Name: use of single molecule fluorescent in situ hybridization (sm-fish) to quantify and localize mrnas in murine oocytes
Uploaded: 2019-04-24T14:00:00.000+00:00
Duration: 8 min 18 s
Description: Watch this Scientific Journal Video about Use of Single Molecule Fluorescent In Situ Hybridization SM-FISH to Quantify and Localize mRNAs in Murine Oocytes at JoVE.com

We thank Dr. Daniel R. Larson for his generous help with the installation and use of the Spot Finding and Tracking Program&#160;13&#160;and the technical support of the University of Nebraska Lincoln Microscopy Core for the confocal microscopy imaging. This study represents a contribution of the University of Nebraska Agricultural Research Division, Lincoln, Nebraska and was supported by UNL Hatch Funds (NEB-26-206/Accession number -232435 and NEB-26-231/Accession number -1013511).

Animal procedures were reviewed and approved by the Institutional Animal Care and Use Committee at the University of Nebraska-Lincoln and all methods were performed in accordance with relevant guidelines and regulations. For this study, CD-1 outbred mice had ad libitum access to normal rodent chow and water; they were maintained in a 12:12 dark: light cycle.
1. Preparation of required media
<ol>
	<li>For base media (OMM), add 100 mM NaCl, 5 mM KCl, 0.5 mM KH2PO4, and 1.7 mM CaCl2-2H2O to 100 mL sterile water. 
	NOTE: OMM medium can be stored for up to 1 month.</li>
	<li>For complete media (OMOPS), add 20 mM 3-morpholinopropane-1-sulfonic acid (MOPS), 1.2 mM MgSO4-7H2O, 0.5 mM glucose, 6 mM L-lactate, 1 mM ala-gln, 0.1 mM taurine, 1x non-essential amino acids (NEAA), 0.01 mM ethylenediaminetetraacetic acid (EDTA), 10 &#181;M alpha lipoic acid, 10 &#181;g/mL undiluted gentamicin, 21 mM 1 M NaOH, 5 mM NaHCO3, 0.2 mM Pyruvate, 0.5 mM Citrate, 4 mg/mL FAF BSA to a 1:10 dilution of OMM in sterile water for a total volume of 100 mL. Sterilize the medium with a 0.22 &#181;M filter. 
	NOTE: OMOPS can be stored for up to 1 week.</li>
	<li>For the holding medium (HM) add 5% fetal bovine serum to OMOPS. Make 2 mL HM per mouse.</li>
	<li>For hyaluronidase solution add 0.1 mg/mL of hyaluronidase derived from bovine testes, to 1 mL HM.</li>
	<li>For the fixation buffer combine 4% paraformaldehyde in 10 mL of 1x PBS along with 0.1% embryo grade polyvinylpyrrolidone (PVP)14.</li>
	<li>To prepare 50 mL wash buffer (WB), add 0.1% non-ionic surfactant and 0.1% PVP to 1x PBS14.</li>
	<li>To Prepare 10 mL permeabilization buffer, add 1% non- ionic surfactant to 1x PBS14. 
	NOTE: The wash and permeabilization buffers described above replace the proprietary buffers in the commercially available kits.</li>
</ol>
2. Collection of ovulated oocytes from female mice
<ol>
	<li>Preparation:
	<ol>
		<li>Stimulate female mice at 5-8 weeks of age by intraperitoneal (IP) injection of 5 IU equine chorionic gonadotropin (eCG) followed by 5 IU human chorionic gonadotropin (hCG) 44-48 h later15,16.</li>
		<li>Keep 35 mm Petri dishes containing 2 mL of HM on a 37 &#176;C warming plate. Pipette one, 100-&#181;L drop of HM containing diluted hyaluronidase followed by three, 50 &#181;L drops of HM without hyaluronidase in a 60 mm petri dish. Place plates containing drops on the 37 &#176;C warming plate prior to use. 
		NOTE: The hyaluronidase drops should be made just prior to the dissection of each oviduct pair to prevent the evaporation and concentration of the components of HM with or without hyaluronidase.</li>
	</ol>
	</li>
	<li>Euthanize mice, 16 h after the IP injection of hCG, using isoflurane overdose followed by cervical dislocation.</li>
	<li>Clean the mouse using 70% ethanol. Expose the abdominal cavity and visualize the female reproductive tract. Hold the ovary with forceps and remove the uterine ligaments and excess adipose tissue from around the ovary. Cut the oviduct from the uterus and place the ovary-oviduct pair in the warm HM in the 35 mm dish.</li>
	<li>Remove the ovary and any surrounding adipose tissue. Tear the swollen ampulla of the oviduct using a &#189; inch 27-gauge needle. Push the oviduct at the site of the tear and the cumulus cell-oocyte complexes (COCs) will be expelled. Transfer the ovulated oocytes, which are presumed to be in metaphase II (MII) of meiosis, to the 100 &#956;L drop containing HM media with hyaluronidase using a mouth pipette (Figure 2).</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59414/59414fig2.jpg" /> 
Figure 2: Parts of the mouth pipettor used to transfer oocytes. (A) mouth piece (B) 0.22 um, 4mm filter (C) aspirator tubing (D) 1000 &#956;L pipet tip (E) 9&#34; Pasteur pipet. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59414/59414fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="5">
	<li>Pipette the MII oocyte-cumulus cell complexes up and down in the hyaluronidase containing HM with the mouth pipette to dislodge cumulus cells. Transfer each oocyte, once they are devoid of cumulus cells to a wash drop containing HM only using the mouth pipette. Repeat this for each wash droplet. Do not transfer fragmented or transparent oocytes15. 
