This work was supported by the National Science Foundation CAREER Award (0953583) and the National Institute of Health NCI/R21-CA116102, NCI/R21-CA125088, NIBIB/R01-EB012065, NCI/R01-CA157766.

In this protocol, one oligonucleotide (RBMB1) was labeled at its 5&#8217;-end with a CF640R reporter dye and has the sequence: 5&#8217;-mCmUmUmC mGmUmC mCmAmC mAmAmA mCmAmC mAmAmC mUmCmC mU mGmAmAmG mGmAmC mGmGmC mAmGmC mGmUmG mCmAmG mCmUmC mUmU-3&#8217;. Self-complementary domains, which drive the formation of the hairpin structure, are in bold. The second oligonucleotide (RBMB2) was labeled at the 3&#8217;-end with an Iowa Black RQ-Sp quencher and has the sequence: 5&#8217;-mGmAmG mCmUmG mCmAmC mGmCmU mGmCmC mGmUmC-3&#8217;.
1. Preparation of RBMBs
<ol>
	<li>Perform a quick spin of the tubes containing the RBMB1 and RBMB2 oligonucleotides, to ensure the dried oligonucleotide is at the bottom of the tube, and resuspend in DNase and RNase-free water to a final concentration of 100 &#956;M. For example, a 10 nmole sample of oligonucleotide should be resuspended in 100 &#956;l of water.</li>
	<li>Measure the exact concentration of RBMB1 and RBMB2 stock samples by UV-vis spectroscopy.
	<ol>
		<li>Mix 3 &#956;l of the RBMB sample with 117 &#956;l DPBS (without calcium and magnesium) in a microcentrifuge tube.</li>
		<li>Blank the spectrophotometer with DPBS and measure the absorbance from 200-800 nm. Note: Peak absorbances at 260 nm and 650 nm should be visible.</li>
		<li>Calculate the stock concentration of the RBMB samples based on the absorbance at 260 nm (or 650 nm for RBMB1), using the equation: 
		Stock Concentration (M) = A260* dilution factor/extinction coefficient where A260 is the absorbance at 260 nm and the Dilution factor is 40. 
		Note: The extinction coefficient for the RBMB can be found on the specification sheet provided by the oligonucleotide manufacturer. The extinction coefficient of CF640R at 650 nm is 103,000.</li>
	</ol>
	</li>
	<li>Mix 20 &#956;l of 100 &#956;M RBMB1, 30 &#956;l of 100 &#956;M RBMB2, 6 &#956;l 10x&#160;phosphate buffer (10x&#160;phosphate buffer: 480 mM K2HPO4, 45 mM KH2PO4, 140 mM NaH2PO4, pH 7.2) and 4 &#956;l DNase and RNase-free water. Incubate the mixture at room temperature for 30 min. Note: This is typically sufficient for approximately 50 studies.</li>
	<li>Prepare a liquid chromatography column using 75 prep grade Superdex to remove any unhybridized RBMB2 oligonucleotides. Use an 8 mL (0.7 x 20 cm) liquid chromatography column with a ~4 ml bed volume to purify the small volume of mixed probes.</li>
	<li>Wash and equilibrate the Superdex with ~50 ml of 1x&#160;phosphate buffer using a syringe pump running at a flow at a flow rate of 0.6 ml/min. Before all of the fluid enters the bed volume, stop the syringe pump, detach it, and let the remaining liquid go through the column by gravity.</li>
	<li>Load the RBMB mixture (60 &#956;l) onto the chromatography column.
	<ol>
		<li>After the RBMB sample has completely entered the bed, slowly add another 250 &#956;l of 1x&#160;phosphate buffer to the top of the bed to ensure that the entire sample has completely entered the column.</li>
		<li>Fill the column to the rim with 1x&#160;phosphate buffer. Close the top, and start the syringe pump - the syringe should be filled with ~50 ml 1x&#160;PBS and run at a flow rate of 0.6 ml/min.</li>
		<li>Once the RBMB nears the bottom of the column &#8211; the RBMB sample can typically be readily visualized due to the color of the dye incorporated into the probe &#8211; collect the flow-through at 2 drops per microcentrifuge tube (1.5 ml). Stop collecting the sample once the color within the microcentrifuge tubes becomes clear.</li>
	</ol>
	</li>
	<li>Combine the tubes containing the colored RBMB sample &#8211; typically 5-6 tubes - and load into a centrifugal filter device (10,000 MW cutoff). Centrifuge the sample at 10,000 RCF for 20 min or until the desired volume. Note: These speeds and times will typically yield a final volume of ~30 &#956;l.</li>
	<li>Measure the exact concentration of the purified RBMB sample by UV-Vis spectroscopy.
	<ol>
		<li>Take 3 &#956;l of the concentrated RBMB sample and mix with 117 &#956;l DPBS in a microcentrifuge tube.</li>
		<li>Blank the spectrophotometer with DPBS and measure the absorbance from 200-800 nm. Note: Peak absorbances at 260 nm and 650 nm should be visible.</li>
		<li>Calculate the stock concentration of the hybridized RBMB based on the absorbance at 650 nm, using the equation: 
		Stock Concentration (M) = A650* dilution factor/extinct coefficient where A650 is the absorbance at 650 nm, the Dilution factor is 40. 
		Note: The extinction coefficient for CF640R at 650nm is 103,000 and the extinction coefficient for Iowa Black RQ is ~20,000. The extinction coefficients are additive and are not sensitive to hybridization. Therefore, the stock RBMB sample has a combined extinction coefficient of approximately 123,000 at 650 nm.</li>
	</ol>
	</li>
	<li>Label the tube appropriately with name and concentration and store at -20 &#176;C for future use.</li>
</ol>
2. Preparation of Poly-d-lysine Coated 8-well Chambered Coverglass
<ol>
	<li>Prepare a 0.2 mg/ml solution of Poly-D-lysine by dissolving 5 mg of Poly-D-lysine (lyophilized powder, g-irradiated) in 25 ml of sterilized water in a sterile environment.</li>
	<li>Add 200 &#956;l of the 0.2 mg/ml Poly-D-lysine solution to each well of an 8-well chambered coverglass, in a sterile environment.</li>
	<li>Incubate at room temperature for 16-18 hr in the cell culture hood.</li>
	<li>Aspirate the Poly-D-lysine and wash the well 3 times with sterile distilled water. Note: The coated chamber slides can be stored at 4 &#176;C.</li>
