Najmeh Nikpour and Fiona Alum provided suggestions and edited this manuscript. Department of Biochemistry at McGill University supported HM and VM with the CIHR Training Initiative in Chemical Biology. This work was supported by Canadian Institute of Health Research (CIHR) grant 119464 (to R. S.).

The protocol below describes the procedure for performing the fluorescence-based RNA editing assay. The assay can be performed in a single PCR tube, 96-well, or 384-well plates depending on the scope of the experiment. Subsequently the fluorescence signal can be read on a suitable real time PCR detection system. The assay here is described in the context of 384-well plates.
1. Culturing T. brucei Cells
<ol>
	<li>Prepare a growth medium for T. brucei procyclic form cells. For 1 L of medium:
	<ol>
		<li>Dissolve 25.4 g SDM-79 powder in 800 ml miliQ water.</li>
		<li>Add 2 g of NaHCO3 and pH to 7.3 with 10 M NaOH.</li>
		<li>Add nanopure water to a final volume of 900 ml, filter sterilize.</li>
		<li>Add Fetal Bovine Serum (FBS), penicillin-streptomycin solution and hemin (2.5 mg/ml) to final concentrations of 10% (v/v), 100 U/ml and 7.5 mg/L respectively.</li>
	</ol>
	</li>
	<li>Grow 300 ml of T. brucei 1.7A wild type (procyclic form) cells at 28 &#176;C, shaking at 70 rpm to a density of 1.5 x 107 cells/ml. NOTE: This should produce 3 ml of active editosome with ~0.5 mg of total protein, sufficient for 600 editing reactions.</li>
	<li>Harvest the cells by centrifugation at 6,000 x g for 10 min at 4 &#176;C.</li>
	<li>Wash the pellet with 50 ml of chilled PBSG buffer (10 mM Na2HPO4, 10 mM NaH2PO4, 145 mM NaCl, and 6 mM glucose), and spin down the cells again by centrifugation at 10,000 x g for 10 min at 4 &#176;C.</li>
</ol>
2. Isolation of Crude Mitochondria
NOTE: All the steps should be performed on ice or at 4 &#176;C to preserve editosome activity.
<ol>
	<li>Resuspend the harvested cells in 30 ml of DTE buffer (1 mM Tris-HCl pH 8.0 and 1 mM EDTA). Use a 40 ml sterile Dounce homogenizer (pre-chilled) to disrupt the cell membrane by stroking up and down at least 10 times on ice.</li>
	<li>Immediately add 4.3 ml of 60% sucrose (w/v; i.e. 1.75 M) to the homogenate to a final concentration of 0.25 M. Centrifuge at 15,800 x g for 10 min at 4 &#176;C, to preferentially bring down mitochondria.</li>
	<li>Resuspend the mitochondrial pellet in 4.6 ml of STM buffer (20 mM Tris-HCl pH 8.0, 250 mM sucrose and 2 mM MgCl2). Add 13.8 &#956;l of 0.1 M CaCl2 and 4 &#956;l of RNase-free DNase I to final concentrations of 0.3 mM and 9 U/ml, respectively. Incubate the mixture for 1 hr on ice.</li>
	<li>Add 4.6 ml of STE buffer (20 mM Tris-HCl pH 8.0, 250 mM sucrose and 2 mM EDTA) to inactivate the DNase I. Centrifuge at 15,800 x g for 10 min at 4 &#176;C.</li>
	<li>Resuspend the pellet in 400 &#956;l of lysis buffer (10 mM Tris-HCl pH 7.2, 10 mM MgCl2, 100 mM KCl, 1 &#181;g/ml pepstatin, 1 mM DTT, and 1x complete EDTA-free protease inhibitor) and transfer to a microfuge tube.</li>
	<li>Add 10% Triton X-100 to a final concentration of 1% and incubate the lysate for 15 min at 4 &#176;C on a tube rotator.</li>
	<li>Clear the mitochondrial lysate by centrifuging twice at 17,000 x g for 15 min at 4&#160;&#176;C; retaining the cleared supernatant each time.</li>
</ol>
3. Editosome Purification
<ol>
	<li>Pour a 10 ml 10%-30% (v/v) linear glycerol gradient (Table 1) in an ultracentrifuge tube using 2x HHE gradient buffer (40 mM HEPES pH 7.9, 20 mM Mg(OAc)2, 100 mM KCl, and 2 mM EDTA) and a gradient maker by following the instruction manual.</li>
	<li>Carefully remove 500 &#956;l of solution from the top of the glycerol gradient and gently load 500 &#956;l of the cleared mitochondrial lysate. Spin at 178,000 x g for 6 hr at 4 &#176;C using an ultracentrifuge.</li>
	<li>Collect 500 &#181;l fractions sequentially from the top to the bottom of the gradient at 4 &#176;C. Then snap freeze the fractions using liquid nitrogen and store at -80 &#176;C until usage.</li>
</ol>
4. RNA Preparation
<ol>
	<li>Anneal the respective DNA template containing sequence complementary to T7 promoter sequence (Table 2) with a T7 promoter oligonucleotide (5&#8217;-TAATACGACTCACTATAGGG-3&#8217;) in a 1:1 molar ratio by heating at 90 &#176;C for 3 min and cooling at RT for at least 10 min.</li>
	<li>Transcribe RNA using an in vitro transcription kit by following the instruction manual.</li>
	<li>Stop the transcription reaction by adding equal volume of 7 M urea dye (7 M urea, 0.05% Xylene Cynol, and 0.05% Bromophenol blue). Run on a filter sterilized 9% denaturing polyacrylamide gel (9% acrylamide, 7 M urea, 1x TBE) .</li>
	<li>Use the ultraviolet (UV) shadowing with a shortwave UV lamp to locate and excise respective RNA. Place the excised gel piece in a microfuge tube and add 400 &#181;l of gel elution buffer (20 mM Tris-HCl pH 7.5, 250 mM NaOAc, 1 mM EDTA and 0.25% SDS). Elute overnight at RT on a tube rotator.</li>
	<li>Precipitate the eluted RNA by adding 1 ml of cold 100% ethanol and incubating either at -80 &#176;C for 30 min or -20 &#176;C overnight.</li>
	<li>Centrifuge at 16,000 x g for 30 min at 4 &#176;C, to pellet down the RNA.</li>
	<li>Wash the pellet with 1 ml of 75% ethanol. Centrifuge at 16,000 x g for 20 min at 4 &#176;C.</li>
	<li>Resuspend the RNA pellet appropriately in RNase free water to achieve the desired concentrations, as shown in Table 2.</li>
</ol>
5. Fluorescence-based RNA Editing Assay
<ol>
	<li>For a single reaction, combine 1 pmol (1 &#181;l) of preA6Rbz and 2.5 pmol (1 &#181;l) of gA6Rbz (1:2.5 molar ratio) in a microfuge tube, incubate at 70 &#186;C for 3 min and let it sit at RT for at least 10 min.</li>
	<li>Meanwhile prepare a master mix using Table 3, without the preA6Rbz and gA6Rbz, for the editing reaction containing 1x HHE buffer (25 mM HEPES pH 7.9, 10 mM Mg(OAc)2, 50 mM KCl and 10 mM EDTA), 1 mM ATP, 5 mM CaCl2, 16 ng/&#181;l of Torula yeast RNA, 0.1% Triton X-100, and the purified editosome.</li>
	<li>Add annealed preA6Rbz and gA6Rbz to complete the master mix.</li>
	<li>Dispense 18 &#181;l of the master mix (Table 3) into wells containing either 2 &#181;l of RNase-free water (wells with no compounds) or 2 &#181;l of 200 &#181;M chemical compounds and include control samples in the plate according to Figure 5.</li>
	<li>Seal the plate with a plate sealer and spin the plate down, to remove any air bubbles. Incubate at 28 &#176;C for 4 hr.</li>
	<li>Add 25 pmol (2 &#181;l) of gA6Rbz competitor to each well. Place a fresh sealer, spin the plate down and place it on a real-time PCR machine. Program the following experiment: 
	Step 1: 85 &#176;C for 5 min; Step 2: 24 &#176;C for 10 min; Step 3: Stop.</li>
	<li>Add 15 pmol (1 &#181;l) of FRET substrate to each well to a final volume of 23 &#181;l. Seal the plate with a fresh sealer. Quickly spin the plate and place it back on the real-time PCR machine.</li>
	<li>Program a new experiment with the following steps: 
	Step 1: 37 &#176;C for 1 min; Step 2: Read; Step 3: Go to Step 1, 40 times; Step 4: Stop.</li>
	<li>Setup the plate by selecting all the wells that require reading and choose the FAM filter. Input volume as 23 &#956;l and start the run.</li>
	<li>Calculate the slope of the values obtained from each well/sample to obtain a kinetic measurement by plotting the slopes on a bar graph for analysis. NOTE: A kinetic read improves the signal-to-noise ratio between the sample and the background; as the background sample would have a slope close to zero. An end point reading has higher background noise.</li>
</ol>