	NOTE: It is important to transfer the oocytes from each drop in the 35 mm dish with as little HM as possible. This is true for every transfer in the protocol. The MII oocytes must not remain in the hyaluronidase containing HM medium for more than one minute.</li>
</ol>
3. SM-FISH Staining of Oocytes
<ol>
	<li>Fix oocytes in an individual well of a 6-well plate containing 500 &#181;L of Fixation Buffer. Submerge 20 oocytes or less in the well. Incubate for 20 min at room temperature. 
	NOTE: Each SM-FISH staining step occurs within an individual well in a 6-well conical plate. Ensure that oocytes are completely submerged in buffers and not floating on top of the buffer. Each step should be performed with 20 oocytes or less in each well.</li>
	<li>Transfer fixed oocytes to 500 &#956;L of wash buffer (WB described in step 1.6) for 10 min each. Repeat 2 more times.</li>
	<li>Incubate oocytes in permeabilization buffer for 30 min at room temperature. 
	NOTE: The permeabilization buffer described in step 1.7 replaces the propriety permeabilization buffer.
	<ol>
		<li>Gather probe sets and quickly spin them down in a microcentrifuge. Warm each probe set for 10 min in a 40 &#176;C water bath or incubator. Cool to the room temperature. 
		NOTE: This step should be performed during the permeabilization incubation</li>
	</ol>
	</li>
	<li>Wash oocytes in 500 &#181;L of WB for 10 min at room temperature.</li>
	<li>Transfer oocytes to 80 &#956;L of Protease III Buffer (available from the kit), that is diluted 1:8 in 1X PBS, for 30 min at room temperature. 
	NOTE: The 80 &#181;L volume adequately covers the bottom of an individual well in a 6-well plate.</li>
	<li>Wash oocytes in 500 &#181;L of WB for 10 min at room temperature.</li>
	<li>Dilute the warmed probe sets for Nanog, Pou5f1, and DapB (a negative control gene), 1:50 in probe diluent. Incubate oocytes in 80 &#956;L of the transcript-specific probe for 2 hours at 40&#176; C. 
	NOTE: Each proprietary probe set is available in one of three fluorescence channels (C1, C2, and C3). The Nanog and Pou5f1 probes were tagged with C2 and C3, respectively.</li>
	<li>Warm the proprietary, Amplifier 1 (AMP 1), Amplifier 2 (AMP2), Amplifier 3 (AMP3) and Amplifier 4-fluorescence (AMP 4-FL) at room temperature. 
	NOTE: This step should be performed during the 2-hour transcript-specific probe incubation.</li>
	<li>Transfer the oocytes to 500 &#956;L of WB and incubate for 10 min at room temperature.</li>
	<li>Incubate oocytes sequentially in amplification buffers.
	<ol>
		<li>Incubate oocytes in 80 &#181;L of AMP1 for 30 min at 40&#176; C. Transfer oocytes to 500 &#181;L of WB for 10 min at room temperature.</li>
		<li>Incubate oocytes in 80 &#181;L of AMP2 for 15 min at 40 &#176;C. Transfer oocytes to 500 &#181;L of WB for 10 min at room temperature.</li>
		<li>Incubate the oocytes in 80 &#181;L of AMP3 for 30 min at 40 &#176;C. Transfer oocytes to 500 &#181;L of WB for 10 min at room temperature. 
		NOTE: The remainder of the protocol is performed in the dark because AMP-FL contains the fluorophore. When working under the dissecting microscope, reduce the light as much as possible.</li>
		<li>Add oocytes to 80 &#956;L of AMP4-FL for 15 min at 40&#176; C. 
		NOTE: AMP4-FL is provided as alternative buffer-A (Alt-A), Alt-B or Alt-C. Select the AMP4-FL buffer dependent on which emission wavelength is desired.</li>
	</ol>
	</li>
	<li>Wash oocytes in 500 &#181;L of WB for 10 min at room temperature. Incubate oocytes in 80 &#181;L of DAPI for 20 min at room temperature. Wash oocytes in 500 &#181;L of WB for 5 min at room temperature.</li>
	<li>Pipette 12 &#181;L of anti-fading mounting media onto the center of a slide without adding bubbles to the reagent. Transfer oocytes with as little WB as possible into the mounting media and apply a coverslip.
	<ol>
		<li>Tilt the coverslip at an angle and slowly and gently place over the liquid on the slide. Avoid pressing the coverslip too hard to prevent distortion of the oocytes and introduction of bubbles.</li>
		<li>Store the slides in a dark box to dry overnight at room temperature. Coat the edges of the slides in clear nail polish to seal coverslip.</li>
	</ol>
	</li>
	<li>Use a standard microscope to find oocytes on the slide and circle with a permanent marker. 
	NOTE: This step is not required but improves locating oocytes on the slide. For best results, image slides within 1 to 5 days as the fluorescent signal will begin to fade.</li>
</ol>
4. Image Processing
<ol>
	<li>Image the 3-dimensional oocytes, using z step confocal microscopy. 
	NOTE: To accurately analyze the images, each z step should be 1.0 &#181;m/slice.</li>
	<li>Save confocal images as a compressed nd2 or individual .TIFF files for each oocyte. Both image types are compatible with the open source image processing program, Fiji.</li>
