Financial support by the Deutsche Forschungsgemeinschaft (DFG, Wa 1386/17-1), the Research Training Group GRK 2039 (funded by DFG) and KIT is gratefully acknowledged.

Caution: Please consult all relevant material safety data sheets (MSDS) before use. Several of the chemicals used in these syntheses are toxic and carcinogenic. Please use all appropriate safety practices that are typically required in organic chemistry laboratories, such as wearing a laboratory coat, safety glasses and gloves.
1. Synthesis of the Dyes
Note: Both dyes can be synthesized by the same types of reaction. Figure 2 shows an overview of these reactions.
<ol>
	<li>Synthesis of 1-(3-azidopropyl)-4-(2-(1-methyl-1H-indol-3-yl)vinyl)pyridin-1-ium iodide (dye 1)
	<ol>
		<li>Synthesis of 1-(3-iodopropyl)-4-methylpyridin-1-ium iodide (Figure 2A, step a)
		<ol>
			<li>Dissolve 466 mg 4-picoline and 5.91 g 1,3-diiodopropane in 10 ml of acetonitrile in a 20 ml headspace vial and seal tightly by a septum cap.</li>
			<li>Heat to 85 &#176;C for 16 hr.</li>
			<li>Allow to cool to RT, then remove the solvent under reduced pressure using a rotatory evaporator.</li>
			<li>Add 20 ml of ethyl acetate to the residual oil and treat the mixture in an 120 W ultrasonic bath for 3 min.</li>
			<li>Collect the formed precipitate by filtration and wash five times with ethyl acetate. Dry the solid product under vacuum O/N.</li>
		</ol>
		</li>
		<li>Synthesis of 1-(3-azidopropyl)-4-methylpyridin-1-ium (Figure 2A, step b)
		<ol>
			<li>Dissolve 900 mg 1-(3-iodopropyl)-4-methylpyridin-1-ium iodide in 12 ml of acetonitrile in a 20 ml headspace vial, add 376 mg of sodium azide, and seal tightly by a septum cap.</li>
			<li>Heat to 85 &#176;C for 16 hr.</li>
			<li>Allow to cool to RT, then remove the solvent under reduced pressure using a rotary evaporator.</li>
			<li>Add 15 ml of dichloromethane to the residue.</li>
			<li>Filter off and discard the resulting precipitate.</li>
			<li>Remove the solvent under reduced pressure using a rotary evaporator to obtain the product as brown oil.</li>
		</ol>
		</li>
		<li>Synthesis of 1-methyl-1H-indole-3-carbaldehyde (Figure 2A, step c)
		<ol>
			<li>Under inert gas (argon), dissolve 1.45 g indole-3-carbaldehyde, 1.52 g potassium carbonate and 2.70 g dimethyl carbonate in 10 ml absolute dimethylformamide in a 50 ml round-bottomed flask equipped with a reflux condenser.</li>
			<li>Stir the mixture at 130 &#176;C for 19 hr.</li>
			<li>Allow to cool to RT, then pour the mixture on ice.</li>
			<li>Extract the aqueous layer three times with 150 ml ethyl acetate.</li>
			<li>Combine the organic layers, wash them with water, dry them over sodium sulfate and remove the solvent at 50 &#176;C under reduced pressure using a rotary evaporator.</li>
		</ol>
		</li>
		<li>Coupling to 1-(3-azidopropyl)-4-(2-(1-methyl-1H-indol-3-yl)vinyl)pyridin-1-ium iodide (dye 1) (Figure 2A, step d)
		<ol>
			<li>Work under argon and under exclusion of moisture. Dissolve 90 mg 1-(3-azidopropyl)-4-methylpyridin-1-ium and 48 mg 1-methyl-1H-indole-3-carbaldehyde in 4 ml ethanol in a 20 ml round-bottomed flask equipped with a reflux condenser.</li>
			<li>Add 0.07 ml piperidine and heat to 80 &#176;C for 4 hr.</li>
			<li>Allow to cool to RT.</li>
			<li>Collect the resulting precipitate by filtration and wash three times with diethylether.</li>
			<li>Add diethylether to the supernatant and collect the resulting precipitate by filtration. Wash three times with diethylether.</li>
			<li>Combine the precipitates.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Synthesis of 1-(3-azidopropyl)-4-(2-(1-methyl-2-phenyl-1H-indol-3-yl)vinyl)quinolin-1-ium iodide (dye 2)
	<ol>
		<li>Synthesis of 1-(3-iodopropyl)-4-methylquinolin-1-ium iodide (Figure 2B, step a)
		<ol>
			<li>Dissolve 715 mg 4-methylquinoline and 5.91 g 1,3-diiodopropane in 10 ml of acetonitrile in a 20 ml headspace vial and seal tightly by a septum cap.</li>
			<li>Heat to 85 &#176;C for 16 hr.</li>
			<li>Allow to cool to RT, then remove the solvent under reduced pressure using a rotary evaporator.</li>
			<li>Add 20 ml of ethyl acetate to the remaining oil and treat the mixture in an 120 W ultrasonic bath for 3 min.</li>
			<li>Collect the formed precipitate by filtration and wash five times with ethyl acetate. Dry the solid product under vacuum O/N.</li>
		</ol>
		</li>
		<li>Synthesis of 1-(3-azidopropyl)-4-methylquinolin-1-ium (Figure 2B, step b)
		<ol>
			<li>Dissolve 900 mg 1-(3-iodopropyl)-4-methylquinolin-1-ium iodide in 12 ml of acetonitrile in a 20 ml headspace vial, add 333 mg of sodium azide, and seal tightly by a septum cap.</li>
			<li>Heat to 85 &#176;C for 16 hr.</li>
			<li>Allow to cool to RT, then remove the solvent under reduced pressure using a rotary evaporator.</li>
			<li>Add 15 ml of dichloromethane to the residue.</li>
			<li>Filter off and discard the resulting precipitate.</li>
