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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
Chem Biol. Author manuscript; available in PMC 2010 May 29.
Published in final edited form as:
Chem Biol. 2009 May 29; 16(5): 557–566.
doi:  10.1016/j.chembiol.2009.03.009
PMCID: PMC2759301
NIHMSID: NIHMS107521

Structural basis for stabilization of the tau pre-mRNA splicing regulatory element by novatrone (mitoxantrone)

Summary

Some familial neurodegenerative diseases are associated with mutations that destabilize a putative stem-loop structure within an intronic region of the tau pre-mRNA and alter the production of tau protein isoforms by alternative splicing. Since stabilization of the stem loop reverses the splicing pattern associated with neurodegeneration, small molecules that stabilize this stem loop would provide new ways to dissect the mechanism of neurodegeneration and treat tauopathies. The anti-cancer drug mitoxantrone was recently identified in a high throughput screen to stabilize the tau pre-mRNA stem loop. Here we report the solution structure of the tau mRNA-mitoxantrone complex, validated by the structure-activity relationship of existing mitoxantrone analogs. The structure describes the molecular basis for their interaction with RNA and provides a rational basis to optimize the activity of this new class of RNA-binding molecules.

Keywords: Tauopathies, Targeting RNA, RNA-small molecule, NMR

Introduction

The microtubule-associated protein tau is expressed predominantly in neurons where it promotes microtubule assembly, reduces microtubule instability, and plays a role in maintaining neuronal integrity and axonal transport (Hirokawa, 1994). Neurofibrillary lesions composed of this protein constitute a defining neuropathological feature of Alzhermer's disease (Spillantini et al., 1998). Filamentous tau protein deposits are also found in other neurodegenerative diseases, many of which are frontotemporal dementias or movement disorders that have been subsumed under the heading of “Pick complex” (Kertesz et al., 1998; Spillantini et al., 1998). Such neurodegenerative disorders associated with tau protein are referred to as tauopathies (Lee et al., 2001). The tau gene consists of 16 exons that are alternatively spliced to generate six major tau isoforms in a developmentally-regulated manner (Andreadis et al., 1992; Goedert et al., 1989). They differ from each other by the presence or absence of 29- or 58-amino acid inserts located in the amino-terminal half of the protein, and a 31-amino acid repeat located in the carboxyl-terminal half. Inclusion or exclusion of the latter by alternative splicing of exon 10 produces tau isoforms with either three (when exon 10 is missing) or four (when exon 10 is present) microtubule-binding repeats (Goode et al., 1994; Gustke et al., 1994); these isoforms are known as 3R and 4R tau, respectively (Goedert et al., 1989). In the normal adult human brain, the relative ratio of 4R to 3R tau is generally close to unity; overproduction of 4R tau appears to be sufficient to lead to dementia.

Mutations in the tau gene cause familial fronto-temporal dementia and parkinsonism linked to chromosome 17 (FTDP-17) (Clark et al., 1998; Dumanchin et al., 1998; Goedert et al., 1999; Hutton et al., 1998; Morris et al., 1999; Poorkaj et al., 1998; Spillantini et al., 1998). Tau mutations within coding regions cluster in the microtubule-binding repeats and alter the ability of tau to interact with microtubules. This occurrence initiates an unclear set of events that lead to the formation of tau filaments in the neurons of patients affected with a variety of neurodegenerative diseases (Dayanandan et al., 1999; Hasegawa et al., 1998; Hong et al., 1998). However, other mutations causing these diseases are located within introns or are synonymous, and cluster near the splice-donor site of the intron following exon 10. Significantly, four mutations at positions +3, +13, +14, and +16 of the intron (with the first nucleotide of the splice-donor site taken as +1, Figure 1A) (Goedert et al., 1999; Hutton et al., 1998; Morris et al., 1999; Spillantini et al., 1998) are found within a putative RNA stem loop that partially masks the splice donor itself. The surprising discovery of these mutations led to the suggestion that the stability of the RNA stem loop affects alternative splicing of exon 10, and that the destabilization of the stem-loop itself alters the balance between different tau isoforms and ultimately leads to neurodegeneration in patients carrying mutations within this RNA structure (Hutton et al., 1998).

