Assaying RNA localization in situ with spatially-restricted nucleobase

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Letter

Assaying RNA localization in situ with spatially-restricted nucleobase oxidation Ying Li, Mahima B. Aggarwal, Kim Nguyen, Ke Ke, and Robert C. Spitale ACS Chem. Biol., Just Accepted Manuscript • DOI: 10.1021/acschembio.7b00519 • Publication Date (Web): 27 Sep 2017 Downloaded from http://pubs.acs.org on September 27, 2017

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ACS Chemical Biology

Assaying RNA localization in situ with spatially-restricted nucleobase oxidation

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Ying Li1, Mahima B. Aggarwal1, Kim Nguyen1, Ke Ke1, Robert C. Spitale*1,2

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(1) Department of Pharmaceutical Sciences, (2) Department of Chemistry. University of

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California, Irvine. Irvine, California. 92697

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Correspondence: [email protected]

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Abstract.

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We report herein a novel chemical-genetic method for assaying RNA localization within living

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cells. RNA localization is critical for normal physiology as well as the onset of cancer and

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neurodegenerative disorders. Despite its importance there is a real lack of chemical methods to

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directly assay RNA localization with high resolution in living cells. Our novel approach relies on

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in situ nucleobase oxidation by singlet oxygen generated from spatially confined fluorophores.

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We demonstrate that our novel method can identify RNA molecules localized within specific

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cellular compartments. We anticipate that this platform will provide the community with a much-

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needed methodology for tracking RNA localization within living cells, and sets the stage for

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systematic large scale analysis of RNA localization in living systems.

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ACS Paragon Plus Environment


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Text. Recent genetic and functional analysis has demonstrated that RNA molecules can

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control many aspects of gene regulation: from transcription to translation.1,

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organization of an RNA molecule can be intimately related to its function. For example,

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chromatin-associated RNAs can regulate transcription.3 Cytosolic RNA molecules can interact

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with messenger RNAs (mRNAs) to control translation and mRNA decay.4 mRNAs can also be

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localized to the outer membrane of the mitochondria for import into the mitochondrial matrix.5

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Each of these facets of RNA localization is critical for RNA function and cellular fitness.

The spatial

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Analysis of RNA localization is critical for studying RNA functions. Current methods for

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RNA localization study primarily focus on a small number of RNAs6 and typically require

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intensive labor7 to achieve slightly larger scale analysis. In addition, these strategies rely highly

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on existing knowledge about the primary sequence of RNAs (in situ hybridization) and limit the

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potential to discover unknown or unconventional RNA localization. Cellular fractionation,

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permitting relatively larger scale analysis (sequencing etc.), inevitably comes with many false-

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positives. During fractionation, the original spatial organization of biomolecules is disrupted and

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the falsely re-associated biomolecules prevent accurate analysis for further study.8,

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development of a method that begins to address these challenges, while assaying RNA

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localization in situ, would have a tremendous impact on our ability to analyze RNA localization.

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The

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Here we demonstrate that RNA localization can be assayed within intact living cells

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through spatially restricted nucleobase oxidation. We employ localized fluorophores, which,

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upon excitation with blue light (Supporting Information, SI), can utilize energy transfer to excite

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nearby ground state triplet oxygen to singlet state, resulting in guanosine oxidation in RNA.

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Nucleophiles in solution can intercept oxidized guanosine residues to tag them for downstream

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study. We demonstrated that this oxidative approach provides subcellular, and even sub-

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organellar resolution for RNA localization. Sub-cellularly tagged RNAs can be enriched and

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profiled by RT-qPCR. The enrichment step significantly reduces background and focuses on


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genes of interest. We anticipate that many labs aiming to explore RNA localization will adopt our

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method: from one gene to many in parallel.

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Tagging RNA with high resolution inside living cells is a formidable challenge. We

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envisioned that singlet oxygen induced guanosine oxidation could be useful in this regard as the

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singlet oxygen has an extremely short half-life and thus a short diffusion radius (~268 nm).10

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Recent analysis on singlet oxygen diffusion inside cells is consistent with this note.11

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Fluorescent dyes are well known for their ability to perform triplet energy transfers to molecular

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oxygen (in the triplet state).12 The resulting singlet oxygen can oxidize guanosine to 8-

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oxoguanosine (8oxoG, Figure 1A). The higher oxidation state of 8oxoG is susceptible to attack

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by a wide variety of nucleophiles in solution.13-15 We therefore hypothesized that restricted

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localization of a fluorophore would induce a high local concentration of singlet oxygen, which

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would permit tagging of guanine residues in RNA in the presence of a nucleophile (amine,

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outlined in Figure 1 B).

