We thank Zhaohua Irene Tang in the W. M. Keck Science Department for the use of her epifluorescence microscope and the Werren lab for donating Nasonia for dissections. This work was supported in part by an NIH-NRSA fellowship (5F32GM105317-02) to AML.

1. Tissue Dissection and Fixation (60 min)
<ol>
	<li>For Drosophila brains, place 3rd instar larvae in a drop of 1x PBS (phosphate buffered saline). Choose large 3rd instar larvae that are actively crawling from vials or bottles that are not overcrowded.
	<ol>
		<li>Use one ultrafine tweezer pair to grab hold of the mouth hooks and another tweezer pair to grab 2/3 down the length of the body (Figure 1A, B). Pull gently on the mouth hooks to expose the brain, ventral ganglia, salivary glands and part of the larval digestive tract. Use the tweezers to separate the brain and ventral ganglia (Figure 1A, B) from the other tissues and place in a droplet of 1x PBT (phosphate buffered saline with Tween) on a plastic Petri plate.</li>
	</ol>
	</li>
	<li>For Nasonia testis dissections, choose male 3-day-old pupae (yellow bodies with red eyes). Male Nasonia have small wing pad lengths relative to females during the pupal stage (Figure 1C).
	<ol>
		<li>Hold the pupa at the top of the abdomen near the thoracic region with one tweezer pair, and using the other tweezer pair, grab the very distal tip of the abdomen and pull out the tear-drop-shaped testes (they will be surrounded by fat body that can be gently shaken away; Figure 1D). Detach any exterior body parts from the testes, which will prevent proper squashing, and place in a droplet of 1x PBT on a plastic Petri plate.</li>
	</ol>
	</li>
	<li>For each slide, dissect a total of four or five tissue samples (i.e., Drosophila larval brains or Nasonia testes). 
	NOTE: More than five samples will lead to overcrowded nuclei and chromosomes.</li>
	<li>Optionally, to achieve some separation of sister chromatids in euchromatic regions of mitotic chromosomes, treat the tissue with a hypotonic solution: transfer brains from 1x PBT to a drop of 0.5% sodium citrate for 5-10 (no more than 10) min. 
	NOTE: Colchicine (a mitotic inhibitor) can be helpful for increasing the number of mitotic figures at metaphase11. However, its use is unnecessary if very large healthy larvae are used, and even undesirable because it can negatively affect the chromosome resolution, spreading and morphology.</li>
	<li>Place a droplet (~20 &#181;l) of fixative solution (2.5% paraformaldehyde in 45% acetic acid) onto the surface of a clean Sigmacote-treated cover slip. 
	NOTE: Make fixative solution fresh for each day of use. Fixative solutions ranging from 1.8-3.7% paraformaldehyde in 45% acetic acid yield the best results for FISH. Using tissues other than brains or testes, or adapting this protocol for immuno-FISH may require experimenting with different fixatives (for a list of different fixatives, see12).</li>
	<li>Carefully transfer each tissue sample from the dissecting buffer (1x PBT) into the fixative droplet with ultrafine tweezers, minimizing the transfer of dissecting buffer into the fixative solution. Position the tissue samples so that they are evenly spaced from one another within the fixative droplet. Incubate the tissues in fixative for 4 min at room temperature.</li>
	<li>Carefully place a poly-lysine-coated slide face down onto the tissue and cover slip. Do not press down at this point but instead allow the two to contact lightly so that the cover slip sticks onto the underside of the slide. Invert the slide so that the cover slip is on top.</li>
	<li>Sandwich the slide/tissue/cover slip inside a folded piece of filter paper. On a stable surface, using the thumb, press very firmly straight down onto the position directly above the cover slip. Be careful to avoid lateral sliding of the cover slip (this will cause smearing of the tissue).</li>
	<li>Submerge the slide/tissue/cover slip into liquid nitrogen (see apparatus in Figure 1E, F), and let stand until the Nitrogen stops boiling (longer is fine). Remove the slide and immediately snap off the cover slip with a fresh razor blade by flicking a corner of the cover slip in an upward direction (avoid scratching the fixed tissue with the razor blade).
