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://www.jove.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://www.jove.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://www.jove.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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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://www.jove.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.

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</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), ...

use-single-molecule-fluorescent-situ-hybridization-sm-fish-to

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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

Detection of Copy Number Alterations Using Single Cell Sequencing

Detection of genomic changes at single cell resolution is important for characterizing genetic heterogeneity and evolution in normal tissues, cancers, and microbial populations. Traditional methods for assessing genetic heterogeneity have been limited by low resolution, low sensitivity, and/or low specificity. Single cell sequencing has emerged as a powerful tool for detecting genetic heterogeneity with high resolution, high sensitivity and, when appropriately analyzed, high specificity. Here we provide a protocol for the isolation, whole genome amplification, sequencing, and analysis of single cells. Our approach allows for the reliable identification of megabase-scale copy number variants in single cells. However, aspects of this protocol can also be applied to investigate other types of genetic alterations in single cells.

Single cell sequencing is an increasingly popular and accessible tool for addressing genomic changes at high resolution. We provide a protocol that uses single cell sequencing to identify copy number alterations in single cells.

Detection of genomic changes at single cell resolution is important for characterizing genetic heterogeneity and evolution in normal tissues, cancers, and ...

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 ...

Generation of Fluorescent Protein Fusions in Candida Species

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. Thus, molecular and genetic tools are needed to facilitate the study of their pathogenesis mechanisms. PCR-mediated gene modification is a straightforward and quick approach to generate epitope-tagged proteins to facilitate their detection. In particular, fluorescent protein (FP) fusions are powerful tools that allow visualization and quantitation of both yeast cells and proteins by fluorescence microscopy and immunoblotting, respectively. Plasmids containing FP encoding sequences, along with nutritional marker genes that facilitate the transformation of Candida species, have been generated for the purpose of FP construction and expression in Candida. Herein, we present a strategy for constructing a FP fusion in a Candida species. Plasmids containing the nourseothricin resistance transformation marker gene (NAT1) along with sequences for either green, yellow, or cherry FPs (GFP, YFP, mCherry) are used along with primers that include gene-specific sequences in a polymerase chain reaction (PCR) to generate a FP cassette. This gene-specific cassette has the ability to integrate into the 3'-end of the corresponding gene locus via homologous recombination. Successful in-frame fusion of the FP sequence into the gene locus of interest is verified genetically, followed by analysis of fusion protein expression by microscopy and/or immuno-detection methods. In addition, for the case of highly expressed proteins, successful fusions can be screened for primarily by fluorescence imaging techniques.

Generation of Fluorescent Protein Fusions in Candida Species

PCR-mediated gene modification can be used to generate fluorescent protein fusions in Candida species, which facilitates visualization and quantitation of yeast cells and proteins. Herein, we present a strategy for constructing a fluorescent protein fusion (Eno1-FP) in Candida parapsilosis.

Candida species, prevalent colonizers of the intestinal and genitourinary tracts, are the cause of the majority of invasive fungal infections in humans. ...

Single Molecule Analysis of Laser Localized Psoralen Adducts

The DNA Damage Response (DDR) has been extensively characterized in studies of double strand breaks (DSBs) induced by laser micro beam irradiation in live cells. The DDR to helix distorting covalent DNA modifications, including interstrand DNA crosslinks (ICLs), is not as well defined. We have studied the DDR stimulated by ICLs, localized by laser photoactivation of immunotagged psoralens, in the nuclei of live cells. In order to address fundamental questions about adduct distribution and replication fork encounters, we combined laser localization with two other technologies. DNA fibers are often used to display the progress of replication forks by immunofluorescence of nucleoside analogues incorporated during short pulses. Immunoquantum dots have been widely employed for single molecule imaging. In the new approach, DNA fibers from cells carrying laser localized ICLs are spread onto microscope slides. The tagged ICLs are displayed with immunoquantum dots and the inter-lesion distances determined. Replication fork collisions with ICLs can be visualized and different encounter patterns identified and quantitated.

Lasers are frequently used in studies of the cellular response to DNA damage. However, they generate lesions whose spacing, frequency, and collisions with replication forks are rarely characterized. Here, we describe an approach that enables the determination of these parameters with laser localized interstrand crosslinks.

The DNA Damage Response (DDR) has been extensively characterized in studies of double strand breaks (DSBs) induced by laser micro beam irradiation in live ...

