This work was supported by a grant from the National Cancer Institute, CA131967.

1. BrdU Incorporation (Terminal Labeling)
<ol>
 <li>Plate cells to about 70% confluence in a 150 mm tissue culture dish 24 hr prior to the addition of BrdU.</li>
 <li>Replace media with fresh complete media containing 20 &mu;g/ml BrdU (Sigma) at appropriate time points prior to harvest. The length of time cells are cultured in media with BrdU will vary with cell type and species, typically the G2 phase lasts between 2 and 5 hr (Figure 1).</li>
</ol>
2. Chromosome Harvest of Monolayer Cells Cultures
<ol>
 <li>Remove culture media from the plate and collect 10 ml in a 15 ml conical centrifuge tube. Discard remaining media.</li>
 <li>Rinse cells once with 10 ml Versine [or HBSS (Sigma)].</li>
 <li>Add 5 ml 0.25% trypsin in Versine (or HBSS) to the plate. Incubate at room temperature until cells are detached from the plate.</li>
 <li>Add trypsinized cell suspension to the tube containing the 10 ml of media from step 2.1.</li>
 <li>Centrifuge at 400 x g for 10 min to pellet the cells. Aspirate supernatant, leaving 0.5 ml of media plus the cell pellet.</li>
 <li>Resuspend cells thoroughly by pipetting with a Pasteur pipette.</li>
 <li>Add 3 drops of hypotonic solution (0.075 M KCl (Sigma) warmed to 37 &deg;C); mix the cell suspension with a Pasteur pipette. Add 0.5 ml hypotonic solution and mix again with a Pasteur pipette. Bring volume of hypotonic solution to 5 ml and mix once again. Incubate at 37 &deg;C for 20-45 min, depending on the cell type.</li>
 <li>Centrifuge at 400 x g for 10 min. Aspirate supernatant, leaving 0.5 ml of media plus the cell pellet.</li>
 <li>Resuspend the cell pellet by gently flicking the tube. The osmotically swollen cells will be fragile at this point, so care should be taken not to disrupt the cytoplasmic membranes.</li>
 <li>Add 3 drops of Carnoy's fixative (1:3 Glacial Acetic Acid: Methanol); mix by gently pipetting with a Pasteur pipette. Add 0.5 ml fixative and mix by gently pipetting with a Pasteur pipette. Bring volume of fixative to 5 ml and mix again by gently pipetting with a Pasteur pipette. Fixed cells can be stored in the dark at -20 &deg;C for several months.</li>
 <li>Centrifuge and aspirate as in step 2.8.</li>
 <li>Resuspend the cell pellet in fresh fixative; estimate the volume of the pellet and add ~10x the volume of Carnoy's fixative.</li>
 <li>Add the cell suspension drop-wise on wet, ice-cold microscope slides; hold the slide at an ~45&deg; angle and allow the cell suspension to flow over the surface of the slide. Lay the slide flat on paper towels and flood the slide with fixative and allow to air dry. Check slides for the presence of mitotic spreads using an inverted microscope.</li>
</ol>
3. RNase Treatment and Ethanol Dehydration
<ol>
 <li>Place 200 &mu;l of 10 &mu;g/ml RNase A (Sigma) in 2x SSC to each slide; incubate at 37 &deg;C for 1 hr.</li>
 <li>Wash slides 3 times with 2x SSC, pH 7.0 at room temperature for 3 min each.</li>
 <li>Dehydrate slides through an EtOH (Sigma) series (70%, 90% and 100%) at room temperature for 3 min each.</li>
 <li>Air dry the slides at room temperature.</li>
</ol>
4. Preparation of Probe Cocktails for BAC Plus Chromosome-specific Centromere Enumeration Probe (CEP) or for Whole Chromosome Paints In situ Hybridization
<ol>
 <li>For the BAC/CEP simultaneous hybridization, prepare two separate probe cocktails for pre-hybridization. We have found that pre-hybridizing the BAC and CEP probes separately under different stringencies results in less background and greater signal intensities. We have also found that individual Fosmids can also be used in place of BAC clones. The cocktail formulation described below represents the volumes for a single slide. If processing multiple slides using the same probes simply increase the volumes appropriately.
 *BAC DNA probe cocktail: 
 11.5 &mu;l hybridization buffer (50% Formamide (Sigma) /2x SSC /Dextran Sulfate (Sigma)) 2 &mu;l dH2O
 2.5 &mu;l BAC/Cot1 DNA in dH2O (see below for labeling protocol) 15 &mu;l total volume 
 *CEP probe cocktail: 
 7 &mu;l CEP hybridization buffer (65% Formamide/2x SSC/Dextran Sulfate) 2 &mu;l dH2O 
 1 &mu;l CEP/Cot1 DNA (purchased prelabeled with Spectrum Orange/Cy3/Texas Red) 10 &mu;l total volume 
 *After denaturing and pre-hybridization (step 5.1) mix BAC probe with CEP probe at a 3:2 ratio 25 &mu;l/slide and mix probe cocktails. 
 OR</li>
 <li>Aliquot 20 &mu;l/slide of Whole Chromosome Paint probe (Cy3/Texas Red) into a tube. Proceed to denaturation and pre-hybridization step.</li>
</ol>
5. Slide and Probe Denaturation and In situ Hybridization
<ol>
 <li>Denature probe cocktail(s) at 75 &deg;C for 10 min. Pre-hybridize at 37 &deg;C for 30 min to allow the Cot1 DNA to hybridize to repetitive sequences and to prevent subsequent hybridization to metaphase chromosomes.</li>
 <li>Denature slides in Coplin jars in 70% Formamide/2x SSC, pH 7.0 at 72 &deg;C for 3 min.</li>
 <li>Immediately dehydrate slides through an EtOH series (70%, 90% and 100%) in Coplin jars at 4 &deg;C for 3 min each.</li>
 <li>Allow the slides to air dry at room temperature.</li>