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The authors have nothing to disclose.

A novel high-throughput screening method to identify inhibitors against the RNA editing complex of Trypanosomes was presented, providing a new tool for drug discovery to counter diseases caused by trypanosomatids. FRET-based ribozyme assay has been extensively used for different purposes20-22; however, we have utilized the capacity of FRET-based ribozyme assay for in vitro monitoring of RNA editing activity19. This assay could potentially be adapted to other types of RNA editing in eukaryotes, such as nucleotide substitution editing of nucleus-encoded RNAs of mammals23.
The novelty of this assay is not in establishing FRET-based ribozyme assays per se, but in developing a method that incorporates this technique into an RNA editing assay that is amenable to high-throughput screening of chemicals against editosome. The ribozyme assay-based method has several advantages over other screens and assays currently utilized for this purpose. Specifically, the technique offers a sensitive and reproducible &#8220;mix-and-measure&#8221; assay relying on a fluorescent substrate as opposed to a radiolabeled one, thereby making it fit for high-throughput screening19. Real-time monitoring of a fluorescence signal after addition of the substrate instead of a simple end-point signal makes it possible to more accurately determine IC50 values of assayed compounds19. Furthermore, it permits testing of the inhibitory effects of compounds on the whole-editosome complex as opposed to individual recombinant proteins. Given the dynamic nature of interactions within protein complexes, it can be suggested that this method allows identification of inhibitors against editosome proteins in a more biologically representative setting24. In addition, targeting the whole-editosome complex provides an opportunity to potentially arrest the progression of RNA editing at various transient steps that have not been previously identified. Thus, the technique will allow gaining a better perspective as to interactions and activities that are crucial to the process of RNA editing. This approach has been found to be successful in the past as exemplified by the elucidation of prokaryotic ribosome complex assembly and function utilizing antibiotics, such as viomycin and erythromycin, acting as inhibitors of the complex24,25.
To ensure the successful completion of the method, certain adjustments in protocols may be necessary. First, selection of the proper glycerol gradient mitochondrial extract fraction is crucial. Here, fractions 7 through 11, corresponding to the ~20S region of the gradient, showed the highest editing activity, with fraction 9 exhibiting maximum activity. Although this fraction was selected for all of the tests presented, it is essential to test all fractions beforehand in order to assess in which fraction the editing activity peaks. Second, to validate fraction selection, it is critical to perform preliminary tests to assess the signal-to-noise ratio by calculating the Z&#8217;-factor value. If proper mitochondrial extract glycerol gradient fractionation has been completed, a Z&#8217;-value of 0.6 can be achieved. Next, it is recommended to re-evaluate the reaction volume of mitochondrial extract necessary to achieve maximal editing activity via titration19.
An important limitation of the technique presented in this paper concerns the laborious and time-consuming process of mitochondrial extract glycerol gradient fractionation by which the editosome complex is purified19. Despite these disadvantages, this technique is preferentially suggested to obtain crude editosome fractions given its low cost versus yield potential, thereby qualifying it for application to high-throughput screening. In contrast, other methods such as TAP-tag purification, which are able to give more purified editosome fractions, are advisable for low to medium throughput assays such as in the case of secondary assays. Moreover, in order to ensure specific inhibitory effect of compounds, it is necessary to include controls to test them against the activity of ribozyme (A6Rbz) in the absence of the editosome. It should be noted that previous studies have highlighted that this method may be ineffective at calculating IC50 values for dye-like compounds due to the possible interference at high compound concentrations19.
To further increase the sensitivity of the HTS technique presented, developing a similar fluorescence-based assay involving pre-cleaved pre-edited ribozyme is beneficial. This would allow bypassing the rate-limiting endonucleolytic cleavage step catalyzed by endonucleases in the editosome complex. Another advantage of this modified assay would be the application as a secondary screening tool to monitor the effect of the inhibitors identified through the primary screen on ExoUase, TUTase and ligation catalytic activities. The currently proposed assay is limited to the detection of inhibitors against the deletion type of RNA editing. Therefore, another avenue to explore would be the modification of the presented assay to enable monitoring of compound effects on the insertion type of RNA editing.

