We thank Anne Dodson and Stephanie Heinrich for advice on how to optimize the smFISH protocol, Xavier Darzacq for help with the analysis platform, Haiyan Huang for suggestions on the statistical analysis. This work was supported by funds from the March of Dimes (5-FY15-99), Pew Charitable Trusts (00027344), Damon Runyon Cancer Research Foundation (35-15) and Glenn Foundation to E&#220;, and a NSF Graduate Research Fellowship Grant No. DGE-1106400 to JC.

NOTE: All the buffers and the media used in this protocol are listed in Table 1. The vendor information for the reagents is listed in the Table of Materials.
Day 1/Day 1-2:
NOTE: For vegetative culture, grow cells to an OD600 of 0.4 to 0.6 in a preferred medium. For meiotic culture, induce cells to undergo meiosis using a preferred method (typically culturing in&#160;an OD600 of 1.85).
1. Sample Fixation and Digestion
<ol>
	<li>Fix a total of ~3.5 OD600 of cells in 3% formaldehyde.
	<ol>
		<li>For vegetative culture, add 5.52 mL of culture to 480 &#181;L of 37% formaldehyde in 15-mL conical tubes.</li>
		<li>For meiotic culture, add 1840 &#181;L of culture to 160 &#181;L of 37% formaldehyde in 2-mL microcentrifuge tubes. Invert ~5 times to mix. 
		CAUTION: Formaldehyde is toxic. Handle and dispose according to institutional regulations.</li>
	</ol>
	</li>
	<li>Place tubes on a roller drum at room temperature for 20 min. For meiotic samples, after fixing at room temperature for 20 min, continue fixing overnight at 4 &#730;C, rotating. 
	NOTE: The overnight fixation increases the reproducibility and quality of the smFISH data obtained for&#160;meiotic samples. Fixation time should be optimized.</li>
	<li>While the samples are fixing, thaw 200 mM Vanadyl Ribonucleoside Complexes (VRC) at 65 &#730;C for at least 10 min.</li>
	<li>Prepare digestion master mix in a 15-mL tube: for 1 sample, mix 425 &#181;L of Buffer B with 40 &#181;L of 200 mM VRC (warmed to 65 &#730;C);&#160;for 5 samples, mix 2125&#160;&#181;L of Buffer B with 200 &#181;L of 200 mM VRC (warmed to 65 &#730;C). Vortex ~5 s to fully resuspend the VRC before adding to the master mix, which will appear light brownish green after VRC addition. 
	NOTE: Addition of VRC during digestion improves the consistency of smFISH results, potentially by inhibiting the nuclease contaminant introduced by the zymolyase mixture, which is purified from crude extract. The amount of VRC required at this step should be optimized.</li>
	<li>For vegetative samples, centrifuge the tubes at ~1057 x g for 3 min. For meiotic samples (after overnight fixation), centrifuge at 21,000 x g for 1.5 min. Decant or aspirate the supernatant to formaldehyde waste. 
	NOTE: All the centrifugation steps are performed at room temperature.</li>
	<li>Resuspend cells in 1.5 mL of cold Buffer B by pipetting up and down or by inverting the tubes and flicking to mix. Transfer vegetative samples to 2-mL tubes after resuspension.</li>
	<li>Centrifuge at 21,000 x g for 1.5 min. Remove the bulk of the liquid by vacuum aspiration or by pipetting, leaving behind ~100 &#181;L.</li>
	<li>Resuspend the cells in 1.5 mL of cold Buffer B.</li>
	<li>Centrifuge at 21,000 x g for 1.5 min. Remove the bulk of the liquid by vacuum aspiration or by pipetting, leaving behind ~100 &#181;L.</li>
	<li>Resuspend cells in 1.5 mL of cold Buffer B. Centrifuge at 21,000 x g for 1.5 min. Aspirate the liquid completely by vacuum or by pipetting.</li>
	<li>Resuspend cells in 425 &#181;L of digestion master mix, and briefly vortex to resuspend. Add 5 &#181;L of 100T 10 mg/mL zymolyase to each tube. 
	NOTE: Vortex the zymolyase every time before adding to the tubes. Add zymolyase to each tube individually, instead of adding to the master mix. Both steps help maintain digestion consistency among tubes because zymolyase precipitates quickly.</li>
	<li>Vortex 2-3 s to mix. Place the tubes on a roller drum, and digest at 30 &#730;C for 15-30 min. For vegetative cells and early meiotic stages, it usually takes ~15 min, and for meiotic prophase and meiotic divisions, it usually takes ~30 min. 
	NOTE: Check on the microscope every 5 min after 15 min, and stop the digestion when ~80% of cells appear non-transparent and non-refractive. From this point on, cells are very fragile. Handle cells gently and avoid using vacuum aspiration or vortexing.</li>
	<li>Centrifuge the tubes at ~376 x g for 3 min. Remove the liquid completely by pipetting.</li>
	<li>Gently resuspend cells with 1 mL of Buffer B by pipetting up and down 1-2 times to mix.</li>
	<li>Centrifuge the tubes at ~376 x g for 3 min. Remove Buffer B liquid by pipetting. Gently resuspend cells in 1 mL of 70% ethanol (diluted with RNase-free water).</li>
	<li>Incubate at room temperature for 3.5-4 hours.</li>
</ol>
2. Hybridization
<ol>
	<li>Bring formamide to room temperature (for 50-mL aliquot, it takes ~30 min in water bath). 
	NOTE: Do not open the formamide bottle until the bottle reaches room temperature to avoid oxidation of formamide. 
	CAUTION: Formamide is toxic. Handle and dispose according to institutional regulations.</li>
	<li>Prepare 10% formamide wash buffer in a 15-mL conical tube.</li>
	<li>Centrifuge the tubes at ~376 x g for 3 min. Remove 500 &#181;L of 70% ethanol by pipetting. Gently pipet up and down to resuspend the remaining cells, and then transfer the cells to low-adhesion tubes. 
	NOTE: Using low-adhesion tubes greatly reduces cell loss during subsequent washes.</li>
	<li>Centrifuge the tubes again at ~376 x g for 3 min. Remove all the ethanol by pipetting.</li>