 <li>Pre-warm slides to 45 &deg;C on slide warmer for the last 10 min of the probe pre-hybridzation (step 5.1). Mix pre-hybridized probe cocktails if needed and add 25 ul to each slide; gently place a coverslip on each slide and seal along all edges with rubber cement. Incubate overnight at 37 &deg;C in a humidified chamber to prevent evaporation.</li>
</ol>
6. Post Hybridization Washes
<ol>
 <li>Wash slides in Coplin jars 3 times in 50% formamide/2x SSC, pH 7.0 at 38-40 &deg;C for 3 min each. The temperature of the washes may vary between probes. If there is high background hybridization you can increase the temperature of the washes. Conversely, if the signal is faint and there is no background then you can decrease the temperature of these washes.</li>
 <li>Wash slides 1 time in PN buffer at room temperature for 3 min, and proceed to the BrdU detection step.</li>
</ol>
7. BrdU Detection
<ol>
 <li>Block slides with PNM buffer for 10 min (200 &mu;l/slide) at room temperature in the dark.</li>
 <li>Drain slides by turning on edge on a paper towel. Add 100 &mu;l of anti-BrdU-FITC (50 &mu;g/ml Roche or Millipore) in PNM buffer for 30 min at 37 &deg;C.</li>
 <li>Wash slides 3 times in PN buffer at room temperature for 3 min each.</li>
 <li>Drain excess PN buffer on slides one at a time by turning on edge on a paper towel. Add 20 &mu;l DAPI/antifade mounting solution (Invitrogen). Cover the slide with a paper towel and press down on the coverslip to force out air bubbles and excess mounting solution. Proceed to image analysis.</li>
</ol>
8. Capturing Images and Quantifying BrdU Incorporation
<ol>
 <li>Images are captured with an Olympus BX61 Fluorescent Microscope and Olympus CCD Camera, 100x objective, automatic filter-wheel and Cytovision software (Applied Imaging). BrdU is captured using an FITC filter; chromosome paints (Texas Red, Metasystems or Cytocell), BAC's (Cy3) and CEP (Vysis Spectrum Orange) probes are captured using a Texas Red or Cy3 filter; DAPI is captured with a DAPI Filter (see Figures 2 and 3).</li>
 <li>Individual chromosomes of interest are identified with BAC/CEP or chromosome-specific paint probes (Figures 2B and 3B). Utilizing the Cytovision software, each chromosome of interest is "cut-out" from the metaphase spread as a whole, a line is drawn through the along the length of the chromosome from short arm to long arm (Figures 2C and 3C). The Cytovision software is used to quantify the pixel intensity profile along the length of each chromosome. The Cytovision software calculates the area occupied by the pixels and quantifies the intensity of the pixels represented by the BrdU or DAPI signals on each isolated chromosome. The average pixel intensity represented by each chromosome is then multiplied by the area occupied by those pixels to obtain the total number of pixels (Table 1 and Figure 4). In addition, this protocol allows for the visualization of the latest replicating regions of chromosomes. Accordingly, the banded pattern of BrdU incorporation allows for the detection of actively replicating regions of chromosomes, and differences in replication timing between chromosome pairs are seen as differences in this banding pattern (Figures 2 and 3). Finally, these differences in binding patterns can be compared to known replication timing maps for each chromosome6,7, which allows for an estimate of the replication timing difference between homologous chromosomes or chromosome arms4. For example, the two chromosome 6's in Figure 3A display a difference in banding pattern of &gt;2 hr when compared to the normal replication timing of chromosome 64. 
 In addition, a similar procedure to the one described here has been successfully used for the study of the asynchronous replication timing between the inactive and active X chromosomes8,9. The active and inactive X chromosomes were identified using X chromosome paints. Images were acquired using Quips mFISH software (Vysis) and the number of pixels occupied by the X chromosomes and the number of pixels occupied by fluorescently labeled BrdU were then calculated using NIH Image (<a href="http://rsb.info.nih.gov/nihimage" target="_blank">http://rsb.info.nih.gov/nih-image/index.html</a>).</li>
</ol>
9. Nick Translation of BAC DNA for Fluorescent Labeling (Vysis)
<ol>
 <li>Add the following probe mix to chilled a microfuge tube: 
 70 &mu;l (4 &mu;g DNA) in dH2O 
 10 &mu;l .2 mM dUTP-Orange or Green 
 20 &mu;l .1 mM dTTP 
 20 &mu;l 10x Nick Translation Buffer 
 40 &mu;l NT dNTP's (minus dTTP) 
 40 &mu;l nick Translation enzyme 
 200 &mu;l total volume 
 Incubate @ 16 &deg;C O/N. Stop reaction by heating at 70 &deg;C for 10 min. Chill on ice for 5 min.</li>
 <li>Precipitate the DNA with ethanol by adding the following to the probe mix: 
 200 &mu;l probe solution 
 48 &mu;l 3 M NaOAc 
 160 &mu;l Cot1DNA (.25 &mu;g/ul) 
 1,200 &mu;l 100% EtOH</li>
 <li>Store at -80 &deg;C for 10 min to O/N.</li>
 <li>Spin in cold at 12,000 x g for 20 min. Decant off EtOH. Wash 1 time with 70% EtOH. Spin as before. Decant off EtOH. Air dry just until EtOH evaporates. Resuspend in 40 &mu;l dH2O for final concentration of 100 ng/&mu;l of BAC DNA.</li>
</ol>