	<li>Download and install the open access Fiji software (https://imagej.net/Fiji/Downloads).&#160;
	<ol>
		<li>Drag nd2 files into Fiji and choose hyperstack. If confocal images were saved as .TIFF files skip to step 4.4. 
		NOTE: When the nd2 file is dropped into Fiji the hyperstack dropdown should appear automatically.</li>
		<li>Click the Image tab, Select Color, and click Split Channels to separate the fluorescent channels of the nd2 file.</li>
		<li>Generate individual .TIFF files for each z-slice of the oocyte in each fluorescent channel. Click the Image tab, select Stacks, and click Stack to Images. Click the Image tab, select Type, and click RGB Color to convert each z slice to an individual RGB color image. 
		NOTE: The RGB color is artificial and can be chosen as desired for each emission wavelength.</li>
		<li>Save each converted image as .TIFF file. Place images from a single oocyte for each fluorescent channel in a new folder to avoid confusion during stitching (step 4.3).</li>
	</ol>
	</li>
	<li>Normalize each .TIFF image for Pou5f1 and Nanog using negative control images (DapB). 
	NOTE: Normalization is performed using a photo editing program. Make sure to remove the same levels of background fluorescence from each control image.</li>
	<li>Open each normalized .TIFF file in Fiji to stitch together all z-slices for each oocyte in each wavelength.
	<ol>
		<li>Click the Plugins tab, select Stitching, and click on Grid/Collection (Figure 3A). Select Sequential Images from the drop-down menu and click OK (Figure 3B).</li>
		<li>Browse the directory and select the folder containing all of the z-slice images for an individual oocyte at one wavelength (see step 4.3.4). Click OK.</li>
		<li>Move the slider at the bottom of the stitched image to the appropriate color channel for the wavelength used and create the final RGB stitched image by clicking Image, selecting Type, and click RGB Color. 
		NOTE: This image will be used for fluorescence quantitation described in step 4.6 below.</li>
	</ol>
	</li>
	<li>Convert the stitched image to 32-bit maximum projected picture. Click Image, select Type, and click 32-bit (Figure 3C). Save this image as a new .TIFF file.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59414/59414fig3.jpg" /> 
Figure 3: Stitching together of confocal z-series images of oocytes. (A) Screenshot showing the Plug-in Grid/Collection tool in Fiji that was used to produce composite images of the oocyte. (B) Sequential Images uses fluorescence overlap between sequential .TIFF files to generate a composite image. (C) The composite image was saved as a 32-bit .TIFF file. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59414/59414fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="7">
	<li>Download and install the Spot Finding and Tracking Program13, which is available from the website for D.R. Larson, an investigator at the National Institutes of Health National Cancer Institute (https://ccr.cancer.gov/Laboratory-of-Receptor-Biology-and-Gene-Expression/daniel-r-larson). Download and install the open access virtual machine for the interactive data language (IDL) operating system which is required to run the Spot Finding and Tracking Program (http://www.spacewx.com/pdf/idlvm.pdf).&#160;&#160;</li>
	<li>Open the 32-bit, stitched image, which was generated in step 4.6 (Figure 4A), in the Spot Finding and Tracking Program. Select the Localize dropdown and click Localize (Figure 4B), which will calculate the number spots found in the image. 
	NOTE: Each spot counted represents an individual mRNA. Band pass and photon threshold settings are shown in the screenshot (Figure 4B). For this protocol, default for each threshold setting was used. Representative positive and background spots are shown (Figure 4C).</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59414/59414fig4.jpg" /> 
Figure 4: Quantification of mRNAs using Spot Finder and Tracking. (A) Individual z-series images were stitched together as described in Figure 3 and saved as a 32-bit maximum projected .TIFF file. (B) Composite image was opened in Spot Finder and Tracking. Localize was used to count the fluorescent spots (red box). Band pass and photon threshold are indicated by the blue box. (C) The blue arrow points to a positive signal (above threshold). The white arrow shows a fluorescent spot below the threshold and, therefore, not counted. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59414/59414fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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R., Omholt, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenomics:+the+next+challenge.">Phenomics: the next challenge.</a> Nature Reviews Genetics. 11, 855 (2010).</li><li>Freimer, N., Sabatti, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Human+Phenome+Project.">The Human Phenome Project.</a> Nature Genetics. 34, 15 (2003).</li></ol>

