SMN-C2/C3 Interacts with the Pre-mRNA of SMN2

SMN-C3 has been documented to selectively regulate splicing of SMN2 along with only a handful other genes, such as STRN3 (21). Therefore, the direct target of SMN-C3 is unlikely to be the global splicing machinery. We hypothesized two potential targets for SMN-C3: (i) a sequence of RNA on or close to exon 7, or (ii) a splicing regulatory protein or protein complex that is specific to exon 7. We therefore carried out a pull-down experiment to determine whether SMN-C3 directly binds to an RNA target using the photo–cross-linking probe, SMN-C2-BD (Fig. 1C), in which a biotin-diazirine bifunctional handle replaces the ethyl group of SMN-C2. SMN-C2-BD is ∼20-fold less active than its parent compound as demonstrated by Western blot analyses of SMN protein in SMNΔ7 mouse astrocytes (SI Appendix, Fig. S1).

To capture the interacting RNA fragment, 293T cells were treated with 2 μM SMN-C2-BD, in the absence or presence of 50× SMN-C3, and then cross-linked by irradiation (365 nm) of the live cells. Total RNA was extracted, pulled down by streptavidin beads, washed, and bound RNA was released under denaturing conditions. The released RNA fraction was reverse transcribed to cDNA by random hexamer extension and analyzed by sequencing. Using the competitively treated cells as reference control, alignment of the sequencing reads of the probe-only sample to the human genome did not reveal an enrichment of exon 7. However, we were able to identify a binding motif for SMN-C2-BD in a genome-wide analysis of the captured pre-mRNA. A de novo motif searching for common sequences was performed within the peaks of the aligned reads using MEME motif discovery algorithm (24). A purine-rich binding motif, GAGGAAGA (Fig. 2A), was discovered with a satisfying E value of 1.3 × 10⁻¹⁵¹.

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Fig. 2.

(A) A purine-rich RNA binding motif of SMN-C2 identified by MEME analysis (24) with an E value of 1.3 × 10⁻¹⁵¹. (B) Sequences of synthetic RNA 15-mers. The coverage of each 15-mer is indicated with a bar above or below the sequences. Exon 7 is highlighted in bold with the putative binding sequence AGGAAG of SMN-C2 and a similar AAGGAG sequence at the 5′-ss in red and green, respectively. (C) Fluorescence polarization assays with SMN-C2 (200 nM) and seven RNA 15-mers that cover the whole exon 7, part of the polypyrimidine tract of intron 6, and the junction of exon 7/intron 7 (n = 2). The K_d values of SMN-C2 binding to oligo-4 and -7 were determined to be 16 ± 2 and 46 ± 3 µM, respectively.

This putative binding sequence highly resembled the +24 to +29 (AGGAAG) region of exon 7 (Fig. 2B). To validate the binding specificity, seven RNA 15-mers were synthesized, which cover the entire exon 7 and adjacent intron 6 and 7 regions (Fig. 2B). Each adjacent oligomer was designed to have a 5-nt overlap. An analog of SMN-C3, SMN-C2, contains a coumarin fluorophore so that fluorescence polarization could be used to measure the binding affinity between SMN-C2 and its target (Fig. 2C). As expected, the sequence containing the putative binding site (oligo-4, Fig. 2 B and C), AGGAAG, showed more than 10-fold decrease in K_d compared with sequences 2, 3, 5, and 6 (Fig. 2 B and C). The lower binding affinity of oligo-3 to SMN-C2 suggests that high purine content alone is probably not an SMN-C2 recognition pattern. The pyrimidine-rich sequence, oligo-1, showed the lowest binding affinity to SMN-C2 (Fig. 2 B and C). It has been shown that SMN-C5 also binds to an RNA duplex of 5′-ss of exon 7 and U1 snRNA (23). Although oligo-7 at the 5′-ss of exon contains an AAGGAG sequence that is similar to the putative binding sequence AGGAAG, the binding affinity of SMN-C2 to oligo-7 (K_d, 46 ± 3 µM) is somewhat lower compared with binding of SMN-C2 to oligo-4 (K_d, 16 ± 2 µM).