</ol>
3. Probe Delivery
Note: The cell system used should contain an integrated gene construct that expresses RNA with at least 4-sequential binding sites for the RBMB in the 3&#8217;-untranslated region (UTR). The target sequences should be complementary to the loop of the RBMB. It is advised that as a negative control, the same cell line be engineered to express the same gene construct, but without the tandem repeats. In this protocol, a human fibrosarcoma cell line, HT1080, was engineered to express gfp RNA with 96-tandem repeats of the RBMB target sequence in the 3&#8217;-UTR. The control cell line was engineered to express wild-type gfp RNA.
<ol>
	<li>Plate cells in a T25 flask at 40-50% confluency one day before probe delivery. Culture the cells with DMEM media supplemented with 1% pen/strep and 10% fetal bovine serum (FBS), and incubate at 37 &#176;C with 5% CO2. </li>
	<li>The next day, turn on the microporator and set up the microporation parameters to 950 V, 2 pulses, 25 msec. Note: These parameters have been optimized for HT1080 cells, but they are cell-type dependent and can be adjusted for other cell types as described in the manufacturer&#8217;s instructions.</li>
	<li>Fill the microporation tube with 4 ml electrolytic buffer and place it on the microporation station.</li>
	<li>Take out the stock sample of purified RBMB and defrost it. Dilute several microliters to a final concentration of 12 &#956;M with 1x&#160;phosphate buffer. 1 &#956;l is needed for each microporation.</li>
	<li>Pipette 1 ml of culture medium with FBS but without antibiotics into a microcentrifuge tube. Note: This will be used to suspend the cells immediately after microporation and can be set aside, near the microporation device, for the time being. The inclusion of antibiotics in culture media will decrease the cell viability after microporation.</li>
	<li>Remove the cell culture media from the engineered HT1080 cells (60-80% confluent), wash the cells with 1 ml Ca2+ and Mg2+-free DPBS once, and incubate with 1 ml trypsin for 1-2 min.</li>
	<li>Stop the trypsinization by adding 1 ml DMEM media supplemented with 10% fetal bovine serum, without antibiotics and phenol red, and transfer the cells into two 1.5 ml microcentrifuge tubes.</li>
	<li>Spin down the cells in the microcentrifuge tube at 200 x g for 5 min. Remove the supernatant, resuspend and combine the cell pellets in a final volume of 1 ml DPBS.</li>
	<li>Take 10 &#956;l from the well-mixed cell suspension and count the cells.</li>
	<li>Pipette the cells up and down several times to make sure they are well dispersed and transfer 300,000 cells with DPBS into a new 1.5 ml microcentrifuge tube and spin down at 200 x g for 5 min.</li>
	<li>Remove the supernatant, being careful not to disturb the cell pellet. Resuspend the pellet in 11 &#956;l resuspension buffer and pipette up and down several times to ensure that the cells are well dispersed. Be sure not to generate any air bubbles. Note: Air bubbles will cause a spark during microporation and result in poor RBMB delivery and cell death.</li>
	<li>Add 1 &#956;l of the diluted RBMB (12 &#956;M) and mix well by pipetting up and down several times. Again be careful not to generate any air bubbles.</li>
	<li>Aspirate 10 &#956;l of the RBMB-cell mixture, using the microporation pipette, and insert the pipette into the microporation tube. Push the start button to start the microporation. Note: There should be no visible air bubbles in the tip.</li>
	<li>When the screen of the microporator shows completion, remove the pipette from the station, expel the 10 &#956;l mixture into the microcentrifuge tube that was prepared earlier, with 1 ml culture medium with FBS but without antibiotics. Mix gently by rocking the tube side-to-side several times.</li>
	<li>Spin the cells at 200 x g for 5 min and wash the cells two more times, in 1 ml phenol red free culture medium with FBS but without antibiotics, to remove any RBMBs that were not delivered into the cells. Resuspend with 400 &#956;l phenol red free culture medium with FBS but without antibiotics.</li>
	<li>Plate the microporated cells into the Poly-D-lysine coated 8-well chambered coverglass at 200 &#956;l per well or at the desired confluency.</li>
	<li>Optional: If it is desirable to image the nucleus, add Hoescht 33342 to the cells at a final concentration of 0.01 mg/ml.</li>
	<li>Place the chambered coverglass into a cell culture incubator. Incubate the cells for 1-2 hr prior to imaging, so that the cells have adequate time to settle down on the coverglass surface. Note: Imaging can be performed as soon as 30 min post-microporation.</li>
</ol>
4. Image Acquisition
<ol>
	<li>Turn on the live cell stage top incubation system and equilibrate it until it reaches 37 &#176;C, 5% CO2, and 75% humidity.</li>
	<li>Turn on the microscope and fluorescent light source and open the Metamorph software. Note: Other similar software packages for microscope control and image acquisition can also be used.</li>
	<li>Apply Immersol oil to the objective.</li>
	<li>Transfer the chambered coverglass with microporated cells to the live cell stage top incubation system. Incubate the stage top system until the temperature and CO2 levels are stabilized.</li>
	<li>Open the Acquire tab, in Metamorph software. Click the Show Live button to find the field, adjust the focus under white light and click the Stop Live button.</li>
	<li>Under the Acquire tab, click and open the stream acquisition pop-up button, and set up the desired movie acquisition parameters. Acquire 150 frames using the Cy5/CF640R filter and save the images to the hard drive.</li>
</ol>