The authors have nothing to declare

A series of minor steps during the protocol will ensure successful fluorescence and accurate counts of mRNAs. First, the protocol must be performed immediately after collection and fixation of the oocytes. Note that PVP is added to the 4% paraformaldehyde fixation buffer to prevent oocytes from sticking to each other. We found that it is necessary to perform the experiment immediately after the collection and fixation of the oocytes. Any delay results in a much lower fluorescence signal that would result in undercounting...

Reverse-transcriptase polymerase chain reaction (PCR) has been the gold standard for mRNA quantitation. Two assays, digital PCR (dPCR)1 and quantitative, real time PCR (qPCR)2 are currently used. Of the two PCR techniques, dPCR has greater sensitivity than qPCR suggesting that it could be used to measure mRNA abundance in single cells. However, in our hands, dPCR analysis of low abundance mRNAs in pools of 5 to 10 oocytes per each experimental sample has produced data with low reproducibility and high variation3. This is likely due to the experimental error associated with RNA extraction and reverse transcription efficiency. RNA sequencing has also been performed using a single mouse and human oocytes4,5. This technique requires cDNA amplification steps required for the library generation which likely increases variability within an experimental group. Furthermore, low abundance transcripts may not be detectable. Although sequencing prices have gone down&#160;the last few years, it can still be cost prohibitive due to the high cost of bioinformatics analyses. Finally, mRNA localization is a dynamic process with spatial changes contributing to protein function6. Therefore, we set out to adopt a technique that would produce accurate and reproducible quantitative measures and localization of individual mRNAs in single oocytes.
Branched DNA coupled to fluorescence in situ hybridization amplifies fluorescence signal rather than amplifying RNA/cDNA enabling detection of single mRNAs in individual cells 7,8,9. The assay is performed through a series of hybridization, amplification (using branched DNA), and fluorescence labeling steps in order to amplify the fluorescence signal7. The technique begins with binding of 18- to 25-base oligonucleotide probe pairs that are complementary to a specific mRNA3,8,10. Fifteen to twenty probe pairs are designed for each transcript ensuring specificity for the target transcript. The mRNA-specific hybridization is followed by pre-amplifier and amplifier probes that form a branched configuration. Approximately, 400 label fluorophores bind to each amplifier, resulting in an 8000-fold increase in fluorescence allowing detection of individual mRNAs (Figure 1)11.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59414/59414fig1.jpg" /> 
Figure 1: Schematic of the SM-FISH protocol. Sequential hybridization of transcript specific probe, branched DNA amplifier and fluorophore to a target mRNA is shown. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/59414/59414fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Previous studies using single molecule fluorescence in situ hybridization (SM-FISH) localized &#946;-actin mRNAs in individual neurons12 and human papillomavirus DNA in cervical cancer cell lines7. The computer software Spot Finding and Tracking Program identifies individual punctate fluorescent signal and has been successfully used to quantify the number of mRNAs in each cell3,13.
Based on the results of mRNA detection in neurons12, we hypothesized that SM-FISH would prove a useful tool to quantitate transcript levels in murine oocytes and embryos including low abundance mRNAs. However, the technique is optimized for the use with adherent fixed cells and formaldehyde fixed paraffin embedded (FFPE) tissue sections. Oocytes cannot adhere to a slide, even when they are coated with Poly-L-lysine. Furthermore, they are more fragile than somatic cells and tissue sections resulting in cell lysis when subjected to some of the proprietary buffers in commercially available kits3. To overcome these challenges, oocytes were fixed and manually transferred between drops of the buffers. Furthermore, permeabilization and wash buffers in the kits were replaced to reduce the cell lysis. Predesigned probes are purchased alongside the FISH kit or specific transcripts can be requested. Each proprietary probe set is available in one of three fluorescence channels (C1, C2, and C3) to allow for multiplexing. In the current experiment, murine oocytes were dual-stained and quantified using a C2 Nanog probe and a C3 Pou5f1 probe. These probes were selected based on the reported expression of Nanog and Pou5f1 in oocytes and embryos. At the conclusion of the hybridization steps, oocytes were placed in drops of anti-fade mounting media for application to histological slides. Confocal images were used to quantify the number of punctate fluorescent signals which represent individual mRNAs. In addition to quantifying the mRNAs, imaging also showed the spatial distribution of the specific mRNA in the cell, which other RNA quantification methods are unable to achieve. This technique proved to have low variability within an experimental group allowing the use of smaller numbers of oocytes in each experimental group to identify significant differences between experimental groups3.