Figure 1Figure 1
A: Sequences and secondary structures of wild type and mutant human tau exon 10 splicing regulatory elements. Destabilizing mutations at the +3, +13, +14 and +16 (in red) positions are linked to familial neurodegenerative diseases and increase inclusion ...

Several studies demonstrated that the intronic mutations found in frontotemporal dementia and parkinsonism decrease the thermodynamic stability of the RNA stem-loop in vitro (Donahue et al., 2006; Hutton et al., 1998; Varani et al., 1999). It was thus postulated that destabilization of this RNA structure partially unmasks the splice site, leading to increase inclusion of exon 10 and therefore to increased production of the 4R tau isoform compared to the 3R isoform (Hutton et al., 1998). More recently, Donahue and colleagues have demonstrated the reverse phenomenon as well; namely, mutations that stabilize the RNA stem loop in vitro substantially decrease exon 10 inclusion (and therefore alter the 4R/3R ratio) in an otherwise wild type tau minigene (Donahue et al., 2006). Evidence supporting the formation of the tau splicing regulatory element (SRE) stem loop structure in vivo was provided by the anti-correlation between the stability of the stem loop and the efficiency of antisense inhibition of exon 10 inclusion (Donahue et al., 2006). These experimental findings and the resulting model suggest that small molecules that stabilize the tau SRE RNA stem loop would provide very useful tools to probe the mechanism of neurodegeneration and perhaps new therapeutic avenues to treat these diseases.

The aminoglycoside antibiotic neomycin, a naturally-occurring RNA-binding molecule, was found to bind the tau SRE RNA major groove and to stabilize the RNA structure (Varani et al., 2000). However, this class of molecules lacks the selectivity that would be necessary for therapeutic or even investigative applications. A high-throughput screening assay for compounds that bind to and stabilize the tau SRE RNA identified the anti-cancer drug mitoxantrone from a library of ~110,000 molecules (MTX, Figure 2, denoted by code number LDN-13978 in (Donahue et al., 2007) and revealed as MTX for the first time here) (Donahue et al., 2007). MTX is used in clinical practice as a drug against breast cancer, acute leukemia and non-Hodgkin's lymphoma (Hagemeister et al., 2005; Tsavaris et al., 2005); it has also been approved in the U.S. for the treatment of patients with progressive multiple sclerosis (Gonsette, 2003). The lead-like activity of MTX in binding to the tau SRE (EC50~700 nM) and its current multiple uses in clinical practice makes it an attractive lead to discover new small molecule regulators of tau mRNA splicing, but its activity must first be improved.

Figure 2
Structure of mitoxantrone (abbreviated as MTX in the text).

Here we report the NMR structure of the complex between MTX and the region of SRE that constitutes the defining structural characteristic of this RNA. MTX intercalates into the partially open bulge region of tau SRE RNA with the side chains forming sequence-specific hydrogen-bonds with the nearby RNA bases and electrostatic interactions with the phosphate backbone. The structure explains the structure-activity relationship (SAR) of a series of MTX analogs and provides the structural basis for the design of more potent and specific regulators of tau pre-mRNA splicing.

Results

Formation and stoichiometry of the MTX-Tau RNA complex

In order to characterize the MTX-RNA complex, we titrated tau SRE RNA with MTX at concentrations suitable for NMR investigations (0.1-1 mM) and recorded spectra for the imino proton region, where the RNA resonances corresponding to Watson-Crick paired nucleotides are well resolved (Figure 3). From the bottom, the spectra correspond to MTX:RNA ratios of: 0.0, 0.5, 1.0, 1.5, 2.0, 3.0. The chemical shifts of the imino resonances in the upper helical region (G+1, U+2 and G+3) were not changed, but only broadened during the titration. For resonances near the bulge loop (G-1 and U0), we observed a decrease in the intensity of the peak corresponding to the free RNA signal as the MTX concentration increases; at the same time, a new set of peaks appeared at distinct positions, that were later assigned to the MTX-bound form of the RNA. The peaks corresponding to the free form of the RNA disappear completely only when the ratio of MTX:RNA is > 2.0 (Figure 3). Multiple set of imino resonances appear when the ratio of MTX:RNA is larger than 1.5; these new peaks must arise from the binding of multiple copies of MTX to the RNA when more than one molecule of MTX per RNA is present. The observation of distinct peaks in slow exchange on the NMR time scale corresponding to free and bound RNA indicates that MTX binds tightly (Kd < 1 μM) at or near the RNA bulge region, consistent with an EC50 of about 700 nM previously reported (Donahue et al., 2007).