<ol>
	<li>Payer, B., Lee, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=X+chromosome+dosage+compensation:+how+mammals+keep+the+balance.">X chromosome dosage compensation: how mammals keep the balance.</a> Annu. Rev. Genet. 42, 733-772 (2008).</li><li>Breger, K. S., Smith, L., Turker, M. S., Thayer, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ionizing+radiation+induces+frequent+translocations+with+delayed+replication+and+condensation.">Ionizing radiation induces frequent translocations with delayed replication and condensation.</a> Cancer Research. 64, 8231-8238 (2004).</li><li>Smith, L., Plug, A., Thayer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Delayed+Replication+Timing+Leads+to+Delayed+Mitotic+Chromosome+Condensation+and+Chromosomal+Instability+of+Chromosome+Translocations.">Delayed Replication Timing Leads to Delayed Mitotic Chromosome Condensation and Chromosomal Instability of Chromosome Translocations.</a> Proc. Natl. Acad. Sci. U.S.A. 98, 13300-13305 (2001).</li><li>Stoffregen, E. P., Donley, N., Stauffer, D., Smith, L., Thayer, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+autosomal+locus+that+controls+chromosome-wide+replication+timing+and+mono-allelicexpression.">An autosomal locus that controls chromosome-wide replication timing and mono-allelicexpression.</a> Hum. Mol. Genet. 20, 2366-2378 (2011).</li><li>Breger, K. S., Smith, L., Thayer, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Engineering+translocations+with+delayed+replication:+evidence+for+cis+control+of+chromosome+replication+timing.">Engineering translocations with delayed replication: evidence for cis control of chromosome replication timing.</a> Hum. Mol. Genet. 14, 2813-2827 (2005).</li><li>Camargo, M., Cervenka, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterns+of+DNA+replication+of+human+chromosomes.+II.+Replication+map+and+replication+model.">Patterns of DNA replication of human chromosomes. II. Replication map and replication model.</a> Am. J. Hum. Genet. 34, 757-780 (1982).</li><li>Cohen, S. M., Cobb, E. R., Cordeiro-Stone, M., Kaufman, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+chromosomal+bands+replicating+early+in+the+S+phase+of+normal+human+fibroblasts.">Identification of chromosomal bands replicating early in the S phase of normal human fibroblasts.</a> Exp. Cell Res. 245 (98), 321-329 (1998).</li><li>Diaz-Perez, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+element(s)+at+the+nontranscribed+Xist+locus+of+the+active+X+chromosome+controls+chromosomal+replication+timing+in+the+mouse.">The element(s) at the nontranscribed Xist locus of the active X chromosome controls chromosomal replication timing in the mouse.</a> Genetics. 171, 663-672 (2005).</li><li>Diaz-Perez, S. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+deletion+at+the+mouse+Xist+gene+exposes+trans-effects+that+alter+the+heterochromatin+of+the+inactive+X+chromosome+and+the+replication+time+and+DNA+stability+of+both+X+chromosomes.">A deletion at the mouse Xist gene exposes trans-effects that alter the heterochromatin of the inactive X chromosome and the replication time and DNA stability of both X chromosomes.</a> Genetics. 174, 1115-1133 (2006).</li><li>Salic, A., Mitchison, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+chemical+method+for+fast+and+sensitive+detection+of+DNA+synthesis+in+vivo.">A chemical method for fast and sensitive detection of DNA synthesis in vivo.</a> Proc. Natl. Acad. Sci. U.S.A. 105, 2415-2420 (2008).</li><li>Schlesinger, S., Selig, S., Bergman, Y., Cedar, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Allelic+inactivation+of+rDNA+loci.">Allelic inactivation of rDNA loci.</a> Genes Dev. 23, 2437-2447 (2009).</li><li>Branzei, D., Foiani, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+checkpoint+response+to+replication+stress.">The checkpoint response to replication stress.</a> DNA Repair (Amst). 8, 1038-1046 (2009).</li></ol>