			<li>Remove the solvent under reduced pressure using a rotary evaporator to obtain the product as brown oil.</li>
		</ol>
		</li>
		<li>Synthesis of 1-methyl-2-phenyl-1H-indole-3-carbaldehyde (Figure 2B, step c)
		<ol>
			<li>Under inert gas (argon), dissolve 1.45 g 2-phenyl-1H-indole-3-carbaldehyde, 0.996 g potassium carbonate and 1.77 g dimethyl carbonate in 10 ml absolute dimethylformamide in a 50 ml round-bottomed flask equipped with a reflux condenser.</li>
			<li>Stir the mixture at 130 &#176;C for 19 hr.</li>
			<li>Allow to cool to RT, then pour the mixture on ice.</li>
			<li>Extract the aqueous layer three times with 150 ml ethyl acetate.</li>
			<li>Combine the organic layers, wash them with water, dry them over sodium sulfate and remove the solvent under reduced pressure using rotary evaporator.</li>
		</ol>
		</li>
		<li>Coupling to 1-(3-azidopropyl)-4-(2-(1-methyl-2-phe nyl-1H-indol-3-yl)vinyl)quinolin-1-ium iodide (Figure 2B, step d)
		<ol>
			<li>Work under argon and under exclusion of moisture. Dissolve 90 mg 1-(3-azidopropyl)-4-methylquinolin-1-ium and 59.7 mg 1-methyl-2-phenyl-1H-indole-3-carbaldehyde in 4 ml ethanol in a 20 ml round-bottomed flask equipped with reflux condenser.</li>
			<li>Add 0.06 ml piperidine and heat to 80 &#176;C for 4 hr.</li>
			<li>Allow to cool to RT.</li>
			<li>Collect the resulting precipitate by filtration and wash with diethylether three times.</li>
			<li>Add diethylether to the supernatant and collect the resulting precipitate by filtration. Wash three times with diethylether.</li>
			<li>Combine the precipitates.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. Synthesis of the DNA Strands
Note: The synthesis of the DNA strands is carried out using the phosphoramidite method on a solid phase, as described by M. Caruthers11 on a DNA synthesizer. The functioning of the synthesizer is tested before the synthetic procedure, and reagents are renewed if necessary.
<ol>
	<li>Prearrangements
	<ol>
		<li>Dissolve the commercially available 2&#39;-O-propargyl-deoxyuridinephosphoramidite (cU) in 1.2 ml of amidite diluent (extra dry acetonitrile). Move solution in the vial to a synthesizer vial and screw into synthesizer.</li>
		<li>Perform a &#34;leak test&#34; (according to manufacturer&#39;s instructions) to make sure that no argon gas leak exists. If the &#34;leak test&#34; fails, screw in the vial more tightly.</li>
		<li>Prime the cU solution by filling the tube that connects to the solid phase synthesis chamber.</li>
		<li>Wash all lines with acetonitrile.</li>
	</ol>
	</li>
	<li>Synthesis of DNA strands
	<ol>
		<li>Use the connected computer to enter the DNA sequence and coupling method, following the prompts from the manufacturer&#39;s protocol. The coupling step of the cU building block takes 168 sec with 7 pulses (each pulse are 16 &#181;l) compared to 40 sec with 7 pulses for a conventional phosphoramidite as building blocks.</li>
		<li>Mount the column containing the CPG (controlled pore glass) as solid phase that is modified with 1 &#181;mol of the first base (DNA synthesis is performed from 3&#39; to 5&#39;) into the synthesizer.</li>
		<li>Start the synthesis on the DNA synthesizer11.</li>
	</ol>
	</li>
	<li>Workup of synthesized DNA strands
	<ol>
		<li>Dry the columns with the synthesized DNA strands under vacuum O/N.</li>
		<li>Use a pincer to open the column and release the CPG into a reaction vial. Add 700 &#181;l of concentrated aqueous ammonia solution to deprotect the oligonucleotide and release the DNA strand from the CPG.</li>
		<li>Close the vial and apply a security lid onto the reaction vessel to prevent it from bursting, and heat to 50 &#176;C for 18 hr.</li>
		<li>Remove ammonia by centrifugation under reduced pressure (100 mbar) at 30 &#176;C for 30 min.</li>
		<li>Start filtration from CPG: Centrifuge vessel at 11,000 x g for 3 min. Take supernatant and transfer it into a centrifugal device with a pore size of 0.45 &#181;m. Centrifuge at 1,000 x g for 4 min.</li>
		<li>Meanwhile add 300 &#181;l of double distilled water to the CPG, vortex for 20 sec and centrifuge at 11,000 x g for 3 min. Transfer the supernatant into the centrifugal device and centrifuge at 1,000 x g for 4 min. Repeat this washing procedure twice.</li>
		<li>Remove the water from the combined aqueous solutions by centrifuging at 0.1 mbar and 25 &#176;C O/N.</li>
	</ol>
	</li>
</ol>
3. &#34;Clicking&#34; Procedure
<ol>
	<li>Add 50 &#181;l doubly distilled water, 25 &#181;l of a sodium ascorbate solution (0.4 M in water), 34 &#181;l tris-[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (0.1 M in DMSO/tBuOH 3:1), 114 &#181;l of the azide (0.01 M in DMSO/tBuOH 3:1) and 17 &#181;l of a tetrakis(acetonitrile) copper(I)hexafluorophosphate solution (0.1 M in DMSO/tBuOH 3:1) to the lyophilized alkyne-modified DNA sample.</li>
	<li>Incubate the sample at 60 &#176;C for 1.5 hr.</li>
	<li>Cool to RT.</li>