Figure 3
NMR titration of tau SRE RNA with increasing concentrations of MTX recorded at 4 °C. From the bottom, spectra correspond to the following ratios of MTX:RNA: 0.0, 0.5, 1.0, 1.5, 2.0, 3.0. The label in the bottom spectrum corresponds to the assignments ...

The observation of two peaks with comparable intensity corresponding to the free and bound RNA resonances at MTX:RNA ratios of about 1 (see e.g. U0, Figure 3) has two possible explanations. MTX could binds as a dimer; or it could form more than one independent interaction with the RNA. The observation of additional peaks in the 1D spectra at ratios above 1:1 strongly suggests that when there are two binding sites, and additional MTX molecule(s) binds with comparable affinity to a second site within the RNA. On the other hand, the reported self-association constant (12400 ±4000 L mol-1) of MTX in aqueous solutions (Davies et al., 2001) and the 1D titration experiment, MTX should be dimeric at NMR concentrations.

We confirmed the dimeric form of free MTX in solution by observing intermolecular NOEs between the H2/3 resonances of one MTX and the H6/7 resonances as well as the side chain resonances (H11/15 and H12/16) of a second copy of the molecule (Davies et al., 2001). These NOE interactions are only possible if the two MTX molecules are arranged in a reversed stacking conformation sitting on each other in a head-to-tail orientation. However, in the 2D NOESY spectra of the complex, we failed to observe any intermolecular NOEs involving two MTX molecules between the H2/3 and H6/7 of MTXs; we also failed to observe NOEs between the H2/3 aromatic resonances and the same side chain resonances (H11/15 and H12/16) that were observed instead for free MTX. All these NOEs would be expected for a dimeric structure and indeed were observed for the free MTX dimer. As a control, we readily observed NOEs between the H6/7 resonances and the side chain H11/15 and H12/16 in the RNA complex; in fact, these NOEs were much stronger compared to the free MTX dimeric structure, most likely because the conformation of the side chain is rigidified in the complex (see below) leading to more efficient NOE transfer. If MTX bound to RNA as a dimer, we would expect to see much stronger NOEs than we observe in the RNA-free MTX dimer between the aromatic H2/3 and H6/7, as well as the side chain H11/15 and H12/16 resonances. Altogether, these results are inconsistent with the binding of MTX as an antiparallel dimer to the tau SRE RNA. Although it is possible that MTX dimerizes forming a parallel stacking arrangement (in this case, intra- and intermolecular NOEs would be very difficult to distinguish), this is highly unlikely because of the resulting electrostatic repulsion. Thus, the titration experiments are better explained by a model where multiple molecules of MTX bind to different locations on the RNA.

Consistent with the conclusion that two MTX molecules bind to the SRE RNA in distinct locations, we observed two distinct sets of intermolecular NOEs when analyzing NOESY spectra. A more numerous and more intense set of intermolecular contacts mapped to the RNA bulge; a second, much less numerous set, corresponded to the minor groove near the upper helix. Binding of MTX to the minor groove of the RNA does not lead to significant chemical shift changes in the RNA, suggesting that this interaction is only superficial and perhaps corresponds to the non-specific electrostatic interactions between the highly cationic MTX and the negatively charged phosphate backbone.

The binding sites of MTX on tau SRE RNA was mapped by chemical shift perturbation analysis as well (Supplemental Tables S1 and S2). Modest chemical shift changes were observed in the upper stem-loop, never larger than 0.12 ppm, where the NOE data indicate that a second molecule of MTX binds. Much more significant chemical shift changes were observed near the bulge region (A-2). The chemical shift change for the H8 of A-2 is 0.16 ppm, the imino resonance of G+17 was not observed in the free form due to exchange with solvent, but readily observed in the complex with MTX. The largest chemical shift change is observed for the imino of G-1 (0.62ppm). The observation of the G+17 imino in the complex, but not in the free form, indicates that this resonance is shielded from solvent in the complex either because the bulge region is stabilized by the binding of MTX, because MTX itself shields this hydrogen, or both. Thus, all available NMR results are consistent with the conclusion that MTX binds to tau SRE RNA in two locations. A first site corresponds to the region near the bulge; binding of MTX here induces significant chemical shift changes, indicating an intimate interaction, and is likely responsible for the stabilization of the stem-loop. A second complex forms in the minor groove close to the upper helical region; this second complex does not induce large conformational and chemical shift changes in the RNA.