	<ol>
		<li>Pre-cooling the slide/tissue/cover slip on a block of dry ice, cover slip up, for 1-2 min prior to submerging in liquid nitrogen will help prevent the slides from cracking.</li>
	</ol>
	</li>
	<li>Immediately place the slide with tissue into a Coplin jar filled with 100% ethanol at room temperature and let stand for at least 5 min (this time can be longer if needed). 
	NOTE: Cold 100% ethanol could also be used.</li>
	<li>Remove the slide with tissue, wick away excess ethanol with a Kimwipe (without touching the fixed tissue), and leave the slide to air dry for 1 hr.
	<ol>
		<li>Proceed directly to step 2 or keep slides dry in low humidity air or in a desiccation chamber for weeks to even months before performing the hybridization.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/52288/52288fig1highres.jpg" /> 
Figure 1: (A) A 3rd instar Drosophila larva (right), with positions indicated for where to grab the mouth hooks (denoted with *) and 2/3 of the way down the larvae to dissect brains; (left) a schematic of a larva a the same developmental stage, depicting the relative position of the brain within the larval head. (B) The brain and ventral ganglia dissected from a 3rd instar Drosophila larva (right) and a schematic of this tissue (left). (C) 3-day-old Nasonia pupa at the yellow body-red eye stage. (D) A testis pair dissected from a 3 day old Nasonia male pupa (right) with positions indicating where to grab the pupa at the posterior of the abdomen (denoted with *) and midway on the body; (left) schematic depicting male and female wasp pupae; male pupae can be distinguished by wings that do not extend past the saggital profile (black arrow), in contrast to females, which have wings that extend past the profile; the relative position of the testis pair is shown in the male pupa. (E) The apparatus&#8212;a string of paperclips&#8212;and (F) method used to immerse slides in a vessel of liquid nitrogen. Multiple paperclip strings can be used for simultaneous immersion of multiple slides.
2. In situ Hybridization (30 min on day 1; 1 hr&#8212;2.5 hr for long probes&#8212;on day 2)
<ol>
	<li>Add 1 &#181;l (100 ng) of each probe to 20 &#181;l of 1.1x hybridization buffer. Pipet probe/hybridization buffer onto the surface of the fixed tissue (avoid touching the tissue).</li>
	<li>Carefully place a cover slip directly onto the probe/hybridization buffer, making sure that the cover slip is centered directly over the tissue. The buffer should migrate to the outer edge of the cover slip, leaving no air bubbles. 
	NOTE: Small bubbles that do not contact tissue do not pose any problems to the procedure. Remove large bubbles by carefully lifting a corner of the cover slip and carefully dropping it back onto the slide.</li>
	<li>Place the slide/tissue/cover slip onto the surface of a block pre-heated to 95 &#176;C (cover slip up). Cover with a large piece of aluminum foil to prevent light exposure. Let the slide incubate at 95 &#176;C for 5 min. 
	NOTE: A typical heat block with holes for tubes can be flipped over to provide a flat surface on which to place the slide/tissue/cover slip.</li>
	<li>Remove the slide, letting it cool slightly until it is warm to the touch. Carefully wrap a piece of stretched Parafilm around the cover slip to seal the liquid beneath it.</li>
	<li>Place the sealed slide inside a humidity chamber and place the chamber into an incubator preheated to 30 &#176;C. Incubate at 30 &#176;C for 4 hr to overnight.
	<ol>
		<li>Create a humidity chamber from an empty tip box or lidded Tupperware container with dampened Kimwipes or paper towels placed at the bottom. 