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of tissues, we have developed numerous modifications and improvements to our original fluorescent in situ hybridization (FISH) protocol. To facilitate throughput and cost effectiveness, all steps, from probe generation to signal detection, are performed using exon 96-well microtiter plates. Digoxygenin (DIG)-labelled antisense RNA probes are produced using either cDNA clones or genomic DNA as templates. After tissue fixation and permeabilization, probes are hybridized to transcripts of interest and then detected using a succession of anti-DIG antibody conjugated to biotin, streptavidin conjugated to horseradish peroxidase (HRP) and fluorescently conjugated tyramide, which in the presence of HRP, produces a highly reactive intermediate that binds to electron dense regions of immediately adjacent proteins. These amplification and localization steps produce a robust and highly localized signal that facilitates both cellular and subcellular transcript localization. The protocols provided have been optimized to produce highly specific signals in a variety of tissues and developmental stages. References are also provided for additional variations that allow the simultaneous detection of multiple transcripts, or transcripts and proteins, at the same time.

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

The described RNA in situ hybridization protocol allows the detection of RNA in whole Drosophila embryos or dissected tissues. Using 96-well microtiter plates and tyramide signal amplification, transcripts can be detected at high resolution, sensitivity, and throughput, and at a relatively low cost.

In our efforts to determine the patterns of expression and subcellular localization of Drosophila RNAs on a genome-wide basis, and in a variety of ...

Visualization of Microbiota in Tick Guts by Whole-mount In Situ Hybridization

Infectious diseases transmitted by arthropod vectors continue to pose a significant threat to human health worldwide. The pathogens causing these diseases, do not exist in isolation when they colonize the vector; rather, they likely engage in interactions with resident microorganisms in the gut lumen. The vector microbiota has been demonstrated to play an important role in pathogen transmission for several vector-borne diseases. Whether resident bacteria in the gut of the Ixodes scapularis tick, the vector of several human pathogens including Borrelia burgdorferi, influence tick transmission of pathogens is not determined. We require methods for characterizing the composition of the bacteria associated with the tick gut to facilitate a better understanding of potential interspecies interactions in the tick gut. Using whole-mount in situ hybridization to visualize RNA transcripts associated with particular bacterial species allows for the collection of qualitative data regarding the abundance and distribution of the microbiota in intact tissue. This technique can be used to examine changes in the gut microbiota milieu over the course of tick feeding and can also be applied to analyze expression of tick genes. Staining of whole tick guts yield information about the gross spatial distribution of target RNA in the tissue without the need for three-dimensional reconstruction and is less affected by environmental contamination, which often confounds the sequencing-based methods frequently used to study complex microbial communities. Overall, this technique is a valuable tool that can be used to better understand vector-pathogen-microbiota interactions and their role in disease transmission.

Visualization of Microbiota in Tick Guts by Whole-mount In Situ Hybridization

Here, we present a protocol to spatially and temporally assess the presence of viable microbiota in tick guts using a modified whole-mount in situ hybridization approach.

Infectious diseases transmitted by arthropod vectors continue to pose a significant threat to human health worldwide. The pathogens causing these ...

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.

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) is a powerful technique to study gene expression in single cells due to its ability to detect ...

Quantitative Analysis of Alternative Pre-mRNA Splicing in Mouse Brain Sections Using RNA In Situ Hybridization Assay

Alternative splicing (AS) occurs in more than 90% of human genes. The expression pattern of an alternatively spliced exon is often regulated in a cell type-specific fashion. AS expression patterns are typically analyzed by RT-PCR and RNA-seq using RNA samples isolated from a population of cells. In situ examination of AS expression patterns for a particular biological structure can be carried out by RNA in situ hybridization (ISH) using exon-specific probes. However, this particular use of ISH has been limited because alternative exons are generally too short to design exon-specific probes. In this report, the use of BaseScope, a recently developed technology that employs short antisense oligonucleotides in RNA ISH, is described to analyze AS expression patterns in mouse brain sections. Exon 23a of neurofibromatosis type 1 (Nf1) is used as an example to illustrate that short exon-exon junction probes exhibit robust hybridization signals with high specificity in RNA ISH analysis on mouse brain sections. More importantly, signals detected with exon inclusion- and skipping-specific probes can be used to reliably calculate the percent spliced in values of Nf1 exon 23a expression in different anatomical areas of a mouse brain. The experimental protocol and calculation method for AS analysis are presented. The results indicate that BaseScope provides a powerful new tool to assess AS expression patterns in situ.

Quantitative Analysis of Alternative Pre-mRNA Splicing in Mouse Brain Sections Using RNA In Situ Hybridization Assay

An in situ hybridization (ISH) protocol that uses short antisense oligonucleotides to detect alternative pre-mRNA splicing patterns in mouse brain sections is described.

Alternative splicing (AS) occurs in more than 90% of human genes. The expression pattern of an alternatively spliced exon is often regulated in a cell ...