To further establish the physical chemical characteristics of the anthraquinone chromophore in the complex, we employed UV/Vis and fluorescence spectroscopy. The experiment differs from the NMR titration because RNA is now titrated into a fixed amount of MTX (10 μM) and not the reverse. The maxima in absorbance of free MTX were located at 608.4 and 659.4 nm (Figure 4). In the first stage of the titration (RNA/MTX<0.1), the intensities of these two absorbance peaks decreased sharply; in the second stage, the intensities of the two new peaks (626 and 681 nm, Figure 4) increased gradually until saturation was reached at a RNA/MTX ratio ≥1.0. Both absorption peaks shift to the red by about 18 and 22 nm, respectively; such bathochromic effects are commonly observed for DNA intercalators (Li et al., 2005). Comparable bathochromic effects were also observed in fluorescence titration experiments (Supplementary Figure S1). The UV/Vis and fluorescence data indicates that MTX binds to tau SRE RNA by partially intercalating or at least forming clear stacking interaction with base ring systems. The two-stage binding mode (first, the free MTX intensity decrease quickly; second, the bound MTX absorption increases gradually) suggests that there are multiple MTX binding sites. The sharp initial decrease in absorbance may be due to RNA condensation caused by charge interactions when multiple MTX molecules, each with 2 positive charges, bind to one RNA molecule and neutralize the negative charges in the phosphate backbone (Kapuscinski et al., 1984; Kapuscinski et al., 1986). In the second stage, as the titration continues, MTX molecules re-distribute among increasing amount of available free RNAs, until eventually most find their preferred binding site are saturated.

Figure 4
Titration of tau SRE RNA with MTX monitored by UV/Vis spectroscopy. The experiments were carried out in buffer solution (10 mM, pH 6.0) containing 10 μM MTX in the presence of different RNA concentrations; RNA:MTX are as follows (a) 0.0; (b) 0.02; ...

The NMR results indicate that there are multiple binding sites for MTX on the RNA. Sharp NMR spectra were obtained at MTX:RNA ratios near 1.0. Under this condition, we observed two sets of resonances near the lower bulge corresponding to the free and bound forms of the RNA, with comparable intensities. Despite the increased complexity in the spectra at 1:1 ratio, the signals corresponding to bound and free components can be distinguished unambiguously at high field (Supplementary Figures S2 and S3), particularly when using partially deuterated nucleotides or 13C/15N labeled RNA in 3-dimensional spectra. Increasing the MTX:RNA ratio (e.g. at ratios of 1.5:1 and 2:1) to saturate binding at the bulge, caused considerable broadening in the aromatic and ribose regions of the NMR spectrum, presumably due to conformational exchange involving the other weaker sites (e.g. the site near the apical loop). Furthermore, there are not sufficient intermolecular NOEs to define the minor groove binding conformation. Thus, we chose the 1:1 ratio as the best condition to determine the structure of the complex; under these conditions, we could only study the most favorable and specific binding site on tau SRE near the bulge immediately adjacent to the splice junction.

NMR Spectral assignments

Spectral assignments were obtained in a straightforward manner starting from the available assignments for the free RNA (supplemental material Figures S2). Some assignments were obtained by observing exchange peaks near the diagonal, others simply by following the conventional procedures that connected both the free and bound RNA peaks (in the MTX binding site) to a single set of peaks corresponding to resonances that do not shift in the complex (supplemental material Figures S3 and S4). Assignments are reported in Supplementaty Table S1. Critical to structure determination was the observation of intermolecular NOEs between the MTX side chains and RNA resonances; some are shown in the 2D NOESY spectrum of Figure 5A; their complete list is reported in Table 2. At 25 °C, we observed numerous intermolecular NOEs between MTX side chains and the RNA, but none was observed with the MTX ring H2/3 resonances. As a result, the location of the anthraquinone chromophore relative to the RNA could not at first be defined unambiguously.