	<li>Add 150 &#181;l Na2EDTA (0.05 M in water) and 450 &#181;l sodium acetate (0.3 M in water) to the DNA sample and transfer to a 10 ml tube.</li>
	<li>Add 10 ml ethanol (100%) and keep at -32 &#176;C for 16 hr.</li>
	<li>Centrifuge vessel for 15 min at 1,000 x g and remove supernatant.</li>
	<li>Wash DNA-pellet with 2 ml cold ethanol (80%).</li>
	<li>Dry pellet under reduced pressure.</li>
</ol>
4. HPLC Purification
Note: Before separating the DNA strands make sure that the HPLC is working properly, enough solvent is available and the column is clean. Rinse column with the starting concentration of buffer/acetonitrile.
<ol>
	<li>Dissolve the crude DNA-pellet in 250 &#181;l of doubly distilled water and transfer to an HPLC vial.</li>
	<li>Start HPLC run of 45 min with a gradient of 0% acetonitrile to 15% of acetonitrile for dye 1 and a gradient of 0% acetonitrile to 17% of acetonitrile for dye 2. Monitor the run by checking 260 nm (DNA absorption) and 459 nm (dye 1) or 542 nm (dye 2). Collect those fractions that show absorption in both wavelength channels.</li>
	<li>Check the collected fractions for the right mass by matrix assisted laser desorption ionization (MALDI) mass spectrometry using a matrix consisting of 3-hydroxypicolinic acid (3HPA) and diammoniumhydrogencitrate.
	<ol>
		<li>Prepare the matrix by mixing 900 &#181;l of a saturated 3-HPA solution in 1:1-mixture of acetonitrile and water with 100 &#181;l of a diammoniumhydrogencitrate solution (100 g/L) in water.</li>
		<li>Pipette 1-2 &#181;l of the DNA probe on the target and let it dry on air.</li>
		<li>Add 0.2 &#181;l of the 3-HPA/diammoniumhydrogencitrate matrix to the sample and mix until it crystallizes.</li>
		<li>Perform MALDI mass spectrometry.</li>
		<li>Combine those HPLC fractions that exhibit the right mass.</li>
	</ol>
	</li>
</ol>
5. Determination of Concentration
Note: The concentration is determined by measuring the absorption at 260 nm using a UV/Vis spectrophotometer, based on the extinction coefficients (&#949;260) of the DNA bases and the dye.
<ol>
	<li>Dissolve the DNA in 100 - 200 &#181;l doubly distilled water.</li>
	<li>Apply 1 &#181;l of the DNA solution on the UV/Vis spectrophotometer.</li>
	<li>Measure the absorption at 260 nm. Repeat three times.</li>
	<li>Take the average value to calculate the concentration of the solution. For the calculation of the extinction coefficients, use &#949;260nm of the four natural bases12 and the &#949;260nm of the purchased building block cU (considered as natural uridine)12 to obtain the &#949;260nm of the whole DNA strand. For dye 1, use &#949;260nm = 10,200 L*mol-1*cm-1 and for dye 2, use &#949;260nm = 13,100 L*mol-1*cm-1. Calculate the concentration of the solution following Lambert-Beers&#39; law based on the extinction coefficients and measured absorbances.13</li>
</ol>
6. Sample Preparation and Spectroscopy
<ol>
	<li>Preparation of single strand samples
	<ol>
		<li>Prepare separately 1 ml of 2.5 &#181;M solutions of each of the modified DNA1 and DNA2 in 200 mM NaCl, 50 mM NaPi buffer, pH = 7.</li>
	</ol>
	</li>
	<li>Preparation of double strand samples
	<ol>
		<li>Prepare 1 ml of a solution containing 2.5 &#181;M of DNA1 and 2.5 &#181;M of DNA2 in 200 mM NaCl, 50 mM NaPi buffer, pH = 7 in a reaction vessel.</li>
		<li>Secure the lid with a safety cap and heat to 90 &#176;C for 10 min. Then turn off heating and allow to cool down to RT O/N.</li>
	</ol>
	</li>
	<li>Absorption spectroscopy 
	Note: To determine the excitation wavelength for the fluorescence spectroscopy absorption spectra are recorded.
	<ol>
		<li>Single strand measurements
		<ol>
			<li>Record a blank measurement containing only 200 mM NaCl, 50 mM NaPi buffer, pH = 7.</li>
			<li>Transfer the prepared solution of DNA1 into a 1 cm quartz glass cuvette and record the absorption. Correct the measurement against the blank data. Determine the maximum value of dye absorption.</li>
			<li>Repeat for DNA2.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Fluorescence spectroscopy
	<ol>
		<li>Single strand measurements
		<ol>
			<li>Record a blank measurement containing only 200 mM NaCl, 50 mM NaPi buffer, pH = 7; using the excitation wavelength of dye 1.</li>
			<li>Transfer the DNA1 solution into a 1 cm quartz glass cuvette.</li>
			<li>Record the fluorescence spectrum.</li>
		</ol>
		</li>
		<li>Double strand measurement
		<ol>
			<li>Transfer the double strand solution into a quartz glass cuvette.</li>
			<li>Record the fluorescence spectrum using the excitation wavelength of dye 1.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Visualization experiments 
	Caution: Appropriate eye protection should be worn to avoid UV damage to the eyes!
	<ol>
		<li>To gain a better understanding of what the recorded spectra are showing, irradiate the cuvettes with handheld UV-lamps (Figure 5). Observe the change in fluorescence color from the single to the double stranded DNA.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