<table><tbody><tr><td>(&amp;plusmn;)-&amp;alpha;-Lipoic acid</td><td>Sigma-Aldrich</td><td>T1395</td><td>Alpha Lipoic Acid</td></tr><tr><td>Albumin, Bovine Serum, Low Fatty Acid</td><td>MP Biomedicals, LLC</td><td>199899</td><td>FAF BSA</td></tr><tr><td>BD 10mL TB Syringe</td><td>Becton, Dickinson and Company</td><td>309659</td><td>10 mL syringe</td></tr><tr><td>BD PrecisionGlide Needle</td><td>Becton, Dickinson and Company</td><td>305109</td><td>27 1/2 gauge needle</td></tr><tr><td>Calcium chloride dihydrate</td><td>Sigma-Aldrich</td><td>C7902</td><td>CaCl2-2H2O</td></tr><tr><td>Citric acid</td><td>Sigma-Aldrich</td><td>C2404</td><td>Citrate</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma-Aldrich</td><td>G6152</td><td>Glucose</td></tr><tr><td>Disodium phosphate</td><td></td><td></td><td>Na2HPO4</td></tr><tr><td>Easy Grip Petri Dish</td><td>Falcon Corning</td><td>351008</td><td>35 mm dish</td></tr><tr><td>Edetate Disodium</td><td>Avantor</td><td>8994-01</td><td>EDTA</td></tr><tr><td>Extra Fine Bonn Scissors</td><td>Fine Science Tools</td><td>14084-08</td><td>Straight, Sharp/Sharp, non-serrated, 13mm cutting edge scissors</td></tr><tr><td>Fetal Bovine Serum</td><td>Atlanta biologicals</td><td>S10250</td><td>FBS</td></tr><tr><td>Gentamicin Reagent Solution</td><td>gibco</td><td>15710-064</td><td>Gentamicin</td></tr><tr><td>GlutaMAX-I (100X)</td><td>gibco</td><td>35050-061</td><td>Glutamax</td></tr><tr><td>Gold Seal Micro Slides</td><td>Gold Seal</td><td>3039</td><td>25 x 75mm slides</td></tr><tr><td>Gonadotropin, From Pregnant Mares&#039; Serum</td><td>Sigma</td><td>G4877</td><td>eCG</td></tr><tr><td>hCG recombinant</td><td>NHPP</td><td>AFP8456A</td><td>hCG</td></tr><tr><td>Hyaluronidase, Type IV-S: From Bovine Testes</td><td>Sigma-Aldrich</td><td>H3884</td><td>Hyaluronidase</td></tr><tr><td>Jewelers Style Forceps</td><td>Integra</td><td>17-305X</td><td>Forceps 4-3/8&quot;, Style 5F, Straight, Micro Fine Jaw</td></tr><tr><td>L-(+)-Lactic Acid, free acid&amp;nbsp;</td><td>MP Biomedicals, LLC</td><td>190228</td><td>L-Lactate</td></tr><tr><td>Magnesium sulfate heptahydrate</td><td>Sigma-Aldrich</td><td>M2773</td><td>MgSO4-7H2O</td></tr><tr><td>MEM Nonessential Amino Acids</td><td>Corning&amp;nbsp;</td><td>25-025-Cl</td><td>NEAA</td></tr><tr><td>Microscope Cover Glass</td><td>Fisher Scientific</td><td>12-542-C</td><td>25 x 25x 0.15 mm cover slips</td></tr><tr><td>Mm-Nanog-O2-C2 RNAscope Probe</td><td>Advanced Cell Diagnostics</td><td>501891-C2</td><td>Nanog Probe</td></tr><tr><td>Mm-Pou5f1-O1-C3 RNAscope Probe</td><td>Advanced Cell Diagnostics</td><td>501611-C3</td><td>Pou5f1 Probe</td></tr><tr><td>MOPS</td><td>Sigma-Aldrich</td><td>M3183</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>P6148</td><td>Paraformaldehyde</td></tr><tr><td>PES 0.22 um Membrane -sterile</td><td>Millex-GP</td><td>SLGP033RS</td><td>0.22 um filters</td></tr><tr><td>Polyvinylpyrrolidone</td><td>Sigma-Aldrich</td><td>P0930</td><td>PVP</td></tr><tr><td>Potassium chloride</td><td>Sigma-Aldrich</td><td>60128</td><td>KCl</td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma-Aldrich</td><td>60218</td><td>KH2PO4</td></tr><tr><td>Prolong Gold antifade reagent</td><td>invitrogen</td><td>P36934</td><td>Antifade reagent without DAPI</td></tr><tr><td>RNAscope DAPI</td><td>Advanced Cell Diagnostics</td><td>320858</td><td>DAPI</td></tr><tr><td>RNAscope FL AMP 1</td><td>Advanced Cell Diagnostics</td><td>320852</td><td>Amplifier 1</td></tr><tr><td>RNAscope FL AMP 2</td><td>Advanced Cell Diagnostics</td><td>320853</td><td>Amplifier 2</td></tr><tr><td>RNAscope FL AMP 3</td><td>Advanced Cell Diagnostics</td><td>320854</td><td>Amplifier 3</td></tr><tr><td>RNAscope FL AMP 4 ALT A</td><td>Advanced Cell Diagnostics</td><td>320855</td><td>Amplifier 4 ALT A</td></tr><tr><td>RNAscope FL AMP 4 ALT B</td><td>Advanced Cell Diagnostics</td><td>320856</td><td>Amplifier 4 ALT B</td></tr><tr><td>RNAscope FL AMP 4 ALT C</td><td>Advanced Cell Diagnostics</td><td>320857</td><td>Amplifier 4 ALT C</td></tr><tr><td>RNAscope Fluorescent Multiplex Detection Reagents Kit</td><td>Advanced Cell Diagnostics</td><td>320851</td><td>FISH Reagent Kit</td></tr><tr><td>RNAscope Probe 3-plex Negative Control Probe</td><td>Advanced Cell Diagnostics</td><td>320871</td><td>Negative Control</td></tr><tr><td>RNAscope Probe 3-plex Positive Control</td><td>Advanced Cell Diagnostics</td><td>320881</td><td>Positive Control</td></tr><tr><td>RNAscope Probe Diluent</td><td>Advanced Cell Diagnostics</td><td>300041</td><td>Probe Diluent</td></tr><tr><td>RNAscope Protease III</td><td>Advanced Cell Diagnositics</td><td>322337</td><td>Protease III</td></tr><tr><td>RNAscope Protease III &amp;amp; IV Reagent Kit</td><td>Advanced Cell Diagnostics</td><td>322340</td><td>FISH Protease Kit</td></tr><tr><td>RNAscope Protease IV</td><td>Advanced Cell Diagnostics</td><td>322336</td><td>Protease IV</td></tr><tr><td>S/S Needle with Luer Hub 30G</td><td>Component Supply Co.</td><td>NE-301PL-50</td><td>blunt 30 gauge needle</td></tr><tr><td>Sodium bicarbonate</td><td>Sigma-Aldrich</td><td>S6297</td><td>NaHCO3</td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich</td><td>S6191</td><td>NaCl</td></tr><tr><td>Sodium hydroxide</td><td>Sigma-Aldrich</td><td>306576</td><td>NaOH</td></tr><tr><td>Sodium pyruvate, &gt;= 99%</td><td>Sigma-Aldrich</td><td>P5280</td><td>Pyruvate</td></tr><tr><td>Solution 6 Well Dish</td><td>Agtechinc</td><td>D18</td><td>6 well dish</td></tr><tr><td>Taurine</td><td>Sigma-Aldrich</td><td>T8691</td><td>Taurine</td></tr><tr><td>Tissue Culture Dish</td><td>Falcon Corning</td><td>353002</td><td>60 mm dish</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100</td><td>Triton X-100</td></tr></tbody></table>

