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 NDC80 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 other yeast strains and growth conditions.


Introduction
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 cells 1,2 . This method allows 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 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 molecule 3 .
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 expression 4 , such as transcription elongation 1,2 , splicing 5,6 , transcriptional bursting 7,8,9 , intracellular allelic expression 10,11,12 , and RNA localization 13,14,15 . Recently, we have used this method to study the expression of two transcript isoforms of the same gene, in which the short transcript isoform (NDC80 ORF ) has its entire sequence shared with the long isoform (NDC80 luti ) 16  . 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. 5. 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. 6. 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. 7. 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 µL. 8. Resuspend the cells in 1.5 mL of cold Buffer B. 9. 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 µL. 10. 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. 11. Resuspend cells in 425 µL of digestion master mix, and briefly vortex to resuspend. Add 5 µL 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. 12. Vortex 2-3 s to mix. Place the tubes on a roller drum, and digest at 30 ˚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 nonrefractive. From this point on, cells are very fragile. Handle cells gently and avoid using vacuum aspiration or vortexing. 13. Centrifuge the tubes at ~376 x g for 3 min. Remove the liquid completely by pipetting. 14. Gently resuspend cells with 1 mL of Buffer B by pipetting up and down 1-2 times to mix. 15. 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). 16. Incubate at room temperature for 3.5-4 hours.

Washing and imaging
1. Bring formamide to room temperature. 2. Prepare 10% formamide wash buffer (FWB) in a 15-mL conical tube. 3. Remove the tubes from the roller drum and place them into a foil-covered box to protect from light. 4. Centrifuge the samples at ~376 x g for 3 min. Remove the supernatant into hazardous waste by pipetting. 5. Resuspend in 1 mL of 10% FWB by gently pipetting up and down 2-3 times. 6. Incubate at 30 ˚C for 30 min (not rotating) in the foil-covered box. 7. Centrifuge the samples at ~376 x g for 3 min. Remove the supernatant to hazardous waste by pipetting, and leave ~50 µL.  15. 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). 16. Transfer the slide to the microscope room in a box covered by aluminum foil. 17. 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 µm, 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.

Image Analysis
1. Analyze the smFISH data with published smFISH analysis tools such as FISH-quant 17 and StarSearch 2 , 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.