The process of RNA editing, a post-transcriptional mRNA modification, was first discovered in trypanosomatids1. Since then, substantial work has been conducted in studying the mechanism behind RNA editing in Trypanosoma brucei2,3. In a series of enzymatic reactions, the editosome, a core complex of about 20 proteins, creates mature mitochondrial mRNAs for multiple components of the energy generating oxidative phosphorylation system. The order of catalytic events is endonucleolytic cleavage, uridylate (U) addition or deletion, and ligation, as dictated by guide RNAs (gRNAs)4.
In addition to the core editosome complex proteins, a number of accessory factors have also been identified5-7. These proteins are mostly seen grouped in independent complexes. However, the order of protein assembly in the core editosome complex and the interaction patterns of the core complex with the accessory complexes are yet to be determined. Targeting the RNA editing process in trypanosomatids may provide chemical dissectors that aid in studying the assembly and function of the editosome complex. Furthermore, functional studies on several editosome proteins have shown essentiality across different life stages, indicating their potential as drug targets8-12. Therefore, the found inhibitors of the editosome may also act as lead compounds against trypanosomatids. This is timely, as drugs currently available against diseases caused by trypanosomatid are toxic, inefficient and expensive13,14.
An efficient and convenient in vitro assay is necessary to explore the chemical universe for specific inhibitors that block RNA editing. Three assays have been developed and used to monitor editosome activities: (a) full round in vitro RNA editing assay15, (b) pre-cleaved in vitro RNA editing assay16,17, and (c) hammerhead ribozyme (HHR)-based assay18. The first two assays rely on direct visualization of the edited product (ATPase 6 mRNA) with the help of radioactivity. The HHR-based assay uses a modified version of the ATPase 6 mRNA that is modeled to behave as a ribozyme upon editing. The functional ribozyme then specifically cleaves a radiolabeled RNA substrate, serving as a reporter. Recently, Moshiri et al. developed a &#8216;mix and measure&#8217; HHR-based in vitro reporter assay to monitor RNA editing where the radiolabeled RNA substrate is replaced with a fluorescence resonance energy transfer (FRET) substrate19. The principle advantages of this assay are: (a) it is a rapid and convenient mix and measure type of assay, as the production of active ribozyme and substrate cleavage occur simultaneously in the same tube in low volume (i.e. 20 &#956;l), (b) it avoids the use of radioactively labeled materials, (c) sensitivity that is afforded by fluorescence instrumentation in a micro-titer plate format, and (d) a high signal to noise ratio. Using this assay, the effect of known RNA editing ligase inhibitors against purified editosome was confirmed19. This experiment validated the assay for rapid identification of RNA editing inhibitors, primarily against whole editosomes from T. brucei.
Figure 1 is a detailed step-by-step schematic of the fluorescence-based in vitro RNA editing assay. This protocol can either be used for monitoring RNA editing in vitro or easily be adapted for screening compound libraries of various scales.

