Name: high-throughput physical mapping of chromosomes using automated in situ hybridization
Uploaded: 2012-06-28T13:00:00
Duration: 8 min 48 s
Description: Watch this Scientific Journal Video about High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization at JoVE.com

This work was supported by the grant from National Institutes of Health 1R21AI094289 to Igor V. Sharakhov. The Xmatrx Automated Slide Staining System and ACCORD PLUS Automated Scanning System were purchased with the help of the Equipment Trust Fund program, Fralin Life Science Institute, Department of Entomology and Department of Biochemistry of Virginia Tech.

1. High-pressure Polytene Chromosome Preparation from Ovarian Nurse Cells
<ol>
	<li>Dissect half-gravid Anopheles females at 25 hours post-blood feeding under a dissection microscope and fix ovaries in fresh modified Carnoy's solution with methanol (100% methanol: glacial acetic acid, 3:1). At this stage the ovaries are at Christophers' III stage of development when follicles have an oval shape and a transparent area with nurse cells within follicles has a round shape13. Put ovaries from approximately five females into 500 &mu;l of modified Carnoy's solution in a 1.5 ml Eppendorf tube and keep them at room temperature for 24 hours. Transfer ovaries to -20 &deg;C for a long-term storage.</li>
	<li>Prepare modified Carnoy's solution with ethanol (100% ethanol: glacial acetic acid, 3:1) and 50% propionic acid just prior to making slides. Place one pair of ovaries on a dust-free slide in one drop of modified Carnoy's solution. Split ovaries into approximately 6 sections with dissecting needles and place them into drops of 50% propionic acid on clean slides under a dissection MZ6 Leica stereomicroscope. Use a separate slide for each section.</li>
	<li>Separate follicles dissecting using needles and wipe out remaining tissue with filter paper or paper towel under a dissection microscope. Add a new drop of 50% propionic acid to the follicles and allow them to sit for 3-5 minutes at room temperature. Place a coverslip on top of the 50% propionic acid droplet. Let the slide stand for approximately 1 minute.</li>
	<li>Wrap the slide with filter paper and plastic. Using a Dremel 200 rotary tool with a Flex-Shaft attachment and soft plastic tip set between 3000-5000 RPM, express the follicles by swirling the tip in circles and lightly pressing the coverslip to evenly spread the nuclei. This step should take approximately 1 minute. Check the spread quality with an Olympus CX41 Phase Microscope using a 20x objective.</li>
	<li>Prepare a sandwich by placing an additional coverslip next to the coverslip covering the chromosomes, and cover them with a second microscope slide. This reduces the chance of crushing the slide in the vise. Wrap the slides with a plastic sheet that come in glass slide containers and filter paper to hold the sandwich and to protect the slide from scratching due to the vise.</li>
	<li>Apply pressure to the slides via mechanical vise. A pressure of 85-120 inch-lbs is sufficient and is achieved by using a torque wrench. This step is necessary for flattening the chromosomes as much as possible.</li>
	<li>Remove the second microscope slide and the additional coverslip. Heat the slide with the coverslip covering the chromosomes to 55 &deg;C on a slide denaturation/hybridization system for 10-15 minutes to further flatten the chromosomes. Dip the slide in liquid nitrogen for at least 15 seconds, and, when bubbling has stopped, proceed to quickly remove the coverslip with a razor blade. Immediately place slides in cold 50% ethanol for 5 minutes. Dehydrate slides in 70%, 90%, 100% ethanol for 5 minutes each. Air dry slides.</li>
</ol>
2. FISH Using an Automated Slide Staining System
Program is set up with the Xmatrx automated slide staining system to run the following steps, except the probe labeling. The detailed nick-translation labeling protocol is described in the documentation provided with DNA polymerase from Fermentas.
<ol>
	<li>Before FISH, prepare fluorescent probes by labeling genomic BAC DNA with a fluorochrome using a nick-translation protocol. Mix 1 &mu;g of DNA, 0.05 mM each of unlabeled dATP, dCTP, and dGTP and 0.015 mM dTTP, 1 &mu;l Cy3- or Cy5-dUTP, 0.05 mg/ml BSA, 5 &mu;l of 10x nick-translation buffer, 20 units of DNA polymerase I, 0.0012 units of DNase I, and nuclease-free water to 25 &mu;l. The DNA polymerase I/DNase I ratio is selected empirically to obtain probes with a size range from 300 to 500 bp. Incubate the mix at 15 &deg;C for 1 hour.</li>
	<li>Put slides and reagents into the Xmatrx automated slide staining system and start the program to run the following steps. Apply 800 &mu;l of 1x PBS for 20 min. Blow slides with air. Perform formalin fixation by applying 450 &mu;l of 4% formalin in 1x PBS for 1 min followed by washes with 100% ethanol for 1 sec twice and for 2 min once. Blow slides with air.</li>
	<li>Heat slides at 45 &deg;C for 2 min to avoid bubbling when applying the probes. Apply 20 &mu;l of the DNA probes, add drops of mineral oil to avoid evaporation of a hybridization solution, and place a coverslip on top. Denature chromosomes and DNA probes by heating slides at 90 &deg;C for 10 min.</li>
	<li>For hybridization, incubate slides at 42 &deg;C for 14 hours with coverslips on. The hybridization solution consists of 2x SSC, 100 mM sodium phosphate, 1x Denhardt's solution, 100 &mu;g/ml of sodium azide, and 10% dextran sulfate in formamide.</li>
