/(included + skipped)×100]
(top); +SEM, or the change in total full-length SMN (SMN1 + SMN2 tran-
scripts including exon 7) (bottom);
(C) Immunoblot analysis of SMN protein levels in cells transfected with
decoy plasmids. Quantitation is shown to the right with the ratio of SMN to
b-actin signal normalized to mock transfected (C) samples; +SEM, n ¼ 3.
∗P , 0.05,
∗∗P , 0.01.
Human Molecular Genetics, 2010, Vol. 19, No. 244913
exon 8 3’ splice site and thereby improve the competitiveness
of the exon 7 3’ splice site for splicing. Contrary to this
interpretation, depletion of the U2 snRNP protein SNRP B"
resulted in an increase in exon 7 skipping (Fig. 4). This differ-
ence may be due to secondary effects of U2 snRNP depletion
in cells or may indicate a functional difference in U2 snRNP
when the B" protein is limiting.
The effect of snRNP abundance on splicing outcome is
more complex in the context of alterations of multiple
snRNP species, as we observed in the iPS cells (Fig. 1) and
SMN knockdown experiments (Fig. 2). Our in vitro results
(Fig. 3) indicate that a decrease in U2, U4 or U5 snRNP abun-
dance causes an increase in exon 7 inclusion. However, in
cells, a decrease in SMN protein always correlates with a
decrease in exon 7 inclusion (Figs 1 and 2). It is possible
that early recognition of the 5’ss by U1 snRNP is the determin-
ing step in splice site selection, and thus U1 snRNP abundance
has a dominant role in exon 7 splicing.
The SMN feedback loop is likely an important regulator of
SMN expression in SMA where SMN1 is mutated or deleted
and thus the equilibrium of the feedback loop is disrupted and
splicing of SMN2 exon 7 is reduced. The ability of a feedback
loop to potentially impact endogenous SMN1 and SMN2 spli-
cing and SMN protein production is demonstrated using the
in vitro splicing assay and in experiments with the U1 snRNA
decoy where both SMN1 and SMN2 exon 7 splicing were
affected by alterations in the snRNP abundance (Figs 3 and
5). However, in cells, splicing of endogenous or minigene
SMN1 exon 7 was not affected by changes in the SMN
protein abundance. This insensitivity may be due to the high
efficiency of SMN1 exon 7 splicing in combination with the
relatively modest reduction in SMN protein and snRNAs. Our
results suggest that the feedback loop could play a regulatory
role when SMN1 is present in a non-diseased state in situations.
Regulation of exon 7 splicing could reflect an indirect effect
of SMN protein on SMN2 exon 7 splicing. Reduction in SMN
protein levels has been reported to result in changes in alterna-
tive splicing of a large number of transcripts (14). Alternative
splicing changes are likely to have an impact on the abundance
or activity of the resulting proteins. A number of protein
factors have been described that alter the splicing of SMN2
exon 7. The reduction in SMN protein may cause a change
in the splicing of one of these regulators and thus indirectly
alter SMN2 exon 7 splicing. Microarray experiments examin-
ing global changes in splicing in SMA mice have not revealed
changes in alternative splicing of any known regulators of spli-
cing (14,16). To address this possibility more directly, we
tested whether depletion of SMN protein in cells causes a
change in known regulators of SMN2 exon 7 splicing and
did not observe quantitative changes in the abundance of a
number of these regulatory proteins including SF2/ASF,
hnRNPA1, hnRNP Q/R or Tra2b1 (Figs 1D and 2B)
(19,20,25,33). However, we cannot rule out the possibility
that the effect of lowering the SMN protein levels on exon 7
inclusion results in part from alterations in the abundance or
activity of other effectors of exon 7 splicing. Nonetheless,
our demonstration that alterations in snRNP levels can alter
exon 7 splicing in vitro suggests that the decrease in exon 7
splicing upon reduction of SMN protein levels is due, at
least in part, to change in snRNP levels in the cell. We also
provide evidence that changes in the relative abundance of
core spliceosomal snRNPs can regulate alternative splicing.
Our results are a first demonstration of feedback regulation
whereby an alteration in SMN protein levels controls
expression of the protein itself by affecting alternative splicing
of its pre-mRNA transcripts. These finding lay the groundwork
for future studies to understand the degree to which an initially
small increase in exon 7 splicing can result in a disproportio-
nately larger increase in SMN protein levels. From a more
broad perspective, our results indicate that the relative abun-
dance of individual snRNPs can regulate alternative splicing.