<table><tbody><tr><td>SDM-79 Medium</td><td>Gibco by Life Technologies</td><td></td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Life Technologies</td><td>12483-020</td><td>heat inactivation at 55 &amp;#176;C for 1 hr</td></tr><tr><td>Hemin, minimum 80%</td><td>Sigma</td><td>H5533-10G</td><td></td></tr><tr><td>Penicillin-Streptomycin Sollution</td><td>Fisher Scientific</td><td>MT-30-002-CI</td><td></td></tr><tr><td>Dnase 1 recombinant, Rnase Free&amp;nbsp;</td><td>Roche</td><td>4716728001</td><td></td></tr><tr><td>T7 RiboMax Express Large&amp;nbsp; scale RNA&amp;nbsp; production system</td><td>Promega</td><td>P1320</td><td></td></tr><tr><td>Kimble Kontes Dounce Tissue Grinders&amp;nbsp;&amp;nbsp;</td><td>Fisher Scientific</td><td>K885300-0040&amp;nbsp;&amp;nbsp;</td><td></td></tr><tr><td>Gradient Master, ver 5.25&amp;nbsp;</td><td>Biocomp</td><td>107-201M</td><td></td></tr><tr><td>Ultra Clear Tube, 13.2 ml</td><td>Beckman Coulter</td><td>344059</td><td></td></tr><tr><td>Optima L-100XP&amp;nbsp; Ultracentrifuge&amp;nbsp;</td><td>Beckman Coulter</td><td>392052</td><td></td></tr><tr><td>SW 41 Ti ROTOR</td><td>Beckman Coulter</td><td>331336</td><td></td></tr><tr><td>MicroSeal &#039;B&#039; Seal, Seals</td><td>Biorad</td><td>MSB1001</td><td></td></tr><tr><td>CFX 384 Touch Real-Time PCR Detection System</td><td>Biorad</td><td>185-5484</td><td></td></tr><tr><td>Acryl/Bis solution (19:1), 40% (w/v)</td><td>Bio Basic</td><td>A0006-500ML</td><td></td></tr><tr><td>Urea, Molecular biology grade, 1 kg</td><td>Life Technologies</td><td>AM9902</td><td></td></tr></tbody></table>

rna catalyst as a reporter for screening drugs against rna editing in trypanosomes

Substantial progress has been made in determining the mechanism of mitochondrial RNA editing in trypanosomes. Similarly, considerable progress has been made in identifying the components of the editosome complex that catalyze RNA editing. However, it is still not clear how those proteins work together. Chemical compounds obtained from a high-throughput screen against the editosome may block or affect one or more steps in the editing cycle. Therefore, the identification of new chemical compounds will generate valuable molecular probes for dissecting the editosome function and assembly. In previous studies, in vitro editing assays were carried out using radio-labeled RNA. These assays are time consuming, inefficient and unsuitable for high-throughput purposes. Here, a homogenous fluorescence-based &#8220;mix and measure&#8221; hammerhead ribozyme in vitro reporter assay to monitor RNA editing, is presented. Only as a consequence of RNA editing of the hammerhead ribozyme a fluorescence resonance energy transfer (FRET) oligoribonucleotide substrate undergoes cleavage. This in turn results in separation of the fluorophore from the quencher thereby producing a signal. In contrast, when the editosome function is inhibited, the fluorescence signal will be quenched. This is a highly sensitive and simple assay that should be generally applicable to monitor in vitro RNA editing or high throughput screening of chemicals that can inhibit the editosome function.

