Name: single molecule fluorescence in situ hybridization (smfish) analysis in budding yeast vegetative growth and meiosis
Uploaded: 2018-05-25T15:00:00
Duration: 9 min 28 s
Description: Watch this Scientific Journal Video about Single Molecule Fluorescence In Situ Hybridization smFISH Analysis in Budding Yeast Vegetative Growth and Meiosis at JoVE.com

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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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.

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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.

Watch this Scientific Journal Video about Single Molecule Fluorescence In Situ Hybridization smFISH Analysis in Budding Yeast Vegetative Growth and Meiosis at JoVE.com

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 ...

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