No conflicts of interest declared.

The preparation of chromosome spreads is a critical step for the successful replication-timing assay described here. The inclusion of a colcemid pretreatment step prior to the hypotonic treatment may aid in the frequency and spreading of mitotic cells. We typically expose cells to colcemdi for 1-3 hr prior to harvest, and use colcemid at a final concentration of 10 ug/ml. However, the inclusion of a colcemid pretreatment step may alter the length of G2 and consequently may alter the apparent replication timing and state ...

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Anti-BrdU-FITC</td>
 <td>Roche Millipore</td>
 <td>11202593001 MAB326F</td>
 <td>50 &mu;g/&mu;l</td>
 </tr>
 <tr>
 <td>Nick Translation Kit</td>
 <td>Abbott Molecular (Vysis)</td>
 <td>07J00-0001</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Spectrum Orange dUTP</td>
 <td>Abbott Molecular (Vysis)</td>
 <td>02N33-050</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CEP</td>
 <td>Abbott Molecular (Vysis)</td>
 <td>Varies</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>LSI/WCP hybridization buffer</td>
 <td>Abbott Molecular (Vysis)</td>
 <td>06J67-011</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>CEP hybridization buffer</td>
 <td>Abbott Molecular (Vysis)</td>
 <td>07J36-001</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Chromosome paints</td>
 <td>MetaSystems Group</td>
 <td>D-14NN-050-TR</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Olympus BX61 Fluorescent Microscope</td>
 <td>Olympus</td>
 <td>BX61TRF-1-5</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microscope imaging software system</td>
 <td>Applied Imaging</td>
 <td>Cytovision 3.93.1</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Digital Camera</td>
 <td>Olympus</td>
 <td>UCMAD3</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>
 IN SITU HYBRIDIZATION RECIPES 
 Formamide Solutions 
 70% Formamide/2x SSC
 35 ml Formamide* (Sigma) 
 10 ml 10x SSC 
 5 ml d2H20 
 pH to 7.0 with HCl (Sigma)
 * It is important to use formamide that has been stored at -20 &deg;C. Prolonged room temperature storage will generate formic acid and the pH will be too low.
 50% Formamide/2x SSC
 25 ml formamide (Sigma) 
 10 ml 10x SSC 
 15 ml dH2O 
 pH to 7.0 with HCl (Sigma)
 20x SSC, 4 L
 702 g NaCl (Sigma) 
 358 g Na Citrate (Sigma) 
 dH2O to volume
 PN Buffer [0.1 M NaP04 0.1% NP_40 (Sigma)]
 Make a 0.1 M solution each of sodium phosphate (Filter sterilize and store in 500 ml aliquots).
 0.1 M NaH2P04 , 1 L
 13.8 g NaH2P04 (Sigma): 
 dH2O to volume
 0.1 M NaH2P04 1 L
 14.2 g NaH2P04 (Sigma) 
 dH2O to volume.
 PN: Adjust pH of 0.1 M Na2HP04 to pH 8.0 with .1 M NaH2P04. Filter sterilize and add 1 ml of NP-40.
 PNM 50 ml
 1.25 g Non-fat dry milk (Sigma) 
 25 ml PN buffer (Recipe above)
 Mix for 15-20 min with constant stirring. Spin 2 times at 400 x g for 10 min. Use supernatant, and make sure not to disturb precipitated milk proteins.</td>
 </tr>
 </tbody>
</table>