Figure 5Figure 5
Two-dimensional NOESY experiment recorded at 4 °C on the complex between MTX and tau SRE RNA. A: Regions of the spectrum corresponding to intra-(MTX) and inter-molecular (MTX and RNA) NOEs are boxed and labeled; B: The intermolecular NOE observed ...
Table 2
Intermolecular distance constraints used to calculate the structure of the complex based on the intermolecular NOEs observed between MTX and tau SRE RNA.

The data at 25 °C are consistent both with the chromophore being inserted into the bulge region and stacked on the RNA bases, and with the chromophore pointing away from the bulge region with only the MTX side chains interacting with the RNA. We deemed this second conformation unrealistic because of the results of the UV/VIS experiments that suggest partial intercalative binding, and because it would expose the hydrophobic chromophore to solvent. However, we sought to establish the correctness of the first conformation unambiguously. Although the peaks become broader at lower temperature, the NOESY spectra could still be assigned at 4 °C with the help of partially deuterated RNA (that decreases the linewidth due to less efficient relaxation) and of 13C/15N labeled RNA. At 4 °C, we observed key intermolecular NOEs between MTX H2/3 and the H1′ of G+17 (Figure 5B), which indicates that MTX was stacked on the base of G+17, consistent with the most likely of the two conformations described above.

Structure of the complex between MTX and tau SRE RNA

Structure calculations were carried out using NIH-Xplor and standard protocols (Clore et al., 2003) using a total of 41 intermolecular NOEs (Table 2). From 100 calculated structures, the top 20 were chosen to represent the ensemble of conformations based on their energy and restraint violation statistics. Statistics of data collection and for the converged structure are shown in Table 1. The superposition of the 10 best structures (Figure 6A) shows the helical stem and bulge where MTX binds are very well defined, but the apical loop remains conformationally flexible. The anthraquinone chromophore of MTX stacks on the base of G+17, consistent with the large chemical shift change for its NH, while the positively charged side chains interact with the major groove of the RNA; the unpaired A-2 nucleotide protrudes outwards into solution (Figure 6B). The length and helical axis of the RNA in the complex were almost unaltered compared to the free RNA structure (Varani et al., 1999).

Figure 6Figure 6
Structure of the tau SRE RNA– MTX complex represented with MOLMOL (Koradi et al., 1996). A: Superposition of the 10 best calculated structures (heavy atoms only). The RNA is shown in blue, and MTX is shown in red. B: The best calculated structure; ...
Table 1
Experimental constraints and structure statistics

Close-up views of the interaction between MTX and RNA taken from the best-scoring structure are shown in Figure 7. Consistent with the UV/VIS results and the protection of the G+17 imino from exchange with solvent, the chromophore of MTX is stacked between the base pairs of G+17–C-3 and C+16–G-1, forming partial stacking interaction with both base pairs. This interaction may be the dominant driving force for the formation of the complex. In addition, a well-ordered network of hydrogen bonds and electrostatic contacts is observed between the MTX side chains and Tau RNA. The distances of all relevant hydrogen bonding and electrostatic contacts are shown in Table 3. These additional interactions increase both the affinity of MTX binding and presumably the stability of the tau SRE RNA structure (Donahue et al., 2007). Although the predicted pKa (http://ibmlc2.chem.uga.edu/sparc/) showed that the two amines (N3 and N4) should be protonated at the experimental pH (pH ~6), we did not include any specific electrostatic term during structure calculation or refinement to avoid introducing any unintended bias in the structure determination process. Thus, the additional electrostatic and hydrogen-bonding interactions observed in the complex, such as the electrostatic interaction between N4 of MTX and phosphate of G+17 and hydrogen bond between O6 of MTX and O4 of U+18 (Figure 7), are derived from the experimental intermolecular NOE constraints (Table 2), although not directly verified from experimental data (such as RNA-MTX scalar couplings).