MATERIALS AND METHODS
Plasmids and constructs
The pCI-SMN2cDNA rescue plasmid was constructed by
reverse transcription of HEK-293T total cellular RNA with
oligodT primers using Superscript III cDNA synthesis kit
(Invitrogen, Carlsbad, CA). SMN2 full-length cDNA was
amplified by PCR using the primers SMNex1XhoI and
SMNex8NotI. The PCR product and the plasmid pCI
(Promega, Madison, WI) were digested with XhoI and NotI
and ligated together using T4 DNA ligase (New England
Biolabs, Ipswich, MA). The resulting plasmid was used as a
template for mutagenesis using QuikChange Lightning site-
directed mutagenesis kit (Stratagene, Santa Clara, CA) and
the primers SMNmisF and SMNmisR, according to the manu-
Primer and RNAi sequences are provided in Supplementary
Material, Table S1.
Cell-free in vitro splicing
Plasmids pCI-SMN1 and pCI-SMN2 (61) were used as tem-
T7SMNex6, that is specific to the 5’ end of SMN exon 6
with a T7 promoter sequence at the 3’ end and a reverse
primer, SMNex8-75R+5’, specific to SMN exon 8 (20).
PCR products were used as templates for in vitro transcription
with T7 RNA polymerase (Promega) to make SMN1 and
SMN2 E678 transcripts. b-globin transcripts were made
using the template, pSP64-HbD6 linearized with BamHI and
transcribed with SP6 RNA polymerase (48).
Transcription reactions were carried out in the presence of
32P-UTP and 7Me-GpppG cap analog (NEB) to make pre-
mRNAs for in vitro splicing analysis. RNAs were purified
by denaturing polyacrylamide gel electrophoresis (PAGE).
RNase H degradation of snRNAs in HeLa nuclear extract dia-
lyzed in buffer D [20 mM HEPES-KOH, pH 8; 100 mM KCl;
0.2 mM EDTA; 20% (v/v) glycerol] was achieved by incubat-
ing extract with antisense oligonucleotides targeting the
snRNA for 15 min at 308C (48). Treated extracts were used
immediately in a 10 ml splicing reaction in which 10 fmol of
RNA was incubated with 3 ml of nuclear extract, 1.3% (w/v)
polyvinyl alcohol (PVA), 0.5 mM ATP, 20 mM creatine
using theforward primer,
4914 Human Molecular Genetics, 2010, Vol. 19, No. 24
phosphate, 1.6 mM MgCl2and 30% (v/v) buffer D at 308C for
3 h (62).
Cell culture, transfections and RNAi
HEK-293T cells were cultured in Dulbecco’s modified Eagle’s
medium supplemented with 10% fetal bovine serum. For
RNAi experiments, 1.5 × 105
six-well dish 24 h prior to treatment. DsiRNAs were syn-
thesized by Integrated DNA Technologies, Inc. (IDT, Coral-
ville, IA). The sequences of the siRNA duplexes are listed
in Supplementary Material, Table S1. The siC sequence was
the Dicector DS scrambled negative control duplex (IDT).
For SMN knockdown experiments, cells were transfected
with 1 mg of SMN2 or SMN1 minigene reporter (20) and
20 nM siRNA duplex using Lipofectamine 2000 (Invitrogen),
according to the manufacturer’s protocol. For rescue of
SMN knockdown, the pCI-SMN2 cDNA rescue plasmid was
also transfected. After 24 h, cells were split 1:2. Cells were
transfected with an additional 20 nM of siRNA 48 h after the
initial treatment and grown for an additional 48 h at which
time total RNA and protein were collected. For the time-
course experiment, cells were treated similarly except that
cells were treated with 20 nM siRNA initially and then every
48 h throughout the time-course. The SMN2 minigene
plasmid was transfected at time 0 h for the 24 and 48 h time
points and at 48 h for the 72 h time point. For RNAi-mediated
depletion of U170K, snRNP B" and SmB, cells were treated
with 20 nM siRNA and total RNA and protein were isolated
after 72 h. All RNA was collected from cells using Trizol
(Invitrogen). Protein samples were prepared by lysing cells
in Laemmli buffer and by heating at 998C for 10 min.
U1 decoy plasmids were transfected into HEK-293T cells
using Optifect (Invitrogen), according to the manufacturer’s
protocol. Total RNA was collected using Trizol (Invitrogen)
48 h post-transfection.
iPS cells were cultured and collected as neurospheres, as
described previously (42).
cells were plated into a
Protein samples were separated by sodium dodecyl sulfate
(SDS)–PAGE and transferred to Immobilon-FL membrane
(Millipore, Billerica, MA). Blots were probed with mouse
monoclonal antibodies specific for mouse and human SMN
(BD Biosciences), b-actin (Sigma, St Louis, MO), hnRNP-Q
(Sigma), Sm proteins (mAb Y12), U1-70K (anti-RNP),
U2-B" snRNP protein (mAb 4G3) (63), SF2/ASF (mAb96),
hnRNP A1 (mAb A1/55) (64) and SRp55 (gifts from Adrian
Krainer) or rabbit polyclonal antibody against SFRS10/
Tra2b (Sigma), followed by Alexafluor 594-conjugated anti-
mouse or anti-rabbit secondary antibody (Invitrogen) or horse-
radish peroxidase (HRP)-conjugated goat anti-mouse or anti-
rabbit secondary antibody. Detection and quantitation of the
signal was performed using a Typhoon 9400 Variable Mode
Imager (GE Healthcare, Waukesha, WI) and ImageQuant T
software for fluorescently detected blots or with Lumi-Light
Western Blotting Substrate (Roche Diagnostics, Indianapolis,
IN) for HRP-labeled blots.