chromosome replicating timing combined with fluorescent in situ hybridization

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The specification of replication timing is thought to be a dynamic process regulated by tissue-specific and developmental cues that are responsive to epigenetic modifications. However, the mechanisms regulating where and when DNA replication initiates along chromosomes remains poorly understood. Homologous chromosomes usually replicate synchronously, however there are notable exceptions to this rule. For example, in female mammalian cells one of the two X chromosomes becomes late replicating through a process known as X inactivation1. Along with this delay in replication timing, estimated to be 2-3 hr, the majority of genes become transcriptionally silenced on one X chromosome. In addition, a discrete cis-acting locus, known as the X inactivation center, regulates this X inactivation process, including the induction of delayed replication timing on the entire inactive X chromosome. In addition, certain chromosome rearrangements found in cancer cells and in cells exposed to ionizing radiation display a significant delay in replication timing of &gt;3 hours that affects the entire chromosome2,3. Recent work from our lab indicates that disruption of discrete cis-acting autosomal loci result in an extremely late replicating phenotype that affects the entire chromosome4. Additional 'chromosome engineering' studies indicate that certain chromosome rearrangements affecting many different chromosomes result in this abnormal replication-timing phenotype, suggesting that all mammalian chromosomes contain discrete cis-acting loci that control proper replication timing of individual chromosomes5.
Here, we present a method for the quantitative analysis of chromosome replication timing combined with fluorescent in situ hybridization. This method allows for a direct comparison of replication timing between homologous chromosomes within the same cell, and was adapted from6. In addition, this method allows for the unambiguous identification of chromosomal rearrangements that correlate with changes in replication timing that affect the entire chromosome. This method has advantages over recently developed high throughput micro-array or sequencing protocols that cannot distinguish between homologous alleles present on rearranged and un-rearranged chromosomes. In addition, because the method described here evaluates single cells, it can detect changes in chromosome replication timing on chromosomal rearrangements that are present in only a fraction of the cells in a population.

An example of the replication timing analysis for human chromosome 6 is shown in Figure 2. Cells containing a deletion of the ASAR6 gene4, located at 6q16.1, were exposed to BrdU for 5 hr, harvested for mitotic cells and processed for FISH with a chromosome 6 paint probe (Vysis) and for BrdU incorporation. Note that there is a significant difference in the BrdU banding pattern between the two 6's, which is consistent with a delay in replication timing of &gt;2 hr for one of the chromosome 6's ...

Watch this Scientific Journal Video about Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization at JoVE.com

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

A quantitative method for the analysis of chromosome replication timing is described. The method utilizes BrdU incorporation in combination with fluorescent in situ hybridization (FISH) to assess replication timing of mammalian chromosomes. This technique allows for the direct comparison of rearranged and un-rearranged chromosomes within the same cell.

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The ...

chromosome-replicating-timing-combined-with-fluorescent-situ

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13.9K 次观看.  Oregon Health & Science University. 染色体复制定时分析的定量方法进行说明。的方法，利用结合BrdU掺入荧光

视频: Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Fluorescence in situ hybridization (FISH) Protocol in Human Sperm

Aneuploidies are the most frequent chromosomal abnormalities in humans. Most of these abnormalities result from meiotic errors during the gametogenic process in the parents. In human males, these errors can lead to the production of spermatozoa with numerical chromosome abnormalities which represent an increased risk of transmitting these anomalies to the offspring.
For this reason, the technique of fluorescence in situ hybridization (FISH) on sperm nuclei has become a protocol widely incorporated in the context of clinical diagnosis. This practice provides an estimate of the frequencies of numerical chromosome abnormalities in the gametes of the patients that seek for genetic reproductive advice.
To date, the chromosomes most frequently included in sperm FISH analysis are chromosomes X, Y, 13, 18 and 21.
This video-article describes, step by step, how to process and fix a human semen sample, how to decondense and denature the sperm chromatin, how to proceed to obtain sperm FISH preparations, and how to visualize the results at the microscope. Special remarks of the most relevant steps are given to achieve the best results.

Fluorescence in situ hybridization FISH Protocol in Human Sperm

This video-article describes, step by step, how to process a semen sample to achieve good-quality fluorescence in situ hybridization on human spermatozoa. Preparations obtained can be used for aneuploidy screening in the context of clinical diagnosis.

Aneuploidies are the most frequent chromosomal abnormalities in humans. Most of these abnormalities result from meiotic errors during the gametogenic ...