This protocol shows the complete procedure to label DNA post-synthetically via CuAAC by azide-modified fluorescent dyes. This includes the synthesis of the dyes and the alkyne-modified DNA as well as the labeling procedure.
The synthesis of the dyes follows four steps. All products can be obtained by a rather simple precipitation due to their positive charge and no time consuming column chromatography is needed. The introduction of the azide functionalities before the central coupling steps sh...

Molecular imaging represents a fundamental technique for understanding biological processes within living cells.1-3 The development of fluorescent nucleic acid based probes for such chemical-biological applications has become an expanding research field. These fluorescent probes need to meet a few requirements to become suitable tools for cell imaging. Firstly, the applied dyes should exhibit fluorescence with high quantum yields, large Stokes&#39; shifts and, most importantly, high photostabilities to allow long-term in vivo imaging. And secondly, they should show a reliable fluorescence readout. Conventional chromophore-quencher-systems are based on the readout of a single fluorescence color by simple changes in fluorescence intensities.4 This approach bears the risk of false positive or false negative results due to autofluorescence of intracellular components or low signal-to-noise ratios due to undesired quenching by other components.4
We recently reported on the concept of &#34;DNA traffic lights&#34; that show dual fluorescence color readouts by using two different chromophores.5-6 The concept is based on the energy transfer (ET) from the donor dye to the acceptor dye which changes the fluorescence color (see Figure 1). This allows a more reliable readout and thereby provides a powerful tool for fluorescent imaging probes. Labelling of oligonucleotides with fluorescent dyes can be achieved by two different approaches. Dyes can be incorporated during the chemical DNA synthesis on a solid phase by using correspondingly modified phosphoramidite building blocks.7 This method is limited to dyes that are stable under standard phosphoramidite and deprotection conditions. As an alternative, post-synthetic modification methodologies were established in oligonucleotide chemistry. Here, we demonstrate the synthesis of one of our new photostable energy transfer pairs8,9 and the post-synthetic labelling of DNA by using copper-catalyzed 1,3-cycloaddition between azides and alkynes (CuAAC).10