		NOTE: Single stranded DNA oligo probes have been designed to be 28-33 bases to achieve a theoretical melting temperature (Tm) of 45-47 &#176;C. These length and Tm ranges reflect the fact that many repetitive sequences that we have studied are AT-rich and therefore have very low GC content. Longer probes will likely have higher Tm values; this may result in higher background hybridization at the standard hybridization temperature of 30 &#176;C. Thus, some troubleshooting with hybridization temperatures may be required to achieve the best results. To find the best hybridization temperature, increase (or decrease) the temperature by 5 &#176;C, incrementally.</li>
	</ol>
	</li>
	<li>Carefully remove the Parafilm from the slide and then carefully remove the cover slip by slowly lifting one corner. Wash the slide three times for 15 min each wash in 0.1x SSC buffer. Cover the Coplin jar with aluminum foil during the washes to minimize light exposure.</li>
	<li>If not using a long biotinylated probe, proceed to step 2.8.
	<ol>
		<li>If using a long biotinylated probe, dry the area around the tissue with a Kimwipe, being careful not to touch the tissue itself. Place 100 &#181;l of blocking solution over the tissue and cover gently with a coverslip, taking care to avoid trapping bubbles. Wrap the slide over the coverslip with Parafilm and place at 37 &#176;C for 30 min.</li>
		<li>Carefully remove the coverslip and blot around the tissue with a Kimwipe. Pipet 100 &#181;l of rhodamine-avidin diluted 1:1,000 in SBT over the tissue and cover gently with a coverslip, taking care to avoid trapping bubbles. Wrap the slide over the coverslip with Parafilm and place at 37 &#176;C for 30 min.</li>
		<li>Carefully remove the coverslip and wash the slide 3 times for 5 min each in 4x SSCT and then 3 times for 5 min each in 0.1x SSC. 
		NOTE: Slides can be washed for longer periods of time.</li>
	</ol>
	</li>
	<li>Remove the slide and blot around the tissue with a dry Kimwipe to remove excess buffer (avoid touching the tissue). Place the slide tissue side up in a dark place for 10-15 min or until the moisture completely dissipates.</li>
	<li>Pipet 11 &#181;l of Vectashield mounting medium (with 4&#8217;,6-diamidino-2-phenylindole&#8212;DAPI) onto the tissue. Carefully place a clean cover slip (not treated with Sigmacote) directly over the center of the mounting medium and tissue. The mounting medium should migrate slowly outward toward the edges of the cover slip. 
	NOTE: If the mounting medium fails to reach the edge of the cover slip on all sides, then an additional 1-2 &#181;l of mounting medium can be applied to one position at the edge of the cover slip to fill in the needed volume. In this case, be sure to wipe away any excess medium from the slide surface before sealing.</li>
	<li>Seal the edges of the cover slip with nail polish. Avoid painting the nail polish over the tissue sample.</li>
	<li>Place the slide upright in a dark place and let the nail polish dry until completely hard (usually 30 min or more). At this point, image the tissue or store at -20 &#176;C for up to 1 week for later imaging.</li>
</ol>
Buffer/Solution Recipes
10x PBS
<ul>
	<li>80 g NaCl</li>
	<li>2.0 g KCl</li>
	<li>14.4 g Na2HPO4</li>
	<li>2.4 g KH2PO4</li>
	<li>pH to 7.4, H2O to 1 L</li>
</ul>
1x PBT
<ul>
	<li>5 ml 10x PBS</li>
	<li>45 ml H2O</li>
	<li>0.1% Tween 20</li>
</ul>
20x SSC
<ul>
	<li>175.3 g NaCl</li>
	<li>88.2 g Na Citrate</li>
	<li>in 800 ml H2O</li>
	<li>pH to 7, H2O to 1 L</li>
</ul>
4x SSCT
<ul>
	<li>200 ml 20x SSC</li>
	<li>799 ml H2O</li>
	<li>0.1% Tween 20</li>
</ul>
0.1x SSC
<ul>
	<li>5 ml 20x SSC</li>
	<li>995 ml H2O</li>
</ul>
Hybridization mix (20 &#181;l; modified from 11)
<ul>
	<li>10 &#181;l formamide&#160;</li>
	<li>4 &#181;l 50% dextran sulfate</li>
	<li>2 &#181;l 20x SSC</li>
	<li>4 &#181;l H2O</li>
</ul>
SBT8 (10 ml)
<ul>
	<li>2 ml 20x SSC</li>
	<li>0.01 g Bovine Serum Albumin (BSA)</li>
	<li>10 &#181;l Tween 20</li>
	<li>7.9 ml H2O</li>
</ul>
Blocking solution8 (10 ml)
<ul>
	<li>0.3 g BSA</li>
	<li>10 &#181;l Tween 20</li>
	<li>2 ml 20x SSC</li>
	<li>8 ml H2O</li>
</ul>
Fixative solution with paraformaldehyde (1 ml)
<ul>
	<li>393.75 &#181;l H2O (add water first)</li>
	<li>450 &#181;l glacial acetic acid</li>
	<li>156.25 &#181;l 16% paraformaldehyde</li>
</ul>