use of single molecule fluorescent in situ hybridization (sm-fish) to quantify and localize mrnas in murine oocytes

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

Upon the completion of the protocol, the result will be individual images from confocal z-series (Figure 4A and Figure 5), stitched images (Figure 4C), and mRNA counts (Figure 4B). When multiplexing is performed, there will also be merged images showing the label for two different mRNAs (Fig...

Watch this Scientific Journal Video about Use of Single Molecule Fluorescent In Situ Hybridization SM-FISH to Quantify and Localize mRNAs in Murine Oocytes at JoVE.com

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

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

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

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

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Universidad de Cartagena UDEC

Research

JoVE Journal

Genetics

10.6K Views.  University of Nebraska-Lincoln. To reproducibly count the numbers of mRNAs in individual oocytes, single molecule RNA fluorescence in situ hybridization (RNA-FISH) was optimized for non-adherent cells. Oocytes were collected, hybridized with the transcript specific probes, and quantified using an image quantification software.

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

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively correlated with maternal age, age alone is not always predictive of the risk of generating an aneuploid embryo. Therefore, gene variants might account for incorrect chromosome segregation during oogenesis. Given that access to human oocytes is limited for research purposes, a series of assays were developed to study human gene function during meiosis I using mouse oocytes. First, messenger RNA (mRNA) of the gene and gene variant of interest are microinjected into prophase I-arrested mouse oocytes. After allowing time for expression, oocytes are synchronously released into meiotic maturation to complete meiosis I. By tagging the mRNA with a sequence of a fluorescent reporter, such as green fluorescent protein (Gfp), the localization of the human protein can be assessed in addition to the phenotypic alterations. For example, gain or loss of function can be investigated by establishing experimental conditions that challenge the gene product to fix meiotic errors. Although this system is advantageous in investigating human protein function during oogenesis, adequate interpretation of results should be undertaken given that protein expression is not at endogenous levels and, unless controlled for (i.e. knocked out or down), murine homologs are also present in the system.

As the genetic variants associated with human disease begin to become uncovered, it is becoming increasingly important to develop systems with which to rapidly evaluate the biological significance of those identified variants. This protocol describes methods for evaluating human gene function during female meiosis I using mouse oocytes.

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively ...

Visualizing RNA Localization in Xenopus Oocytes

RNA localization is a conserved mechanism of establishing cell polarity. Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to spatially restrict gene expression of Vg1 protein. Tight control of Vg1 distribution in this manner is required for proper germ layer specification in the developing embryo. RNA sequence elements in the 3' UTR of the mRNA, the Vg1 localization element (VLE) are required and sufficient to direct transport. To study the recognition and transport of Vg1 mRNA in vivo, we have developed an imaging technique that allows extensive analysis of trans-factor directed transport mechanisms via a simple visual readout. 
To visualize RNA localization, we synthesize fluorescently labeled VLE RNA and microinject this transcript into individual oocytes. After oocyte culture to allow transport of the injected RNA, oocytes are fixed and dehydrated prior to imaging by confocal microscopy. Visualization of mRNA localization patterns provides a readout for monitoring the complete pathway of RNA transport and for identifying roles in directing RNA transport for cis-acting elements within the transcript and trans-acting factors that bind to the VLE (Lewis et al., 2008, Messitt et al., 2008). We have extended this technique through co-localization with additional RNAs and proteins (Gagnon and Mowry, 2009, Messitt et al., 2008), and in combination with disruption of motor proteins and the cytoskeleton (Messitt et al., 2008) to probe mechanisms underlying mRNA localization.