<table border="0" cellpadding="0" cellspacing="0" width="885">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:409px;">synthesis</td>
			<td style="width:180px;"></td>
			<td style="width:132px;"></td>
			<td style="width:164px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">4-Picoline</td>
			<td>Sigma Aldrich</td>
			<td>239615</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">1,3-Diiodopropane</td>
			<td>Sigma Aldrich</td>
			<td>238414</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Acetonitrile</td>
			<td>Fisher Scientific</td>
			<td>10660131</td>
			<td>HPLC grade</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ethyl acetate</td>
			<td>Fisher Scientific</td>
			<td>10456870</td>
			<td>technical grade</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium azide</td>
			<td>Sigma Aldrich</td>
			<td>71290</td>
			<td>p.a. grade</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dichloromethane</td>
			<td>Fisher Scientific</td>
			<td>10626642</td>
			<td>technical grade</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Indole-3-carboxaldehyde; 98%</td>
			<td>ABCR</td>
			<td>AB112969</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Potassium carbonate, 99+%</td>
			<td>Acros</td>
			<td>424081000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">dimethylcarbonate</td>
			<td>Sigma Aldrich</td>
			<td>517127</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">N,N-Dimethylformamide, 99.8%, Extra Dry over Molecular Sieve</td>
			<td>Acros</td>
			<td>348435000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium sulfate</td>
			<td>Bernd Kraft</td>
			<td>12623.46</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ethanol, 99.5%</td>
			<td>Acros</td>
			<td>397690010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Piperidine, 99%</td>
			<td>Acros</td>
			<td>147181000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Diethylether</td>
			<td>Fisher Scientific</td>
			<td>10407830</td>
			<td>technical grade</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">2-Phenylindole-3-carboxaldehyde; 97%</td>
			<td>ABCR</td>
			<td>AB125050</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">4-Methylquinoline</td>
			<td>ABCR</td>
			<td>AB117222</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DNA synthesis</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Expedite 8909 Nucleic Acid Synthesizer</td>
			<td>Applied Biosystems</td>
			<td>&#160;-</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMT-dA(bz) Phosphoramidite</td>
			<td>Sigma Aldrich</td>
			<td>A111081</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMT-dT Phosphoramidite</td>
			<td>Sigma Aldrich</td>
			<td>T111081</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMT-dG(dmf) Phosphoramidite</td>
			<td>Sigma Aldrich</td>
			<td>G11508</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">DMT-dC(bz) Phosphoramidite</td>
			<td>Sigma Aldrich</td>
			<td>C11108</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Amidite Diluent for DNA synthesis</td>
			<td>Sigma Aldrich</td>
			<td>L010010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Ultrapure Acetonitrile for DNA synthesis</td>
			<td>Sigma Aldrich</td>
			<td>L010400</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cap A</td>
			<td>Sigma Aldrich</td>
			<td>L840000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cap B</td>
			<td>Sigma Aldrich</td>
			<td>L850000</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">CPG dT Column 1.0 &#181;mole</td>
			<td>Proligo Reagents</td>
			<td>T461010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">CPG dA(bz) Column 1.0 &#181;mole</td>
			<td>Proligo Reagents</td>
			<td>A461010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">CPG dG(ib) Column 1.0 &#181;mole</td>
			<td>Proligo Reagents</td>
			<td>G461010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">CPG dC(bz) Column 1.0 &#181;mole</td>
			<td>Proligo Reagents</td>
			<td>C461010</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">ammonia (aqueous solution)&#160;</td>
			<td>Fluka Analytical</td>
			<td>318612</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">centrifugal devices nanosep 0.45 &#181;m</td>
			<td>Pall</td>
			<td>ODGHPC34</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">5-(Benzylthio)-1H-tetrazole (Activator)</td>
			<td>Sigma Aldrich</td>
			<td>75666</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">2&#39;-O-propargyl deoxyuridinephosphoramidite</td>
			<td>Chem Genes</td>
			<td>ANP-7754</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">workup</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">vacuum concentrator</td>
			<td>Christ</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">clicking procedure</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tetrakis(acetonitrile)copper(I) hexafluorophosphate</td>
			<td>Sigma Aldrich</td>
			<td>346276</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Sodium acetate</td>
			<td>Sigma Aldrich</td>
			<td>S2889</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">(+)-Sodium L-ascorbate</td>
			<td>Sigma Aldrich</td>
			<td>A7631</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EDTA disodium salt</td>
			<td>Sigma Aldrich</td>
			<td>E5134</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">TBTA-ligand</td>
			<td>&#160;-</td>
			<td>&#160;-</td>
			<td>synthesized according to a literature procedure [1]</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">HPLC</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">HPLC-system</td>
			<td>Shimadzu</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">MALDI-Biflex-IV spectrometer</td>
			<td>Bruker Daltonics</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">LC-318 C18 column</td>
			<td>Supelcosil via Sigma Aldrich</td>
			<td>58368</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">determination of concentration</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">ND 1000 Spectrophotometer</td>
			<td>nanodrop</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">sample preparation and spectroscopy</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cary 100 Bio</td>
			<td>Varian</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Fluoromax-3 fluorimeter</td>
			<td>Jobin-Yvon</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;"></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:409px;">[1] R. Chan Timothy, R. Hilgraf, K. B. Sharpless, V. Fokin Valery, Org Lett 2004, 6, 2853-2855.</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