	<li>For stringency washes, heat slides at 42 &deg;C for 2 min, remove coverslips, wash slides in 2x SSC for 1 sec 4 times. Blow slides with air. Apply 800 &mu;l of 0.4x SSC at 42 &deg;C for 10 min 2 times. Wash slides in 2x SSC at 25 &deg;C for 10 min.</li>
	<li>Perform chromosome staining by applying 50 &mu;l of 1 &mu;M YOYO-1 in 1x PBS. Apply drops of mineral oil to avoid evaporation of the staining solution, put on coverslips, and incubate at 25 &deg;C for 10 min. Remove coverslips, wash slides in 2x SSC for 1 sec 4 times. Blow slides with air. Apply 15 &mu;l of ProLong Gold antifade reagent. Put on coverslips.</li>
</ol>
3. Slide Reading with an Automatic Fluorescent Imaging System
This section details the use of the Duet software, which comes standard with the ACCORD PLUS automated scanning system. Instructions begin after turning on in this order: Olympus U-RFL-T power supply for a halogen bulb, computer Dell precision T3500, microscope Olympus BX61 with a connected camera Olympus U-CMAD3.
<ol>
	<li>For setting up 10x Pre-Scan, open the Duet software in the ACCORD PLUS automated scanning system. Click "Online" button. Enter new Case ID and assign a Slide ID. Click the dot labeled "BF." This is the brightfield option. Set a scan choice to "10x Prescan." Use "2500x circle," "10000x circle" or "rectangle." Click "Set&amp;run" for 10x Pre-Scan.</li>
	<li>Click the "OK" button to run 10x Pre-Scan. Follow the prompts to adjust the scanning properly. Click "Finish" to start the scan. Press the "Main" tab to go back to the main screen. Click "Offline" button. Find the Case ID and Slide ID that was assigned and click "Offline Scan." In the black box at the top left area click on an arrow and select "10x Prescan." Using the arrows (&lt; || &Delta; &gt; a.k.a. "back," "pause," "play," "forward" buttons), go through the scanned images.</li>
	<li>After finding an image of interest, double click on the screen the middle of the target region and press "Snap." This will target an image for capture later on. After selecting all targets, click "Classify." Select "10x Prescan." Right click an image and get the chance to classify the images. Select "Polytene."</li>
	<li>Set up 40x Pre-Scan in the Duet software. Click "Main" to go back to the main menu. Select "Online" again. Select a slide. Change "BF" to "FL." Change the task name to "Revisit-X40-RG." Change the section right below the last setting to "Revisit-ALL." Click "Set&amp;Run." Press "OK." Follow the prompts again to set up the automation. Click the "Start matching views" button to match 10x and 40x images. Click "Finish" to start the scan. Once done, click "Classify" and look at the images.</li>
</ol>
4. Representative Results
Figure 1 graphically depicts the scheme of the high-pressure chromosome preparation. This step involves the process of squashing and flattening chromosomes using a Dremel tool and mechanical vise, as well as chromosome visualization using a phase-contrast microscope. Figure 2 illustrates the hybridization of a fluorescently labeled probe to target DNA on the chromosome slide preparations using an automated slide staining system, the use of an automated scanning microscope for visualizing and mapping the probes after the FISH experiment, and placing and orienting genomic scaffolds on the chromosomes.
Preparations of ovarian nurse cell polytene chromosomes from females of An. gambiae were made using the high-pressure technique. This method does not damage or change most of the chromosome structure (Figure 3). It flattens bended chromosomes and, thus, reveals hidden fine bands that are not seen on regular preparations (Figure 4).
The probes used in this protocol are genomic BAC DNA clones obtained from a ND-TAM BAC library generated from An. gambiae PEST strain DNA. The genomic DNA for this library was extracted from newly hatched first instar larvae of both sexes. Figure 5 shows the results of FISH using BAC clones hybridized to polytene chromosomes of An. gambiae. This procedure was performed using the Xmatrx automated slide staining system. Using a cytogenetic photomap for An. gambiae10, two BAC clones, 102B24 (GenBank: BH372701, BH372694) and BAC 142O19 (GenBank: BH368703, BH368698), were localized in subdivisions 16C and 16D of the 2R arm, respectively. However, the BLAST search against the An. gambiae PEST strain AgamP3 assembly 14 identified the sequences homologous to 102B24 and 142O19 in subdivisions 16B and 17A of the 2R arm, respectively (Table 1). Therefore, the correspondence between the genomic coordinates and the cytogenetic subdivisions can now be adjusted according to our mapping data. The BLAST search against the An. gambiae M form and S form genome assemblies found that the two BAC clones are located in different contigs, but in the same scaffolds within each form (Table 1). The identified contigs can now be associated with specific chromosomal locations. Moreover, the identified scaffolds can now be properly oriented within the cytological subdivisions 16CD. Interestingly, the distances between the 102B24 and 142O19 sequences in the scaffolds are 1,892,981 bp, 1,658,391 bp, and 1,688,426 bp in the PEST strain, M-form, and S-form genome assemblies, respectively. Because, the PEST strain is a hybrid between the M and S forms, this difference is likely due to incorrect assembly of the PEST genome.
<table border="1">
	<tbody>
		<tr>
			<td>BAC 
			(accession)</td>
			<td>PEST-strain 
			AgamP3 
			 