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J., Pinto, M., Schanze, K., Tan, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+synthesis+of+an+oligonucleotide-poly(phenylene+ethynylene)+conjugate+with+a+precise+one-to-one+molecular+ratio.">Direct synthesis of an oligonucleotide-poly(phenylene ethynylene) conjugate with a precise one-to-one molecular ratio.</a> Angew Chem Int Ed Engl. 44 (17), 2572-2576 (2005).</li><li>Lu, J., Tsourkas, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+individual+microRNAs+in+single+mammalian+cells+in+situ.">Imaging individual microRNAs in single mammalian cells in situ.</a> Nucleic Acids Res. 37 (14), e100 (2009).</li><li>Raj, A., Peskin, C. S., Tranchina, D., Vargas, D. Y., Tyagi, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stochastic+mRNA+synthesis+in+mammalian+cells.">Stochastic mRNA synthesis in mammalian cells.</a> PLoS Biol. 4 (10), e309 (2006).</li></ol>

Dr. Mark Behlke and Dr. Ling Huang are employed by IDT which offers oligonucleotides for sale similar to some of the compounds described in the manuscript. IDT is, however, not a publicly traded company and they personally do not own any shares/equity in IDT.

The ability to image single engineered RNA transcripts in living cells using a conventional wide-field microscope requires a bright and photostable fluorescent signal to be associated with each RNA transcript and a low fluorescent background emanating from unbound fluorescent probes. In this method, a bright fluorescent signal is achieved by hybridizing multiple (up to 96) oligonucleotide-based fluorescent probes, i.e. RBMBs, onto each RNA transcript. However, as few as four bindings sites are sufficient16<...

The expression and regulation of RNA transcripts is a complex and dynamic process that is largely responsible for controlling cell behavior and fate.&#160; Although the importance of RNA in dictating cell function has been known for some time, it is often difficult to draw clear connections between the two since most RNA analysis tools lack the spatial and temporal resolution required to capture important regulatory events such as transcription bursting, RNA trafficking, and localized RNA processing. This has led to the advent of several techniques that allow individual RNA transcripts to be visualized in living cells in real-time1. Perhaps, the most prominent of these techniques utilizes a GFP-MS2 fusion protein to target RNA that has been engineered to contain tandem repeats of the MS2 binding site in the 3&#8217;-UTR2,3. By bringing multiple GFP molecules into close proximity, individual RNA transcripts appear as bright fluorescent spots via fluorescence microscopy. The GFP-MS2 system has provided unprecedented insight into RNA behavior, including direct visualization and measurements of transcriptional bursting4,5, detection of unique sub-cellular localization and processing6-9, and real-time imaging of RNA transport10. However, despite the tremendous potential of the GFP-MS2 system, unbound GFP-MS2 fusion proteins can create a high background fluorescent signal that limits the versatility and dynamic range of this technique.&#160; Several approaches have been developed to limit this background signal, including subcellular compartmentalization of unbound GFP-MS210, protein fragment complementation11, and alternative RNA binding proteins and targets12.&#160; However, all of these approaches remain sensitive to the relative and total expression of the RNA target and GFP-MS2 fusion protein.

