RNA used to be considered a simple and straightforward molecule in cells. The three major classes of RNA, i.e., transfer RNA, ribosomal RNA, and messenger RNA (mRNA), have generally not been thought to be subjected to regulation by signaling pathways, or to have major roles in disease processes. However, a rapidly emerging concept over the past few years is that transcription and other cell signaling pathways are regulated by a diverse array of noncoding RNAs, such as microRNAs, termini-associated RNAs (Han et al., “Promoter-associated RNA Is Required for RNA-directed Transcriptional Gene Silencing in Human Cells,” Proc Natl Acad Sci USA 104:12422-12427 (2007)), and other noncoding RNAs. Additionally, mRNA is no longer viewed as a simple intermediate between DNA and protein, but instead is now known to be subjected to wide range of post-transcriptional processing events, including diverse types of splicing reactions, nonsense-mediated decay, RNA editing, exo- and endonucleolytic degradation, polyadenylation, and deadenylation. Another intriguing aspect of RNA biology is the finding that trinucleotide repeat-containing mRNAs exert specific gain-of-function toxicities associated with their accumulation at certain intracellular sites (Ranum et al., “Myotonic Dystrophy: RNA Pathogenesis Comes Into Focus,” Am. J. Hum. Genet. 74:793 (2004)). In addition to these different regulatory pathways, recent studies indicate that RNAs traffic through different parts of the cell during RNA maturation. For example, nascent RNA transcripts are likely trafficked to specific intracellular sites in the nucleus for processing events, such as splicing, nonsense-mediated decay, or for packaging into transport granules. After nuclear export, some RNAs have been localized to RNA-enriched intracellular structures including RNA granules, stress granules, and processing bodies (P-bodies) (Kiebler et al., “Neuronal RNA Granules: Movers and Makers,” Neuron 51:685-690 (2006)). The diversity of these RNA regulatory mechanisms makes it clear that RNA is regulated by a complex and intricate network of regulatory mechanisms and intracellular structures that have a critical role in gene expression.
RNA regulatory pathways are particularly prominent in neurons. For example, RNA splicing is more highly regulated and is more complex in neurons than in any other cell type (Dredge et al., “The Splice of Life: Alternative Splicing and Neurological Disease,” Nat Rev Neurosci. 2:43 (2001)). Similarly, RNA editing and trinucleotide repeat-containing mRNA diseases are especially prominent in neurons despite the widespread expression of these transcripts (Keegan et al., “Adenosine Deaminases Acting on RNA (ADARs): RNA-editing Enzymes,” Genome Biol. 5:209 (2004)). A recent analysis of 1,328 noncoding RNAs (>200 nt) revealed that 64% were expressed in the brain, many of which had strikingly specific patterns of expression in discrete brain structures (Mercer et al., “Specific Expression of Long Noncoding RNAs in the Mouse Brain,” Proc Natl Acad Sci USA 105:716-721 (2008)).
One form of RNA regulation that has received considerable attention is “local” RNA translation. A compelling argument for a fundamental role for RNA localization in cells was presented in a recent landmark study, in which high-throughput gene-specific in situ hybridization in Drosophila embryos showed that 71% of cellular RNAs exhibit specific and often striking intracellular localizations (Lecuyer et al., “Global Analysis of mRNA Localization Reveals a Prominent Role in Organizing Cellular Architecture and Function,” Cell 131:174-187 (2007)). RNA localization is also a feature of neurons, where mRNAs are enriched in axons and dendrites. Local translation of these mRNAs may have evolved to accommodate the highly spatially polarized nature of neuronal morphology, which typically involves axons and dendrites that can extend distances of tens to thousands of micrometers from the cell body. In local translation, mRNAs are translated directly within dendrites and axons, often within particularly small domains such as a 1-2 μm long dendritic spine or small domain within an axonal growth cone (Steward et al., “Compartmentalized Synthesis and Degradation of Proteins in Neurons,” Neuron 40:347-59 (2003)). RNAs are dynamically transported between RNA granule structures and P-bodies during synaptic stimulation, which likely regulates mRNA translation (Zeitelhofer et al., “Dynamic Interaction Between P-bodies and Transport Ribonucleoprotein Particles in Dendrites of Mature Hippocampal Neurons,” J Neurosci. 28:7555-7562 (2008)). Only a small subset of the total mRNA population is trafficked to axons or dendrites, and it appears that discrete 3′ UTR sequences are required for signal-dependent translation of distinct pools of RNAs (Sutton et al., “Local Translational Control in Dendrites and its Role in Long-term Synaptic Plasticity,” J Neurobiology 64:116-131 (2005)). Local translation bypasses the time-consuming process of propagating a signal to the nucleus, followed by protein synthesis and subsequent transport of the newly synthesized protein to the specific site of receptor activation (Steward et al., “Compartmentalized Synthesis and Degradation of Proteins in Neurons,” Neuron 40:347-359 (2003)). Thus, neurons display complex, time-dependent, and spatially specific regulation of local mRNA translation to accommodate their functional and morphologic demands.