			Coordinates</td>
			<td>M-form 
			contigs 
			 
			Coordinates</td>
			<td>M-form 
			scaffolds 
			 
			Coordinates</td>
			<td>S-form 
			contigs 
			 
			Coordinates</td>
			<td>S-form 
			scaffolds 
			 
			Coordinates</td>
		</tr>
		<tr>
			<td>102B24 
			(BH372694)</td>
			<td>2R:AAAB0100 
			8844_26 (16B) 
			 
			43,681,342- 
			43,681,874</td>
			<td>ABKP020247 
			53.1 
			 
			32,513- 
			33,045</td>
			<td>EQ090167.1 
			 
			1,858,377- 
			1,858,909</td>
			<td>ABKQ010123 
			32.1 
			 
			4,299- 
			4,832</td>
			<td>EQ099711.1 
			 
			215,891- 
			216,424</td>
		</tr>
		<tr>
			<td>102B24 
			(BH372701)</td>
			<td>2R:AAAB0100 
			8844_28 (16B) 
			 
			43,788,213- 
			43,788,969</td>
			<td>ABKP020247 
			51.1 
			 
			24,816- 
			25,575</td>
			<td>EQ090167.1, 
			 
			1,772,683- 
			1,773,442</td>
			<td>ABKQ010123 
			35.1 
			 
			27,782- 
			28,540</td>
			<td>EQ099711.1, 
			 
			305,514- 
			306,272</td>
		</tr>
		<tr>
			<td>142O19 
			(BH368698)</td>
			<td>2R:AAAB0100 
			 
			8805_5 (17A) 
			45,428,580- 
			45,429,264</td>
			<td>ABKP020247 
			01.1 
			 
			2,499- 
			3,185</td>
			<td>EQ090167.1 
			 
			315,806- 
			316,492</td>
			<td>ABKQ010123 
			76.1 
			 
			49,242- 
			49,942</td>
			<td>EQ099711.1 
			 
			1,791,625- 
			1,792,325</td>
		</tr>
		<tr>
			<td>142O19 
			(BH368703)</td>
			<td>2R:AAAB0100 
			8805_8 (17A) 
			 