<ol>
	<li>Blattes, R., Kas, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+in+situ+hybridization+(FISH)+on+diploid+nuclei+and+mitotic+chromosomes+from+Drosophila+melanogaster+larval+tissues.">Fluorescent in situ hybridization (FISH) on diploid nuclei and mitotic chromosomes from Drosophila melanogaster larval tissues.</a> Cold Spring Harbor Protocols. 2009 (9), (2009).</li><li>Dimitri, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+in+situ+hybridization+with+transposable+element+probes+to+mitotic+chromosomal+heterochromatin+of+Drosophila.">Fluorescent in situ hybridization with transposable element probes to mitotic chromosomal heterochromatin of Drosophila.</a> Methods in Molecular Biology. 260, 29-39 (2004).</li><li>Pardue, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+situ+hybridization+to+polytene+chromosomes+in+Drosophila+using+digoxigenin-labeled+probes.">In situ hybridization to polytene chromosomes in Drosophila using digoxigenin-labeled probes.</a> Cold Spring Harbor Protocols. 2011 (8), 1003-1006 (2011).</li><li>Hoskins, R. A., 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=Heterochromatic+sequences+in+a+Drosophila+whole-genome+shotgun+assembly.">Heterochromatic sequences in a Drosophila whole-genome shotgun assembly.</a> Genome Biology. 3 (12), (2002).</li><li>Hoskins, R. A., 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=Sequence+finishing+and+mapping+of+Drosophila+melanogaster+heterochromatin.">Sequence finishing and mapping of Drosophila melanogaster heterochromatin.</a> Science. 316 (58331), 1625-1628 (2007).</li><li>Treangen, T. J., Salzberg, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Repetitive+DNA+and+next-generation+sequencing:+computational+challenges+and+solutions.">Repetitive DNA and next-generation sequencing: computational challenges and solutions.</a> Nature Reviews Genetics. 13 (1), 36-46 (2012).</li><li>He, B., 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=Mapping+the+pericentric+heterochromatin+by+comparative+genomic+hybridization+analysis+and+chromosome+deletions+in+Drosophila+melanogaster.">Mapping the pericentric heterochromatin by comparative genomic hybridization analysis and chromosome deletions in Drosophila melanogaster.</a> Genome Research. 22 (12), 2507-2519 (2012).</li><li>Pimpinelli, S., Bonaccorsi, S., Fanti, L., Gatti, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+in+situ+hybridization+(FISH)+of+mitotic+chromosomes+from+Drosophila+larval+brain.">Fluorescent in situ hybridization (FISH) of mitotic chromosomes from Drosophila larval brain.</a> Cold Spring Harbor Protocols. 2010 (3), (2010).</li><li>Larracuente, A. M., Noor, M. A., Clark, A. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Translocation+of+Y-linked+genes+to+the+dot+chromosome+in+Drosophila+pseudoobscura.">Translocation of Y-linked genes to the dot chromosome in Drosophila pseudoobscura.</a> Molecular Biology and Evolution. 27 (7), 1612-1620 (2010).</li><li>Ferree, P. M., Barbash, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Species-specific+heterochromatin+prevents+mitotic+chromosome+segregation+to+cause+hybrid+lethality+in+Drosophila.">Species-specific heterochromatin prevents mitotic chromosome segregation to cause hybrid lethality in Drosophila.</a> PLoS Biology. 7 (10), e1000234 (2009).</li><li>Williams, B. C., Karr, T. L., Montgomery, J. M., Goldberg, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+l(1)zw10+gene+product,+required+for+accurate+mitotic+chromosome+segregation,+is+redistributed+at+anaphase+onset.">The Drosophila l(1)zw10 gene product, required for accurate mitotic chromosome segregation, is redistributed at anaphase onset.</a> The Journal of Cell Biology. 118 (4), 759-773 (1992).</li><li>Gatti, M., Bonaccorsi, S., Pimpinelli, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Looking+at+Drosophila+Mitotic+Chromosomes.">Looking at Drosophila Mitotic Chromosomes.</a> Methods in Cell Biology. 44, 371-391 (1994).</li><li>Werren, J. 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The authors declare that they have no competing financial or any other conflict of interest.