Next, we attempted to confirm the binding site of SMN-C2 using an RNA footprinting experiment in the context of a longer RNA sequence containing the entire exon 7. Iron or copper chelates in the presence of hydrogen peroxide have previously been used as an artificial chemical ribonuclease to probe the nucleic acid binding sites of molecules (25). Conjugation of the metal ion chelate to a nucleic acid binding molecule leads to oxidative cleavage of the phosphate sugar backbone adjacent to the binding site (26). Therefore, a conjugate of SMN-C2 and 1,10-phenanthroline, SMN-C2–Phen (Fig. 3A), was synthesized with a two-carbon linker (27). To eliminate a potential bias by the phenanthroline moiety in binding to RNA, another copper ligand, Gly–Gly–His (28), conjugate was also synthesized, SMN-C2–GGH (Fig. 3A). In the presence of sodium ascorbate and H₂O₂, a 5′-³²P–labeled 120-nt RNA, which spans exon 7 (54 nt) and adjacent intron 6 and 7 regions, was treated with submicromolar concentrations (40∼400 nM) of the SMN-C2–Phen- and SMN-C2–GGH–copper complexes. As predicted, both SMN-C2–Phe–copper and SMN-C2–GGH–copper exclusively cleaved the RNA three nucleotides away (Fig. 3B) from the putative binding site (29), AGGAAG. This sequence-specific cleavage data confirmed the binding site of SMN-C2.

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Fig. 3.

(A) Structures of SMN-C2–Phen and SMN-C2–GGH complexed with copper. (B) SMN-C2–Phen–Cu and SMN-C2–GGH–Cu exclusively cleave position +32 of exon 7. 1∼3, cleavage reaction with 400, 40, and 0 nM SMN-C2–Phen–Cu; 4∼6, cleavage reaction with 400, 40, and 0 nM SMN-C2–GGH–Cu; 7, RNase T1 ladder (G); 8, alkaline hydrolysis ladder; 9, purified 5′-³²P–labeled RNA. The ribonuclease reaction consisted of 60 mM Hepes, pH 8.0, 20 mM MgCl₂, 0.2 μg/mL yeast tRNA, 0.5 M KCl, 10 nM 5′-³²P–labeled RNA, 0.1% H₂O₂, 1 mM sodium ascorbate, and copper-complexed chemical probe at different concentrations. See SI Appendix for the full sequence of RNA.

SMN-C2 Does Not Disrupt the Secondary Structure of SMN2 Pre-mRNA upon Binding

After the AGGAAG binding motif was identified for SMN-C2/C3, the next step was to determine whether any structural alterations are induced on exon 7 by SMN-C2 binding. The selective 2′-hydroxyl acylation analyzed by primer extension (SHAPE) method was used to interrogate the local nucleotide conformation of exon 7. SHAPE uses a 2′-OH alkylation reagent, 2-methylnicotinic acid imidazolide (NAI), to modify the backbone ribose of an RNA sequence. A base-paired nucleotide is less accessible than an unpaired one to NAI modification; as a consequence, reverse transcriptase stops at the ribose-NAI adduct to form a truncated product at the primer extension stage (30). The resulting band intensity of an unpaired nucleotide is higher than a paired one by PAGE analysis of the radiolabeled cDNA fragments.

A 140-nt RNA clone of SMN2 exon 7 and adjacent regions was used as a template. This 140-nt RNA was imbedded within a stabilizing cassette for in vitro SHAPE (30). The in vitro SHAPE-directed modeling revealed the following: (i) a stem-loop structure (loop 2, Fig. 4 A and B) at the 5′-ss of exon 7 that was previously reported in literature as TSL2 (17); (ii) a secondary structure at the 3′-ss of exon 7 consisting of two bulges and a loop (Fig. 4 A and B), designated previously as TSL1 (17); and (iii) a distal stem-loop structure on intron 7 (SI Appendix, Fig. S2 for full-length SHAPE profile). Addition of SMN-C2 (50 μM) did not change the overall secondary structure of exon 7 as determined by in vitro SHAPE (Fig. 4A, lane 2). Importantly, no band was observed for the sequence complementary to the AGGAAG binding motif in the in vitro SHAPE analysis, suggesting that binding of SMN-C2 to AGGAAG does not displace the complementary strand. However, the intensity of the bands corresponding to three nucleotides in loop 1 and bulge 2 increased compared with DMSO control (Fig. 4A), suggesting these nucleotides adopt a more flexible conformation in the presence of SMN-C2.