			45,560,099- 
			45,574,323</td>
			<td>ABKP020220 
			17.1 
			 
			30,971- 
			32,469</td>
			<td>EQ090167.1 
			200,518- 
			202,016</td>
			<td>ABKQ010123 
			79.1 
			 
			27,760- 
			28,508</td>
			<td>EQ099711.1 
			 
			1,903,569- 
			1,904,317</td>
		</tr>
	</tbody>
</table>
Table 1. BLAST results of the BAC-end sequences against the genome assemblies of An. gambiae.
<img alt="Figure 1" src="/files/ftp_upload/4007/4007fig1.jpg" /> 
Figure 1. Schematic representation of high-pressure chromosome preparation. Mosquito ovaries are shown at the correct stage of development.
<img alt="Figure 2" src="/files/ftp_upload/4007/4007fig2.jpg" /> 
Figure 2. A scheme representing automated FISH, slide scanning, and chromosome mapping of genomic scaffolds.
<img alt="Figure 3" src="/files/ftp_upload/4007/4007fig3.jpg" /> 
Figure 3. A spread of polytene chromosome 3 of An. gambiae obtained using the high-pressure technique. 3L and 3R mark left and right arms at their telomeres. PH and IH indicate pericentric and intercalary heterochromatin, respectively.
<img alt="Figure 4" src="/files/ftp_upload/4007/4007fig4.jpg" /> 
Figure 4. Comparison of An. gambiae polytene chromosomes prepared using the traditional (top) and high-pressure (bottom) technique. The images cover subdivisions 29A-30E of arm 3R (A) and 43D-46D of arm 3L (B).
<img alt="Figure 5" src="/files/ftp_upload/4007/4007fig5.jpg" /> 
Figure 5. FISH of BAC clones to polytene chromosomes of An. gambiae. A) Hybridization of 102B24 (red signal) with the 2R arm. B) Dual-color FISH of 102B24 (red signal) and 142O19 (blue signal) to subdivisions 16C and 16D of the stretched 2R arm, respectively. Arrows indicate signals of hybridization of BAC clones labeled with Cy3 (red) and Cy5 (blue). C-the centromeric region. a/+ shows the heterozygote 2La inversion. Chromosomes were counterstained with the fluorophore YOYO-1.

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No conflicts of interest declared.