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We focused on the fluorophore eosin, as it has been demonstrated to afford singlet

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oxygen efficiently.16 To first demonstrate the ability to form adducts with guanine, in a singlet-

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oxygen dependent manner, we exposed single stranded synthetic DNA oligonucleotides (15 nt),

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which contain four guanosine residues to blue light in the presence of eosin and amino PEG

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(Figure 2A). We observed adducts formed only in the presence of blue light and only on an

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oligonucleotide that contained guanine (Figure 2B). We also characterized the time-dependent

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nature of the adduct formation and observed the reaction reaches a limit after ~15 minutes of

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blue light exposure (Figure 2C). To demonstrate the formation of a biotin adduct we exposed

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isolated total RNA to blue light with eosin and amino-PEG2-biotin and observed biotinylation by

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transfer blot (Figure 2D). Biotinylation was inhibited in the presence of sodium azide, a known

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singlet oxygen quencher (Figure 2E).17 Overall these data suggest that singlet oxygen,

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generated from fluorophore excitation, can oxidize guanine and permit biotinylation for potential

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downstream enrichment.



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We next focused on demonstrating that RNA tagging could be accomplished within living

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cells. HEK293T cells were incubated with eosin and propargyl amine (PA), a smaller amine

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nucleophile, which should easily traverse into cells. Extracted RNA was appended with a biotin

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tag through azide-biotin using conventional Cu(I)-catalyzed azide-alkyne cycloaddition

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(CuAAC). We were able to observe significant amounts of tagged RNA when cells were

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exposed to blue light in the presence of eosin and PA, even at concentrations as low as 100 µM

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for eosin (Figure 2F). We also observed significant amounts of protein and DNA labeling under

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these conditions, suggesting that our methodology could be used to track DNA and protein

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localization in situ (SI, Figure S1). Our results demonstrate that our tagging approach is capable

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of working inside cells.

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As a starting point for verifying the resolution of RNA tagging, we focused on the nucleus

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and cytoplasm. These two compartments are experimentally feasible and have several known

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positive controls. To localize fluorophores at specific subcellular regions we hypothesized we

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could tether dyes to Halo-tag fusion proteins, which are localized to specific regions of the cells

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(Figure 3A).18 We synthesized a dibromofluorescein-halo ligand (DBF), a synthetically easier

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eosin derivative that is more hydrophilic and has slightly lower yield of singlet oxygen generation

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(SI).19 We reasoned DBF would have lower background and higher-resolution tagging than

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eosin.

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We utilized Halo-tag constructs localized in the nucleus (histone protein H2B) and

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cytoplasm (NF-kappa-B p65) (Figure 3A; SI). Imaging experiments revealed very specific

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localization of Halo-DBF. Exposure to blue light, followed by CuAAC to visualize propargyl

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amine, afforded well-overlapping signals with Halo-DBF. (Figure 3B and 3C). Negative controls

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(- hv; -PA; -DBF) showed lack of adduct formation with PA (Figure S2).

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To test the specificity of enrichment we profiled tagged RNAs by RT-qPCR. We chose

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the following controls for nuclear-localized RNAs: (1) 7SK, a ncRNA transcribed from RNA

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polymerase III (pol. 3) that remains tightly associated with chromatin;20 (2) U6 nuclear snRNA,


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which is transcribed from pol. 3 and remains in the nucleus;21 (3) U1 snRNA, which is

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transcribed by pol. 2, exported into the cytoplasm and recycled back into the nucleus.22 For the

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cytoplasm the following control RNAs were profiled: (1) GAPDH and actin mRNA, which are

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transcribed by pol. 2 and exported into the cytoplasm;23 (2) 18S rRNA, which is transcribed by

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pol. 1 in the nucleolus and exported into the cytoplasm.24

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We first focused on 7SK for enrichment from Halo-H2B expressing cells. Increased

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exposure time to blue light irradiation, in the presence of PA, resulted in an exponential increase

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in enrichment over background, with 5-minute short exposure giving reasonable enrichment

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(Figure S3). Enrichment experiments based on fixed and permeabilized cells were also

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performed but afford less spatial resolution despite high yields of adduct formation (Figure S4).