To demonstrate the necessary steps required for setting up a large-scale screen, Figures 2-5 are representative control experiments related to the quality of the assay. These are essential control experiments for a consistent assay over several days of screening or for comparison of different screens.
Assessing the Fluorescence Signal-to-noise Ratio
To ensure the stability and quality of the &#64258;uorescein-labeled oligoribonucleotide substrate in a large-scale setup, Z&#8217;-factor, defined as the difference between the assay background and the maximum signal was calculated using the active ribozyme molecule (A6Rbz). Assays with Z&#8217;-factor &#62;0.5 are considered acceptable for high-throughput screen. Figure 2 shows representative data using the FRET substrate labeled with 5&#8217; fluorescein (FAM; emitter) and 3&#8217; N,N&#8217;-tetramethylrhodamine (TAMRA; quencher) (A6Rbz_F/T). This substrate produced a Z&#8217;-factor of 0.64 when calculated using 72 replicates in the presence and absence of A6Rbz.
In this assay an alternative substrate using Iowa Black dark quencher (A6Rbz_F/Ib) can yield a better signal-to-noise ratio compared to TAMRA. This is because Iowa Black, unlike TAMRA, emits absorbed energy as heat and not light. The Z&#8217;-factor obtained using the FAM/Iowa Black (F/Ib)-labeled substrate was 0.68. The improvement in the Z&#8217;-factor for the F/Ib-labeled substrate over TAMRA-labeled substrate is because of its relatively lower background. These representative results show that both substrates are viable options for use in a high-throughput screen.
Determining the Editing Activities of Glycerol Gradient Fractions
To determine and select the most active editosome fractions for experiments, the glycerol gradient fractions were tested for in vitro editing using the fluorescence-based assay (Step V). These data show (Figure 3) fractions 7-12 as the most active fractions (&#8805;50% editing activity; with the most active fraction as 100%). These fractions can be combined when more editosome is required.
Calculating the Z&#8217;-factor when the Editosome Fractions were Combined
To determine the effect of combinig the editosome fraction, the Z&#8217;-factor value of 0.6 was calculated when the most active fractions (F8+F9+F10) were used as the source of editosome. To calculate the Z&#8217;-factor 72 replicates were tested in the presence and absence of the editosome (Figure 4). The F/Ib substrate was used in this experiment. These data show that based on the Z&#8217;-factor, combining the glycerol gradient fractions have minimal effect on the quality of the assay.
Representative Control Experiment for the Assay
Representative data comparing different control samples were used to analyze the variables in the assay. As shown in Figure 5, reactions missing any RNA editing components (reactions 1-4) do not cleave the substrate. Cleavage of the substrate as measured by the fluorescence and relative editing activity is only observed in the presence of all components of the editing reactions in the absence of an inhibitor (reaction 5) or in the presence of all editing components and an inactive compound (reaction 6). In contrast, an inhibitory compound can block RNA editing and is used as a positive control (reaction 7). Here, Mordant black (MrB) and C53 compounds were used as the positive and negative controls, respectively.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51712/51712fig1highres.jpg" width="500" /> 
Figure 1. Hammerhead ribozyme-based in vitro RNA editing assay. A) Step-by-step schematic representation of the fluorescence-based in vitro editing assay: (a) Hybridization of the pre-edited hammerhead ribozyme (pre-edited A6Rbz) with its guide RNA (gA6Rbz). (b) Recognition and interaction of the RNA duplex by the purified editosome complex. (c) Deletion RNA editing catalyzed by the editosome. (d) Dissociation of the edited A6Rbz from the editosome and gA6Rbz by heating at 85 &#176;C and addition of guide RNA competitor (gA6Rbz_comp). (e) Hybridization of the FRET substrate with the active A6 hammerhead ribozyme (active A6Rbz). (f) Detection of FAM signals following cleavage of the FRET substrate by the active ribozyme. B) The pre-edited hammerhead ribozyme (pre-edited A6Rbz) is shown in association with gA6Rbz that specifies the deletion of three Us (double-headed arrow) from the editing site (ES). As a result of RNA editing in the presence of functional editosome the inactive ribozyme is edited into its active form (edited A6Rbz) that can now cleave the FRET substrate (cleavage site indicated by an arrow). The conserved 5&#8242;-CUGA-3&#8242; of the edited A6Rbz in the catalytic core essential for ribozyme activity (edited site) is highlighted (This figure has been modified from Moshiri19). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51712/51712fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51712/51712fig2highres.jpg" width="500" /> 
Figure 2. Signal-to-noise ratio comparison between FRET substrates harboring different quenchers. Ribozyme (A6Rbz) activity using FAM/TAMRA (F/T) and FAM/Iowa Black FQ (F/Ib) substrates. The y-axis represents fluorescence arbitrary unit (FAU) per minute.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51712/51712fig3highres.jpg" width="500" /> 
Figure 3. RNA editing activity of select fractions obtained from glycerol gradient centrifugation of the mitochondrial lysate. Fraction #9 (F9) has the highest activity. The y-axis represents relative editing activity in percentage, considering F9 as 100%.
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51712/51712fig4highres.jpg" width="500" /> 
Figure 4. Z&#8217;-factor calculation using the most active fractions (F8+F9+F10) as the editosome source. Experimental variation was obtained from 72 replicates of each type of reaction. Z&#8217;-factor was calculated as 0.6. The y-axis represents relative editing activity.
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51712/51712fig5highres.jpg" width="500" /> 
Figure 5. Representative experiment for the fluorescence-based RNA editing assay. Reactions were performed in a final volume of 20 &#181;l per well and CFX384 TouchTM real-time PCR detection system was used for measuring the fluorescence signal. The graph presents various controls in addition to a complete editing reaction that contains all components for the assay (#5). The fluorescence was measured in the presence of FRET substrate alone (#1) to assess the integrity of the substrate. Sample #2 was performed in the absence of the editosome to monitor any ribozyme activity prior to editing. To assess the effect of denatured editosome on the substrate, the fluorescence was measured in the presence of the editosome and FRET substrate only (#3). To test the guide-directed editing specificity, a sample in the absence of gA6Rbz was used (#4). A non-inhibitory (C53, #6) and an inhibitory compound (MrB, #7) were used to test the effect on the editing assay. While RNA editing is inhibited significantly by MrB, C53 has negligible effect. The error bars correspond to an experimental variation (standard deviation) between 10 replicates. The y-axis represents relative editing activity in percentage, considering the complete reaction (#5) as 100%.
<table border="0" cellpadding="0" cellspacing="0" width="614">
	<tbody>
		<tr>
			<td></td>
			<td>10%</td>
			<td>30%</td>
		</tr>
		<tr>
			<td>2x HHE gradient buffer</td>
			<td>5 ml</td>
			<td>5 ml</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>1 ml</td>
			<td>3 ml</td>
		</tr>
		<tr>
			<td>DEPC H2O</td>
			<td>4 ml</td>
			<td>2 ml</td>
		</tr>
		<tr>
			<td>1 M DTT</td>
			<td>10 &#181;l</td>
			<td>10 &#181;l</td>
		</tr>
	</tbody>
</table>
Table 1. 10% &#38; 30% Glycerol Solution (10ml each).
<table border="0" cellpadding="0" cellspacing="0" width="600">
	<tbody>
		<tr>
			<td colspan="2">Pre-edited A6 Ribozyme (preA6Rbz)</td>
		</tr>
		<tr>
			<td>preA6Rbz DNA template</td>
			<td>5&#8217;-ACATTTGATCTATTGTTTCGTCCTCACGGACTCA TCAAAAGTCACAACTTTCCCTTTCTCTCCTCCCCCTAACCTTT CCCCCTATAGTGAGTCGTATTA-3&#8217;</td>
		</tr>
		<tr>
			<td>preA6Rbz RNA</td>
			<td>5&#8217;-GGGAAAGGUUAGGGGGAGGAGAGAAAGGGAAA GUUGUGACUUUUGAUGAGUCCGUGAGGACGAAACAAUAGA UCAAAUGU-3&#8217;</td>
		</tr>
		<tr>
			<td>RNA stock conc.</td>
			<td>1 &#956;M</td>
		</tr>
		<tr>
			<td colspan="2">Guide A6 Ribozyme (gA6Rbz)</td>
		</tr>
		<tr>
			<td>gA6RBz DNA template</td>
			<td>5&#8242;-AAAAAAAAAAAAAAAAATAATTATCATATCACTGT CAAGGGAAAGTTGTGAGGGTGATGAGTCCGTGTATATCCCC CTATAGTGAGTCGTATTA-3&#8242;</td>
		</tr>
		<tr>
			<td>gA6Rbz RNA</td>
			<td>5&#8242;-GGGAUAUACACGGACUCAUCACCCUCACAACUU UCCCUUGACAGUGAUAUGAUAAUUAUUUUUUUUUUUUUUUUU-3&#8217;</td>
		</tr>
		<tr>
			<td>RNA stock conc.</td>
			<td>2.5 &#956;M</td>
		</tr>
		<tr>
			<td colspan="2">Guide A6 Ribozyme Competitor (gA6RBz_comp)</td>
		</tr>
		<tr>
			<td>gA6Rbz_comp DNA template</td>
			<td>5&#8242;-GGATATACACGGACTCATCACCCTCACAACTTTC CCTTGACAGTGATATGATAATTATTTTTTTTTTTTTTTTTCCCTAT AGTGAGTCGTATTA-3&#8242;</td>
		</tr>
		<tr>
			<td>gA6Rbz_comp RNA</td>
			<td>5&#8217;-AAAAAAAAAAAAAAAAAUAAUUAUCAUAUCACUG UCAAGGGAAGUUGUGAGGGUGAUGAGUCCGUGUAUAUCC-3&#8217;</td>
		</tr>
		<tr>
			<td>RNA stock conc.</td>
			<td>12.5 &#956;M</td>
		</tr>
		<tr>
			<td colspan="2">Active A6 Ribozyme (A6RBz)</td>
		</tr>
		<tr>
			<td>A6Rbz DNA template</td>
			<td>5&#8242;-ACATTTGATCTATTGTTTCGTCCTCACGGACTCAT CAGTCACAACTTTCCCTTTCTCTCCTCCCCCTAACCTTTCCCC CTATAGTGAGTCGTATTA-3&#8242;</td>
		</tr>
		<tr>
			<td>A6Rbz RNA</td>
			<td>5&#8217;-GGGAAAGGUUAGGGGGAGGAGAGAAAGGGAAA GUUGUGACUGAUGAGUCCGUGAGGACGAAACAAUAGAUCA AAUGU-3&#8217;</td>
		</tr>
		<tr>
			<td>RNA stock conc.</td>
			<td>1 &#956;M</td>
		</tr>
		<tr>
			<td colspan="2">FRET substrate</td>
		</tr>
		<tr>
			<td>TAMRA quencher</td>
			<td>5&#8217;-FAM-GAUCUAUUGUCUCACA-TAMRA-3&#39; (Eurogentec)</td>
		</tr>
		<tr>
			<td>Iowa Black quencher</td>
			<td>5&#8217;-FAM- GAUCUAUUGUCUCACA-Iowa Black-3&#39; (IDT)</td>
		</tr>
		<tr>
			<td>RNA stock conc.</td>
			<td>15 &#956;M (for both)</td>
		</tr>
	</tbody>
</table>
Table 2. DNA Templates and RNA Substrates.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Composition</td>
			<td>Editing reaction (&#956;l)</td>
		</tr>
		<tr>
			<td>10x HHE</td>
			<td>1.5</td>
		</tr>
		<tr>
			<td>0.1 M CaCl2</td>
			<td>1</td>
		</tr>
		<tr>
			<td>100 mM ATP</td>
			<td>0.2</td>
		</tr>
		<tr>
			<td>10% Tritonx-100</td>
			<td>0.2</td>
		</tr>
		<tr>
			<td>500 ng/&#956;l Torula Yeast RNA&#160;</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Editosome</td>
			<td>5</td>
		</tr>
		<tr>
			<td>RNase free H2O</td>
			<td>7.1</td>
		</tr>
		<tr>
			<td>1 &#956;M preA6Rbz</td>
			<td>1</td>
		</tr>
		<tr>
			<td>2.5 &#956;M gA6Rbz</td>
			<td>1</td>
		</tr>
		<tr>
			<td>Total</td>
			<td>18</td>
		</tr>
	</tbody>
</table>
Table 3. Reaction Composition and Master Mix.