	<li>Add 1 mL of 10% formamide wash buffer, and gently pipet up and down 2-3 times to resuspend the cells.</li>
	<li>Allow the cells to sit at room temperature for ~20 min while preparing the Hybridization Solution. For 1 sample, mix&#160;50 &#181;L of Hybridization Buffer (room temp.), 5 &#181;L of 200 mM VRC (warmed to 65 &#730;C), and 1 &#181;L of each probe (200 nM final). For 5 samples, mix 250 &#181;L of Hybridization Buffer (room temp.), 25 &#181;L of 200 mM VRC (warmed to 65 &#730;C), and 5 &#181;L of each probe (200 nM final).
	<ol>
		<li>Thaw Hybridization Buffer (frozen at -20 &#730;C) to room temperature before opening the tube to prevent formamide oxidation.</li>
		<li>After adding the 200 mM VRC, vortex for 5-10 s before adding the probes, which are designed and purchased commercially, and reconstituted according to the manufacturer&#39;s instructions.</li>
		<li>If two probes are co-incubated, add 1 &#181;L of the 1:10 dilution of the probe #1 stock, as well as 1 &#181;L of the 1:10 dilution of the probe #2 stock, to make a final dilution of ~1:500 for either&#160;probe. The concentration needed for the best signal-to-noise ratio needs to be optimized (See Discussion for details).</li>
	</ol>
	</li>
	<li>Centrifuge the samples at ~376 x g for 3 min. Remove the supernatant into hazardous waste by pipetting.</li>
	<li>Add at least 50 &#181;L of&#160;Hybridization Solution to each tube. Flick the tubes to mix.</li>
	<li>Incubate at 30 &#730;C on a roller drum for at least 16 hours in the dark.</li>
</ol>
Day 2/Day 3
3. Washing and imaging
<ol>
	<li>Bring formamide to room temperature.</li>
	<li>Prepare 10% formamide wash buffer (FWB) in a 15-mL conical tube.</li>
	<li>Remove the tubes from the roller drum and place them into a foil-covered box to protect from light.&#160;</li>
	<li>Centrifuge the samples at ~376 x g for 3 min.&#160;Remove the supernatant into hazardous waste by pipetting.</li>
	<li>Resuspend in 1 mL of 10% FWB by gently pipetting up and down 2-3 times.</li>
	<li>Incubate at 30 &#730;C for 30 min (not rotating) in the&#160;foil-covered box.</li>
	<li>Centrifuge the samples at ~376 x g for 3 min. Remove the supernatant to hazardous waste by pipetting, and leave ~50 &#181;L.</li>
	<li>Meanwhile, prepare DAPI/FWB in a 15-mL conical tube: For&#160;1 sample, mix 1000 &#181;L of 10% formamide wash buffer with 1 &#181;L of 5 mg/mL DAPI. For 10 samples, mix 10 mL of 10% formamide wash buffer with 10 &#181;L of 5 mg/mL DAPI. Resuspend in 1 mL of DAPI/FWB by gently pipetting up and down&#160;2-3 times.</li>
	<li>Incubate at 30 &#730;C for 30 min (not rotating) in the&#160;foil-covered box.</li>
	<li>Thaw anti-bleach reagents on ice.</li>
	<li>Centrifuge the samples at ~376 x g for 3 min. Remove the supernatant completely by pipetting. If needed, centrifuge again to remove all of the supernatant.</li>
	<li>For samples that are not imaged immediately, resuspend the pellet in 50 &#181;L of GLOX Buffer without enzymes. Pipet up and down 3-4 times to mix.
	<ol>
		<li>Keep all the unimaged samples in the foil-covered box at 4 &#730;C until ready to image. When ready to image, centrifuge the samples at ~376 x g for 2 min and remove the supernatant completely by pipetting. Resuspend in GLOX with enzymes as below.</li>
	</ol>
	</li>
	<li>For samples being imaged immediately, add 15-20 &#181;L of GLOX buffer with enzymes.&#160;Gently pipet up and down to mix. 
	NOTE: The volume added can&#160;vary&#160;depending on the size of the cell pellet. We recommend resuspending the pellet in 15 &#181;L of GLOX buffer with enzymes and check the cell density on microscope. If cells are too dense (cells clumping on top of one another), add extra GLOX buffer with enzymes. If cells are too sparse, centrifuge the sample and remove ~5 &#181;L of buffer.</li>
	<li>Pipet 5 &#181;L onto a coverslip (18 mm x 18 mm, No. 1), and put the coverslip on a slide.</li>
	<li>Put a laboratory wipe on top of the slide where the coverslip is placed. Gently press on the laboratory wipe to set slide (should see liquid coming off from all four edges of the coverslip).</li>
	<li>Transfer the slide to the microscope room in a box covered by aluminum foil.</li>
	<li>Image using a wide-field fluorescence microscope with high magnification (60-100X) and high numerical aperture. 
	NOTE: To collect the maximal number of photons emitted by the smFISH probes, confocal microscopes are not recommended, as they significantly limit the amount of light to be collected. Here, the data were collected on a microscope equipped with a 100X, 1.4 NA objective, using filters CY5 (EX632/22, EM679/34) imaged at 1.3 s, 100% T; TRITC (EX542/27, EM597/45), 1.3 s, 100% T; and DAPI (EX390/18, EM435/48), 0.05-0.1 s, 32%-50% T. A bright-field reference image should also be acquired. Acquire 15-25 slices with a step size of 0.1-0.2 &#181;m, from entirely below the focus of the field of view to entirely above, in order to ensure that all of the RNA spots are accounted for.</li>
</ol>
4. Image Analysis
<ol>
	<li>Analyze the smFISH data with published smFISH analysis tools such as FISH-quant17&#160;and StarSearch2, or with custom-made programs, depending on the exact requirements of the experiment. 
	NOTE: A good analysis pipeline should allow the users to determine a threshold to separate the true RNA spots from the background, and output statistics such as the X, Y, and Z (optional) coordinates and the intensity of each spot, the number of spots in each cell, and possibly fitting parameters as a way to filter out false detections.</li>
</ol>