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of specific mRNA transcripts within whole mount specimen. This technique (adapted from Albertson and Yelick, 2005) was used in an upper level undergraduate Comparative Vertebrate Biology laboratory classroom at Syracuse University. The first two thirds of the Comparative Vertebrate Biology lab course gave students the opportunity to study the embryology and gross anatomy of several organisms representing various chordate taxa primarily via traditional dissections and the use of models. The final portion of the course involved an innovative approach to teaching anatomy through observation of vertebrate development employing molecular techniques in which WISH was performed on zebrafish embryos. A heterozygous fibroblast growth factor 8 a (fgf8a) mutant line, ace, was used. Due to Mendelian inheritance, ace intercrosses produced wild type, heterozygous, and homozygous ace/fgf8a mutants in a 1:2:1 ratio. RNA probes with known expression patterns in the midline and in developing anatomical structures such as the heart, somites, tailbud, myotome, and brain were used. WISH was performed using zebrafish at the 13 somite and prim-6 stages, with students performing the staining reaction in class. The study of zebrafish embryos at different stages of development gave students the ability to observe how these anatomical structures changed over ontogeny. In addition, some ace/fgf8a mutants displayed improper heart looping, and defects in somite and brain development. The students in this lab observed the normal development of various organ systems using both external anatomy as well as gene expression patterns. They also identified and described embryos displaying improper anatomical development and gene expression (i.e., putative mutants).
For instructors at institutions that do not already own the necessary equipment or where funds for lab and curricular innovation are limited, the financial cost of the reagents and apparatus may be a factor to consider, as will the time and effort required on the part of the instructor regardless of the setting. Nevertheless, we contend that the use of WISH in this type of classroom laboratory setting can provide an important link between developmental genetics and anatomy. As technology advances and the ability to study organismal development at the molecular level becomes easier, cheaper, and increasingly popular, many evolutionary biologists, ecologists, and physiologists are turning to research strategies in the field of molecular biology. Using WISH in a Comparative Vertebrate Biology laboratory classroom is one example of how molecules and anatomy can converge within a single course. This gives upper level college students the opportunity to practice modern biological research techniques, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of ...

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures are lengthy and technically demanding with multiple important steps that collectively contribute to the quality of the final result. This protocol describes in detail several key quality control steps for optimizing probe labeling and performance. Overall, our protocol provides a detailed description of the critical steps necessary to reproducibly obtain high quality results. First, we describe the generation of digoxygenin (DIG) labeled RNA probes via in vitro transcription of DNA templates generated by PCR. We describe three critical quality control assays to determine the amount, integrity and specific activity of the DIG-labeled probes. These steps are important for generating a probe of sufficient sensitivity to detect endogenous mRNAs in a whole mouse embryo. In addition, we describe methods for the fixation and storage of E8.5-E11.5 day old mouse embryos for in situ hybridization. Then, we describe detailed methods for limited proteinase K digestion of the rehydrated embryos followed by the details of the hybridization conditions, post-hybridization washes and RNase treatment to remove non-specific probe hybridization. An AP-conjugated antibody is used to visualize the labeled probe and reveal the expression pattern of the endogenous transcript. Representative results are shown from successful experiments and typical suboptimal experiments.

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

This whole mount in situ hybridization protocol discusses critical steps that ensure reproducible high quality results for gene expression studies in E8.5-E11.5 day old mouse embryos.

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures ...

In Situ Hybridization for the Precise Localization of Transcripts in Plants

With the advances in genomics research of the past decade, plant biology has seen numerous studies presenting large-scale quantitative analyses of gene expression. Microarray and next generation sequencing approaches are being used to investigate developmental, physiological and stress response processes, dissect epigenetic and small RNA pathways, and build large gene regulatory networks1-3. While these techniques facilitate the simultaneous analysis of large gene sets, they typically provide a very limited spatiotemporal resolution of gene expression changes. This limitation can be partially overcome by using either profiling method in conjunction with lasermicrodissection or fluorescence-activated cell sorting4-7. However, to fully understand the biological role of a gene, knowledge of its spatiotemporal pattern of expression at a cellular resolution is essential. Particularly, when studying development or the effects of environmental stimuli and mutants can the detailed analysis of a gene's expression pattern become essential. For instance, subtle quantitative differences in the expression levels of key regulatory genes can lead to dramatic phenotypes when associated with the loss or gain of expression in specific cell types. 
Several methods are routinely used for the detailed examination of gene expression patterns. One is through analysis of transgenic reporter lines. Such analysis can, however, become time-consuming when analyzing multiple genes or working in plants recalcitrant to transformation. Moreover, an independent validation to ensure that the transgene expression pattern mimics that of the endogenous gene is typically required. Immunohistochemical protein localization or mRNA in situ hybridization present relatively fast alternatives for the direct visualization of gene expression within cells and tissues. The latter has the distinct advantage that it can be readily used on any gene of interest. In situ hybridization allows detection of target mRNAs in cells by hybridization with a labeled anti-sense RNA probe obtained by in vitro transcription of the gene of interest. 
Here we outline a protocol for the in situ localization of gene expression in plants that is highly sensitivity and specific. It is optimized for use with paraformaldehyde fixed, paraffin-embedded sections, which give excellent preservation of histology, and DIG-labeled probes that are visualized by immuno-detection and alkaline-phosphatase colorimetric reaction. This protocol has been successfully applied to a number of tissues from a wide range of plant species, and can be used to analyze expression of mRNAs as well as small RNAs8-14.