As an alternative to the GFP-MS2 systems, molecular beacons have also been used to detect engineered RNA transcripts with tandem repeats of the complementary binding site in the 3&#8217;-UTR13,14. Molecular beacons are hairpin-forming oligonucleotide probes that are labeled at one end with a quencher and at the other end with a fluorescent reporter. When not bound to target RNA the fluorescent reporter and quencher remain in close proximity, resulting in a low-fluorescent state. Upon hybridization, the fluorescent reporter and quencher are forced apart and fluorescence is restored. Similar to the GFP-MS2 system, the binding of multiple molecular beacons onto a single RNA transcript results in a bright fluorescent spot that can be identified by fluorescence microscopy; however, the background fluorescence is expected to be much lower, due to the quenched configuration of unbound molecular beacons. Unfortunately, despite the clever activation mechanism that is incorporated into the molecular beacon design, there is now growing evidence that unhybridized molecular beacons do not remain in the hairpin conformation when introduced into living cells.&#160; Consequently, they generate a false-positive signal that significantly reduces the signal-to-background. To overcome this shortcoming, we recently developed a new synthetic probe for imaging RNA in living cells, ratiometric bimolecular beacons (RBMBs; Figure 1A)15,16. RBMBs are composed of two 2&#8217;-O-methyl oligonucleotide strands that form a hybrid structure with features from both short hairpin RNA (shRNA) and molecular beacons.&#160; The loop and fluorescence activation mechanism is similar to the molecular beacon, while the long double stranded domain with a 3&#8217;-UU overhang is more characteristic of shRNA.&#160; The shRNA features are designed to drive nuclear export, which we have found increases intracellular lifetime to &#62;24 hr, with minimal observable degradation, and prevents non-specific opening of the loop. As a result RBMBs exhibit a significantly higher signal-to-background than molecular beacons.

It should be noted that RBMBs cannot be prepared using DNA oligonucleotides, since DNA-based probes do not possess the same nuclear export capabilities as RNA-based probes. Structurally, the RBMB loop is typically designed to be between 15 and 21 bases long, to create a balance between specificity and selectivity upon RNA hybridization.&#160; The short stem that forms from the two self-complementary domains is usually designed to be 4 bases.&#160; If a longer stem is selected, the rate of hybridization between the RBMB loop and target RNA is significantly slowed17,18. Conversely, when a shorter stem sequence is selected the melting temperature is often too low to sustain the stem-loop structure at 37 &#176;C, leading to a high background signal. The specificity of the RBMB is also reduced as the stem length is shortened. Since the sequence of the RBMB stem-loop can also influence RBMB performance, it must be carefully selected19. In particular, ideal loop sequences should have minimal secondary structure, hybridize to RNA sequences with minimal secondary structure, avoid protein binding sites, and avoid off-target binding. Predictions of both RBMB and RNA secondary structure can be obtained using software such as mfold20. Complementary off-target sites can identified using a nucleotide Basic Local Assignment Search Tool (BLAST).&#160; However, because of inconsistencies in model predictions and the difficulty in identifying protein-biding sites, the specificity of all RBMBs must ultimately be validated experimentally.

If desirable, an unquenched reference dye that is insensitive to the hybridization state can be added to the RBMB15.&#160; The addition of a reference dye can provide a marker for probe delivery and be used for ratiometric imaging, if more accurate measurements of total cellular fluorescence are required.&#160; The reference dyes allows measurements to be adjusted for differences in background owing to cell-to-cell variations in delivery.&#160; However, when imaging individual RNA transcripts the reference dye is not necessary. Notably, some reference dyes can interfere with the export of RBMBs from the nucleus, leading to a slightly higher background signal in the nucleus.

When designed properly, RBMBs can be used to image individual RNA transcripts in single living cells, if the target RNA is engineered to contain at least four RBMB binding sites (Figure 1B)16. RBMBs can be efficiently delivered into a wide range of cells types via microporation, with little to no effect on cell viability21, and quantitative measurements of gene expression can be acquired within 30 min.&#160; Moreover, the methodology is fairly insensitive to RBMB concentration and target RNA levels, since fluorescence from unbound RBMBs is efficiently quenched. Here we provide a detailed description of the methodology used to prepare and purify RBMBs, as well as a general procedure for the delivery of RBMBs into live-cells via microporation and the imaging of single RNA transcripts in real-time.