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A., van Oudenaarden, A., 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=Imaging+individual+mRNA+molecules+using+multiple+singly+labeled+probes.">Imaging individual mRNA molecules using multiple singly labeled probes.</a> Nat Methods. 5 (10), 877-879 (2008).</li><li>Rahman, S., Zenklusen, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-molecule+resolution+fluorescent+in+situ+hybridization+(smFISH)+in+the+yeast+S.+cerevisiae.">Single-molecule resolution fluorescent in situ hybridization (smFISH) in the yeast S. cerevisiae.</a> Methods Mol Biol. 1042, 33-46 (2013).</li><li>Gaspar, I., Ephrussi, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strength+in+numbers:+quantitative+single-molecule+RNA+detection+assays.">Strength in numbers: quantitative single-molecule RNA detection assays.</a> Wiley Interdiscip Rev Dev Biol. 4 (2), 135-150 (2015).</li><li>Vargas, D. 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M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mating+type+switching+in+yeast+controlled+by+asymmetric+localization+of+ASH1+mRNA.">Mating type switching in yeast controlled by asymmetric localization of ASH1 mRNA.</a> Science. 277 (5324), 383-387 (1997).</li><li>Park, H. Y., Trcek, T., Wells, A. L., Chao, J. A., Singer, R. 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The authors have no conflict of interest to disclose.