The most critical step of the high-pressure procedure is the proper squashing of ovarian nurse cells isolated at Christophers' III stage of ovarian development13. Improper squashing can lead to insufficiently spread chromosomes, which can cause problems when trying to determine probe locations after FISH. If the chromosomes are over-squashed, they can be become broken or elongated to the point where banding patterns are lost. Production of multiple slides should lead to a consistency when attempting to squash the slides using the Dremel tool, which will increase overall slide production efficiency. The high-pressure technique was first developed for freshly isolated salivary glands of D. melanogaster9. However, ovaries of mosquitoes are routinely preserved in modified Carnoy's fixative solution (3 methanol: 1 glacial acetic acid by volume) before they are used for chromosomal preparations. Therefore, we modified the existing high-pressure protocol to make it suitable for the fixed ovarian nurse cell polytene chromosomes of the malaria mosquito An. gambiae. Because the high pressure is applied using a precision vise possessing a highly parallel work surface of the entire slide, it takes significantly less time to prepare the chromosome squash than using a traditional tapping technique with a pencil's eraser.
Other important steps include sufficiently soaking follicles in 50% propionic acid and heating the slide. Both of these steps are essential in helping to flatten the chromosomes. If they are neglected, chromosomes can appear shiny after dehydration, which potentially leads to an overabundance of background that can be mistaken for a signal in FISH. The high-pressure squashing technique also works well using the same solutions when making mitotic chromosome preparations. The additional spreading and flattening helps to better express the chromosomes from their nuclei. Adding equal pressure should flatten them accordingly to allow for measurements and potentially better resolution of bands visible from staining. The details about isolating mitotic chromosomes from mosquitoes are given elsewhere15. Although regular squash preparations are sufficient for many purposes including FISH 16 and immunostaining17, the high-pressure method not only lowers potential variance from one slide to the next, but also increases overall chromosome quality, leading to higher detail when mapping chromosomes. This procedure can also be used for preparing polytene chromosome membrane slides for laser-capture microdissection.
Limitations to the high-pressure method include slide breakage and overstretching of the chromosomes. Slide breakage caused by placing too much stress on the slide via the vise is possible, but is limited by using the pressures denoted in the article. For overstretching of the chromosomes, if the Dremel tool is applied to the slide for an extended period of time, there is the possibility that the chromosomes can become too stretched out and lose resolution. This can be remedied by applying the tool for a brief period of time, checking under the microscope, and applying more time with the Dremel tool if the chromosomes are insufficiently spread.
A traditional FISH protocol includes a number of washing and incubation steps, which are usually 5-20 min long and require almost full attention of a researcher for the whole duration of the experiment. Moreover, the number of slides that can be handled manually is usually limited to a few slides in a given experiment. In contrast, an automated slide staining system performs all steps (including washing, incubation, probe application, denaturation, hybridization, coverslip application and removal) automatically. This frees up to 6-8 working hours for a researcher in a FISH experiment. Moreover, an automated FISH system can significantly increase the throughput. For example, the Xmatrx system is capable of processing up to 40 assays on single preparation slides and up to 80 assays on dual preparation slides simultaneously. Among limitations of this system is that it is not efficient for a small number of FISH experiments as it requires preparing large volumes of solutions. In addition, the FISH protocol programmed in the system may require modifications and adjustments for new applications.
A slide scanning system is an automated version of a fluorescent microscope. Automated stage moving and a simple microscope control panel free a researcher's time and make operating the microscope extremely simple. For instance, the software in the ACCORD PLUS scanning system allows for easy capturing of multiple channels of fluorescence, and easy to manage image acquisition. An example of this is the inclusion of z-stack capturing, rather than taking a single image; the software captures a configurable z-stack of images to ensure that at least one image remains in focus. Although this system is made more for multiple image acquisition (a lot of cells on a single slide), it still makes finding chromosomes on the slide much easier than navigating through the slide manually.
Both the automated slide staining system and the scanning system are routinely used for FISH diagnostics of chromosomal abnormalities in human cells by cytogenetics clinics. In this study, we developed a protocol that utilizes these automated systems for a basic research application. The automated high-throughput physical mapping protocol can facilitate rapid development of physical chromosome maps for any species of interest. Although, in this report, we used polytene chromosomes for physical mapping, mitotic chromosomes can also be utilized with these systems. It would be useful for the protocol to be adjusted to mitotic chromosomes because the majority of organisms do not develop readable polytene chromosomes. However, the polytene chromosomes, when available, can provide useful information about the correspondence of functional genome domains with the chromosome structure at the highest resolution. The organizational principles of polytene chromosomes, such as patterns of bands and interbands, have recently been likened to that of regular nonpolytene (interphase) chromosomes18. Therefore, the detailed physical mapping performed on high-pressure chromosome preparations has the potential to link DNA sequences to specific chromosomal structures such as bands, interbands, puffs, centromeres, telomeres, and heterochromatin; thus, creating chromosome-based genome assemblies.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent/Equipment</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments </td>
 </tr>
 <tr>
 <td>22x22 mm microscope coverslips</td>
 <td>Fisher</td>
 <td>12-544-10</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>18x18 mm microscope coverslips</td>
 <td>Fisher</td>
 <td>12-553-402 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Acetic acid</td>
 <td>Fisher</td>
 <td>A491-212</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Methanol</td>
 <td>Fisher </td>
 <td>A412-4</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Propionic acid </td>
 <td>Sigma-Aldrich</td>
 <td>402907</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dremel 200 rotary tool with a Flex-Shaft attachment</td>
 <td>Rand</td>
 <td>&nbsp;</td>
 <td>3" Multi-purpose Mini Bench Grinder (with rate limiter)</td>
 </tr>
 <tr>
 <td>Specially designed 200 &mu;l plastic tips (beaded plastic edges) for Dremel tool</td>
 <td>Pipetman</td>
 <td>F171300</td>
 <td>Cut and heat fat end of pipet tips</td>
 </tr>
 <tr>
 <td>Tin coated rapid vise</td>
 <td>Avenger Gold Toolmaker</td>
 <td>MTC-200-1</td>
 <td>Precision ground square and parallel to within 0.00025</td>
 </tr>
 <tr>
 <td>Torque wrench</td>
 <td>Craftsman</td>
 <td>44593</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>ThermoBrite Slide Denaturation/Hybridization System</td>
 <td>Abbott Molecular</td>
 <td>30-144110</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>25 mm barrier slides</td>
 <td>Abbott Molecular </td>
 <td>XT108-SL</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>25 mm coverslips</td>
 <td>Abbott Molecular </td>
 <td>XT122-90X</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Xmatrx Automated Slide Staining System</td>
 <td>Abbott Molecular</td>
 <td>08L46-001</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>MZ6 Leica stereomicroscope</td>
 <td>Leica</td>
 <td>VA-OM-E194-354</td>
 <td>A different stereomicroscope can be used</td>
 </tr>
 <tr>
 <td>Olympus CX41 Phase Microscope</td>
 <td>Olympus</td>
 <td>CX41RF-5</td>
 <td>A different phase microscope can be used</td>
 </tr>
 <tr>
 <td>ACCORD PLUS Automated Scanning System</td>
 <td>BioView (USA) </td>
 <td>BV-5000-ACCP</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>10x PBS</td>
 <td>Invitrogen </td>
 <td>P5493</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>10% NBF (neutral buffered formalin)</td>
 <td>Sigma-Aldrich</td>
 <td>HT501128</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>99% formamide</td>
 <td>Fisher Scientific</td>
 <td>BP227500</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dextran sulfate sodium salt</td>
 <td>Sigma</td>
 <td>D8906</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>20x SSC buffer</td>
 <td>Invitrogen </td>
 <td>AM9765</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sodium phosphate</td>
 <td>Sigma-Aldrich</td>
 <td>S3264</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>50x Denhardt's solution</td>
 <td>Sigma-Aldrich</td>
 <td>D2532</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sodium azide</td>
 <td>Sigma-Aldrich</td>
 <td>S2002</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>1 mM YOYO-1 iodide (491/509) solution in DMSO</td>
 <td>Invitrogen </td>
 <td>Y3601</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>ProLong Gold antifade reagent </td>
 <td>Invitrogen </td>
 <td>P36930</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>dATP, dCTP, dGTP, dTTP</td>
 <td>Fermentas</td>
 <td>R0141, R0151, R0161, R0171</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Cy3-dUTP, Cy5-dUTP</td>
 <td>GE Healthcare</td>
 <td>PA53022, PA55022</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>BSA</td>
 <td>Sigma</td>
 <td>A3294</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>DNA polymerase I</td>
 <td>Fermentas</td>
 <td>EP0041</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>DNase I </td>
 <td>Fermentas</td>
 <td>EN0521</td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

