Name: single molecule fluorescence in situ hybridization (smfish) analysis in budding yeast vegetative growth and meiosis
Uploaded: 2018-05-25T15:00:00.000+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.

<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.

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 ...

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

Nanyang Technological University

Research

JoVE Journal

Genetics

19.2K Views.  University of California, Berkeley. 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.

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

Detection of Inter-chromosomal Stable Aberrations by Multiple Fluorescence In Situ Hybridization (mFISH) and Spectral Karyotyping (SKY) in Irradiated Mice

Ionizing radiation (IR) induces numerous stable and unstable chromosomal aberrations. Unstable aberrations, where chromosome morphology is substantially compromised, can easily be identified by conventional chromosome staining techniques. However, detection of stable aberrations, which involve exchange or translocation of genetic materials without considerable modification in the chromosome morphology, requires sophisticated chromosome painting techniques that rely on in situ hybridization of fluorescently labeled DNA probes, a chromosome painting technique popularly known as fluorescence in situ hybridization (FISH). FISH probes can be specific for whole chromosome/s or precise sub-region on chromosome/s. The method not only allows visualization of stable aberrations, but it can also allow detection of the chromosome/s or specific DNA sequence/s involved in a particular aberration formation. A variety of chromosome painting techniques are available in cytogenetics; here two highly sensitive methods, multiple fluorescence in situ hybridization (mFISH) and spectral karyotyping (SKY), are discussed to identify inter-chromosomal stable aberrations that form in the bone marrow cells of mice after exposure to total body irradiation. Although both techniques rely on fluorescent labeled DNA probes, the method of detection and the process of image acquisition of the fluorescent signals are different. These two techniques have been used in various research areas, such as radiation biology, cancer cytogenetics, retrospective radiation biodosimetry, clinical cytogenetics, evolutionary cytogenetics, and comparative cytogenetics.

Detection of Inter-chromosomal Stable Aberrations by Multiple Fluorescence In Situ Hybridization mFISH and Spectral Karyotyping SKY in Irradiated Mice

The present protocol describes the usefulness of multiple fluorescence in situ hybridization (mFISH) and spectral karyotyping (SKY) in identifying inter-chromosomal stable aberrations in the bone marrow cells of mice after exposure to total body irradiation.

Ionizing radiation (IR) induces numerous stable and unstable chromosomal aberrations. Unstable aberrations, where chromosome morphology is substantially ...

Analysis of the Development of a Morphological Phenotype as a Function of Protein Concentration in Budding Yeast

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of phenotype development as a function of protein concentration at population and single-cell levels in Saccharomyces cerevisiae. Although this protocol is based on the overexpression of a protein, it can easily be adapted for morphological phenotypes dependent on suppression of protein expression. Our lab is interested in studying the signaling properties of the endocytic adaptor protein epsin. To that purpose we used a dominant negative approach in which we over-expressed the conserved Epsin N-Terminal Homology (ENTH) domain in order to interfere with the functions of endogenous epsin-2 (Ent2 or YLR206W). We observed that overexpression of the ENTH domain of Ent2 (ENTH2) in wild type cells led to a cell division defect that is dependent on the mislocalization of a family of scaffolding proteins, septins.

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of phenotype development as a function of protein concentration at population and single-cell levels in Saccharomyces cerevisiae.

Gene deletion and protein overexpression are common methods for studying functions of proteins. In this article, we describe a protocol for analysis of ...

Biology

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 ...

Analysis of Lipid Droplet Content in Fission and Budding Yeasts using Automated Image Processing

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are used as models in lipid metabolic research or for industrial lipid production. Lipid droplets are highly dynamic storage bodies and their cellular content represents a convenient readout of the lipid metabolic state. Fluorescence microscopy is a method of choice for quantitative analysis of cellular lipid droplets, as it relies on widely available equipment and allows analysis of individual lipid droplets. Furthermore, microscopic image analysis can be automated, greatly increasing overall analysis throughput. Here, we describe an experimental and analytical workflow for automated detection and quantitative description of individual lipid droplets in three different model yeast species: the fission yeasts Schizosaccharomyces pombe and Schizosaccharomyces japonicus, and the budding yeast Saccharomyces cerevisiae. Lipid droplets are visualized with BODIPY 493/503, and cell-impermeable fluorescent dextran is added to the culture media to help identify cell boundaries. Cells are subjected to 3D epifluorescence microscopy in green and blue channels and the resulting z-stack images are processed automatically by a MATLAB pipeline. The procedure outputs rich quantitative data on cellular lipid droplet content and individual lipid droplet characteristics in a tabular format suitable for downstream analyses in major spreadsheet or statistical packages. We provide example analyses of lipid droplet content under various conditions that affect cellular lipid metabolism.

Here, we present a MATLAB implementation of automated detection and quantitative description of lipid droplets in fluorescence microscopy images of fission and budding yeast cells.

Lipid metabolism and its regulation are of interest to both basic and applied life sciences and biotechnology. In this regard, various yeast species are ...

Immunology and Infection

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 ...

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 ...

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and development. Cytokinesis involves a series of events that are well coordinated in time and space. Cytokinesis involves the formation of an actomyosin ring at the division site, followed by ring constriction, membrane furrow formation and extra cellular matrix remodeling. The fission yeast, Schizosaccharomyces pombe (S.&#160;pombe)&#160;is a well-studied model system that has revealed with substantial clarity the initial events in cytokinesis. However, we do not understand clearly how different cytokinetic events are coordinated spatiotemporally. To determine this, one needs to analyze the different cytokinetic events in great details in both time and in space. Here we describe a microscopy approach to examine different cytokinetic events in live cells. With this approach it is possible to time different cytokinetic events and determine the time of recruitment of different proteins during cytokinesis. In addition, we describe protocols to compare protein localization, and distribution at the site of cell division. This is a basic protocol to study cytokinesis in fission yeast and can also be used for other yeasts and fungal systems.

The fission yeast, Schizosaccharomyces pombe&#160;is an excellent model system to study cytokinesis, the final stage in cell division. Here we describe a microscopy approach to analyze different cytokinetic events in live fission yeast cells.

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

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 ...

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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Chapters in this video

Detection of Inter-chromosomal Stable Aberrations by Multiple Fluorescence In Situ Hybridization (mFISH) and Spectral Karyotyping (SKY) in Irradiated Mice

Analysis of the Development of a Morphological Phenotype as a Function of Protein Concentration in Budding Yeast

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Analysis of Lipid Droplet Content in Fission and Budding Yeasts using Automated Image Processing

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

Microscopy of Fission Yeast Sexual Lifecycle

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

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

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