The protocol presented here is derived from other published smFISH protocols2,3,21,22,23, and is specifically optimized for the yeast strain background SK1. The parameters optimized included the cell number, fixation duration, centrifugation speed and duration, digestion duration, digestion buffer, probe concentration, and hybridization buffer. Other parameters such as hybridization temperature and duration were not optimized. In this section, we share a few notes that would help adapt this protocol to any yeast strain background and the growth conditions of interest.
The cell wall of budding yeast is one major challenge against obtaining high-quality smFISH images in budding yeast because the cell wall prevents probe penetration. Incomplete digestion of the cell wall by zymolyase leads to inefficient hybridization of the probes and high cell-to-cell variability in signal. However, over-digestion can make&#160;cells too fragile, leading to significant cell loss during the washing steps and cell bursting during slide preparation for imaging. Thus, optimizing the duration of digestion is crucial. We recommend conducting a pilot smFISH experiment by digesting the same sample for different amounts of time. In our case, we obtained the best smFISH data if we stopped the digestion when ~80% of cells became non-refractive. Typically, for the same growth condition, different genetic mutant strains can be digested with similar timing. However, the duration of digestion differs when compared among different growth conditions and different stages of meiosis in budding yeast. Once the timing is determined, the quality of smFISH is reproducible. Note that for mutant strains with defective cell wall synthesis and/or composition, digestion may be completed in less than 15 min. Use of different batches of zymolyase may also slightly change the digestion timing.
Optimal probe concentration is needed to achieve a high signal-to-noise ratio. We designed and purchased our smFISH probes commercially. Good probes (often 20-mers) should have a percentage of GC ranging&#160;from 35% to 45%, a minimum spacing of 2 base pairs between probes, and low cross-reactivity. We used a web-based probe designer from the manufacturer to generate a list of probes, and if there were enough probes to choose from, we would use the BLAST algorithm in the Saccharomyces Genome Database to eliminate the probes with more than 17 base pairs overlapped with other genomic regions. We chose the more photostable fluorescent dye (CAL Fluor 590) for the probe set that anneals to the unique region of NDC80luti (CF 590) because in this set, the number of probes that satisfied all of the above criteria (20 probes) was lower than the minimum number recommended by the manufacturer (~25 probes). After reconstituting the probe solution following the manufacturer&#39;s instructions, we made a 1:10 dilution of the stock solution and stored the diluted solution in 5-&#181;L aliquots at -20 &#730;C. Each aliquot was used only one time. For optimization, one should make serial dilutions from the 1:10 diluted solution and test which concentration yields the best signal-to-noise ratio. We recommend making a dilution series of 1:250, 1:500, 1:1000, and 1:2000 from the original stock. In our case, a 1:500 dilution factor resulted in the best signal-to-noise ratio, for both the CF 590 and Q 670 probe sets.
Optimal fixation time is also critical for successful smFISH in budding yeast. We found that overnight fixation at 4 &#730;C, rather than fixing at room temperature for 20 min, significantly improved the consistency and quality of the smFISH results for meiotic samples. Although we have not tested how overnight fixation might impact vegetative samples, fixation at room temperature worked well for this growth condition. Thus, to optimize the smFISH protocol for new strains or growth conditions, we recommend starting with short fixation time at room temperature and increasing the fixation time if high variability of signal is encountered.
Compared with other published protocols3,22,23, our protocol uses&#160;the RNase inhibitor VRC during digestion and&#160;a higher concentration of VRC during hybridization. Addition of VRC in these two steps improved the consistency of the smFISH results, possibly by better preserving RNA molecules against nuclease activity, which may be introduced by the zymolyase mix (the enzyme is purified from crude extracts and may contain RNase contaminants). Thus, we recommend using the amount of VRC as listed in our protocol or even higher concentrations of VRC for optimization.
A significant portion of the cells can be lost during the washing steps in both Buffer B and the formamide wash buffer. To reduce cell loss, one could increase the centrifugation speed. In our case, using a high speed (21,000 x g) to pellet our samples during the Buffer B washes significantly reduced&#160;cell loss. However, cells become very fragile after zymolyase digestion, so changing centrifugation speed is not recommended. Instead, we suggest using the low-adhesion tubes from USA Scientific, which greatly help pellet the cells during the washes in the formamide wash buffer. Overall, our protocol can consistently generate a monolayer of cells dense enough for efficient imaging. Typically, 7 fields of view should yield &#62;130 cells suitable for quantification.
Lastly, it is important to determine the optimal parameters for and the outputs needed from image analysis. To detect smFISH spots, published analysis programs commonly filter the raw images using Gaussian kernel to remove background signal, and ask the users to determine the signal-to-noise ratio to use for each set of images2,17. Unfortunately, there is not currently a single standard by which the proper parameters are determined, and so some empirical testing of these different settings is necessary. To set the parameters needed for each of these steps, one needs to iteratively input different sets of parameters and check how well the results obtained from each set matches that from manual counting in a few representative cells. Once one set of parameters is found, it can be used for most of the images obtained despite different growth conditions and genetic backgrounds.
In addition, one can test if maximum-intensity projection of the smFISH images can be performed prior to quantification22. This step simplifies the spot detection algorithm and significantly reduces imaging time, though at the cost of some potentially useful information about individual spots, such as their subcellular localization. In our case, the number of mRNA molecules produced by the NDC80 gene was low enough that the spots were well separated after this processing (not uncommon for transcripts in budding yeast3,7). In cases where colocalization analysis is critical, the analysis pipeline needs to determine the location of each smFISH spot in each channel to assess colocalization. Depending on the specific questions being asked, other information such as the intensity of each spot may also need to be extracted from the pipeline for further analysis. Optimizing the key steps in the protocol and the image analysis pipeline is crucial in obtaining high quality of smFISH data to study the question of interest.