synthesis of wavelength-shifting dna hybridization probes by using photostable cyanine dyes

In this protocol, we demonstrate a method for the synthesis of 2&#39;-alkyne modified deoxyribonucleic acid (DNA) strands by automated solid phase synthesis using standard phosphoramidite chemistry. Oligonucleotides are post-synthetically labeled by two new photostable cyanine dyes using copper-catalyzed click-chemistry. The synthesis of both donor and acceptor dye is described and is performed in three consecutive steps. With the DNA as the surrounding architecture, these two dyes undergo an energy transfer when they are brought into close proximity by hybridization. Therefore, annealing of two single stranded DNA strands is visualized by a change of fluorescence color. This color change is characterized by fluorescence spectroscopy but can also be directly observed by using a handheld ultraviolet (UV) lamp. The concept of a dual fluorescence color readout makes these oligonucleotide probes excellent tools for molecular imaging especially when the described photostable dyes are used. Thereby, photobleaching of the imaging probes is prevented, and biological processes can be observed in real time for a longer time period.

Absorption and fluorescence spectra of the single and double stranded DNA are recorded as shown in Figure 4.
The recorded absorption spectra (Figure 4 right) show absorption maxima &#955;max at 465 nm for single-stranded DNA1 (dye 1) and 546 nm for single-stranded DNA2 (dye 2). The annealed DNA1_2 (dye 1 &#38; dye 2) shows maxima at both 469 nm and 567 nm. Both absorption maxima show ...

Watch this Scientific Journal Video about Synthesis of Wavelength-shifting DNA Hybridization Probes by Using Photostable Cyanine Dyes at JoVE.com

Synthesis of Wavelength-shifting DNA Hybridization Probes by Using Photostable Cyanine Dyes

Photostable cyanine dyes are attached to oligonucleotides to monitor hybridization by energy transfer.

In this protocol, we demonstrate a method for the synthesis of 2&#39;-alkyne modified deoxyribonucleic acid (DNA) strands by automated solid phase ...

synthesis-wavelength-shifting-dna-hybridization-probes-using

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11.0K Views.  Karlsruhe Institute of Technology. Photostable cyanine dyes are attached to oligonucleotides to monitor hybridization by energy transfer.

Video: Synthesis of Wavelength-shifting DNA Hybridization Probes by Using Photostable Cyanine Dyes