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Fig. 4.

(A) In vitro SHAPE experiment with NAI and a 140-nt-long RNA template containing exon 7. 1, DMSO; 2, SMN-C2 (50 μM); 3, SMN-C2 (5 μM); 4, lacking of NAI; 5∼8, ladders generated by addition of ddATP, ddTTP, ddCTP, and ddGTP during primer extension. PAGE was carried out on a TBE–urea sequencing gel at 60 W for 3 h. Red asterisks indicate increased band intensity with 50 μM SMN-C2. See SI Appendix for the full sequence of the RNA template. (B) In vitro and in-cell SHAPE-directed modeling of exon 7 and adjacent regions. For in vitro RNA model, SHAPE stabilizing cassette (orange) and nucleotides 1∼19 (blue) are shown in sketch. For in-cell RNA model, nucleotide numbering is aligned with in vitro SHAPE template. Nucleotides 1∼18 and 120∼140 were omitted. Significant reactivity changes are indicated in red and green asterisks for in vitro and in-cell SHAPE, respectively. The secondary structures that were previously named TSL1 and TSL2 (17) are enclosed in blue boxes. (C) Differential in-cell SHAPE reactivity in SMN2 minigene-transfected 293T cells for 10 μM SMN-C2 and DMSO in TSL1. SHAPE reactivity in single-nucleotide resolution and its SD were calculated by ShapeMapper software (32). Green asterisks indicate significant SHAPE reactivity change induced by 10 μM SMN-C2.

In vitro RNA secondary structure can be different from that in a cellular context primarily because of the equilibrium between naked RNA and RNA/RNA-binding protein complexes (31). Therefore, an in-cell SHAPE-mutation profiling (MaP) experiment was carried out to investigate structural differences in exon 7 upon treatment with SMN-C2. First, 293T cells were transfected with an SMN2 minigene to enrich the target RNA. The cells were subsequently treated with SMN-C2, a less active analog, SMN-C2-Ac (SI Appendix, Fig. S3) or DMSO, and NAI. Total RNA was then reverse transcribed with a gene-specific primer. Replacing Mg²⁺ with Mn²⁺ in the buffer system results in an error-prone reverse transcription at the NAI-addition position, creating mutations (32). Finally, a region of 276 nt containing exon 7 was amplified by PCR with high fidelity for sequencing analysis. Analogous to in vitro SHAPE, a high mutation rate or high SHAPE reactivity translates to an unpaired nucleotide or a more flexible conformation [see SI Appendix, Fig. S4, for SHAPE reactivity and mutation rate calculated by ShapeMapper software (32) at single-nucleotide resolution].

The secondary structure within this 276-nt amplicon was predicted by matching the SHAPE reactivity for each nucleotide with a software package, SuperFold (SI Appendix, Figs. S5 and S6) (32). Specifically, the TSL2 region remains unchanged in the in-cell model but TSL1 undergoes some base pair rearrangement around bulge 2 and loop 1 (Fig. 4B). RNA-binding proteins, such as Tra2β1 (15) and hnRNP A1 (14), are known to bind to subsequences within TSL1 and may contribute to the base pair rearrangement. Despite this arrangement, the base pairing between AGGAAG and polypyrimidine tract is retained in the cellular context (Fig. 4B). Like the in vitro SHAPE experiment, treatment with 10 μM SMN-C2 results in significant SHAPE reactivity changes only in TSL1, but not TSL2 (SI Appendix, Fig. S7). An increase of SHAPE reactivity in position 34 and a decrease in position 41 near the AGGAAG binding site, compared with DMSO, indicates a more flexible conformation in position 34 and a more constrained one in position 41 (Fig. 4C). SHAPE reactivity of a less active analog, SMN-C2-Ac, resembles that of DMSO at 10 μM and does not show significant changes in positions 34 and 41 (Dataset S1). In the 276-nt sequence, there is only one other secondary structure upstream of TSL1 which showed significant SHAPE reactivity changes (SI Appendix, Fig. S7). Thus, under both in vitro and in-cell conditions, the base-paired SMN-C2–binding site AGGAAG on exon 7 and its complementary sequence in the polypyrimidine tract on intron 6 forms a stem-loop structure, TSL1, at the junction of intron 6 and exon 7. SMN-C2 treatment only affected the SHAPE reactivity of some unpaired nucleotides on TSL1 but not TSL2 or other areas on exon 7. These changes revealed by SHAPE indicated that SMN-C2 does not disrupt the secondary structure of TSL1, but tunes the conformation of the nucleotides in the bulge and loop of TSL1.