Figure 7
Close-up view of the tau SRE RNA-MTX interaction. The carbon atoms of MTX are shown in yellow and the RNA in green with Pymol (DeLano, 2002). Some key inter molecular NOEs between the H1′ of G+17 and H2/3 of MTX are shown in blue dashed line.
Table 3
The distance of hydrogen bond and electrostatic contacts observed between MTX and tau SRE RNA

In the complex, the N3 and N4 amines in the MTX side chains form hydrogen bonds with the O6 of the G-1 and N7 of G+17 bases, respectively (Figure 7). The MTX N4 amine can form hydrogen bonds with the base of G+17, and at the same time electrostatic contacts with the phosphate of G+17. One hydroxyl group (O6) of MTX is in hydrogen bonding contact with the O4 of U+18, while another hydroxyl group (O5) is hydrogen bonded with the phosphate of G-1. Although the hydrogen bond between the imino N2 group and the phosphate of C+16 is not likely to provide specificity for a particular RNA sequence, the hydrogen bond between the imino group N1 and O6 of G-1 is specific for the G-1 base. The two hydroxyl groups in the chromophore can also form hydrogen bonds with the O2′ of C+16 and A-2, as supported by weak NOE interaction with A-2, providing selectivity over DNA and a distinct mode of binding. Although the intercalation of the anthraquinone chromophore into RNA is not believed to be sequence specific, the hydrogen bonds between the side-chain N3/4 of MTX and the G-1 and G+17 bases can provide specificity for adjacent GC base pairs in a structural context permissive to binding, as generated by the bulged A-2 nucleotide.

Structural-activity Relationship (SAR) for MTX analogues

To explore the SAR of MTX binding to tau SRE RNA, a series of MTX analogs (Table 4) were obtained from NCI and tested for their ability to stabilize the tau RNA stem loop, using a FRET assay that employs a tau RNA oligonucleotide with a fluorophore on the 3′ end and a quencher on the 5′ end of the stem-loop (Donahue et al., 2007). Satisfactorily, the NMR structure of the complex is for the most part consistent with the results of Table 4. In the structure, the hydrophilic groups of MTX (O1/2, O5/6 and N3/4) are involved in a well-defined hydrogen bonding network, as described above. All observable activity is eliminated when these hydrophilic groups are substituted with non polar groups (compound 4 in Table 4). The N3/4 amine groups form hydrogen bonds with the guanine bases from the two GC base pairs abetting the unpaired A-2 nucleotide. Even a change in the N3/4 amine to ether groups, as in 1, leads to a >70-fold decrease in the EC50. The O1/2 hydroxyl groups involved in hydrogen bonds with the hydroxyl O2′ of C+16 and A-2 contribute less to RNA binding, because their substitution to hydrogen only reduces the EC50 about 6 times (3). To our surprise, modifying the O5/6 hydroxyl groups to the better hydrogen bonding donor amines (2), did not increase binding, but instead decreased the EC50 more than 7,000 times.

Table 4
SAR for MTX analogs

Discussion

Modulating RNA with small molecule drugs would provide the opportunity to regulate the function of RNA in chronic and infectious disease conditions (Thomas et al., 2008). An inviting target for small molecules that bind to RNA is the splicing regulatory element within the tau pre-mRNA that controls the ratio of the two major tau protein isoforms. Mutations that destabilize the tau stem loop structure increase inclusion of exon 10 and are associated with neurodegenerative tauopathies. Since mutations that stabilize the RNA can reduce exon 10 inclusion, stabilization of the stem-loop with a small molecule would also reduce exon 10 inclusion. Thus, if we were able to regulate alternative splicing of tau exon 10, we would be better able to define its precise role in a variety of neurodegenerative disease conditions and perhaps explore new therapeutic options as well.

Unfortunately, there is no precedent for the successful clinical or even pre-clinical development of RNA-binding small molecules outside of the ribosome, a mainstay target of antibacterial treatment. Among well-known antibacterials targeting the ribosome are aminoglycoside antibiotics such as neomycin. In a previous study, we showed that this drug binds in the major groove of the tau SRE RNA. However, these molecules have poor specificity and are synthetically challenging; they have proven to be unattractive leads to the development of new RNA-binding drugs because of the lack of selectivity that would be necessary for therapeutic efficacy. In the search for new small molecules that interfere with tau pre-mRNA splicing, the anticancer drug mitoxanthone (MTX) was found to bind to the tau SRE and stabilize its structure in a screen of over 100,000 molecules (Donahue et al., 2007). Here we report the structure of its complex with tau SRE RNA and a molecular rationale for its binding activity.