Dynamic regulation of gene expression drives the development of an organism, as well as its response to environmental stress, infection, and changes in metabolism. Studies that focus on transcriptional regulation rely on technologies that measure RNA abundance. One such method, termed single molecule fluorescence in situ hybridization (smFISH), is used for detection of individual RNA molecules in single cells1,2. This method allows&#160;measurement of the cell-to-cell variability in gene expression and determination of intracellular RNA localization.
In the most commonly used smFISH technique, detection of an individual RNA molecule requires multiple short DNA probes (often ~48 20-mer probes) that are complementary to the target RNA&#160;and are conjugated to the same fluorescent dye. Binding of a single fluorescent probe results in weak signal, but the signal from the ensemble of all the probes is robust. This feature greatly improves the signal-to-noise ratio because even though a single probe can exhibit off-target binding, such signal is expected to be very weak compared to that of the target RNA molecule3. Within error of detection, the number of RNA molecules can be counted and compared across various growth conditions and among different mutants.
Since its first development, smFISH has been adapted to study various aspects of gene expression4, such as transcription elongation1,2, splicing5,6, transcriptional bursting7,8,9, intracellular allelic expression10,11,12, and RNA localization13,14,15. Recently, we have used this method to study the expression of two transcript isoforms of the same gene, in which the short transcript isoform (NDC80ORF) has its entire sequence shared with the long isoform (NDC80luti)16. To uniquely identify the two mRNA isoforms, we used two probe sets: one set is specific to the unique sequence of NDC80luti, and the other set, conjugated to another fluorescent dye, binds to the common region of the two isoforms. The&#160;NDC80luti RNA is identified as a colocalized spot with both fluorescent signals, whereas the NDC80ORF RNA is the one that contains only the signal from the common probe set. Since the number of the NDC80ORF transcripts is calculated by a &#34;subtraction&#34; method, high efficiency of probe hybridization and a high signal-to-noise ratio are necessary to confidently identify smFISH spots and reduce error propagation.
This paper describes an optimized protocol for performing smFISH in the budding yeast Saccharomyces cerevisiae. In this protocol, the cell number, fixation time, digestion time, digestion buffer, probe concentration, and hybridization buffer used for the smFISH experiments were optimized for the SK1 strain background of S. cerevisiae undergoing vegetative growth or meiosis. However, we also note in this manuscript: (1) the method to check the quality of smFISH data after image acquisition, and (2) the steps in the protocol that may require additional optimization for different strain backgrounds and growth conditions.