SMN-C2, SMN2 Pre-mRNA Exon 7, and FUBP1 Form a Ternary Structure

The interaction of SMN-C2/C3 with its binding motif, AGGAAG, does not explain the selectivity of SMN-C3 as a pre-mRNA splicing modulator. There are more than 15 sequences of AGGAAG in the SMN2 gene alone, in addition to a larger number within the whole human genome. To look for additional cofactors that may contribute to selectivity, a proteomic analysis was performed by photo–cross-linking cell lysates in the presence of 2 μM biotin-diazirine bifunctional probe, SMN-C2-BD, followed by immunoblotting for biotinylated proteins (33, 34). After ammonium sulfate fractionation, the cell lysate was further resolved by two-dimensional SDS/PAGE. Western blots with anti-biotin antibody revealed two associated proteins, FUBP1 and hnRNP A1 (SI Appendix, Fig. S8). FUBP1 and hnRNP A1 are both known to play a role in pre-mRNA splicing. To confirm specificity for SMN-C2/C3, a protein pull-down was carried out using SMN-C2-BD with or without unlabeled SMN-C3 competitor (Fig. 5A). FUBP1 was consistently competed with SMN-C3, whereas hnRNP A1 only showed a basal signal level compared with 1% of the input control (Fig. 5A). The cross-linking of hnRNP A1 likely results from the high abundance of this protein in the nucleus. This result suggests that FUBP1 is a potential protein target for SMN-C3. The pull-down of FUBP1 is dependent on endogenous RNA as evidenced by a reduced FUBP1 signal when treating the protein lysate with a mixture of RNase A/T1 (Fig. 5A).

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Fig. 5.

(A) Biotin–streptavidin pull-down with SMN-C2-BD (2 μM)–cross-linked cell lysate visualized by Western blots with anti-FUBP1 and anti-hnRNP A1 antibody. The 20 or 80 μM SMN-C3 was used as a competitor. RNase A (10 µg/mL)/T1 (25 U/mL) mix was used to digest the endogenous RNA. The quantification of bands in Western blots was by the ratio of pull-down signal over 1% input signal in the same blot (n = 2). (B) Cellular thermal shift assay (CETSA) with THP-1 cells and a serial dilution of SMN-C3. The dose–response was observed optimal at 76 °C. FUBP1 that remained in the supernatant was visualized by Western blot with anti-FUBP1 antibody, and the bands were integrated by ImageStudio (n = 2). (C) Fluorescence polarization with SMN-C2 (200 nM), purified recombinant FUBP1 (20 μM) or hnRNP A1 (20 µM) and a 15-mer oligo-4 (used in Fig. 2C) or a 120-nt RNA that contains exon 7 (used in Fig. 3B). (D) EMSA with 10 pmol of 3′-biotin–labeled RNA (500 nt containing exon 7) and a serial dilution of SMN-C3. The gel was visualized by Northern blot with streptavidin–HRP (n = 2). (E) Dose–response of SMN-C3 (24 h) for end-point RT-PCR of FUBP1/KHSRP dual knockdown in SMN2 minigene-transfected 293T cells compared with a random siRNA control. FL SMN and Δ7 SMN were amplified by PCR with minigene vector-specific primers and resolved in a denaturing TBE–urea PAGE.