DNA ISH is frequently used to map specific sequences to chromosomes. We have described a simple method for DNA ISH optimized for high copy number, heterochromatic sequences. Rather than using washes in a formamide solution, which is a requirement in other existing DNA ISH protocols, we place tissue-mounted slides directly on a pre-heated block to denature DNA. This method circumvents the use of large amounts of formamide. One critical step for producing crisp hybridization signals is to use freshly made fixation solution...

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. Probes to single copy regions within euchromatin can be generated through a handful of approaches, including nick-translation or end-labeling of long DNA products1,2 and the incorporation of deoxygenin (DIG)-attached nucleotides and their recognition through a wide variety of group-conjugated antibodies1-3. Visualization of euchromatic sequences in few or single copy number requires the use of either single, large probes with high specific activity or a cocktail of multiple, smaller probes that collectively enhance signal.
In contrast, highly repetitive sequences found in heterochromatin, such as satellite DNAs, are easier targets for DNA ISH because they normally exist as tens to thousands of repeats clustered in single chromosome regions known as blocks. Transposable elements also can be found at high copy numbers at distinct chromosomal loci2. In these cases, single probes with low specific activity can effectively label heterochromatic sequences due to their hybridization at multiple sites. Probes to repetitive sequences can be synthesized commercially as short oligonucleotides (30-50 bp) and chemically conjugated with any of multiple different fluorescent groups. Mapping repetitive sequences within heterochromatin by using genome-sequencing technologies is difficult due to challenges encountered in building scaffolds within highly repetitive satellite blocks4-6,7. Currently, ISH stands as the most effective way of mapping these sequences at the sub-chromosome level. This strategy is important for mapping large numbers of repetitive sequences that are being uncovered by ongoing genome and transcriptome sequencing studies.
The efficiency and ease of mapping repetitive sequences on slide-mounted chromosomes would be greatly enhanced by a simplified protocol for DNA ISH. For example, existing protocols for DNA ISH involve multiple washes of hybridized tissues in formamide solution2,8, thus adding substantially to the required time for mapping sequences and also producing large amounts of chemical waste for this costly reagent. Here we describe a revised DNA ISH method that circumvents the need for formamide washes and utilizes basic laboratory equipment and reagents. This method was originally designed for the rapid mapping of highly repetitive DNA sequences in heterochromatic regions of Drosophila larval neuroblasts by using commercially synthesized oligos that are conjugated with fluorescence dyes. However, this method also works for mapping repetitive sequences by using larger probes synthesized through other means9,10 and across multiple different tissue and chromosome types. Additionally, this method can be used to map euchromatic sequences by using longer or multiple, short probes within the euchromatic sequence of interest.