The activity of MTX as an anticancer drug has led to some interest in understanding the molecular basis of its interaction with DNA, but so far these studies have been based on molecular modeling (Islam et al., 1985; Kostjukov et al., 2007) and not a formal structure. In addition to providing a structural basis to further optimizing the affinity and specificity of MTX for tau pre-mRNA stem-loop structure, the present structure also sheds more insight into the general mechanism of the interaction of MTX with nucleic acids in general.

MTX binds the bulge region of tau SRE RNA by partially intercalating in the space created by the extruded base A-2 and by forming hydrogen bonds to two adjacent GC base pairs (G-1–C+16 and G+17–C-3) and one uracil residue (U+18) nearby that are specific to these bases. Although free MTX self-associates forming a dimer, a variety of spectroscopic observations indicate that MTX binds to tau SRE RNA as a monomer and has a second, slightly weaker binding site in the minor groove of the RNA near the apical stem-loop. The second binding site is most probably due to superficial electrostatic interactions between the positively charged MTX and the RNA backbone, because chemical shift changes in the RNA upon binding are very small. In contrast to the main complex near the bulge, we cannot establish this second binding conformation due to the paucity of intermolecular NOEs observed for it.

The partial intercalation of the chromophore and electrostatic contacts with the RNA do not lead to sequence-specific binding, but require nonetheless a specific structure, a bulged nucleotide. Moreover, the hydrogen bonds between the MTX side chain amine N3/4 and the base O6 carbonyls of G-1 and N7 of G+17 suggest that MTX prefers to recognize GC base pairs (G-1–C+16 and C-3–G+17). Although these interactions are clearly sequence specific, the flexibility of the side chains allows the positions of the hydrogen-bonding groups to adapt themselves to other RNA sequences and bind to them with comparable effectiveness. However, the structure suggests that by rigidifying these amines in a more constrained structure, it should be possible to optimize hydrogen bonding interactions between MTX and the GC base pairs and thereby generate MTX derivatives that are specific for the tau SRE structure.

In addition to the side chain amines, the hydroxyl group O6 of MTX is in hydrogen-bonding contact with the O4 of U+18, a result well established by one of the strongest intermolecular NOEs observed between the H18 of MTX and the H5 of U+18. This hydrogen-bonding interaction also contributes to the specific recognition of U+18. Therefore, the length of the MTX side chain between O6 and the chromophore may be varied to enforce stronger hydrogen bonds with U+18 to select against other RNAs. We plan to exploit these suggestions in future studies to improve the binding characteristics of MTX derivatives with the tau stem-loop and to generate MTX variants that selectively bind to this structure.

Significance

Modulating RNA structure with small molecule drugs would provide the opportunity to regulate the function of RNA in chronic and infectious disease conditions. An inviting target for small molecules that bind to RNA is the splicing regulatory element within the tau pre-mRNA that controls the ratio of the two major tau protein isoforms. Mutations that destabilize the tau SRE RNA increase inclusion of exon 10 into the mature tau protein and are associated with neurodegenerative tauopathies. Since mutations that stabilize the RNA reduce exon 10 inclusion, stabilization of the stem-loop with small molecules would also reduce exon 10 inclusion and provide a new way to define its precise role in a variety of neurodegenerative disease conditions and to explore new therapeutic options. The anticancer drug mitoxanthone (MTX) was recently identified in a high-throughput screen to stabilize the tau splicing regulatory element RNA. Here we report the structure of tau pre-mRNA in complex with MTX at the bulge region, which identifies a new class of RNA-small molecule interactions. This structure provides a basis to optimize the potency and specificity of MTX analogues and sheds light into the general mechanism of the interaction of MTX with nucleic acids.

Experimental Procedures

RNA preparation

A model oligonucleotide mimic of the wild type tau SRE containing nucleotides from G-5 to C+19 (Figure 1B) was used in all experiments. RNA oligonucleotides were synthesized by in vitro transcription using T7 polymerase and synthetic DNA templates (IDT) with commercially available unlabeled, selectively deuterated and 13C/15N labeled nucleotides (Cambridge Isotope Inc.). The selectively deuterated nucleotides were used to simplified the NMR spectra by only containing H2/H6/H8 and H1′/2″ resonances; all other positions were deuterated. All RNA samples were purified as described (Price et al., 1998). Microdialysis was used to bring the final buffer to 10 mM sodium-phosphate (pH 6.0) with 0.1 mM EDTA. Lyophilized RNAs were re-dissolved in 5% D2O/95% H2O or 100% D2O as needed. Sample used for residual dipolar coupling (RDC) measurements were mixed with Pf1 phage (ASLA Ltd) and dialyzed against the same buffer at a final phage concentration of 12-15 mg/ml. Mitoxantrone (> 97%) was purchased from Sigma and used without further purification.