RNA was collected using Trizol reagent (Invitrogen). Reverse
transcription was performed using Superscript III cDNA syn-
thesis kit (Invitrogen) with oligo dT primers except for snRNA
quantitation experiments in which RNA was reverse-
transcribed using random primers. PCR with GoTaq polymer-
ase (Promega) was carried out for 25 amplification cycles in
reactions containing (a-32P)-dCTP. Exon 7 splicing of
hSMN2 RNA from transgenic mouse tissues was analyzed
using human-specific primers E4-33to55-F and E8-15to36-R
(30). Endogenous SMN1 and SMN2 RNA transcripts from
human cells were amplified using primers SMNex6Xho and
rescue and pCI-SMN1 and SMN2 minigenes were amplified
using pCI-FwdB and pCI-Rev. Following PCR of endogenous
SMN1 and SMN2, reactions were treated with DdeI for 1 h,
which digests at a unique site within SMN2 exon 8 that is
not present in SMN1. Products were separated on 6% native
polyacrylamide gels. Quantitation is based on phosphorimage
analysis (Typhoon 9400; GE Healthcare).
Real-Time PCR experiments for snRNA quantitation were
carried out on an Applied Biosystems (ABI) 7500 Real-Time
PCR System using Power SYBR Green (Applied Biosystems,
Foster City, CA). Each snRNA was measured in triplicate.
Absolute quantification was performed using the ABI 7500
detection software with correction for amplification efficiency
based on an exponential model of PCR (65,66).
Cells (?1 × 106) were harvested in 1 ml of IP-500 buffer
[500 mM NaCl, 10 mM Tris-Cl, pH 7.4, 0.1% Triton X-100,
50 mM NaF, 0.2 mM sodium vanadate, protease inhibitor cock-
tail (Sigma)] and sonicated. Lysates were centrifuged, and
supernatant was collected and used in the assay. Protein G
Dynabeads were incubated with anti-Sm antibody (Y12,
Abcam, Cambridge, MA) in IP-100 buffer (100 mM NaCl,
10 mM Tris–Cl pH 7.4, 0.1% Triton X-100, 50 mM NaF,
0.2 mM sodium vanadate) for 4 h at 48C. Beads (5 ml) were
subsequently washed with IP-100 and added to 400 ml of
cell lysate and rotated for 4 h at 48C. Proteinase K was then
added to the lysates and incubated at 378C for 20 min followed
by phenol extraction and ethanol precipitation of RNA. For
input control, cell lysate without beads was treated in a
RNA linker ligation
For linker-ligated RT–PCR, the 3’ end of RNA was ligated to
linker 1 (IDT, Coralville, IA, USA), according to manufac-
turer’s instructions for miRCat (IDT), except for the substi-
tution of Ligation Enhancer with PVA, and truncated RNA
ligase 2 (NEB) to promote polar ligation. Modban primer
was used for reverse transcription and PCR.
Transgenic mice containing human SMN2 and the mouse Smn
knockout are as described previously (41). Genotyping and
Human Molecular Genetics, 2010, Vol. 19, No. 244915
SMN2 copy number determination were performed as
described previously. Mouse genotypes are hSMN2+/+;
mSmn+/+or hSMN2+/+and mSmn2/2. Mice were maintained
in accordance with the Cold Spring Harbor Laboratory Animal
Care and Use regulations. Mouse tissues were snap-frozen in
liquid nitrogen and stored at 2708C. RNA was extracted
from livers using Trizol reagent (Invitrogen) according to
the manufacturer’s protocol. To prepare protein lysates,
100 mg of liver tissue was sonicated in 900 ml of RIPA
buffer [1× phosphate-buffered saline, 0.25% (w/v) sodium
deoxycholate, 0.1% (w/v) SDS, 1 mM EDTA and complete
mini protease inhibitor cocktail (Roche)].
Supplementary Material is available at HMG online.
We thank Adrian Krainer for the mouse tissue samples and
antibodies, Xavier Roca for the U1 decoy constructs and
Clive Svendsen for the iPS cells used in this study. We are
grateful to David Horowitz, Judy Potashkin, Mallory Havens
and Shan-Qing Gu for comments on this manuscript and
Anthony Hinrich for technical assistance.
Conflict of Interest statement. None declared.
This work was supported by Families of SMA and the
Funding to pay the Open Access publication charges for this
article was provided by NINDS/NIH.
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