<table><tbody><tr><td>Poly-L-lysine coated slides (regular slides also can be used)</td><td>Sigma Aldrich</td><td></td><td></td></tr><tr><td>Ultrafine tweezers (5 gauge)</td><td>Dumont</td><td></td><td></td></tr><tr><td>22 x 22 mm cover slips</td><td>Fisher</td><td></td><td>Sigmacote-treated by immersion for 15 sec, blotting dry, and wiping away all traces of Sigmacote so that cover slip is clear</td></tr><tr><td>Sigmacote</td><td>Sigma</td><td></td><td></td></tr><tr><td>Filter paper</td><td></td><td></td><td>75 - 150 mm</td></tr><tr><td>Paraffin wax paper</td><td></td><td></td><td></td></tr><tr><td>Heat block with thermometer</td><td></td><td></td><td></td></tr><tr><td>Dry incubator</td><td></td><td></td><td></td></tr><tr><td>Razor blades</td><td></td><td></td><td></td></tr><tr><td>Humidity chamber</td><td></td><td></td><td>empty pipette tip box or Tupperware, lined with moistened paper towels or Kimwipes</td></tr><tr><td>Coplin jars</td><td></td><td></td><td>with slide grooves</td></tr><tr><td>Aluminum foil</td><td></td><td></td><td></td></tr><tr><td>Pasteur pipettes</td><td></td><td></td><td></td></tr><tr><td>1.5 ml microfuge tubes</td><td></td><td></td><td></td></tr><tr><td>Nail polish</td><td></td><td></td><td>clear or colored</td></tr><tr><td>P20 micropipette and plastic tips</td><td></td><td></td><td></td></tr><tr><td>Paperclips</td><td></td><td></td><td>20 - 25 standard metal paperclips linked to form a chain</td></tr><tr><td colspan="4">Reagents</td></tr><tr><td>16% EM grade paraformaldehyde</td><td>Electron Microscopy Reagents</td><td></td><td></td></tr><tr><td>Acetic acid</td><td>Sigma</td><td></td><td></td></tr><tr><td>Liquid nitrogen</td><td></td><td></td><td></td></tr><tr><td>100% Ethanol, chemical grade</td><td></td><td></td><td></td></tr><tr><td colspan="4">Commercially synthesized, fluorescently labeled oligos</td></tr><tr><td>Long biotinylated probe</td><td>Invitrogen; Alternative steps 2.7.1-2.7.3</td><td></td><td>&lt;em&gt;e.g.,&lt;/em&gt; nick translated and biotinylated with BioNick from Invitrogen</td></tr><tr><td>Rhodamine-Avidin</td><td>Roche; Alternative steps 2.7.1-2.7.3</td><td></td><td>for detection of long biotinylated probe</td></tr><tr><td>Hybridization buffer</td><td>Recipe above</td><td></td><td></td></tr><tr><td>4x SSCT</td><td>Recipe above</td><td></td><td>saline-sodium citrate + Tween</td></tr><tr><td>0.1x SSC</td><td>Recipe above</td><td></td><td>saline-sodium citrate</td></tr><tr><td>Blocking solution</td><td>Recipe above</td><td></td><td></td></tr><tr><td>SBT</td><td>Recipe above</td><td></td><td>SSC, bovine serum albumin, Tween</td></tr><tr><td>1x PBT</td><td>Recipe above</td><td></td><td>phosphate-buffered saline + Tween</td></tr><tr><td>1x PBS</td><td></td><td></td><td>phosphate-buffered saline</td></tr><tr><td>Hypotonic solution</td><td></td><td></td><td>0.5% sodium citrate in H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Formamide</td><td>Sigma Aldrich</td><td></td><td></td></tr><tr><td>Vectashield mounting medium with DAPI</td><td>Vector laboratories</td><td></td><td></td></tr></tbody></table>

simple method for fluorescence dna in situ hybridization to squashed chromosomes

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. This approach is particularly effective at mapping highly repetitive sequences to heterochromatic regions, where computational approaches face prohibitive challenges. Here we describe a streamlined protocol for DNA ISH that circumvents formamide washes that are standard steps in other DNA ISH protocols. Our protocol is optimized for hybridization with short single strand DNA probes that carry fluorescent dyes, which effectively mark repetitive DNA sequences within heterochromatic chromosomal regions across a number of different insect tissue types. However, applications may be extended to use with larger probes and visualization of single copy (non-repetitive) DNA sequences. We demonstrate this method by mapping several different repetitive sequences to squashed chromosomes from Drosophila melanogaster neural cells and Nasonia vitripennis spermatocytes. We show hybridization patterns for both small, commercially synthesized probes and for a larger probe for comparison. This procedure uses simple laboratory supplies and reagents, and is ideal for investigators who have little experience with performing DNA ISH.

To demonstrate this method, we hybridized a set of small commercially synthesized oligos that were chemically modified with fluorescent conjugates (Figure 2) and a longer biotinylated probe (made through nick translation of a PCR product; Figure 2B), to chromosomes from several different tissue types (see Table 1). The target sequences included satellite repeats located in pericentromeric (heterochromatic) regions of mitotic chromosomes from D. melanogaster larv...

Watch this Scientific Journal Video about Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes at JoVE.com

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

Here, we present a simple method for performing fluorescence DNA in situ hybridization (DNA ISH) to visualize repetitive heterochromatic sequences on slide-mounted chromosomes. The method requires minimal reagents and it is versatile for use with short or long probes, different tissues, and detection with fluorescence or non-fluorescence-based signals.