Selective Labelling of Cell-surface Proteins using CyDye DIGE Fluor Minimal Dyes

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of almost all intracellular signaling. Moreover, they play an important role in environmental adaptation and drug treatment, and are often involved in disease pathogenesis and pathology (1). Protein-protein interactions are intrinsic to signaling pathways, and to gain more insight in these complex biological processes, sensitive and reliable methods are needed for studying cell surface proteins. Two-dimensional (2-D) electrophoresis is used extensively for detection of biomarkers and other targets in complex protein samples to study differential changes. Cell surface proteins, partly due to their low abundance (1 2% of cellular proteins), are difficult to detect in a 2-D gel without fractionation or some other type of enrichment. They are also often poorly represented in 2-D gels due to their hydrophobic nature and high molecular weight (2). In this study, we present a new protocol for intact cells using CyDye  DIGE Fluor minimal dyes for specific labeling and detection of this important group of proteins. The results showed specific labeling of a large number of cell surface proteins with minimal labeling of intracellular proteins. This protocol is rapid, simple to use, and all three CyDye DIGE Fluor minimal dyes (Cy 2, Cy 3 and Cy 5) can be used to label cell-surface proteins. These features allow for multiplexing using the 2-D Fluorescence Difference Gel Electrophoresis (2-D DIGE) with Ettan  DIGE technology and analysis of protein expression changes using DeCyder  2-D Differential Analysis Software. The level of cell-surface proteins was followed during serum starvation of CHO cells for various lengths of time (see Table 1). Small changes in abundance were detected with high accuracy, and results are supported by defined statistical methods.

A simple and specific method was demonstrated for fluorescent labeling and enhanced detection of cell surface proteins without a fractionation step. Differential abundance in cell surface proteins was analyzed using two-dimensional (2-D) electrophoresis and Ettan&#x2122; DIGE technology.

Surface  proteins are central to the cell's ability to react to its environment and to interact with neighboring cells. They are known to be inducers of ...

Generation of RNA/DNA Hybrids in Genomic DNA by Transformation using RNA-containing Oligonucleotides

Synthetic short nucleic acid polymers, oligonucleotides (oligos), are the most functional and widespread tools of molecular biology. Oligos can be produced to contain any desired DNA or RNA sequence and can be prepared to include a wide variety of base and sugar modifications. Moreover, oligos can be designed to mimic specific nucleic acid alterations and thus, can serve as important tools to investigate effects of DNA damage and mechanisms of repair. We found that Thermo Scientific Dharmacon RNA-containing oligos with a length between 50 and 80 nucleotides can be particularly suitable to study, in vivo, functions and consequences of chromosomal RNA/DNA hybrids and of ribonucleotides embedded into DNA. RNA/DNA hybrids can readily form during DNA replication, repair and transcription, however, very little is known about the stability of RNA/DNA hybrids in cells and to which extent these hybrids can affect the genetic integrity of cells. RNA-containing oligos, therefore, represent a perfect vector to introduce ribonucleotides into chromosomal DNA and generate RNA/DNA hybrids of chosen length and base composition. Here we present the protocol for the incorporation of ribonucleotides into the genome of the eukaryotic model system yeast /Saccharomyces cerevisiae/. Yet, our lab has utilized Thermo Scientific Dharmacon RNA-containing oligos to generate RNA/DNA hybrids at the chromosomal level in different cell systems, from bacteria to human cells.

This work shows how to form an RNA/DNA hybrid at the chromosomal level and reveal transfer of genetic information from RNA to genomic DNA in yeast cells.

Synthetic short nucleic acid polymers, oligonucleotides (oligos), are the most functional and widespread tools of molecular biology. Oligos can be ...

DNA Microarrays: Sample Quality Control, Array Hybridization and Scanning

Microarray expression profiling of the nervous system provides a powerful approach to identifying gene activities in different stages of development, different physiological or pathological states, response to therapy, and, in general, any condition that is being experimentally tested1. Expression profiling of neural tissues requires isolation of high quality RNA, amplification of the isolated RNA and hybridization to DNA microarrays. In this article we describe protocols for reproducible microarray experiments from brain tumor tissue2. We will start by performing a quality control analysis of isolated RNA samples with Agilent's 2100 Bioanalyzer "lab-on-a-chip" technology. High quality RNA samples are critical for the success of any microarray experiment, and the 2100 Bioanalyzer provides a quick, quantitative measurement of the sample quality. RNA samples are then amplified and labeled by performing reverse transcription to obtain cDNA, followed by in vitro transcription in the presence of labeled nucleotides to produce labeled cRNA. By using a dual-color labeling kit, we will label our experimental sample with Cy3 and a reference sample with Cy5. Both samples will then be combined and hybridized to Agilent's 4x44 K arrays. Dual-color arrays offer the advantage of a direct comparison between two RNA samples, thereby increasing the accuracy of the measurements, in particular for small changes in expression levels, because the two RNA samples are hybridized competitively to a single microarray. The arrays will be scanned at the two corresponding wavelengths, and the ratio of Cy3 to Cy5 signal for each feature will be used as a direct measurement of the relative abundance of the corresponding mRNA. This analysis identifies genes that are differentially expressed in response to the experimental conditions being tested.