Visualizing RNA Localization in Xenopus Oocytes

Visualization of in vivo RNA transport is accomplished by microinjection of fluorescently labeled RNA transcripts into Xenopus oocytes, followed by confocal microscopy.

RNA localization is a conserved mechanism of establishing cell polarity.  Vg1 mRNA localizes to the vegetal pole of Xenopus laevis oocytes and acts to ...

Biology

Defining the Program of Maternal mRNA Translation during In vitro Maturation using a Single Oocyte Reporter Assay

Events associated with oocyte nuclear maturation have been well described. However, much less is known about the molecular pathways and processes that take place in the cytoplasm in preparation for fertilization and acquisition of totipotency. During oocyte maturation, changes in gene expression depend exclusively on the translation and degradation of maternal messenger RNAs (mRNAs) rather than on transcription. Execution of the translational program, therefore, plays a key role in establishing oocyte developmental competence to sustain embryo development. This paper is part of a focus on defining the program of maternal mRNA translation that takes place during meiotic maturation and at the oocyte-to-zygote transition. In this method paper, a strategy is presented to study the regulation of translation of target mRNAs during in vitro oocyte maturation. Here, a Ypet reporter is fused to the 3&#39; untranslated region (UTR) of the gene of interest and then micro-injected into oocytes together with polyadenylated mRNA encoding for mCherry to control for injected volume. By using time-lapse microscopy to measure reporter accumulation, translation rates are calculated at different transitions during oocyte meiotic maturation. Here, the protocols for oocyte isolation and injection, time-lapse recording, and data analysis have been described, using the Ypet/interleukin-7 (IL-7)-3&#39; UTR reporter as an example.

This protocol describes a reporter assay to study the regulation of mRNA translation in single oocytes during in vitro maturation.

Events associated with oocyte nuclear maturation have been well described. However, much less is known about the molecular pathways and processes that ...

Developmental Biology

Detection of Axonally Localized mRNAs in Brain Sections Using High-Resolution In Situ Hybridization

mRNAs are frequently localized to vertebrate axons and their local translation is required for axon pathfinding or branching during development and for maintenance, repair or neurodegeneration in postdevelopmental periods. High throughput analyses have recently revealed that axons have a more dynamic and complex transcriptome than previously expected. These analysis, however have been mostly done in cultured neurons where axons can be isolated from the somato-dendritic compartments. It is virtually impossible to achieve such isolation in whole tissues in vivo. Thus, in order to verify the recruitment of mRNAs and their functional relevance in a whole animal, transcriptome analyses should ideally be combined with techniques that allow the visualization of mRNAs in situ. Recently, novel ISH technologies that detect RNAs at a single-molecule level have been developed. This is especially important when analyzing the subcellular localization of mRNA, since localized RNAs are typically found at low levels. Here we describe two protocols for the detection of axonally-localized mRNAs using a novel ultrasensitive RNA ISH technology. We have combined RNAscope ISH with axonal counterstain using fluorescence immunohistochemistry or histological dyes to verify the recruitment of Atf4 mRNA to axons in vivo in the mature mouse and human brains.

Detection of Axonally Localized mRNAs in Brain Sections Using High-Resolution In Situ Hybridization

RNA in situ hybridization (ISH) enables the visualization of RNAs in cells and tissues. Here we show how combination of RNAscope ISH with immunohistochemistry or histological dyes can be successfully used to detect mRNAs localized to axons in sections of mouse and human brains.

mRNAs are frequently localized to vertebrate axons and their local translation is required for axon pathfinding or branching during development and for ...

Neuroscience

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Development of the lower urogenital tract (LUT) is an intricate process. This complexity is evidenced during formation of the prostate from the fetal male urethra, which relies on androgenic signals and epithelial-mesenchymal interactions1,2. Understanding the molecular mechanisms responsible for prostate development may reveal growth mechanisms that are inappropriately reawakened later in life to give rise to prostate diseases such as benign prostatic hyperplasia and prostate cancer.
The developing LUT is anatomically complex. By the time prostatic budding begins on 16.5 days post conception (dpc), numerous cell types are present. Vasculature, nerves and smooth muscle reside within the mesenchymal stroma3. This stroma surrounds a multilayered epithelium and gives rise to the fetal prostate through androgen receptor-dependent paracrine signals4. The identity of the stromal androgen receptor-responsive genes required for prostate development and the mechanism by which prostate ductal epithelium forms in response to these genes is not fully understood. The ability to precisely identify cell types and localize expression of specific factors within them is imperative to further understand prostate development. In situ hybridization (ISH) allows for localization of mRNAs within a tissue. Thus, this method can be used to identify pattern and timing of expression of signaling molecules and their receptors, thereby elucidating potential prostate developmental regulators.
Here, we describe a high throughput ISH technique to identify mRNA expression patterns in the fetal mouse LUT using vibrating microtome-cut sections. This method offers several advantages over other ISH protocols. Performing ISH on thin sections adhered to a slide is technically difficult; cryosections frequently have poor structural quality while both cryosections and paraffin sections often result in weak signal resolution. Performing ISH on whole mount tissues can result in probe trapping. In contrast, our high throughput technique utilizes thick-cut sections that reveal detailed tissue architecture. Modified microfuge tubes allow easy handling of sections during the ISH procedure. A maximum of 4 mRNA transcripts can be screened from a single 17.5dpc LUT with up to 24 mRNA transcripts detected in a single run, thereby reducing cost and maximizing efficiency. This method allows multiple treatment groups to be processed identically and as a single unit, thereby removing any bias for interpreting data. Most pertinently for prostate researchers, this method provides a spatial and temporal location of low and high abundance mRNA transcripts in the fetal mouse urethra that gives rise to the prostate ductal network.