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

DNA in situ hybridization (DNA ISH) is a commonly used method for mapping sequences to specific chromosome regions. This approach is particularly ...

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18.0K Views.  University of Rochester. Here, we present a simple method for performing fluorescence DNA in situ hybridization (DNA ISH) to visualize repetitive heterochromatic sequences on slide-mounted chromosomes. The method requires minimal reagents and it is versatile for use with short or long probes, different tissues, and detection with fluorescence or non-fluorescence-based signals.

Video: Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

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 ...

Genetics

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

Immunology and Infection

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological advantages provided by the giant salivary gland polytene chromosomes of third instar larvae.  In this tissue the chromosomes undergo many rounds of replication in the absence of cell division giving rise to approximately 1000 copies.  The DNA remains aligned after each replicative cycle resulting in greatly enlarged chromosomes that provide a unique opportunity to correlate chromatin morphology with the localization of specific proteins.  Consequently, there has been a high level of interest in defining the epigenetic modifications present at different genes and at different stages of the transcription process. An important tool for such studies is the labeling of polytene chromosomes with antibodies to the enzyme, transcription factor, or histone modification of interest. This video protocol illustrates the squash technique used in the Johansen laboratory to prepare Drosophila polytene chromosomes for antibody labeling.

This video protocol illustrates the squash technique used in the Johansen laboratory to prepare Drosophila polytene chromosomes for antibody labeling.

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological ...

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological processes. In particular, appropriate regulation of calcium activity patterns during embryogenesis is necessary for many aspects of vertebrate neural development, including proper neural tube closure, synaptogenesis, and neurotransmitter phenotype specification. While the observation that calcium activity patterns can differ in both frequency and amplitude suggests a compelling mechanism by which these fluxes might transmit encoded signals to downstream effectors and regulate gene expression, existing population-level approaches have lacked the precision necessary to further explore this possibility. Furthermore, these approaches limit studies of the role of cell-cell interactions by precluding the ability to assay the state of neuronal determination in the absence of cell-cell contact. Therefore, we have established an experimental workflow that pairs time-lapse calcium imaging of dissociated neuronal explants with a fluorescence in situ hybridization assay, allowing the unambiguous correlation of calcium activity pattern with molecular phenotype on a single-cell level. We were successfully able to use this approach to distinguish and characterize specific calcium activity patterns associated with differentiating neural cells and neural progenitor cells, respectively; beyond this, however, the experimental framework described in this article could be readily adapted to investigate correlations between any time-series activity profile and expression of a gene or genes of interest.

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

We present a two-part protocol that combines fluorescent calcium imaging with in situ hybridization, allowing the experimenter to correlate patterns of calcium activity with gene expression profiles on a single-cell level.

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological ...

Developmental Biology

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 ...

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.

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.

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

Combined Immunofluorescence and DNA FISH on 3D-preserved Interphase Nuclei to Study Changes in 3D Nuclear Organization

Fluorescent in situ hybridization using DNA probes on 3-dimensionally preserved nuclei followed by 3D confocal microscopy (3D DNA FISH) represents the most direct way to visualize the location of gene loci, chromosomal sub-regions or entire territories in individual cells. This type of analysis provides insight into the global architecture of the nucleus as well as the behavior of specific genomic loci and regions within the nuclear space. Immunofluorescence, on the other hand, permits the detection of nuclear proteins (modified histones, histone variants and modifiers, transcription machinery and factors, nuclear sub-compartments, etc). The major challenge in combining immunofluorescence and 3D DNA FISH is, on the one hand to preserve the epitope detected by the antibody as well as the 3D architecture of the nucleus, and on the other hand, to allow the penetration of the DNA probe to detect gene loci or chromosome territories 1-5. Here we provide a protocol that combines visualization of chromatin modifications with genomic loci in 3D preserved nuclei.

Here we describe a protocol for simultaneous detection of histone modifications by immunofluorescence and DNA sequences by DNA FISH followed by 3D microscopy and analyses (3D immuno-DNA FISH).

Fluorescent in situ hybridization using DNA probes  on 3-dimensionally preserved nuclei followed by 3D confocal microscopy (3D DNA  FISH) represents the ...