Watch this Scientific Journal Video about RNA Catalyst as a Reporter for Screening Drugs against RNA Editing in Trypanosomes at JoVE.com

RNA Catalyst as a Reporter for Screening Drugs against RNA Editing in Trypanosomes

A highly sensitive ribozyme-based assay, applicable to high-throughput screening of chemicals targeting the unique process of RNA editing in trypanosomatid pathogens, is described in this paper. Inhibitors can be used as tools for hypothesis-driven analysis of the RNA editing process and ultimately as therapeutics.

Substantial progress has been made in determining the mechanism of mitochondrial RNA editing in trypanosomes. Similarly, considerable progress has been ...

rna-catalyst-as-reporter-for-screening-drugs-against-rna-editing

Nanyang Technological University

Research

JoVE Journal

Biology

8.4K Views.  McGill University. A highly sensitive ribozyme-based assay, applicable to high-throughput screening of chemicals targeting the unique process of RNA editing in trypanosomatid pathogens, is described in this paper. Inhibitors can be used as tools for hypothesis-driven analysis of the RNA editing process and ultimately as therapeutics.

Video: RNA Catalyst as a Reporter for Screening Drugs against RNA Editing in Trypanosomes

In Vitro Transcription Assays and Their Application in Drug Discovery

In vitro transcription assays have been developed and widely used for many years to study the molecular mechanisms involved in transcription. This process requires multi-subunit DNA-dependent RNA polymerase (RNAP) and a series of transcription factors that act to modulate the activity of RNAP during gene expression. Sequencing gel electrophoresis of radiolabeled transcripts is used to provide detailed mechanistic information on how transcription proceeds and what parameters can affect it. In this paper we describe the protocol to study how the essential elongation factor NusA regulates transcriptional pausing, as well as a method to identify an antibacterial agent targeting transcription initiation through inhibition of RNAP holoenzyme formation. These methods can be used a as platform for the development of additional approaches to explore the mechanism of action of the transcription factors which still remain unclear, as well as new antibacterial agents targeting transcription which is an underutilized drug target in antibiotic research and development.