Green fluorescent protein (GFP) has revolutionized biomedical research and biotechnology. As a result of GFP, studies that address the trafficking and processing of proteins in relation to specific intracellular organelles and sites within the cell has become commonplace. Additionally, the roles of organelles and even subdomains within organelles have been elucidated. However, besides the spatiotemporal localization of proteins within cells, GFP has been used to provide information regarding protein-protein interactions, cellular viscosity, and protein degradation (Zhang et al., “Creating New Fluorescent Probes for Cell Biology,” Nature Rev Mol Cell Biol 3:906-918 (2002); Neher et al., “Latent ClpX-Recognition Signals Ensure LexA Destruction After DNA Damage,” Genes & Development 17:1084-1089 (2003)). GFP has also been used in studies of protein folding and has found innumerable uses in biotechnology as a critical component of various cell-based and in vivo assays. GFP and related proteins have been harnessed to generate new classes of FRET probes that report the intracellular localization and concentration of intracellular molecules (Zhang et al., “Creating New Fluorescent Probes for Cell Biology,” Nature Rev Mol Cell Biol 3:906-918 (2002)). Although bioconjugate chemistry methods to make proteins fluorescent have been available for a long time, these approaches require that proteins be modified with agents such as fluorescein isothiocyanate, and then microinjected into cells. Because GFP and GFP-tagged proteins are genetically encodable, they can be expressed from transfected DNA, making the preparation of cells that express fluorescently labeled protein accessible to virtually any biomedical research laboratory. In addition to the ease with which GFP-tagged proteins can be prepared, GFP has been used extensively in live-cell imaging because GFP photobleaches less rapidly than proteins tagged with traditional small molecule fluorescent dyes.
Although GFP has proven to be a valuable tool for studying the cell biology of proteins, similar simple and straightforward technologies for RNA and small molecule visualization are not available. If there were an RNA visualization technology analogous to GFP, it would by a major enabling technology that would permit a wide variety of important questions to be studied. Questions regarding the specific real-time localization of mRNAs, for example, during its processing in the nucleus and nucleolus, and following export to the cytosol and trafficking to neuronal growth cones, spines, axons, nuclei, organelles, etc., could be addressed, especially in terms of specific spliced forms of mRNAs, differentially edited mRNAs, and trinucleotide repeat-containing mRNAs. Furthermore, the timing of mRNA trafficking in response to extracellular signals could be addressed, such as the role of mRNA trafficking to dendritic spines during synaptic plasticity or in growth cones during axon turning. The role of regulated mRNA degradation in dendrites and axons could also be assessed. RNA visualization technologies are not, however, limited to mRNA. In addition to microRNAs, recent studies demonstrate the existence of large numbers of Piwi-interacting RNAs, promoter-associated small RNAs (PASRs), termini-associated small RNAs (TASRs) (Han et al., Promoter-associated RNA is Required for RNA-directed Transcriptional Gene Silencing in Human Cells,” Proc Natl Acad Sci USA 104:12422-12427 (2007)), as well as a plethora of other small noncoding RNAs (Hannon et al., “The Expanding Universe of Noncoding RNAs,” Cold Spring Harb Symp Quant Biol 71:551-564 (2006)) whose function is mysterious, but which appear poised to have the same impact on molecular biology as the discovery of microRNAs. Clearly, a wide variety of fundamental questions are waiting to be addressed.