Data collection and analysis

Most NMR data were acquired on Bruker 500MHz and 600MHz Avance spectrometers equipped with cryo-probes, and a 750 MHz DMX spectrometer equipped with a conventional HCN probe. The D2O NOESY on the complex was collected at Pacific Northwest National Laboratory (PNNL, Richland, WA) on a Varian 800 MHz spectrometer equipped with a cryo-probe. In-Phase-Anti-Phase HSQC spectra (IPAP) (Andersson et al., 1998) for RDC measurement were collected at 500 MHz. Data were processed with nmrPipe (Delaglio et al., 1995) and analyzed with Sparky (Goddard et al.). The spectra for the H2O sample were recorded at 4 °C and those for the spectra in D2O were recorded both at 25 °C and 4 °C.

The assignments for the complex were initiated by standard 2D NOESY walks that were extended from the part of the structure that did not change upon binding to MTX into the section where changes were observed (see also supplemental material Figure S2 and S3); the assignments were facilitated by 3D NOESY-HSQC experiments due to the fact that 13C shifts did not change considerably upon MTX binding (Varani et al., 1996). The bound resonances were further confirmed by observing exchange peaks in 2D NOESY spectra (supplemental material Figure S3); the partial deuterated sample was used to reduce the overlap in the ribose region (supplemental material Figure S4). The complete proton assignments of free and bound tau SRE RNA are listed in supplemental material Table S1 and the assignments of MTX are listed in supplemental material Table S3. Base pairs were established from the imino peask observed in 2D water NOESY collected with Watergate water suppression (Dingley et al., 1998; Pervushin et al., 1998). Dihedral angle restraints were obtained from 2D TOCSY experiments,.as described (Varani et al., 1996), and the C2′-endo constraints for the sugar pucker were used only when both H1′-H2′ and H1′-H3′ coupling are both large.

Structure calculations

NOEs involving non-exchangeable protons were classified as strong (1.8-3.2 Å), medium (2.2-4.2 Å), weak (2.5-5.5 Å) or very weak (3.0-7.0 Å) based on cross-peak intensities at 100 and 200 ms mixing times (2D NOESY experiments) and 120 ms (3D NOESY- HSQC). NOEs involving exchangeable protons were characterized as strong (1.8-3.4 Å), medium (1.8-4.5 Å), weak (1.8-6.0 Å) and very weak (1.8-6.5 Å) based on their NOE cross-peak intensities at 100 and 200 ms mixing times (Chen et al., 2006).

A single extended starting structure was generated with Xplor-NIH (Schwieters et al., 2003) and the structure calculations were carried out by Xplor-NIH with the same strategy by employing the means of conformational database potential (Clore et al., 2003). RDC-derived restraints were added only after the majority of converged structures were found, and only RDCs obtained from well ordered residues devoid of conformational exchange were used in refinement (Leeper et al., 2005).

Optical spectroscopy studies

UV/Vis absorbance spectra were obtained on a Beckman DU-650 spectrophotometer equipped with a quartz micro-colorimetric vessel of 1 cm path length.

Activity measurement of MTX analogues

MTX analogues were obtained from the NCI and were tested for their ability to stabilize the tau RNA stem-loop structure using a fluorescence resonance energy transfer (FRET) assay as previously described (Donahue et al., 2007) in which the 5′ end of the tau RNA oligomer is linked to dabsyl and the 3′ end is linked to fluorescein.

Supplementary Material

01

Acknowledgments

We thank EMSL-PNNL for access to NMR instruments and Dr Nancy Isern (PNNL) for help with data acquisition; Dr Thomas Leeper for assistance with the NMR spectroscopy; Ms Zahra Shajani, Mi-Kyung Lee and Amy Davidson, for help with RNA preparation; Dr Michael F Bardaro for help with RDC data collection; Dr Lev Elson-Schwab for the help with fluorescent data collection. This work was supported in part by grants from NIH.

Footnotes

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