We demonstrate the use of DNA microarrays for expression profiling of the nervous system. We describe RNA quality control, sample labeling, and array hybridization and scanning.

Microarray expression profiling of the nervous system provides a powerful approach to identifying gene activities in different stages of development, ...

Determination of Lipid Raft Partitioning of Fluorescently-tagged Probes in Living Cells by Fluorescence Correlation Spectroscopy (FCS)

In the past fifteen years the notion that cell membranes are not homogenous and rely on microdomains to exert their functions has become widely accepted. Lipid rafts are membrane microdomains enriched in cholesterol and sphingolipids. They play a role in cellular physiological processes such as signalling, and trafficking1,2 but are also thought to be key players in several diseases including viral or bacterial infections and neurodegenerative diseases3.
Yet their existence is still a matter of controversy4,5. Indeed, lipid raft size has been estimated to be around 20 nm6, far under the resolution limit of conventional microscopy (around 200 nm), thus precluding their direct imaging. Up to now, the main techniques used to assess the partition of proteins of interest inside lipid rafts were Detergent Resistant Membranes (DRMs) isolation and co-patching with antibodies. Though widely used because of their rather easy implementation, these techniques were prone to artefacts and thus criticized7,8. Technical improvements were therefore necessary to overcome these artefacts and to be able to probe lipid rafts partition in living cells.
Here we present a method for the sensitive analysis of lipid rafts partition of fluorescently-tagged proteins or lipids in the plasma membrane of living cells. This method, termed Fluorescence Correlation Spectroscopy (FCS), relies on the disparity in diffusion times of fluorescent probes located inside or outside of lipid rafts. In fact, as evidenced in both artificial membranes and cell cultures, probes would diffuse much faster outside than inside dense lipid rafts9,10. To determine diffusion times, minute fluorescence fluctuations are measured as a function of time in a focal volume (approximately 1 femtoliter), located at the plasma membrane of cells with a confocal microscope (Fig. 1). The auto-correlation curves can then be drawn from these fluctuations and fitted with appropriate mathematical diffusion models11.
FCS can be used to determine the lipid raft partitioning of various probes, as long as they are fluorescently tagged. Fluorescent tagging can be achieved by expression of fluorescent fusion proteins or by binding of fluorescent ligands. Moreover, FCS can be used not only in artificial membranes and cell lines but also in primary cultures, as described recently12. It can also be used to follow the dynamics of lipid raft partitioning after drug addition or membrane lipid composition change12.

Determination of Lipid Raft Partitioning of Fluorescently-tagged Probes in Living Cells by Fluorescence Correlation Spectroscopy FCS

A technique to probe the lipid raft partitioning of fluorescent proteins at the plasma membrane of living cells is described. It takes advantage of the disparity in diffusion times of proteins located inside or outside of lipid rafts. Acquisition can be performed dynamically in control conditions or after drug addition.

In the past fifteen years the notion that cell membranes are not homogenous and rely on microdomains to exert their functions has become widely accepted. ...

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

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

Detection of Viral RNA by Fluorescence in situ Hybridization FISH

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

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

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

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

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

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

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

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 ...

Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3&#8217;-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.

Ratiometric bimolecular beacons (RBMBs) can be used to image single engineered RNA transcripts in living cells. Here, we describe the preparation and purification of RBMBs, delivery of RBMBs into cells by microporation and fluorescent imaging of single RNA transcripts in real-time.

The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the ...

Studying DNA Looping by Single-Molecule FRET

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into nucleosomes. Thermodynamics of dsDNA bending has been studied by a method called cyclization which relies on DNA ligase to covalently join short sticky ends of a dsDNA. However, ligation efficiency can be affected by many factors that are not related to dsDNA looping such as the DNA structure surrounding the joined sticky ends, and ligase can also affect the apparent looping rate through mechanisms such as nonspecific binding. Here, we show how to measure dsDNA looping kinetics without ligase by detecting transient DNA loop formation by FRET (Fluorescence Resonance Energy Transfer). dsDNA molecules are constructed using a simple PCR-based protocol with a FRET pair and a biotin linker. The looping probability density known as the J factor is extracted from the looping rate and the annealing rate between two disconnected sticky ends. By testing two dsDNAs with different intrinsic curvatures, we show that the J factor is sensitive to the intrinsic shape of the dsDNA.

This study presents a detailed experimental procedure to measure looping dynamics of double-stranded DNA using single-molecule Fluorescence Resonance Energy Transfer (FRET). The protocol also describes how to extract the looping probability density called the J factor.

Bending of double-stranded DNA (dsDNA) is associated with many important biological processes such as DNA-protein recognition and DNA packaging into ...

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.

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.