Robust 3D DNA FISH Using Directly Labeled Probes

3D DNA FISH has become a major tool for analyzing three-dimensional organization of the nucleus, and several variations of the technique have been published. In this article we describe a protocol which has been optimized for robustness, reproducibility, and ease of use. Brightly fluorescent directly labeled probes are generated by nick-translation with amino-allyldUTP followed by chemical coupling of the dye. 3D DNA FISH is performed using a freeze-thaw step for cell permeabilization and a heating step for simultaneous denaturation of probe and nuclear DNA. The protocol is applicable to a range of cell types and a variety of probes (BACs, plasmids, fosmids, or Whole Chromosome Paints) and allows for high-throughput automated imaging. With this method we routinely investigate nuclear localization of up to three chromosomal regions.

We describe a robust and versatile protocol for analyzing nuclear architecture by 3D DNA FISH using directly labeled fluorescent probes.

3D DNA FISH has become a major tool for  analyzing three-dimensional organization of the nucleus, and several variations  of the technique have been ...

Fluorescence In Situ Hybridization on DNA Halo Preparations to Reveal Whole Chromosomes, Telomeres and Gene Loci

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These structures are collectively known as the nucleoskeleton and include the nuclear lamina, the nucleoli, and nuclear bodies. Although many variants of fluorescence in situ hybridization (FISH) exist to study the genome and its organization, these are often limited by resolution and provide insufficient information on the genome&#39;s association with nuclear structures. The DNA halo method uses high salt concentrations and nonionic detergents to generate DNA loops that remain anchored to structures within nuclei through attachment regions within the genome. Here, soluble nuclear proteins, such as histones, lipids, and DNA not tightly bound to the nuclear matrix, are extracted. This leads to the formation of a halo of unattached DNA surrounding a residual nucleus which itself contains DNA closely associated with internal nuclear structures and extraction-resistant proteins. These extended DNA strands enable increased resolution and can facilitate physical mapping. In combination with FISH, this method has the added advantage of studying genomic interactions with all the structures that the genome is anchored by. This technique, termed HALO-FISH, is highly versatile whereby DNA halos can be coupled with nucleic acid probes to reveal gene loci, whole chromosomes, alpha satellite, telomeres and even RNA. This technique provides an insight into nuclear organization and function in normal&#160;cells and in disease progression such as with cancer.

Combining DNA halo preparations with fluorescence in situ hybridization enables high-resolution analysis of genomic interactions with the nucleoskeleton. Attached genome leads to hybridized fluorescent signals located within the residual extracted nuclei, whereas non-attached genome is in the halo of DNA surrounding the residual nuclei.

The genome is associated with several structures inside cell nuclei, in order to regulate its activity and anchor it in specific locations. These ...

HCR-DNA FISH: A Fluorescence In Situ Hybridization Technique to Detect Viral DNA in Infected Cells

Source: Wei Liu&#160;et al., <a href="https://jove.unicartagenaproxy.elogim.com/v/58950"> Merkel Cell Polyomavirus Infection and Detection</a>, J. Vis. Exp. (2019).
In this video, the cells infected with MCPyV virions were subjected to in situ hybridization chain reaction to detect MCPyV specific genomes. Further, the DNA-HCR technique was combined with FISH to visualize the MCPyV DNA in infected human skin cells.

HCR-DNA FISH: Technika fluorescencyjnej hybrydyzacji in situ do wykrywania wirusowego DNA w zakażonych komórkach

Source: Wei Liu&#160;et al.,  Merkel Cell Polyomavirus Infection and Detection, J. Vis. Exp. (2019).
In this video, the cells infected with MCPyV virions ...

Simple Method for Fluorescence DNA In Situ Hybridization to Squashed Chromosomes

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

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

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

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Combined Immunofluorescence and DNA FISH on 3D-preserved Interphase Nuclei to Study Changes in 3D Nuclear Organization

Robust 3D DNA FISH Using Directly Labeled Probes

Fluorescence In Situ Hybridization on DNA Halo Preparations to Reveal Whole Chromosomes, Telomeres and Gene Loci

HCR-DNA FISH: Technika fluorescencyjnej hybrydyzacji in situ do wykrywania wirusowego DNA w zakażonych komórkach