In Situ Hybridization for the Precise Localization of Transcripts in Plants

The in situ hybridization protocol described here allows a direct localization of mRNA and small RNA expression at the cellular level with high sensitivity and specificity. The procedure is optimized for paraffin-embedded plant tissue sections, is applicable to a wide range of plants and tissues, and can be completed within ten days.

With the advances in genomics research of the past decade, plant biology has seen numerous studies presenting large-scale quantitative analyses of gene ...

Detection of Viral RNA by Fluorescence in situ Hybridization (FISH)

Viruses that infect cells elicit specific changes to normal cell functions which serve to divert energy and resources for viral replication. Many aspects of host cell function are commandeered by viruses, usually by the expression of viral gene products that recruit host cell proteins and machineries. Moreover, viruses engineer specific membrane organelles or tag on to mobile vesicles and motor proteins to target regions of the cell (during de novo infection, viruses co-opt molecular motor proteins to target the nucleus; later, during virus assembly, they will hijack cellular machineries that will help in the assembly of viruses). Less is understood on how viruses, in particular those with RNA genomes, coordinate the intracellular trafficking of both protein and RNA components and how they achieve assembly of infectious particles at specific loci in the cell. The study of RNA localization began in earlier work. Developing lower eukaryotic embryos and neuronal cells provided important biological information, and also underscored the importance of RNA localization in the programming of gene expression cascades. The study in other organisms and cell systems has yielded similar important information. Viruses are obligate parasites and must utilise their host cells to replicate. Thus, it is critical to understand how RNA viruses direct their RNA genomes from the nucleus, through the nuclear pore, through the cytoplasm and on to one of its final destinations, into progeny virus particles 1.
FISH serves as a useful tool to identify changes in steady-state localization of viral RNA. When combined with immunofluorescence (IF) analysis 22, FISH/IF co-analyses will provide information on the co-localization of proteins with the viral RNA3. This analysis therefore provides a good starting point to test for RNA-protein interactions by other biochemical or biophysical tests 4,5, since co-localization by itself is not enough evidence to be certain of an interaction. In studying viral RNA localization using a method like this, abundant information has been gained on both viral and cellular RNA trafficking events 6. For instance, HIV-1 produces RNA in the nucleus of infected cells but the RNA is only translated in the cytoplasm. When one key viral protein is missing (Rev) 7, FISH of the viral RNA has revealed that the block to viral replication is due to the retention of the HIV-1 genomic RNA in the nucleus 8. 
Here, we present the method for visual analysis of viral genomic RNA in situ. The method makes use of a labelled RNA probe. This probe is designed to be complementary to the viral genomic RNA. During the in vitro synthesis of the antisense RNA probe, the ribonucleotide that is modified with digoxigenin (DIG) is included in an in vitro transcription reaction. Once the probe has hybridized to the target mRNA in cells, subsequent antibody labelling steps (Figure 1) will reveal the localization of the mRNA as well as proteins of interest when performing FISH/IF.

Detection of Viral RNA by Fluorescence in situ Hybridization FISH

A fluorescence in situ hybridization (FISH) method was developed to visually detect viral genomic RNA using fluorescence microscopy. A probe is made with specificity to the viral RNA that can then be identified using a combination of hybridization and immunofluorescence techniques. This technique offers the advantage of identifying the localization of the viral RNA or DNA at steady-state, providing information on the control of intracellular virus trafficking events.

Viruses  that infect cells elicit specific changes to normal cell functions which serve  to divert energy and resources for viral replication. Many ...

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place over the next 5 years. For example, the sequencing of the genomes for 15 malaria mosquitospecies is currently being done using an Illumina platform3,4. This Anopheles species cluster includes both vectors and non-vectors of malaria. When the genome assemblies become available, researchers will have the unique opportunity to perform comparative analysis for inferring evolutionary changes relevant to vector ability. However, it has proven difficult to use next-generation sequencing reads to generate high-quality de novo genome assemblies5. Moreover, the existing genome assemblies for Anopheles gambiae, although obtained using the Sanger method, are gapped or fragmented4,6.
Success of comparative genomic analyses will be limited if researchers deal with numerous sequencing contigs, rather than with chromosome-based genome assemblies. Fragmented, unmapped sequences create problems for genomic analyses because: (i) unidentified gaps cause incorrect or incomplete annotation of genomic sequences; (ii) unmapped sequences lead to confusion between paralogous genes and genes from different haplotypes; and (iii) the lack of chromosome assignment and orientation of the sequencing contigs does not allow for reconstructing rearrangement phylogeny and studying chromosome evolution. Developing high-resolution physical maps for species with newly sequenced genomes is a timely and cost-effective investment that will facilitate genome annotation, evolutionary analysis, and re-sequencing of individual genomes from natural populations7,8.
Here, we present innovative approaches to chromosome preparation, fluorescent in situ hybridization (FISH), and imaging that facilitate rapid development of physical maps. Using An. gambiae as an example, we demonstrate that the development of physical chromosome maps can potentially improve genome assemblies and, thus, the quality of genomic analyses. First, we use a high-pressure method to prepare polytene chromosome spreads. This method, originally developed for Drosophila9, allows the user to visualize more details on chromosomes than the regular squashing technique10. Second, a fully automated, front-end system for FISH is used for high-throughput physical genome mapping. The automated slide staining system runs multiple assays simultaneously and dramatically reduces hands-on time11. Third, an automatic fluorescent imaging system, which includes a motorized slide stage, automatically scans and photographs labeled chromosomes after FISH12. This system is especially useful for identifying and visualizing multiple chromosomal plates on the same slide. In addition, the scanning process captures a more uniform FISH result. Overall, the automated high-throughput physical mapping protocol is more efficient than a standard manual protocol.