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As such, performing the singlet oxygen induced tagging in living cells results in lower yield of

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tagging, but much better resolution. These results are consistent with the observation that many

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singlet oxygen quenchers exist in the cells, which would decrease the labeling radius.25 We

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therefore proceeded with these conditions in living cells for all the subsequent experiments.

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We then performed more systematic analysis of enrichment patterns for both nuclear

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and cytoplasmic localizations. As shown in Figure 3D, RNAs predicted to be in the nucleus were

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highly enriched in the Halo-H2B expressing cells. RNAs predicted to be in the cytoplasm were

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highly enriched in the Halo-P65 expressing cells. Notably, the two RNAs that do not exit the

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nucleus (7SK and U6) have very striking differences between the two conditions. 7SK, which is

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tightly bound to chromatin, had the largest difference for fold enrichment. U1 snRNA, which is

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exported then re-imported, still had low enrichment in Halo-p65 expressing cells. Consistent

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with its known localization it was highly enriched in the Halo-H2B cells.

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As predicted, GAPDH and actin mRNAs had substantial enrichment in the Halo-P65

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cells and very little enrichment in the Halo-H2B cells (Figure 3E). Perhaps the most remarkable

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result is the lack of 18S rRNA tagging in the Halo-H2B construct: this is impressive because the

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18S rRNA is a highly abundant RNA and it is transcribed within the nucleolus, a sub



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compartment in the nucleus not associated with chromatin (where H2B is localized). This RT-

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qPCR result is consistent with the lack of DBF and Cy5 signal in the nucleolus (Figure 3B).

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These results strongly suggest that localized RNA tagging can be achieved in living cells with

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locally induced singlet oxygen in the presence of propargyl amine.

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The results above suggested that singlet oxygen induced tagging is a high-resolution

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method for analyzing RNA localization in situ. We next sought to further test the resolution limits

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of our approach. We focused on the nucleolus and wondered if localization of DBF inside the

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nucleolus would permit selective tagging of RNAs therein. We focused on three RNAs: (1) 18S

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rRNA which is transcribed within the nucleolus and then exported into the cytoplasm; (2) U13

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snRNA, which is specifically localized within the nucleolar compartment;26 (3) 7SK which is

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known to remain tightly bound to chromatin.

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We constructed a Halo-tag protein fusion with fibrillarin (Fib), a component of the small

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nucleolar ribonucleoprotein (snRNP) that is highly enriched in the nucleolus.27 Imaging of the

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Halo-Fib fusion showed that DBF is highly accumulated within the nucleolus, in sharp contrast

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to Halo-H2B fusion (Figure 4A). The PA signal also suggested confined RNA tagging within the

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nucleolus for Halo-Fib (Figure 4A). We compared the RT-qPCR profiles for the three RNAs from

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cells transfected with Halo-Fib and Halo-H2B. As predicted, U13 and 18S are highly enriched in

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Halo-Fib expressing cells and 7SK is highly enriched in the Halo-H2B expressing cells (Figure

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4B). Together, these observations strongly support that the resolution of our approach is at sub-

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organellar level.

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In this Letter, we have developed a framework for assaying RNA localization within intact

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living cells. We have utilized localized fluorophores to convert triplet oxygen to singlet oxygen

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for spatially-restricted RNA oxidation. We have demonstrated that oxidized RNAs can be tagged

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with nucleophiles for subsequent enrichment and profiling by RT-qPCR. Our approach is of high

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resolution as RNAs can be tagged within sub-organeller compartments.


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We have designed our approach to interface with the Halo tag proteins, which are now

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commercially available for many human proteins. As such, it can be widely applied for

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systematic analysis of RNA localization. In addition, the approach of RT-qPCR paves the way to

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RNA sequencing for potential discovery of RNAs localized within specific subcellular

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compartments. We anticipate the results herein will provide a roadmap for a wide-scale and

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systematic analysis of RNA localization (live cell differentiation, etc). Such studies are currently

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underway in our lab.

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Acknowledgements

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We thank members of the Spitale lab for their careful reading and critique of the

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manuscript. Spitale lab is supported by start up funds from the University of California, Irvine,

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and the NIH (1DP2GM119164 RCS). RCS is a Pew Biomedical Scholar.