The most commonly used technique to study mRNA localization is in situ hybridization (Levsky et al., “Fluorescence in situ Hybridization: Past, Present and Future,” J Cell Sci. 116:2833-2838 (2003)). This is a well-established technique, but is not a homogeneous assay and does not allow RNA to be monitored in the same cell at different time points. To achieve real-time, single cell in vivo RNA visualization, one technique has been to synthesize RNA in vitro using fluorescent nucleotides and then microinject them into cells (Zhang et al., “Creating New Fluorescent Probes for Cell Biology,” Nat Rev Mol Cell Biol 3:906-918 (2002)). This approach is technically difficult and has low throughput. Another approach is to use molecular beacons, which are oligonucleotides that are dual labeled with a fluorophore and a quencher (Tyagi et al., “Imaging Native Beta-actin mRNA in Motile Fibroblasts,” Biophys J. 87:4153-4162 (2004)). The beacon adopts a stem-loop structure that is nonfluorescent due to the proximity of the fluorophore and quencher at the base of the stem. When a target mRNA that exhibits complementarity to the loop hybridizes to the beacon, the stem is disrupted, resulting in separation of the fluorophore and quencher and subsequent fluorescence. However, transfected beacons exhibit nonspecific nuclear sequestration (Tyagi et al., “Imaging Native Beta-actin mRNA in Motile Fibroblasts,” Biophys J. 87:4153-4162 (2004)), and each mRNA requires a custom-designed beacon for visualization. Because of the inherent difficulties of these synthetic approaches, numerous groups have attempted to develop genetically encoded reporters of RNA localization in cells. However, because none have shown the requisite simplicity and specificity that are required for general use, RNA imaging still remains a largely inaccessible technology.
The most widely used technique is the GFP-MS2 system (Bertrand et al., “Localization of ASH1 mRNA Particles in Living Yeast,” Molecular Cell 2:437-445 (1998)). This approach uses two components: MS2, a viral protein, fused to GFP; and MS2-binding elements, which are RNA sequences, inserted into the 3′ UTR of RNAs of interest. GFP-MS2 and MS2-element-containing RNAs, or “fusion RNAs,” are expressed in cells from transfected DNA. GFP-MS2 binds to the MS2 element-tagged RNA in cells, and fluorescence signals in these cells should represent RNA-GFP complexes. Because unbound GFP-MS2 molecules diffuse throughout the cytosol there would be, in principle, a high fluorescence background. To alleviate this problem, a nuclear localization signal (NLS) is incorporated in the GFP-MS2 fusion protein so that most of the GFP-MS2 moves into the nucleus (Bertrand et al., “Localization of ASH1 mRNA Particles in Living Yeast,” Molecular Cell 2:437-445 (1998)). Unfortunately, the consequence of this is that GFP-MS2-RNA complexes are subjected to two trafficking signals: one encoded within the RNA and another being the NLS within GFP-MS2. The presence of two trafficking signals confounds interpretation of the intracellular movements of the tagged mRNA. An additional drawback is that the NLS causes the GFP-MS2 to accumulate in the nucleus, resulting in intense nuclear fluorescence signals and thereby preventing the analysis of nuclear-localized RNAs. Since much RNA biology occurs in the nucleus, such as nonsense mediated decay, RNA editing, nuclear export of RNA, splicing, pioneer RNA translation, and microRNA processing, nuclear accumulation of GFP-MS2 impedes the analysis of these events. Thus, even though GFP-MS2 has utility, it is not adequate for the needs of the research community.