In Vitro Transcription Assays and Their Application in Drug Discovery

In this manuscript, we describe a protocol to functionally examine transcription and the inhibitory activity of antibacterial agents targeting bacterial transcription.

In vitro transcription assays have been developed and widely used for many years to study the molecular mechanisms involved in transcription. This process ...

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An In Vitro Assay to Detect tRNA-Isopentenyl Transferase Activity

N6-isopentenyladenosine RNA modifications are functionally diverse and highly conserved among prokaryotes and eukaryotes. One of the most highly conserved N6-isopentenyladenosine modifications occurs at the A37 position in a subset of tRNAs. This modification improves translation efficiency and fidelity by increasing the affinity of the tRNA for the ribosome. Mutation of enzymes responsible for this modification in eukaryotes are associated with several disease states, including mitochondrial dysfunction and cancer. Therefore, understanding the substrate specificity and biochemical activities of these enzymes is important for understanding of normal and pathologic eukaryotic biology. A diverse array of methods has been employed to characterize i6A modifications. Herein is described a direct approach for the detection of isopentenylation by Mod5. This method utilizes incubation of RNAs with a recombinant isopentenyl transferase, followed by RNase T1 digestion, and 1-dimensional gel electrophoresis analysis to detect i6A modifications. In addition, the potential adaptability of this protocol to characterize other RNA-modifying enzymes is discussed.

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N6-isopentenyladenosine RNA modifications are functionally diverse and highly conserved among prokaryotes and eukaryotes. One of the most highly conserved ...

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An Oligonucleotide-based Tandem RNA Isolation Procedure to Recover Eukaryotic mRNA-Protein Complexes

RNA-binding proteins (RBPs) play key roles in the post-transcriptional control of gene expression. Therefore, biochemical characterization of mRNA-protein complexes is essential to understanding mRNA regulation inferred by interacting proteins or non-coding RNAs. Herein, we describe a tandem RNA isolation procedure (TRIP) that enables the purification of endogenously formed mRNA-protein complexes from cellular extracts. The two-step protocol involves the isolation of polyadenylated mRNAs with antisense oligo(dT) beads and subsequent capture of an mRNA of interest with 3&#39;-biotinylated 2&#39;-O-methylated antisense RNA oligonucleotides, which can then be isolated with streptavidin beads. TRIP was used to recover in vivo crosslinked mRNA-ribonucleoprotein (mRNP) complexes from yeast, nematodes and human cells for further RNA and protein analysis. Thus, TRIP is a versatile approach that can be adapted to all types of polyadenylated RNAs across organisms to study the dynamic re-arrangement of mRNPs imposed by intracellular or environmental cues.

A tandem RNA isolation procedure (TRIP) for recovery of endogenously formed mRNA-protein complexes is described. Specifically, RNA-protein complexes are crosslinked in vivo, polyadenylated RNAs are isolated from extracts with oligo(dT) beads, and particular mRNAs are captured with modified RNA antisense oligonucleotides. Proteins bound to mRNAs are detected by immunoblot analysis.

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Capture and Identification of RNA-binding Proteins by Using Click Chemistry-assisted RNA-interactome Capture (CARIC) Strategy

A comprehensive identification of RNA-binding proteins (RBPs) is key to understanding the posttranscriptional regulatory network in cells. A widely used strategy for RBP capture exploits the polyadenylation [poly(A)] of target RNAs, which mostly occurs on eukaryotic mature mRNAs, leaving most binding proteins of non-poly(A) RNAs unidentified. Here we describe the detailed procedures of a recently reported method termed click chemistry-assisted RNA-interactome capture (CARIC), which enables the transcriptome-wide capture of both poly(A) and non-poly(A) RBPs by combining the metabolic labeling of RNAs, in vivo UV cross-linking, and bioorthogonal tagging.

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A detailed protocol for applying the click chemistry-assisted RNA-interactome capture (CARIC) strategy to identify proteins binding to both coding and noncoding RNAs is presented.

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There are more than 100 chemically distinct modifications of RNA, two thirds of which consist of methylations. Interest in RNA modifications, and especially methylations, has re-emerged due to the important roles played by the enzymes that write and erase them in biological processes relevant to disease and cancer. Here, a sensitive in vitro assay for accurate analysis of RNA methylation writer activity on synthetic or in vitro transcribed RNAs is provided. This assay uses a tritiated form of S-adenosyl-methionine, resulting in direct labeling of methylated RNA with tritium. The low energy of tritium radiation makes the method safe, and pre-existing methods of tritium signal amplification, make it possible to quantify and to visualize the methylated RNA without the use of antibodies, which are commonly prone to artifacts. While this method is written for RNA methylation, few tweaks make it applicable to the study of other RNA modifications that can be radioactively labeled, such as RNA acetylation with 14C acetyl coenzyme A. Overall, this assay allows to quickly assess RNA methylation conditions, inhibition with small molecule inhibitors, or the effect of RNA or enzyme mutants, and provides a powerful tool to validate and expand results obtained in cells.

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The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been harnessed by researchers to study important biologically relevant problems. The unparalleled power of the CRISPR/Cas genome editing method allows researchers to precisely edit any locus of their choosing, thereby facilitating an increased understanding of gene function. Several methods for editing the C. elegans genome by CRISPR/Cas9 have been described previously. Here, we discuss and demonstrate a method which utilizes in vitro assembled ribonucleoprotein complexes and the dpy-10 co-CRISPR marker for screening. Specifically, in this article, we go through the step-by-step process of introducing premature stop codons into the C. elegans rbm-3.2 gene by homology-directed repair using this method of CRISPR/Cas9 editing. This relatively simple editing method can be used to study the function of any gene of interest and allows for the generation of homozygous-edited C. elegans by CRISPR/Cas9 editing&#160;in less than two weeks.