Tissue-specific miRNA Expression Profiling in Mouse Heart Sections Using In Situ Hybridization

micro-RNAs (miRNAs) are single-stranded RNA transcripts that bind to messenger RNAs (mRNAs) and inhibit their translation or promote their degradation. To date, miRNAs have been implicated in a large number of biological and disease processes, which has signified the need for the reliable detection methods of miRNA transcripts. Here, we describe a detailed protocol for digoxigenin-labeled (DIG) Locked Nucleic Acid (LNA) probe-based miRNA detection, combined with protein immunostaining on mouse heart sections. First, we performed an in situ hybridization technique using the probe to identify miRNA-182 expression in heart sections from control and cardiac hypertrophy mice. Next, we performed immunostaining for cardiac Troponin T (cTnT) protein, on the same sections, to co-localize miRNA-182 with the cardiomyocyte cells. Using this protocol, we were able to detect miRNA-182 through an alkaline phosphatase based colorimetric assay, and cTnT through fluorescent staining. This protocol can be used to detect the expression of any miRNA of interest through DIG-labeled LNA probes, and relevant protein expression on mouse heart tissue sections.

Tissue-specific miRNA Expression Profiling in Mouse Heart Sections Using In Situ Hybridization

micro-RNAs (miRNAs) are short and highly homologous RNA sequences, serving as post-transcriptional regulators of messenger RNAs (mRNAs). Current miRNA detection methods vary in sensitivity and specificity. We describe a protocol that combines in situ hybridization and immunostaining for concurrent detection of miRNA and protein molecules on mouse heart tissue sections.

micro-RNAs (miRNAs) are single-stranded RNA transcripts that bind to messenger RNAs (mRNAs) and inhibit their translation or promote their degradation. To ...

Using In Vitro and In-cell SHAPE to Investigate Small Molecule Induced Pre-mRNA Structural Changes

In the process of drug development of RNA-targeting small molecules, elucidating the structural changes upon their interactions with target RNA sequences is desired. We herein provide a detailed in vitro and in-cell selective 2&#39;-hydroxyl acylation analyzed by primer extension (SHAPE) protocol to study the RNA structural change in the presence of an experimental drug for spinal muscular atrophy (SMA), survival of motor neuron (SMN)-C2, and in exon 7 of the pre-mRNA of the SMN2 gene. In in vitro SHAPE, an RNA sequence of 140 nucleotides containing SMN2 exon 7 is transcribed by T7 RNA polymerase, folded in the presence of SMN-C2, and subsequently modified by a mild 2&#39;-OH acylation reagent, 2-methylnicotinic acid imidazolide (NAI). This 2&#39;-OH-NAI adduct is further probed by a 32P-labeled primer extension and resolved by polyacrylamide gel electrophoresis (PAGE). Conversely, 2&#39;-OH acylation in in-cell SHAPE takes place in situ with SMN-C2 bound cellular RNA in living cells. The pre-mRNA sequence of exon 7 in the SMN2 gene, along with SHAPE-induced mutations in the primer extension, was then amplified by PCR and subject to next-generation sequencing. Comparing the two methodologies, in vitro SHAPE is a more cost-effective method and does not require computational power to visualize results. However, the in vitro SHAPE-derived RNA model sometimes deviates from the secondary structure in a cellular context, likely due to the loss of all interactions with RNA-binding proteins. In-cell SHAPE does not need a radioactive material workplace and yields a more accurate RNA secondary structure in the cellular context. Furthermore, in-cell SHAPE is usually applicable for a larger range of RNA sequences (~1,000 nucleotides) by utilizing next-generation sequencing, compared to in vitro SHAPE (~200 nucleotides) that usually relies on PAGE analysis. In case of exon 7 in SMN2 pre-mRNA, the in vitro and in-cell SHAPE derived RNA models are similar to each other.

Detailed protocols for both in vitro and in-cell selective 2&#39;-hydroxyl acylation analyzed by primer extension (SHAPE) experiments to determine the secondary structure of pre-mRNA sequences of interest in the presence of an RNA-targeting small molecule are presented in this article.

In the process of drug development of RNA-targeting small molecules, elucidating the structural changes upon their interactions with target RNA sequences ...

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

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Detection of Copy Number Alterations Using Single Cell Sequencing

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

Generation of Fluorescent Protein Fusions in Candida Species

Single Molecule Analysis of Laser Localized Psoralen Adducts

High Resolution Fluorescent In Situ Hybridization in Drosophila Embryos and Tissues Using Tyramide Signal Amplification

Visualization of Microbiota in Tick Guts by Whole-mount In Situ Hybridization

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

Quantitative Analysis of Alternative Pre-mRNA Splicing in Mouse Brain Sections Using RNA In Situ Hybridization Assay

Tissue-specific miRNA Expression Profiling in Mouse Heart Sections Using In Situ Hybridization

Using In Vitro and In-cell SHAPE to Investigate Small Molecule Induced Pre-mRNA Structural Changes