<table><tbody><tr><td>BSA, RNase-free (50 mg/mL)</td><td>Ambion</td><td>AM2616</td><td>Store at -20 &amp;deg;C.</td></tr><tr><td>Catalase</td><td>Sigma</td><td>C3515</td><td>Store at 4 &amp;deg;C for short-term. Vortex before use.</td></tr><tr><td>DAPI</td><td>Sigma</td><td>D9564</td><td>Store at -20 &amp;deg;C after reconstitution in water. Protect from light.</td></tr><tr><td>50% Dextran Sulfate</td><td>Milli Pore</td><td>S4030</td><td>Store at room temperature. Very viscous liquid. Handle with patience.</td></tr><tr><td>&lt;em&gt;E. coli&lt;/em&gt; tRNA</td><td>Sigma</td><td>R4251</td><td>Store at -20 &amp;deg;C in aliquots after reconstitution in water.</td></tr><tr><td>Ethanol (100%, 200 proof)</td><td>various</td><td></td><td>Flammable.</td></tr><tr><td>37% Formaldehyde</td><td>Fisher</td><td>F79-500</td><td>Store at room temperature. Toxic. Use and dispose with caution.</td></tr><tr><td>Formamide</td><td>Ambion</td><td>AM9342</td><td>Store at 4 &amp;deg;C. Open after the temperature equilibrates to room temperature in order to prevent oxidation. Toxic. Use and dispose with caution.&amp;nbsp;</td></tr><tr><td>Glucose</td><td>various</td><td></td><td>Store at 4 &amp;deg;C after dissolving in nuclease-free water.</td></tr><tr><td>Glucose oxidase</td><td>Sigma</td><td>G2133</td><td>Store at -20 &amp;deg;C in aliquots after reconstitution in 50 mM NaOAc, pH 5.</td></tr><tr><td>Nuclease-free water</td><td>Ambion</td><td>AM9932</td><td>Store at room temperature.</td></tr><tr><td>Potassium phosphate (monobasic and dibasic)</td><td>various</td><td></td><td>Store at room temperature.</td></tr><tr><td>Probe library, diluted in TE, pH 8.0&amp;nbsp;</td><td>Biosearch Technologies</td><td></td><td>Store at -20 &amp;deg;C in aliquots (5 &amp;micro;L) after reconstitution in TE pH 8.0, following the instructions from the manufacturer. Protect from light.&amp;nbsp;</td></tr><tr><td>Sorbitol</td><td>various</td><td></td><td>Store at room temperature.</td></tr><tr><td>20X SSC</td><td>Ambion</td><td>AM9763</td><td>Store at room temperature.</td></tr><tr><td>TE, pH 8.0</td><td>Ambion</td><td>AM9849</td><td>Store at room temperature.</td></tr><tr><td>1 M Tris, pH 8.0</td><td>Ambion</td><td>AM9856</td><td>Store at room temperature.</td></tr><tr><td>Trolox</td><td>Sigma</td><td>238813</td><td>Store at -20 &amp;deg;C in aliquots.</td></tr><tr><td>200 mM Vanadyl ribonucleoside complex (VRC)</td><td>NEB</td><td>S1402S</td><td>Store at -20 &amp;deg;C in aliquots after reconstitution, following the instructions from the manufacturer.</td></tr><tr><td>Zymolyase 100T</td><td>MP Biomedicals</td><td>08320932</td><td>Store at -20 &amp;deg;C in aliquots after reconstitution in water.</td></tr><tr><td>Low-adhesion tubes</td><td>USA Scientific</td><td>1415-2600</td><td>Store at room temperature.</td></tr></tbody></table>

single molecule fluorescence in situ hybridization (smfish) analysis in budding yeast vegetative growth and meiosis

Single molecule fluorescence in situ hybridization (smFISH) is a powerful technique to study gene expression in single cells due to its ability to detect and count individual RNA molecules. Complementary to deep sequencing-based methods, smFISH provides information about the cell-to-cell variation in transcript abundance and the subcellular localization of a given RNA. Recently, we have used smFISH to study the expression of the gene&#160;NDC80&#160;during meiosis in budding yeast, in which two transcript isoforms exist and the short transcript isoform has its entire sequence shared with the long isoform. To confidently identify each transcript isoform, we optimized known smFISH protocols and obtained high consistency and quality of smFISH data for the samples acquired during budding yeast meiosis. Here, we describe this optimized protocol, the criteria that we use to determine whether high quality of smFISH data is obtained, and some tips for implementing this protocol in&#160;other yeast strains and growth conditions.