Representative Results
To evaluate how well the smFISH protocol worked (outlined in Figure 1), we designed a set of 54 probes that tile the open reading frame of the NDC80 gene (Figure 2A, top). The probe located at the most 5' end is referred to as Probe 1; the next one, Probe 2; and the third one, Probe 3, etc. The 27 probes assigned an odd number (Probe 1, 3, 5…) are all conjugated to the CAL Fluor 590 (CF590) fluorescent dye; and the 27 probes assigned an even number (Probe 2, 4, 6…), conjugated to the Quasar 670 (Q670) fluorescent dye. Hence, these alternating probe sets are often referred to as "odd/even" probes. Upon hybridization, both probe sets should label the same transcript.
After spot detection, we used a few measurements to judge the quality of the smFISH data set. The first one was the degree and quality of colocalization for the odd/even probes. In our case, 88% of all the smFISH spots colocalized (Figure 2A and 2B), with greater than 95% of the spots paired within 2 pixels of each other (Figure 2C, paired), which is within the expected value given any chromatic and detection aberration between the two fluorescent channels. By comparison, fewer than 10% of unpaired spots had a nearest-neighbor distance of less than 2 pixels, showing that the probability of misidentifying a spot pair is low ( Figure 2C). The 12% of unpaired spots were evenly divided between the two channels ( Figure 2B, compare CF590-only with Q670-only); and thus, we concluded that for this 2.4 kb gene with a range of expression between zero to 45 transcripts per cell, ~94% of RNA molecules were accurately detected in each fluorescent channel. If the smFISH protocol were suboptimal, one would observe (1) a greater fraction of the smFISH spots with only one of the two fluorescent signals (non-colocalized), and/or (2) cells with a very low signal-to-noise ratio or no signal at all. We next asked if the detection of the smFISH signal was biased with respect to the total number of RNA molecules per cell. In a population, the total number of a particular RNA in each cell lies in a distribution, with some cells harboring more RNA molecules than others. A good smFISH protocol should robustly detect the RNA regardless whether a high number or low number of RNA molecules is present in each cell. To test this, for each cell, we calculated the fraction of the smFISH spots with colocalized signals and the fraction with only one of the two fluorescent signals. After grouping the cells that had the same number of total spots per cell, we calculated the average fraction of colocalized (paired) or non-colocalized (CF590-only or Q670-only) spots in a given group, and graphed this average as a function of the total number of spots per cell (Figure 3). For each category of spots, the fractions were similar across the entire range of the total number of spots per cell. For instance, comparing the cells with a total of 20 fluorescent spots per cell versus those with a total of 30 spots, the average fractions of the colocalized spots were similar. This result suggested that our protocol could detect a range of RNA molecules in a cell (up to at least ~40 molecules per cell). If the protocol worked suboptimally, one might observe a bias. For example, the fraction of the spots with only one of the two fluorescent signals might increase as the total number of spots per cell increased.
Using this optimized protocol, we examined the expression of the NDC80 gene, which encodes a kinetochore protein, in different growth conditions. The NDC80 gene expresses two mRNA isoforms: The long undecoded transcript isoform, NDC80 luti , has a ~400 base pairs extension at the 5' end in comparison to the short NDC80 ORF isoform, but both transcripts share the coding region of the NDC80 gene (See schematic in Figure 4A). We designed two sets of probes: The CF 590 set binds to the unique 5' region of NDC80 luti ; and the Q 670 set binds to the common region of NDC80 luti and NDC80 ORF . The NDC80 luti transcripts were detected as the smFISH spots where the signals from both probe sets colocalized, whereas the NDC80 ORF transcripts were those with the signal only from Q 670. Due to the size of the unique segment of NDC80 luti , we could only tile 20 oligonucleotide probes along this region, which is fewer than the recommended 30 to 48 probes. Thus, we first determined if this probe set could robustly detect RNA in our optimized protocol. For either fluorescent channel, we graphed the signal-to-noise ratio (SNR, defined as the variance of the pixels surrounding a spot relative to the spot intensity) against the signal detected for each smFISH spot, and generated a scatterplot for all the spots identified in the entire field of view (example images in Figure 4A and plots in Figure 4B). Two distinct populations were clearly identified for both the Q 670 and the CF 590 probe sets (Optimized, Figure 4B), suggesting that the true smFISH spots (inside the grey area) could be separated from the background signal. Note that in the suboptimal condition, such separation was less obvious (Suboptimal, Figure 4B).
We used these two probe sets to study the expression of NDC80 luti and NDC80 ORF during vegetative growth and meiosis. In vegetative cells, robust signal from the Q 670 probe set was detected, but not from the CF 590 probe set ( Figure 5A, vegetative), agreeing with the observation by northern blotting that only the short isoform was expressed during vegetative growth 16 . This result also suggested that the CF 590 probe set is specific, yielding low background in our optimized smFISH protocol. In contrast, robust signal from both probe sets was detected in meiotic prophase, and the majority of the spots had colocalized signal (Figure 5A, meiotic prophase). Together with northern blot analysis (Figure 5B), this observation confirmed that the long isoform NDC80 luti was expressed specifically in meiosis. These two types of mRNAs were detected in the cytoplasm (outside of the DAPI region), suggesting that both were exported from the nucleus, consistent with the ribosome profiling data 18 .
To determine if sufficient data were collected to accurately account for the biological variation intrinsic to our data set, we performed bootstrap analysis using the statistics of each cell, which included (1) the fraction of the colocalized spots per cell and (2) the fraction of the spots with one of the two fluorescent signals (CF 590 only and Q 670 only). In this analysis, a program randomly selected one cell from the 437 quantified cells for 500 iterations. Next, the mean and the variance of the respective statistics associated with these 500 selected cells were calculated. This process was then repeated for randomly selecting two cells from all the cells, without replacement; and then for randomly selecting three cells, etc. until one half of the total data set size was reached. The variance in the data set plateaued after ~40 cells, suggesting that after this point most of the variation in the mean is intrinsic to the data rather than an artifact of undersampling (Figure 6, inset). This effect became more obvious when we graphed the sample variance divided by the sample mean, as a function of the number of cells sampled in each iteration cycle (Figure 6). With the data shown, the change in variance became diminishingly small after ~60 cells. The number of cells quantified in an smFISH experiment should exceed the minimum number of cells required to achieve a stable mean and a plateau of the variance. In our case, we quantified over 95 cells per sample, per replicate. The total number of cells (> 400 cells) well exceeded the minimum number of cells (~60 cells) required to reflect the population mean.
With a custom-made Matlab program 16 , vegetative cells were found to have a median of 5 NDC80 ORF transcripts per cell, whereas in meiotic prophase cells, the median significantly dropped to 4 transcripts per cell (two-tailed Wilcoxon Rank Sum test, p = 0.026) (Figure 7). The median number of NDC80 luti transcripts per cell was 21 transcripts, and 100% of the cells expressed the NDC80 luti transcripts. We graphed the fraction acquired under optimized versus suboptimal conditions. Strains UB8144 (optimized condition) and UB1337 (suboptimal condition) were induced to undergo meiosis and fixed during meiotic prophase. These two strains harbor different synchronization systems to induce meiosis, but the use of either system does not affect smFISH quality (data not shown). In the suboptimal condition, cells were fixed at room temperature for 20 min (no overnight fixation), digested with zymolyase without VRC supplemented, and hybridized in a lower concentration of VRC. Images are shown as the maximum-intensity projections of z-stacks. DNA was stained with DAPI (shown in blue). Scale bar: 5 µm. (B) Scatterplots displaying the signal-to-noise ratio (SNR) and the signal of each smFISH spot detected in the field of view presented in Figure 4A, for either fluorescent channel, as well as for the optimized or suboptimal conditions. In the optimized condition, two populations of the smFISH spots were present. The true smFISH spots were located inside the grey area, separating from the background signal. In the suboptimal condition, individual mRNAs were hard to distinguish by eye, and the separation between the true spots and background after running the spot detection software was less obvious. Please click here to view a larger version of this figure. level was determined by anti-V5 immunoblot at the indicated time points. Hxk1p, loading control for immunoblot. As strain UB8144, this strain also harbors the pGAL-NDT80 GAL4-ER synchronization system 19,20 . Cells were transferred to sporulation medium at 0 hour and released from pachytene arrest by β-estradiol addition 6 hours later. The NDC80 luti transcript was robustly detected in S/meiotic prophase, whereas

Discussion
The protocol presented here is derived from other published smFISH protocols 2,3,21,22,23 , 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 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 wheñ 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 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 NDC80 luti (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's instructions, we made a 1:10 dilution of the stock solution and stored the diluted solution in 5-µL aliquots at -20 ˚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 ˚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 protocols 3,22,23 , our protocol uses the RNase inhibitor VRC during digestion and 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 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 >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 images 2,17 . 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 quantification 22 . 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 yeast 3,7 ). 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.

Disclosures
The authors have no conflict of interest to disclose.