<table><tbody><tr><td>Nuclease-free water</td><td>Life Technologies</td><td>AM9932</td><td>no DEPC-treated</td></tr><tr><td>DPBS</td><td>Life Technologies</td><td>14190-144</td><td>No Ca&lt;sup&gt;2+&lt;/sup&gt; and Mg&lt;sup&gt;2+&lt;/sup&gt;</td></tr><tr><td>Cary 100 UV-Vis spectrophotometer</td><td>Agilent Technologies</td><td></td><td></td></tr><tr><td>Potassium phosphate dibasic</td><td>Fisher Scientific</td><td>P290-500</td><td></td></tr><tr><td>Potassium phosphate monobasic</td><td>Fisher Scientific</td><td>P284-500</td><td></td></tr><tr><td>Sodium phosphate monobasic</td><td>Fisher Scientific</td><td>S381-500</td><td></td></tr><tr><td>Microcentrifuge tubes</td><td>Eppendorf</td><td>22364111</td><td>1.5 ml</td></tr><tr><td>Chromatography column</td><td>Kimble Chase Life Science</td><td>420400-0720</td><td>0.7 x 20 cm</td></tr><tr><td>Superdex 75 prep grade</td><td>GE healthcare</td><td>17-1044-01</td><td></td></tr><tr><td>Syringe pump</td><td>Braintree Scientific</td><td>BS-300</td><td></td></tr><tr><td>Syringe</td><td>BD</td><td>301035</td><td>60 ml, Luer-lok tip</td></tr><tr><td>Centrifugal filter units</td><td>Millipore</td><td>UFC501096</td><td>MW 10,000 cutoff</td></tr><tr><td>Centrifuge 5418</td><td>Eppendorf</td><td></td><td></td></tr><tr><td>Poly-D-lysine</td><td>Sigma-Aldrich</td><td>P7280-5MG</td><td>lyophilized powder, g-irradiated</td></tr><tr><td>8-well chambered coverglass</td><td>Fisher Scientific</td><td>155409</td><td>Working volume 0.2-0.5 ml</td></tr><tr><td>HT1080</td><td>ATCC</td><td>CCL-121</td><td>Human Fibrosarcoma cell line</td></tr><tr><td>Cell culture flask</td><td>Corning</td><td>430639</td><td>25 cm&lt;sup&gt;2&lt;/sup&gt;</td></tr><tr><td>DMEM</td><td>Life Technologies</td><td>11965084</td><td>High glucose</td></tr><tr><td>DMEM without phenol red</td><td>Life Technologies</td><td>21063029</td><td>High glucose</td></tr><tr><td>Fetal bovine serum</td><td>Sigma-Aldrich</td><td>F2442-500ML</td><td></td></tr><tr><td>Penecillin/streptomycin</td><td>Life Technologies</td><td>15140122</td><td></td></tr><tr><td>Trypsin</td><td>Life Technologies</td><td>25300054</td><td>0.05% Trypsin-EDTA</td></tr><tr><td>Neon transfection system</td><td>Life Technologies</td><td>MPK5000</td><td></td></tr><tr><td>Neon transfection system kit</td><td>Life Technologies</td><td>MPK1096</td><td></td></tr><tr><td>Hemacytometer</td><td>Fisher Scientific</td><td>02-671-10</td><td></td></tr><tr><td>Hoescht 33342</td><td>Life Technologies</td><td>H1399</td><td></td></tr><tr><td>IX-81 Inverted fluorescence microscope</td><td>Olympus</td><td></td><td></td></tr><tr><td>SOLA light engine</td><td>Lumencor</td><td></td><td></td></tr><tr><td>Metamorph</td><td>Molecular Devices</td><td></td><td>Software controlling microscope</td></tr><tr><td>Immersol oil 518F</td><td>Fisher Scientific</td><td>12-624-66B</td><td></td></tr></tbody></table>

real-time imaging of single engineered rna transcripts in living cells using ratiometric bimolecular beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Shortly following the microporation of HT1080 cells in the presence of RBMBs, individual RNA transcripts that were engineered to contain multiple RBMB binding sites in their 3&#8217;-UTR appear as bright fluorescent spots when imaged by wide-field fluorescence microscopy (Figure 2). While individual RNA transcripts with as few as four RBMB binding sites can be imaged in live-cells, the more RBMB binding sites in the 3&#8217;-UTR, the stronger the fluorescent signal.&#160; Moreover, the more RBMBs that ar...

Watch this Scientific Journal Video about Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons at JoVE.com

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

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Biology

11.7K Views.  University of Pennsylvania. Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

Video: Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed extensive studies of cap-dependent translation and its regulation, high resolution kinetic characterization of this translation pathway is still lacking. Recently, we developed an in vitro assay to measure cap-dependent translation kinetics with single-molecule resolution. The assay is based on fluorescently labeled antibody binding to nascent epitope-tagged polypeptide. By imaging the binding and dissociation of antibodies to and from nascent peptide&#8211;ribosome&#8211;mRNA complexes, the translation progression on individual mRNAs can be tracked. Here, we present a protocol for establishing this assay, including mRNA and PEGylated slide preparations, real-time imaging of translation, and analysis of single molecule trajectories. This assay enables tracking of individual cap-dependent translation events and resolves key translation kinetics, such as initiation and elongation rates. The assay can be widely applied to distinct translation systems and should broadly benefit in vitro studies of cap-dependent translation kinetics and translational control mechanisms.

An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Tracking individual translation events allows for high-resolution kinetic studies of cap-dependent translation mechanisms. Here we demonstrate an in vitro single-molecule assay based on imaging interactions between fluorescently labeled antibodies and epitope-tagged nascent peptides. This method enables single-molecule characterization of initiation and peptide elongation kinetics during active in vitro cap-dependent translation.

Cap-dependent protein synthesis is the predominant translation pathway in eukaryotic cells. While various biochemical and genetic approaches have allowed ...