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Genome assemblies based on massively parallel DNA sequencing technologies are usually highly fragmented. The development of physical chromosome maps can potentially improve genome assemblies. Here, we demonstrate innovative approaches to chromosome preparation, fluorescent in situ hybridization, and imaging that significantly increase throughput of the physical map development.

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place ...

Analysis of Single-cell Gene Transcription by RNA Fluorescent In Situ Hybridization (FISH)

Adhesion of Plasmodium falciparum infected erythrocytes (IE) to human endothelial receptors during malaria infections is mediated by expression of PfEMP1 protein variants encoded by the var genes.
The haploid P. falciparum genome harbors approximately 60 different var genes of which only one has been believed to be transcribed per cell at a time during the blood stage of the infection. How such mutually exclusive regulation of var gene transcription is achieved is unclear, as is the identification of individual var genes or sub-groups of var genes associated with different receptors and the consequence of differential binding on the clinical outcome of P. falciparum infections. Recently, the mutually exclusive transcription paradigm has been called into doubt by transcription assays based on individual P. falciparum transcript identification in single infected erythrocytic cells using RNA fluorescent in situ hybridization (FISH) analysis of var gene transcription by the parasite in individual nuclei of P. falciparum IE1.
Here, we present a detailed protocol for carrying out the RNA-FISH methodology for analysis of var gene transcription in single-nuclei of P. falciparum infected human erythrocytes. The method is based on the use of digoxigenin- and biotin- labeled antisense RNA probes using the TSA Plus Fluorescence Palette System2 (Perkin Elmer), microscopic analyses and freshly selected P. falciparum IE. The in situ hybridization method can be used to monitor transcription and regulation of a variety of genes expressed during the different stages of the P. falciparum life cycle and is adaptable to other malaria parasite species and other organisms and cell types.

Analysis of Single-cell Gene Transcription by RNA Fluorescent In Situ Hybridization FISH

Fluorescent in situ hybridization (FISH) to identify mRNA transcripts in individual cells allows analysis of polygenic activity such as the simultaneous transcription of more than one member of the var multigene family in Plasmodium falciparum infected erythrocytes 1. The technique is adaptable and can be used on different types of genes, cells and organisms.

Adhesion of Plasmodium falciparum infected erythrocytes (IE) to human endothelial receptors during malaria infections is mediated by expression of PfEMP1 ...

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Assessing the expression pattern of a gene, as well as the subcellular localization properties of its transcribed RNA, are key features for understanding its biological function during development. RNA in situ hybridization (RNA-ISH) is a powerful method used for visualizing RNA distribution properties, be it at the organismal, cellular or subcellular levels 1. RNA-ISH is based on the hybridization of a labeled nucleic acid probe (e.g. antisense RNA, oligonucleotides) complementary to the sequence of an mRNA or a non-coding RNA target of interest 2. As the procedure requires primary sequence information alone to generate sequence-specific probes, it can be universally applied to a broad range of organisms and tissue specimens 3. Indeed, a number of large-scale ISH studies have been implemented to document gene expression and RNA localization dynamics in various model organisms, which has led to the establishment of important community resources 4-11. While a variety of probe labeling and detection strategies have been developed over the years, the combined usage of fluorescently-labeled detection reagents and enzymatic signal amplification steps offer significant enhancements in the sensitivity and resolution of the procedure 12. Here, we describe an optimized fluorescent in situ hybridization method (FISH) employing tyramide signal amplification (TSA) to visualize RNA expression and localization dynamics in staged Drosophila embryos. The procedure is carried out in 96-well PCR plate format, which greatly facilitates the simultaneous processing of large numbers of samples.

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Here we describe a whole-mount fluorescent in situ hybridization (FISH) protocol for determining the expression and localization properties of RNAs expressed during embryogenesis in the fruit fly, Drosophila melanogaster.