To confirm binding of SMN-C3 to FUBP1, a cellular thermal shift assay (CETSA) (Fig. 5B) was conducted (35). In CETSA, small molecules bind and protect their specific cellular protein targets from unfolding and aggregation when the cell suspension is heated (36). THP-1 cells were treated with SMN-C3 at various concentrations or DMSO control and heated at 76 °C. After cell lysis, cell debris and aggregated protein were removed by centrifugation. The amount of FUBP1 in the supernatant increased 58% in the presence of 25 μM SMN-C3 compared with DMSO control and was dose dependent as evidenced by Western blot with an anti-FUBP1 antibody (Fig. 5B). These data suggest SMN-C3 and FUBP1 interact in live cells.

Recombinant FUBP1 was then expressed and purified to further probe its interaction with SMN-C2 in vitro. A fluorescence polarization experiment using recombinant FUBP1 and SMN-C2 (Fig. 5C) revealed no apparent binding between up to 20 μM FUBP1 and 200 nM SMN-C2. In contrast, FUBP1 induced higher polarization for SMN-C2 in the presence of either a 15-mer containing the AGGAAG binding motif (oligo-4) or a 120-nt RNA covering exon 7 and adjacent regions (Fig. 5C). This result suggests a ternary complex is formed by FUBP1, SMN-C2, and exon 7. To compare, 20 µM hnRNP A1 did not induce a significant polarization compared with FUBP1 in the presence of oligo-4, but demonstrated a similar polarization increase with the longer 120-nt RNA. This is likely due to the fact the longer sequence covers both SMN-C2 and multiple hnRNP A1 binding sites (16). To confirm the ternary interaction, a 500-nt RNA covering exon 7 was in vitro transcribed, 3′-labeled with biotin, and subjected to an electrophoretic mobility shift assay (EMSA) visualized by Northern blot with streptavidin–HRP conjugate. SMN-C3 increased the bound fraction of the RNA (10 pmol) in the presence of a low concentration of recombinant FUBP1 (120 nM) in a dose–response manner (Fig. 5D). At this concentration of FUBP1, no protein-bound RNA was observed in EMSA without SMN-C3. Taken together, we conclude that SMN-C3 increases the binding affinity of FUBP1 and exon 7 by forming a ternary structure.

FUBP1 and KHSRP Are SMN2 Splicing Activators

Next, the role of FUBP1 in SMN2 splicing was investigated. FUBP1 has four KH domains that can bind either single-stranded DNA or RNA. The optimal recognition site of multiple KH-domain protein can be long and secondary structure dependent. For example, a multi-KH domain protein, vigilin, has a 75-nt-long recognition sequence (37). In the literature, FUBP1 has been reported as both a pre-mRNA splicing activator and repressor (38⇓–40). In the case of triadin exon 10 splicing, FUBP1 binds to an AU-rich motif downstream of a stem-loop structure at the 3′-ss and inhibits the second step of splicing, that is, nucleophilic attack of the exon 3′-OH on the 3′-splice site to release the lariat (40). Due to the high homology (61%) of FUBP1 and KH-type splicing regulatory protein (KHSRP) (SI Appendix, Fig. S9), especially at the conserved RNA binding KH domains, we hypothesized that the two proteins likely play a similar role in modulating SMN2 splicing by SMN-C3. Indeed, photo–cross-linking pull-down experiments confirmed that SMN-C2-BD can also specifically bind to KHSRP and is competed by SMN-C3 in a dose–response manner as analyzed by Western blotting with an anti-KHSRP antibody (SI Appendix, Fig. S10). KHSRP also contains four KH domains and has been shown to regulate pre-mRNA splicing through cooperation with other splicing regulatory proteins (41, 42). The third and fourth KH domains of KHSRP have been shown to independently interact with different regions of the AU-rich elements and recognize a broad set of mRNAs (43⇓–45).

To further explore their roles, we knocked down FUBP1 and KHSRP individually or in combination with siRNAs in SMN2 minigene-transfected 293T cells. The ratio of full-length SMN (FL SMN) to that which lacks exon 7 (Δ7 SMN) was analyzed by end-point RT-PCR with a pair of minigene vector-specific primers. Although FUBP1 and KHSRP alone did not significantly affect SMN2 splicing (SI Appendix, Fig. S11), dual knockdown of both genes decreased the basal FL SMN mRNA level and shifted the dose–response curve of SMN-C3 (Fig. 5E). In summary, FUBP1 and KHSRP are both activators for exon 7 splicing.