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.

Watch this Scientific Journal Video about High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization at JoVE.com

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.

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 ...

high-throughput-physical-mapping-chromosomes-using-automated-situ

Research

JoVE Journal

Biology

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding location often influences function of a splicing factor.  However, as with any sampling strategy the chance of identifying an RNA bound to a splicing factor is proportional to its cellular abundance.  We have developed a novel in vitro approach for surveying binding specificity on otherwise transient pre-mRNA. This approach utilizes a specifically designed oligonucleotide pool that tiles across introns, exons, splice junctions, or other pre-mRNA.  The pool is subjected to some kind of molecular selection. Here, we demonstrate the method by separating the oligonucleotide into a bound and unbound fraction and utilize a two color array strategy to record the enrichment of each oligonucleotide in the bound fraction. The array data generates high-resolution maps with the ability to identify sequence-specific and structural determinates of ribonucleoprotein (RNP) binding on pre-mRNA.  A unique advantage to this method is its ability to avoid the sampling bias towards mRNA associated with current IP and SELEX techniques, as the pool is specifically designed and synthesized from pre-mRNA sequence. The flexibility of the oligonucleotide pool is another advantage since the experimenter chooses which regions to study and tile across, tailoring the pool to their individual needs.  Using this technique, one can assay the effects of polymorphisms or mutations on binding on a large scale or clone the library into a functional splicing reporter and identify oligonucleotides that are enriched in the included fraction.  This novel in vitro high-resolution mapping scheme provides a unique way to study RNP interactions with transient pre-mRNA species, whose low abundance makes them difficult to study with current in vivo techniques.  



A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding ...

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 ...