Other related approaches have been described which have important limitations. One recent strategy involves the expression of GFP as two separate halves, each fused to half of the protein eIF4A, an RNA-binding protein (Tyagi, S. “Splitting or Stacking Fluorescent Proteins to Visualize mRNA in Living Cells,” Nat Methods 4:391-392 (2007)). RNAs containing the eIF4A binding site nucleate the binding of the eIF4A halves, which results in the juxtaposition of each GFP half and the subsequent formation of a stable GFP complex. Since the GFP complex requires ˜30 min to mature into a fluorescent species (Merzlyak et al., “Bright Monomeric Red Fluorescent Protein with an Extended Fluorescence Lifetime,” Nat Methods 4:555-557 (2007)), this method does not allow for visualization of nascent RNAs. Additionally, once formed, the fluorescent complex can dissociate spontaneously or after RNA degradation, resulting in high levels of background cytoplasmic fluorescence.
A highly desirable strategy would be to have an RNA sequence that would be fluorescent without the aid of an additional binding protein. However, a potential alternative is to tag RNAs with fluorescent dye-binding RNA sequences. Short RNA sequences that bind other molecules have been termed “aptamers.” Using SELEX, RNA aptamers that bind fluorescent dyes such as fluorescein have been described (Holeman, et al., “Isolation and Characterization of Fluorophore-binding RNA Aptamers,” Folding & Design. 3:423-431 (1998); Sando et al., “Transcription Monitoring using Fused RNA with a Dye-binding Light-up Aptamer as a Tag: a Blue Fluorescent RNA,” Chem Commun (Camb) 33:3858-3860 (2008)). However, these RNAs have not found use in live-cell experiments because both bound and unbound dye are fluorescent and have nearly identical emission spectrum properties. Thus, if one had cells that expressed the fluorescein-binding aptamers, and then applied fluorescein to the media, the signal from unbound fluorescein would overwhelm the signal from the fluorescein bound to RNA. Similarly, malachite green, although fluorescent when bound to cognate aptamers (Babendure et al., “Aptamers Switch on Fluorescence of Triphenylmethane Dyes,” J Am. Chem. Soc. 125:14716-14717 (2003)), cannot be used in living cells since malachite green is intensely fluorescent when it interacts with cell membranes (Guidry, G. “A Method for Counterstaining Tissues in Conjunction with the Glyoxylic Acid Condensation Reaction for Detection of Biogenic Amines,” J Histochem Cytochem. 47:261-264 (1999)). Additionally, malachite green generates radicals that are cytotoxic (Beermann et al., “Chromophore-assisted Laser Inactivation of Cellular Proteins,” Methods in Cell Biology 44:715-732 (1994)) and rapidly destroy the RNA aptamer itself (Grate et al., “Laser-mediated, Site-specific Inactivation of RNA Transcripts,” Proc. Natl. Acad. Sci. USA 96:6131-6136 (1999); Stojanovic et al., “Modular Aptameric Sensors,” J Am Chem Soc. 126:9266-9270 (2004)). These unfortunate features of malachite green have prevented the implementation of these otherwise potentially useful malachite green aptamers in cell imaging.
Other dyes are also problematic. There are numerous molecules that exhibit fluorescence upon binding nucleotides. Ethidium bromide and Hoechst dyes are probably the best known, but these molecules bind oligonucleotides relatively nonspecifically. Consequently, when used in a cellular environment or in vitro medium containing both target and non-target nucleic acid molecules, ethidium bromide and Hoechst dyes will generate a fluorescent signal even in the absence of the target nucleic acid molecule. Cyanine dyes are robustly fluorescent, and they exhibit their fluorescence even when they bind to indiscriminately to membrane components. Cyanine dyes, therefore, suffer from the same problems as malachite green.
It would be desirable, therefore, to generate a fluorophore that generates a sufficiently low background fluorescence in the absence of aptamer binding, whereby the aptamer-bound fluorophore complex generates an enhanced or modified fluorescence signal that is readily distinguishable from the fluorescence signal, if any, of the unbound fluorophore.
The present invention is directed to overcoming these and other deficiencies in the art.
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USPTO Backgrounds