A High Throughput in situ Hybridization Method to Characterize mRNA Expression Patterns in the Fetal Mouse Lower Urogenital Tract

Here, we describe an efficient high throughput in situ hybridization (ISH) method for visualizing patterns of mRNA expression in developing fetal mouse prostate tissue sections. The method can be easily adapted to visualize mRNA expression patterns in other mouse tissues or in tissues from other species.

Development of the lower urogenital tract (LUT) is an intricate process. This complexity is evidenced during formation of the prostate from the fetal male ...

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 ...

Meiotic Spindle Assessment in Mouse Oocytes by siRNA-mediated Silencing

Errors in chromosome segregation during meiotic division in gametes can lead to aneuploidy that is subsequently transmitted to the embryo upon fertilization. The resulting aneuploidy in developing embryos is recognized as a major cause of pregnancy loss and congenital birth defects such as Down&#8217;s syndrome. Accurate chromosome segregation is critically dependent on the formation of the microtubule spindle apparatus, yet this process remains poorly understood in mammalian oocytes. Intriguingly, meiotic spindle assembly differs from mitosis and is regulated, at least in part, by unique microtubule organizing centers (MTOCs). Assessment of MTOC-associated proteins can provide valuable insight into the regulatory mechanisms that govern meiotic spindle formation and organization. Here, we describe methods to isolate mouse oocytes and deplete MTOC-associated proteins using a siRNA-mediated approach to test function. In addition, we describe oocyte fixation and immunofluorescence analysis conditions to evaluate meiotic spindle formation and organization.

Here, we present a protocol for specific siRNA-mediated mRNA depletion followed by immunofluorescence analysis to evaluate meiotic spindle assembly and organization in mouse oocytes. This protocol is suitable for in vitro depletion of transcripts and functional assessment of different spindle and/or MTOC-associated factors in oocytes.

Errors in chromosome segregation during meiotic division in gametes can lead to aneuploidy that is subsequently transmitted to the embryo upon ...

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

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

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

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

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

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

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

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

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

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

Impact of Cytoskeleton on Nuclear Shape and Condensate Dynamics in Mouse Oocytes

Capturing Cytoskeleton-Based Agitation of the Mouse Oocyte Nucleus Across Spatial Scales

A major challenge in understanding the causes of female infertility is to elucidate mechanisms governing the development of female germ cells, named oocytes. Their development is marked by cell growth and subsequent divisions, two critical phases that prepare the oocyte for fusion with sperm to initiate embryogenesis. During growth, oocytes reorganize their cytoplasm to position the nucleus at the cell center, an event predictive of successful oocyte development in mice and humans and, thus, their embryogenic potential. In mouse oocytes, this cytoplasmic reorganization was shown to be driven by the cytoskeleton, the activity of which generates mechanical forces that agitate, reposition, and penetrate the nucleus. Consequently, this cytoplasmic-to-nucleoplasmic force transmission tunes the dynamics of nuclear RNA-processing organelles known as biomolecular condensates. This protocol provides an experimental framework to document, with high temporal resolution, the impact of the cytoskeleton on the nucleus across spatial scales in mouse oocytes. It details the imaging and image analysis steps and tools necessary to evaluate i) cytoskeletal activity in the oocyte cytoplasm, ii) cytoskeleton-based agitation of the oocyte nucleus, and iii) its effects on biomolecular condensate dynamics in the oocyte nucleoplasm. Beyond oocyte biology, the methods elaborated here can be adapted for use in somatic cells to similarly address cytoskeleton-based tuning of nuclear dynamics across scales.

Author Spotlight: Elucidating the Dynamics of Mechano-Transduction and Nuclear Agitation in Mouse Oocytes

This protocol provides an experimental framework to document the physical impact of the cytoskeleton on nuclear shape and the internal membrane-less organelles in the mouse oocyte system. The framework can be adapted for use in other cell types and contexts.

A major challenge in understanding the causes of female infertility is to elucidate mechanisms governing the development of female germ cells, named ...

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

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Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

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

Capturing Cytoskeleton-Based Agitation of the Mouse Oocyte Nucleus Across Spatial Scales