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The bacterial Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Streptococcus pyogenes CRISPR-associated protein (Cas) system has been ...

Evaluation of the Efficacy And Toxicity of RNAs Targeting HIV-1 Production for Use in Gene or Drug Therapy

Small RNA therapies targeting post-integration steps in the HIV-1 replication cycle are among the top candidates for gene therapy and have the potential to be used as drug therapies for HIV-1 infection. Post-integration inhibitors include ribozymes, short hairpin (sh) RNAs, small interfering (si) RNAs, U1 interference (U1i) RNAs and RNA aptamers. Many of these have been identified using transient co-transfection assays with an HIV-1 expression plasmid and some have advanced to clinical trials. In addition to measures of efficacy, small RNAs have been evaluated for their potential to affect the expression of human RNAs, alter cell growth and/or differentiation, and elicit innate immune responses. In the protocols described here, a set of transient transfection assays designed to evaluate the efficacy and toxicity of RNA molecules targeting post-integration steps in the HIV-1 replication cycle are described. We have used these assays to identify new ribozymes and optimize the format of shRNAs and siRNAs targeting HIV-1 RNA. The methods provide a quick set of assays that are useful for screening new anti-HIV-1 RNAs and could be adapted to screen other post-integration inhibitors of HIV-1 replication.

Methods to evaluate the efficacy and toxicity of RNA molecules targeting post-integration steps of the HIV-1 replication cycle are described. These methods are useful for screening new molecules and optimizing the format of existing ones.

Small RNA therapies targeting post-integration steps in the HIV-1 replication cycle are among the top candidates for gene therapy and have the potential ...

Immunology and Infection

In Vitro Transcribed RNA-based Luciferase Reporter Assay to Study Translation Regulation in Poxvirus-infected Cells

Every poxvirus mRNA transcribed after viral DNA replication has an evolutionarily conserved, non-templated 5&#39;-poly(A) leader in the 5&#39;-UTR. To dissect the role of 5&#39;-poly(A) leader in mRNA translation during poxvirus infection we developed an in vitro transcribed RNA-based luciferase reporter assay. This reporter assay comprises of four core steps: (1) PCR to amplify the DNA template for in vitro transcription; (2) in vitro transcription to generate mRNA using T7 RNA polymerase; (3) Transfection to introduce in vitro transcribed mRNA into cells; (4) Detection of luciferase activity as the indicator of translation. The RNA-based luciferase reporter assay described here circumvents issues of plasmid replication in poxvirus-infected cells and cryptic transcription from the plasmid. This protocol can be used to determine translation regulation by cis-elements in an mRNA including 5&#39;-UTR and 3&#39;-UTR in systems other than poxvirus-infected cells. Moreover, different modes of translation initiation like cap-dependent, cap-independent, re-initiation, and internal initiation can be investigated using this method.

We present a protocol to study mRNA translation regulation in poxvirus-infected cells using in vitro Transcribed RNA-based luciferase reporter assay. The assay can be used for studying translation regulation by cis-elements of an mRNA, including 5&#8217;-untranslated region (UTR) and 3&#8217;-UTR. Different translation initiation modes can also be examined using this method.

Every poxvirus mRNA transcribed after viral DNA replication has an evolutionarily conserved, non-templated 5&#39;-poly(A) leader in the 5&#39;-UTR. To ...

A Nonsequencing Approach for the Rapid Detection of RNA Editing

RNA editing is a process that leads to posttranscriptional sequence alterations in RNAs. Detection and quantification of RNA editing rely mainly on Sanger sequencing and RNA sequencing techniques. However, these methods can be costly and time-consuming. In this protocol, a portable microtemperature gradient gel electrophoresis (&#181;TGGE) system is used as a nonsequencing approach for the rapid detection of RNA editing. The process is based on the principle of electrophoresis, which uses high temperatures to denature nucleic acid samples as they move across a polyacrylamide gel. Across a range of temperatures, a DNA fragment forms a gradient of fully double-stranded DNA to partially separated strands and then to entirely separated single-stranded DNA. RNA-edited sites with distinct nucleotide bases produce different melting profiles in &#181;TGGE analyses. We used the &#181;TGGE-based approach to characterize the differences between the melting profiles of four edited RNA fragments and their corresponding nonedited (wild-type) fragments. Pattern Similarity Scores (PaSSs) were calculated by comparing the band patterns produced by the edited and nonedited RNAs and were used to assess the reproducibility of the method. Overall, the platform described here enables the detection of even single base mutations in RNAs in a straightforward, simple, and cost-effective manner. It is anticipated that this analysis tool will aid new molecular biology findings.

Rapid detection and reliable quantification of RNA editing events at a genomic scale remain challenging and currently rely on direct RNA sequencing methods. The protocol described here uses microtemperature gradient gel electrophoresis (&#181;TGGE) as a simple, quick, and portable method of detecting RNA editing.

RNA editing is a process that leads to posttranscriptional sequence alterations in RNAs. Detection and quantification of RNA editing rely mainly on Sanger ...

RNA Catalyst as a Reporter for Screening Drugs against RNA Editing in Trypanosomes

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Evaluation of the Efficacy And Toxicity of RNAs Targeting HIV-1 Production for Use in Gene or Drug Therapy

In Vitro Transcribed RNA-based Luciferase Reporter Assay to Study Translation Regulation in Poxvirus-infected Cells

A Nonsequencing Approach for the Rapid Detection of RNA Editing