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Supporting Information Supporting information accompanying this manuscript is available online. These include

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NMR spectra and additional biochemical methods. This material is available free of charge via

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the internet at http://pubs.acs.org.

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Figures.

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Figure 1. Fluorophore-mediated RNA oxidation. (A) Schematic of fluorophore-mediated oxidation of guanosine, followed by nucleophilic tagging. (B) Schematic of localized RNA oxidation, followed by enrichment and profiling.


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Figure 2. Fluorophore-mediated RNA oxidation tagging. (A) Schematic of eosin-mediated oxidation and RNA tagging by an alkyl amine. B = biotin. (B) Phosphoroimaging gel shift demonstrating guanosine-dependent oxidation and nucleophilic adduct formation with amino PEG (1 kDa). G = guanosine (C) Phosphoroimaging gel of time-dependent guanosine tagging (0-20 min). (D) Demonstration of eosin-mediated tagging with amino-PEG2-biotin, in vitro. (E) Sodium azide, a singlet oxygen quencher, decreases the signal of eosin-mediated RNA tagging. (F) Demonstration of eosin-mediated RNA tagging inside living cells (final concentration of eosin: 100, 200 500 µM). EB = ethidium bromide loading control. Biotin = streptavidin blot.



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Figure 3. Spatially restricted tagging of RNA inside living cells. (A) Schematic of localized DBF to Halo fusion proteins. (B) Imaging of Halo-H2B fusions. DBF is denoted by green signal. PA is denoted by red signal. (C) Imaging of Halo-P65 fusions. DBF is denoted by green signal. PA is denoted by red signal. (D & E) RT-qPCR from Halo-H2B and Halo-P65 experiments. Enrichment was calculated against a no-hv negative control with ∆∆Ct method. N=3; biological duplicates.


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Figure 4. Spatially restricted tagging of RNA inside the nucleolus. (A & B) Imaging of HaloFib and Halo-H2B constructs. Yellow arrows mark the nucleolus. (C) RT-qPCR of U13, 7SK, and 18S RNA. Enrichment was calculated against a no-hvnegative control with ∆∆Ct method. N=3; biological duplicates.

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Methods. Additional methods can be found within the supplementary information. Synthesis. Synthesis and spectra for all compounds is found within the supplementary information. Cell lines and culture conditions. Hek293T cells were cultured in DMEM supplemented with 10% FBS, 1% penicillin and streptomycin and grown at 37ºC, 5% CO2. Cloning and plasmids. H2B-Halo plasmid (a kind gift from Dr. Luke D. Lavis laboratory; Jenelia Research Campus, HHMI, Virginia, USA) was used as a template for amplifying Halo tag sequence by polymerase chain reaction (PCR) with primers oKN283F (5’-aattt ACCGGT ATGGGATCCG AAATCGGTAC TGGCTTTCCA TTCGACCC-‘3) and oKN284R (5’TCAATGGTAC CGCCGGAAAT TTCTAGCGTC GACAGCCA-‘3) using KOD Hotstart DNA polymerase (EMD Millipore Corp.) Halo tag fragment was cloned into pTagRFP-C1-Fibrillarin backbone (Addgene ID # 70649) and replaced RFP sequence via AgeI and KpnI resulting pKN326 or Halo-Fib plasmid. The plasmid sequence was verified by Sanger sequencing analysis (Genwiz, Inc.) Photo-oxidation labeling of cellular RNA. The cell culture dishes were coated with poly-Dlysine (50 µg/ml) for 6 h at 37ºC and washed three times with autoclaved water to remove the excessive amount of poly-D-lysine. HEK293T cells were seeded at equal density (106 cells/plate in 10-cm plates). HEK293T cells were transiently transfected for 24-36 h with appropriate halo fusion protein plasmid (5 µg) on the following day using jetPRIME transfection reagent according to manufacturer’s manual (Polyplus Transfection, France). Cell media was replaced with 5 µM Halo-DBF (compound 7) in HBSS, diluted from 5 mM DMSO stocks. Cells were incubated with the ligand for 15 min at 37ºC, 5% CO2 and washed twice with full media for 25 min. Cells were then incubated with 1 mM propargyl amine (PA) in fresh HBSS media for 5 min at 37ºC, 5% CO2, irradiated for 15 min with 500 nm light at room temperature and incubated for 5 min in dark. RNA fluorescence imaging via CuAAC. The cell culture dishes and glass cover slips were coated with poly-D-lysine (50 µg/ml) for 6 h at 37ºC and washed three times with autoclaved water to remove the excessive amount of poly-D-lysine. HEK293T cells were seeded at equal density (2.5×105 cells/well in 6-well plates) and were transiently transfected with 1 µg appropriate halo fusion protein plasmid on the following day using jetPRIME transfection reagent according to manufacturer’s manual (Polyplus Transfection, France) for 24-36 h. Cells were incubated with the ligand (5 µM in HBSS from 5 mM stock in DMSO) for 15 min at 37ºC, 5% CO2 and replaced with fresh full media for 25 min twice. Cells were then incubated with 1 mM propargyl amine (PA) in fresh HBSS media for 5 min at 37ºC, 5% CO2, irradiated for 15 min with 500 nm light at room temperature and incubated for 5 min in dark. Cells were washed three times with DPBS, fixed and permeabilized for 30 min at room temperature with 3.7% paraformaldehyde and 0.1% Triton-X100. Cells were then washed three times (7 min/each) on orbital shaker with DPBS, blocked with BSA (1 mg/ml in DPBS, 0.45% NaCl and 0.025% NaN3) for 30 min at room temperature, washed twice for 5 min with DPBS, and incubated with 200 µL of click solution (1 mM CuSO4, 2 mM THPTA ligand, 10 mM NaAsc, and 15 µM azide-Cy5) for 1 hour at 37ºC in the dark. Cells were washed three times for 5 min/each on an orbital shaker with DPBS-0.1% Triton-X100 and one with DPBS. Cells were stained with Hoechst 333242 (1:2000,