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

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

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

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

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

Quantitative and Automated High-throughput Genome-wide RNAi Screens in C. elegans

RNA interference is a powerful method to understand gene function, especially when conducted at a whole-genome scale and in a quantitative context. In C. elegans, gene function can be knocked down simply and efficiently by feeding worms with bacteria expressing a dsRNA corresponding to a specific gene 1. While the creation of libraries of RNAi clones covering most of the C. elegans genome 2,3 opened the way for true functional genomic studies (see for example 4-7), most established methods are laborious. Moy and colleagues have developed semi-automated protocols that facilitate genome-wide screens 8. The approach relies on microscopic imaging and image analysis. 
	Here we describe an alternative protocol for a high-throughput genome-wide screen, based on robotic handling of bacterial RNAi clones, quantitative analysis using the COPAS Biosort (Union Biometrica (UBI)), and an integrated software: the MBioLIMS (Laboratory Information Management System from Modul-Bio) a technology that provides increased throughput for data management and sample tracking. The method allows screens to be conducted on solid medium plates. This is particularly important for some studies, such as those addressing host-pathogen interactions in C. elegans, since certain microbes do not efficiently infect worms in liquid culture.
	We show how the method can be used to quantify the importance of genes in anti-fungal innate immunity in C. elegans. In this case, the approach relies on the use of a transgenic strain carrying an epidermal infection-inducible fluorescent reporter gene, with GFP under the control of the promoter of the antimicrobial peptide gene nlp 29 and a red fluorescent reporter that is expressed constitutively in the epidermis. The latter provides an internal control for the functional integrity of the epidermis and nonspecific transgene silencing9. When control worms are infected by the fungus they fluoresce green. Knocking down by RNAi a gene required for nlp 29 expression results in diminished fluorescence after infection. Currently, this protocol allows more than 3,000 RNAi clones to be tested and analyzed per week, opening the possibility of screening the entire genome in less than 2 months.

Quantitative and Automated High-throughput Genome-wide RNAi Screens in C. elegans

We describe a protocol using C. elegans and RNAi feeding libraries that allows automated measurement of multiple parameters such as fluorescence, size and opacity of individual worms in a population. We give one example of a screen to identify genes involved in anti-fungal innate immunity in C. elegans.

RNA interference is a powerful method to understand gene function, especially when conducted at a whole-genome scale and in a quantitative context. In C. ...

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 ...

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 ...

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.

Assessing  the expression pattern of a gene, as well as the subcellular localization  properties of its transcribed RNA, are key features for ...

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for using FISH to quantify the number of mRNAs in single yeast cells. Cells can be grown in any condition of interest and then fixed and made permeable. Subsequently, multiple single-stranded deoxyoligonucleotides conjugated to fluorescent dyes are used to label and visualize mRNAs. Diffraction-limited fluorescence from single mRNA molecules is quantified using a spot-detection algorithm to identify and count the number of mRNAs per cell. While the more standard quantification methods of northern blots, RT-PCR and gene expression microarrays provide information on average mRNAs in the bulk population, FISH facilitates both the counting and localization of these mRNAs in single cells at single-molecule resolution.

Visualization and Analysis of mRNA Molecules Using Fluorescence In Situ Hybridization in Saccharomyces cerevisiae

This protocol describes an experimental procedure for performing Fluorescence in situ Hybridization (FISH) for counting mRNAs in single cells at single-molecule resolution.

The Fluorescence in situ Hybridization (FISH) method allows one to detect nucleic acids in the native cellular environment. Here we provide a protocol for ...

MicroRNA In situ Hybridization for Formalin Fixed Kidney Tissues

In this article we describe a method for colorimetric detection of miRNA in the kidney through in situ hybridization with digoxigenin tagged microRNA probes. This protocol, originally developed by Kloosterman and colleagues for broad use with Exiqon miRNA probes1, has been modified to overcome challenges inherent in miRNA analysis in kidney tissues. These include issues such as structure identification and hard to remove residual probe and antibody. Use of relatively thin, 5 mm thick, tissue sections allowed for clear visualization of kidney structures, while a strong probe signal was retained in cells. Additionally, probe concentration and incubation conditions were optimized to facilitate visualization of microRNA expression with low background and nonspecific signal. Here, the optimized protocol is described, covering the initial tissue collection and preparation through the mounting of slides at the end of the procedure. The basic components of this protocol can be altered for application to other tissues and cell culture models.

MicroRNA In situ Hybridization for Formalin Fixed Kidney Tissues

This article describes an in situ hybridization protocol optimized for colormetric detection of microRNA expression in formalin fixed kidney sections.

In this article we describe a method for colorimetric detection of miRNA in the kidney through in situ hybridization with digoxigenin tagged microRNA ...