Biochemistry

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk population methods (Northern blot, PCR, and RNAseq) that measure mRNA levels lack the ability to provide information on cell-to-cell variation in responses. Precise single cell and allelic visualization and quantification is possible via single molecule RNA fluorescence in situ hybridization (smFISH). RNA-FISH is performed by hybridizing target RNAs with labeled oligonucleotide probes. These can be imaged in medium/high throughput modalities, and, through image analysis pipelines, provide quantitative data on both mature and nascent RNAs, all at the single cell level. The fixation, permeabilization, hybridization and imaging steps have been optimized in the lab over many years using the model system described herein, which results in successful and robust single cell analysis of smFISH labeling. The main goal with sample preparation and processing is to produce high quality images characterized by a high signal-to-noise ratio to reduce false positives and provide data that are more accurate. Here, we present a protocol describing the pipeline from sample preparation to data analysis in conjunction with suggestions and optimization steps to tailor to specific samples.&#160;

Single molecule RNA fluorescence in situ hybridization (smFISH) is a method to accurately quantify levels and localization of specific RNAs at the single cell level. Here, we report our validated lab protocols for wet-bench processing, imaging and image analysis for single cell quantification of specific RNAs.

Gene transcription is an essential process in cell biology, and allows cells to interpret and respond to internal and external cues. Traditional bulk ...

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding location often influences function of a splicing factor.  However, as with any sampling strategy the chance of identifying an RNA bound to a splicing factor is proportional to its cellular abundance.  We have developed a novel in vitro approach for surveying binding specificity on otherwise transient pre-mRNA. This approach utilizes a specifically designed oligonucleotide pool that tiles across introns, exons, splice junctions, or other pre-mRNA.  The pool is subjected to some kind of molecular selection. Here, we demonstrate the method by separating the oligonucleotide into a bound and unbound fraction and utilize a two color array strategy to record the enrichment of each oligonucleotide in the bound fraction. The array data generates high-resolution maps with the ability to identify sequence-specific and structural determinates of ribonucleoprotein (RNP) binding on pre-mRNA.  A unique advantage to this method is its ability to avoid the sampling bias towards mRNA associated with current IP and SELEX techniques, as the pool is specifically designed and synthesized from pre-mRNA sequence. The flexibility of the oligonucleotide pool is another advantage since the experimenter chooses which regions to study and tile across, tailoring the pool to their individual needs.  Using this technique, one can assay the effects of polymorphisms or mutations on binding on a large scale or clone the library into a functional splicing reporter and identify oligonucleotides that are enriched in the included fraction.  This novel in vitro high-resolution mapping scheme provides a unique way to study RNP interactions with transient pre-mRNA species, whose low abundance makes them difficult to study with current in vivo techniques.  



A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding ...

Measuring the Kinetics of mRNA Transcription in Single Living Cells

The transcriptional activity of RNA polymerase II (Pol II) is a dynamic process and therefore measuring the kinetics of the transcriptional process in vivo is of importance. Pol II kinetics have been measured using biochemical or molecular methods.1-3 In recent years, with the development of new visualization methods, it has become possible to follow transcription as it occurs in real time in single living cells.4 Herein we describe how to perform analysis of Pol II elongation kinetics on a specific gene in living cells.5, 6 Using a cell line in which a specific gene locus (DNA), its mRNA product, and the final protein product can be fluorescently labeled and visualized in vivo, it is possible to detect the actual transcription of mRNAs on the gene of interest.7, 8 The mRNA is fluorescently tagged using the MS2 system for tagging mRNAs in vivo, where the 3'UTR of the mRNA transcripts contain 24 MS2 stem-loop repeats, which provide highly specific binding sites for the YFP-MS2 coat protein that labels the mRNA as it is transcribed.9 To monitor the kinetics of transcription we use the Fluorescence Recovery After Photobleaching (FRAP) method. By photobleaching the YFP-MS2-tagged nascent transcripts at the site of transcription and then following the recovery of this signal over time, we obtain the synthesis rate of the newly made mRNAs.5 In other words, YFP-MS2 fluorescence recovery reflects the generation of new MS2 stem-loops in the nascent transcripts and their binding by fluorescent free YFP-MS2 molecules entering from the surrounding nucleoplasm. The FRAP recovery curves are then analyzed using mathematical mechanistic models formalized by a series of differential equations, in order to retrieve the kinetic time parameters of transcription.

RNA polymerase II transcriptional kinetics are measured on specific genes in living cells. mRNAs transcribed from the gene of interest are fluorescently tagged and using Fluorescence Recovery After Photobleaching (FRAP) the in vivo kinetics of transcriptional elongation are obtained.

The transcriptional activity of RNA polymerase II (Pol II) is a dynamic process and therefore measuring the kinetics of the transcriptional process in ...