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Trihydrochloride, Trihydrate - 10 mg/mL solution in water from Thermo Fisher Scientific) for 10 min, washed twice with DPBS for 5 min and mounted using VectaShield (Vector Labs). Slides were imaged via fluorescence confocal microscopy using a 63x oil immersion objective on a Leica 700 Carl Zeiss microscope. RT-qPCR for biotinylated RNA. 2 µg of biotinylated RNA from each sample were subjected to cDNA synthesis with reverse primer for each gene using the PrimeScriptTM Reverse Transcriptase from Takara. 3 µl out of a 20-µl reaction was diluted with 87-µl nucleus free water and set aside as INPUT. To the remaining cDNA reaction, Dynabeads MyOne Streptavidin C1 (Invitrogen, Cat# 65002) that had been blocked with BSA (1 mg/mL) and yeast tRNA (1 mg/mL) at 4 ºC overnight.[5] The beads were washed three times with 4 M wash buffer (100 mM Tris, pH7.0, 4 M NaCl, 10 mM EDTA, 0.2% (vol/vol) Tween-20) and re-suspended in bead binding buffer (100 mM Tris, pH7.0, 1 M NaCl, 10 mM EDTA, 0.2% (vol/vol) Tween-20).[6] Blocked beads were added to enrich biotinylated RNA-cDNA hybrids. The mixture was incubated at room temperature for 30 min on shaker. Using DynaMag side magnet to retain the streptavidin beads, the flow-through was removed. The beads were washed twice with 4 M wash buffer at room temperature, twice with 4 M wash buffer incubated at 50 ºC for 3 min, and twice with PBS (1X, pH7.4). The beads were then treated with RNase H mixture [1X RNAse H buffer, 12.5 mM D-biotin (Life Technologies, Cat# B20656), 0.1 U/µl RNAse H (New England Biolabs, Cat# M0297S), RNAse A/T1 cocktail at 40 ng/µL (Thermal Scientific, Cat# EN0551)] for 30 min at 37ºC on shaker. Each RNAse reaction was added 1 µL of DMSO and incubated at 95-98ºC for 5 min. Using DynaMag side magnet to retain the beads, released cDNA was collected and purified using DNA Clean & Concentration columns from Zymo Research (Cat# D4003) according to the manufacturer instructions. Purified cDNA was eluted with 34µl nucleus free water, further underwent a 1:20 dilution and set aside as ENRICH. INPUT (2 µl) and ENRICH (2 µl) were subjected to RT-qPCR using SYBR® Advantage® qPCR Premix from Takara and Biorad CFX connectTM real time system. The fold of enrichment was calculated against a no-hv negative control: 2^((CtENRICH-nohv –CtINPUT-nohv)-(CtENRICH-CtINPUT)).



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Assaying RNA localization in situ with spatially-restricted nucleobase

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