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Single Molecule Fluorescence In Situ Hybridization smFISH Analysis in Budding Yeast Vegetative Growth and Meiosis

This single molecule fluorescence in situ hybridization protocol is optimized to quantify the number of RNA molecules in budding yeast during vegetative growth and meiosis.

Single Molecule Fluorescence In Situ Hybridization (smFISH) Analysis in Budding Yeast Vegetative Growth and Meiosis

Single molecule fluorescence in situ hybridization (smFISH) is a powerful technique to study gene expression in single cells due to its ability to detect ...

single-molecule-fluorescence-situ-hybridization-smfish-analysis

Research

JoVE Journal

Genetics

19.2K Views.  University of California, Berkeley. Dit enkel molecuul fluorescentie in situ hybridisatie protocol is geoptimaliseerd voor het kwantificeren van het aantal molecules van RNA in ontluikende gist tijdens de vegetatieve groei en meiose.

Video: Single Molecule Fluorescence In Situ Hybridization smFISH Analysis in Budding Yeast Vegetative Growth and Meiosis

Use of Single Molecule Fluorescent In Situ Hybridization (SM-FISH) to Quantify and Localize mRNAs in Murine Oocytes

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), quantitative, real-time RT-PCR (RT-qPCR) and RNA sequencing. When these techniques are performed using a single oocyte or embryo, low-copy mRNAs are not reliably detected. To overcome this problem, oocytes or embryos can be pooled together for analysis; however, this often leads to high variability amongst samples. In this protocol, we describe the use of fluorescence in situ hybridization (FISH) using branched DNA chemistry. This technique identifies the spatial pattern of mRNAs in individual cells. When the technique is coupled with Spot Finding and Tracking&#160;computer software, the abundance of mRNAs in the cell can also be quantified. Using this technique, there is reduced variability within an experimental group and fewer oocytes and embryos are required to detect significant differences between experimental groups. Commercially available branched-DNA SM-FISH kits have been optimized to detect mRNAs in sectioned tissues or adherent cells on slides. However, oocytes do not effectively adhere to slides and some reagents in the kit were too harsh resulting in oocyte lysis. To prevent this lysis, several modifications were made to the FISH kit. Specifically, oocyte permeabilization and wash&#160;buffers designed for the immunofluorescence of oocytes and embryos replaced the proprietary buffers. The permeabilization, washes, and incubations with probes and amplifier were performed in 6-well plates and oocytes were placed on slides at the end of the protocol using mounting media. These modifications were able to overcome the limitations of the commercially available kit, in particular, the oocyte lysis. To accurately and reproducibly count the number of mRNAs in individual oocytes, computer software was used. Together, this protocol represents an alternative to PCR and sequencing to compare the expression of specific transcripts in single cells.

Use of Single Molecule Fluorescent In Situ Hybridization SM-FISH to Quantify and Localize mRNAs in Murine Oocytes

To reproducibly count the numbers of mRNAs in individual oocytes, single molecule RNA fluorescence in situ hybridization (RNA-FISH) was optimized for non-adherent cells. Oocytes were collected, hybridized with the transcript specific probes, and quantified using an image quantification software.

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), ...

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk of conceiving an aneuploid fetus increases dramatically and this phenomenon is known as the maternal age effect. One requirement for accurate chromosome segregation during the meiotic divisions is maintenance of sister chromatid cohesion during the extended prophase period that oocytes experience. Cytological evidence in both humans and model organisms suggests that meiotic cohesion deteriorates during the aging process. In addition, segregation errors in human oocytes are most prevalent during meiosis I, consistent with premature loss of arm cohesion. The use of model organisms is critical for unraveling the mechanisms that underlie age-dependent loss of cohesion. Drosophila melanogaster offers several advantages for studying the regulation of meiotic cohesion in oocytes. However, until recently, only genetic tests were available to assay for loss of arm cohesion in oocytes of different genotypes or under different experimental conditions. Here, a detailed protocol is provided for using fluorescence in situ hybridization (FISH) to directly visualize defects in arm cohesion in prometaphase I and metaphase I arrested Drosophila oocytes. By generating a FISH probe that hybridizes to the distal arm of the X chromosome and collecting confocal Z stacks, a researcher can visualize the number of individual FISH signals in three dimensions and determine whether sister chromatid arms are separated. The procedure outlined makes it possible to quantify arm cohesion defects in hundreds of Drosophila oocytes. As such, this method provides an important tool for investigating the mechanisms that contribute to cohesion maintenance as well as the factors that lead to its demise during the aging process.