Dual-Color End-Labeling and Vesicle Encapsulation of RNAs for smFRET-TIRF Studies

Fluorescent End-Labeling and Encapsulation of Long RNAs for Single-Molecule FRET-TIRF Microscopy

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) excels in studying dynamic biomolecules by allowing precise observation of their conformational changes over time. To monitor RNA dynamics with smFRET, we developed a method to covalently label RNAs at their termini with a FRET pair of fluorophores. This direct end-labeling strategy targets the 5&#39;-phosphate by carbodiimide (EDC)/N-hydroxysuccinimide (NHS) activation and the 3&#39;-ribose by periodate oxidation, which can be adapted to other RNAs regardless of their size and sequence to study them independently of artificial modifications. Furthermore, the 5&#39;-EDC/NHS activation is of general interest to all nucleic acids with a 5&#39;-phosphate. The use of commercially available chemicals eliminates the need to synthesize RNA-specific probes.
Total Internal Reflection Fluorescence (TIRF) microscopy requires the surface-immobilized molecules of interest to be within the evanescent field to be illuminated. A sophisticated way of keeping the RNA molecules within the evanescent field is to encapsulate them in phospholipid vesicles. Encapsulation benefits from the best of both worlds, tethering the molecule to the surface while enabling free diffusion of the molecule. We ensure that each vesicle contains only a single RNA molecule, enabling single-molecule imaging. Upon dual-end labeling and encapsulation of the RNA of interest, smFRET measurements offer a dynamic and detailed view of RNA behavior.

This article describes the dual-color labeling of long RNAs at termini positions and their surface immobilization via encapsulation in phospholipid vesicles for single-molecule FRET TIRF microscopy applications. Combining these techniques enables precise visualization and analysis of RNA dynamics at the single-molecule level.

Single-molecule F&#246;rster Resonance Energy Transfer (smFRET) excels in studying dynamic biomolecules by allowing precise observation of their ...

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method makes it possible to count the numbers of protein molecules produced in one cell during sequential, five-minute time windows. It requires a fluorescence microscope with laser excitation power density of ~0.5 to 1 kW/cm2, which is sufficiently sensitive to detect single fluorescent protein molecules in live cells. The fluorescent reporter used in this method consists of three parts: a membrane targeting sequence, a fast-maturing, yellow fluorescent protein and a protease recognition sequence. The reporter is translationally fused to the N-terminus of a protein of interest. Cells are grown on a temperature-controlled microscope stage. Every five minutes, fluorescent molecules within cells are imaged (and later counted by analyzing fluorescence images) and subsequently photobleached so that only newly translated proteins are counted in the next measurement.
Fluorescence images resulting from this method can be analyzed by detecting fluorescent spots in each image, assigning them to individual cells and then assigning cells to cell lineages. The number of proteins produced within a time window in a given cell is calculated by dividing the integrated fluorescence intensity of spots by the average intensity of single fluorescent molecules. We used this method to measure expression levels in the range of 0-45 molecules in single 5 min time windows. This method enabled us to measure noise in the expression of the &lambda; repressor CI, and has many other potential applications in systems biology.

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage CoTrAC

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC), to image the production of protein molecules in live cells with single-molecule precision without perturbing the protein's functionality. This method has been used to follow the stochastic expression dynamics of a transcription factor, the &lambda; repressor CI 1.

We describe a fluorescence microscopy method, Co-Translational Activation by Cleavage (CoTrAC) to image the production of protein molecules in live cells ...

Metabolic Labeling of Newly Transcribed RNA for High Resolution Gene Expression Profiling of RNA Synthesis, Processing and Decay in Cell Culture

The development of whole-transcriptome microarrays and next-generation sequencing has revolutionized our understanding of the complexity of cellular gene expression. Along with a better understanding of the involved molecular mechanisms, precise measurements of the underlying kinetics have become increasingly important. Here, these powerful methodologies face major limitations due to intrinsic properties of the template samples they study, i.e. total cellular RNA. In many cases changes in total cellular RNA occur either too slowly or too quickly to represent the underlying molecular events and their kinetics with sufficient resolution. In addition, the contribution of alterations in RNA synthesis, processing, and decay are not readily differentiated.
We recently developed high-resolution gene expression profiling to overcome these limitations. Our approach is based on metabolic labeling of newly transcribed RNA with 4-thiouridine (thus also referred to as 4sU-tagging) followed by rigorous purification of newly transcribed RNA using thiol-specific biotinylation and streptavidin-coated magnetic beads. It is applicable to a broad range of organisms including vertebrates, Drosophila, and yeast. We successfully applied 4sU-tagging to study real-time kinetics of transcription factor activities, provide precise measurements of RNA half-lives, and obtain novel insights into the kinetics of RNA processing. Finally, computational modeling can be employed to generate an integrated, comprehensive analysis of the underlying molecular mechanisms.

Total cellular RNA provides a poor template for studying short-term changes in RNA synthesis and decay as well as the kinetics of RNA processing. Here, we describe metabolic labeling of newly transcribed RNA with 4-thiouridine followed by thiol-specific biotinylation and purification of newly transcribed RNA allowing to overcome these limitations.

The development of whole-transcriptome microarrays and next-generation sequencing has revolutionized our understanding of the complexity of cellular gene ...