Using Fluorescence In Situ Hybridization FISH to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

This manuscript presents a detailed method for generating X-chromosome arm probes and performing fluorescence in situ hybridization (FISH) to examine the state of sister chromatid cohesion in prometaphase and metaphase I arrested Drosophila oocytes. This protocol is suitable for determining whether meiotic arm cohesion is intact or disrupted in different genotypes.

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk ...

Developmental Biology

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 ...

Biochemistry

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Biology

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

Combined DNA-RNA Fluorescent In situ Hybridization (FISH) to Study X Chromosome Inactivation in Differentiated Female Mouse Embryonic Stem Cells

Fluorescent in situ hybridization (FISH) is a molecular technique which enables the detection of nucleic acids in cells. DNA FISH is often used in cytogenetics and cancer diagnostics, and can detect aberrations of the genome, which often has important clinical implications. RNA FISH can be used to detect RNA molecules in cells and has provided important insights in regulation of gene expression. Combining DNA and RNA FISH within the same cell is technically challenging, as conditions suitable for DNA FISH might be too harsh for fragile, single stranded RNA molecules. We here present an easily applicable protocol which enables the combined, simultaneous detection of Xist RNA and DNA encoded by the X chromosomes. This combined DNA-RNA FISH protocol can likely be applied to other systems where both RNA and DNA need to be detected.

Combined DNA-RNA Fluorescent In situ Hybridization FISH to Study X Chromosome Inactivation in Differentiated Female Mouse Embryonic Stem Cells

Fluorescent in situ hybridization (FISH) allows the detection of nucleic acids in their native environment within cells. We here describe a protocol for the combined, simultaneous detection of RNA and DNA by means of FISH, which can be used to study X chromosome inactivation in mouse embryonic stem cells.

Fluorescent in situ hybridization (FISH) is a molecular technique which enables the detection of nucleic acids in cells. DNA FISH is often used in ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...

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 ...

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often not seen by imaging fixed cells. Budding yeast has proved to be an important model organism to study meiotic chromosome segregation because many meiotic genes are highly conserved. Time-lapse microscopy of meiosis in budding yeast allows the monitoring of different meiotic mutants to show how the mutation disrupts meiotic processes. However, many proteins function at multiple points in meiosis. The use of loss-of-function or meiotic null mutants can therefore disrupt an early process, blocking or disturbing the later process and making it difficult to determine the phenotypes associated with each individual role. To circumvent this challenge, this protocol describes how the proteins can be conditionally depleted from the nucleus at specific stages of meiosis while monitoring meiotic events using time-lapse microscopy. Specifically, this protocol describes how the cells are synchronized in prophase I, how the anchor away technique is used to deplete proteins from the nucleus at specific meiotic stages, and how time-lapse imaging is used to monitor meiotic chromosome segregation. As an example of the usefulness of the technique, the kinetochore protein Ctf19 was depleted from the nucleus at different time points during meiosis, and the number of chromatin masses was analyzed at the end of meiosis II. Overall, this protocol can be adapted to deplete different nuclear proteins from the nucleus while monitoring the meiotic divisions.

Use of Time-Lapse Microscopy and Stage-Specific Nuclear Depletion of Proteins to Study Meiosis in S. cerevisiae

Time-lapse microscopy is a valuable tool for studying meiosis in budding yeast. This protocol describes a method that combines cell-cycle synchronization, time-lapse microscopy, and conditional depletion of a target protein to demonstrate how to study the function of a specific protein during meiotic chromosome segregation.

Time-lapse fluorescence microscopy has revolutionized the understanding of meiotic cell-cycle events by providing temporal and spatial data that is often ...

Single Molecule Fluorescence In Situ Hybridization (smFISH) Analysis in Budding Yeast Vegetative Growth and Meiosis

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