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH allows the measurement of cell-to-cell variability in mRNA copy number of genes of interest, as well as the subcellular location of the transcripts. The main steps involved are fixation of the bacterial cell culture, permeabilization of cell membranes, and hybridization of the target transcripts with sets of commercially available short fluorescently-labeled oligonucleotide probes. smFISH can allow the imaging of the transcripts of multiple genes in the same cell, with limitations imposed by the spectral overlap between different fluorescent markers. Following completion of the protocol illustrated below, cells can be readily imaged using a microscope coupled with a camera suitable for low-intensity fluorescence. These images, together with cell contours obtained from segmentation of phase contrast frames, or from cell membrane staining, allow the calculation of the mRNA copy number distribution of a sample of cells using open-source or custom-written software. The labeling method described here can also be applied to image transcripts with stochastic optical reconstruction microscopy (STORM).

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

This manuscript describes a method for labeling individual messenger RNA (mRNA) transcripts with fluorescently-labeled DNA probes, for use in single-molecule fluorescence in situ hybridization (smFISH) experiments in E. coli. smFISH is a visualization method that allows the simultaneous detection, localization, and quantification of single mRNA molecules in fixed individual cells.

A method is described for labeling individual messenger RNA (mRNA) transcripts in fixed bacteria for use in single-molecule fluorescence in situ ...

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell imaging studies. Methods for detecting RNAs have expanded greatly since seminal studies linked site-specific mRNA localization to gene expression regulation. Dynamic mRNA processes can now be visualized via approaches that detect mRNAs, coupled with microscopy set-ups that are fast enough to capture the dynamic range of molecular behavior. The molecular beacon technology is a hybridization-based approach capable of direct detection of endogenous transcripts in living cells. Molecular beacons are hairpin-shaped, internally quenched, single-nucleotide discriminating nucleic acid probes, which fluoresce only upon hybridization to a unique target sequence. When coupled with advanced fluorescence microscopy and high-resolution imaging, they enable one to perform spatial and temporal tracking of intracellular movement of mRNAs. Although this technology is the only method capable of detecting endogenous transcripts, cell biologists have not yet fully embraced this technology due to difficulties in designing such probes for live cell imaging. A new software application, PinMol, allows for enhanced and rapid design of probes best suited to efficiently hybridize to mRNA target regions within a living cell. In addition, high-resolution, real-time image acquisition and current, open source image analysis software allow for a refined data output, leading to a finer evaluation of the complexity underlying the dynamic processes involved in the mRNA&#39;s life cycle.
Here we present a comprehensive protocol for designing and delivering molecular beacons into Drosophila melanogaster egg chambers. Direct and highly specific detection and visualization of endogenous maternal mRNAs is performed via spinning disc confocal microscopy. Imaging data is processed and analyzed using object detection and tracking in Icy software to obtain details about the dynamic movement of mRNAs, which are transported and localized to specialized regions within the oocyte.

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Here, we present a protocol for the visualization, detection, analysis and tracking of endogenous mRNA trafficking in live Drosophila melanogaster egg chamber using molecular beacons, spinning disc confocal microscopy, and open-source analysis software.

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell ...

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization

Single-molecule fluorescence in situ hybridization (smFISH) allows for counting the absolute number of mRNAs in individual cells. Here, we describe an application of smFISH to measure the rates of transcription and mRNA degradation in Escherichia coli. As smFISH is based on fixed cells, we perform smFISH at multiple time points during a time-course experiment, i.e., when cells are undergoing synchronized changes upon induction or repression of gene expression. At each time point, sub-regions of an mRNA are spectrally distinguished to probe transcription elongation and premature termination. The outcome of this protocol also allows for analyzing intracellular localization of mRNAs and heterogeneity in mRNA copy numbers among cells. Using this protocol many samples (~50) can be processed within 8 h, like the amount of time needed for just a few samples. We discuss how to apply this protocol to study the transcription and degradation kinetics of different mRNAs in bacterial cells.

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization

This protocol describes an application of single-molecule fluorescence in situ hybridization (smFISH) to measure the in vivo kinetics of mRNA synthesis and degradation.

Single-molecule fluorescence in situ hybridization (smFISH) allows for counting the absolute number of mRNAs in individual cells. Here, we describe an ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

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An In Vitro Single-Molecule Imaging Assay for the Analysis of Cap-Dependent Translation Kinetics

Single Cell Analysis Of Transcriptionally Active Alleles By Single Molecule FISH

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Measuring the Kinetics of mRNA Transcription in Single Living Cells

Fluorescent End-Labeling and Encapsulation of Long RNAs for Single-Molecule FRET-TIRF Microscopy

Single-molecule Imaging of Gene Regulation In vivo Using Cotranslational Activation by Cleavage (CoTrAC)

Metabolic Labeling of Newly Transcribed RNA for High Resolution Gene Expression Profiling of RNA Synthesis, Processing and Decay in Cell Culture

Method for Labeling Transcripts in Individual Escherichia coli Cells for Single-molecule Fluorescence In Situ Hybridization Experiments

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization