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><tbody><tr><td colspan="4">&lt;strong&gt;synthesis&lt;/strong&gt;</td></tr><tr><td>4-Picoline</td><td>Sigma Aldrich</td><td>239615</td><td></td></tr><tr><td>1,3-Diiodopropane</td><td>Sigma Aldrich</td><td>238414</td><td></td></tr><tr><td>Acetonitrile</td><td>Fisher Scientific</td><td>10660131</td><td>HPLC grade</td></tr><tr><td>Ethyl acetate</td><td>Fisher Scientific</td><td>10456870</td><td>technical grade</td></tr><tr><td>Sodium azide</td><td>Sigma Aldrich</td><td>71290</td><td>p.a. grade</td></tr><tr><td>Dichloromethane</td><td>Fisher Scientific</td><td>10626642</td><td>technical grade</td></tr><tr><td>Indole-3-carboxaldehyde; 98%</td><td>ABCR</td><td>AB112969</td><td></td></tr><tr><td>Potassium carbonate, 99+%</td><td>Acros</td><td>424081000</td><td></td></tr><tr><td>dimethylcarbonate</td><td>Sigma Aldrich</td><td>517127</td><td></td></tr><tr><td>&lt;em&gt;N,N&lt;/em&gt;-Dimethylformamide, 99.8%, Extra Dry over Molecular Sieve</td><td>Acros</td><td>348435000</td><td></td></tr><tr><td>Sodium sulfate</td><td>Bernd Kraft</td><td>12623.46</td><td></td></tr><tr><td>Ethanol, 99.5%</td><td>Acros</td><td>397690010</td><td></td></tr><tr><td>Piperidine, 99%</td><td>Acros</td><td>147181000</td><td></td></tr><tr><td>Diethylether</td><td>Fisher Scientific</td><td>10407830</td><td>technical grade</td></tr><tr><td>2-Phenylindole-3-carboxaldehyde; 97%</td><td>ABCR</td><td>AB125050</td><td></td></tr><tr><td>4-Methylquinoline</td><td>ABCR</td><td>AB117222</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;DNA synthesis&lt;/strong&gt;</td></tr><tr><td>Expedite 8909 Nucleic Acid Synthesizer</td><td>Applied Biosystems</td><td>&amp;nbsp;-</td><td></td></tr><tr><td>DMT-dA(bz) Phosphoramidite</td><td>Sigma Aldrich</td><td>A111081</td><td></td></tr><tr><td>DMT-dT Phosphoramidite</td><td>Sigma Aldrich</td><td>T111081</td><td></td></tr><tr><td>DMT-dG(dmf) Phosphoramidite</td><td>Sigma Aldrich</td><td>G11508</td><td></td></tr><tr><td>DMT-dC(bz) Phosphoramidite</td><td>Sigma Aldrich</td><td>C11108</td><td></td></tr><tr><td>Amidite Diluent for DNA synthesis</td><td>Sigma Aldrich</td><td>L010010</td><td></td></tr><tr><td>Ultrapure Acetonitrile for DNA synthesis</td><td>Sigma Aldrich</td><td>L010400</td><td></td></tr><tr><td>Cap A</td><td>Sigma Aldrich</td><td>L840000</td><td></td></tr><tr><td>Cap B</td><td>Sigma Aldrich</td><td>L850000</td><td></td></tr><tr><td>CPG dT Column 1.0&amp;nbsp;&amp;micro;mole</td><td>Proligo Reagents</td><td>T461010</td><td></td></tr><tr><td>CPG dA(bz) Column 1.0&amp;nbsp;&amp;micro;mole</td><td>Proligo Reagents</td><td>A461010</td><td></td></tr><tr><td>CPG dG(ib) Column 1.0&amp;nbsp;&amp;micro;mole</td><td>Proligo Reagents</td><td>G461010</td><td></td></tr><tr><td>CPG dC(bz) Column 1.0&amp;nbsp;&amp;micro;mole</td><td>Proligo Reagents</td><td>C461010</td><td></td></tr><tr><td>ammonia (aqueous solution)&amp;nbsp;</td><td>Fluka Analytical</td><td>318612</td><td></td></tr><tr><td>centrifugal devices nanosep 0.45&amp;nbsp;&amp;micro;m</td><td>Pall</td><td>ODGHPC34</td><td></td></tr><tr><td>5-(Benzylthio)-1H-tetrazole (Activator)</td><td>Sigma Aldrich</td><td>75666</td><td></td></tr><tr><td>2&#039;-O-propargyl deoxyuridinephosphoramidite</td><td>Chem Genes</td><td>ANP-7754</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;workup&lt;/strong&gt;</td></tr><tr><td>vacuum concentrator</td><td>Christ</td><td></td><td></td></tr><tr><td>&lt;strong&gt;clicking procedure&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Tetrakis(acetonitrile)copper(I) hexafluorophosphate</td><td>Sigma Aldrich</td><td>346276</td><td></td></tr><tr><td>Sodium acetate</td><td>Sigma Aldrich</td><td>S2889</td><td></td></tr><tr><td>(+)-Sodium L-ascorbate</td><td>Sigma Aldrich</td><td>A7631</td><td></td></tr><tr><td>EDTA disodium salt</td><td>Sigma Aldrich</td><td>E5134</td><td></td></tr><tr><td>TBTA-ligand</td><td>&amp;nbsp;-</td><td>&amp;nbsp;-</td><td>synthesized according to a literature procedure&lt;sup&gt;1&lt;/sup&gt;</td></tr><tr><td colspan="4">&lt;strong&gt;HPLC&lt;/strong&gt;</td></tr><tr><td>HPLC-system</td><td>Shimadzu</td><td></td><td></td></tr><tr><td>MALDI-Biflex-IV spectrometer</td><td>Bruker Daltonics</td><td></td><td></td></tr><tr><td>LC-318 C18 column</td><td>Supelcosil via Sigma Aldrich</td><td>58368</td><td></td></tr><tr><td colspan="4">&lt;strong&gt;determination of concentration&lt;/strong&gt;</td></tr><tr><td>ND 1000 Spectrophotometer</td><td>nanodrop</td><td></td><td></td></tr><tr><td colspan="4">&lt;strong&gt;sample preparation and spectroscopy&lt;/strong&gt;</td></tr><tr><td>Cary 100 Bio</td><td>Varian</td><td></td><td></td></tr><tr><td>Fluoromax-3 fluorimeter</td><td>Jobin-Yvon</td><td></td><td></td></tr><tr><td colspan="4">&lt;sup&gt;1&lt;/sup&gt; R. Chan Timothy, R. Hilgraf, K. B. Sharpless, V. Fokin Valery, &lt;em&gt;Org Lett&lt;/em&gt; 2004, &lt;strong&gt;6&lt;/strong&gt;, 2853-2855.</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 ...

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11.1K 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

An Optimized Protocol for Electrophoretic Mobility Shift Assay Using Infrared Fluorescent Dye-labeled Oligonucleotides

Electrophoretic Mobility Shift Assays (EMSA) are an instrumental tool to characterize the interactions between proteins and their target DNA sequences. Radioactivity has been the predominant method of DNA labeling in EMSAs. However, recent advances in fluorescent dyes and scanning methods have prompted the use of fluorescent tagging of DNA as an alternative to radioactivity for the advantages of easy handling, saving time, reducing cost, and improving safety. We have recently used fluorescent EMSA (fEMSA) to successfully address an important biological question. Our fEMSA analysis provides mechanistic insight into the effect of a missense mutation, G73E, in the highly conserved HMG transcription factor SOX-2 on olfactory neuron type diversification. We found that mutant SOX-2G73E protein alters specific DNA binding activity, thereby causing olfactory neuron identity transformation. Here, we present an optimized and cost-effective step-by-step protocol for fEMSA using infrared fluorescent dye-labeled oligonucleotides containing the LIM-4/SOX-2 adjacent target sites and purified SOX-2 proteins (WT and mutant SOX-2G73E proteins) as a biological example.

We describe here an optimized protocol of fluorescent Electrophoretic Mobility Shift Assays (fEMSA) using purified SOX-2 proteins together with infrared fluorescent dye-labeled DNA probes as a case study to tackle an important biological question.

Electrophoretic Mobility Shift Assays (EMSA) are an instrumental tool to characterize the interactions between proteins and their target DNA sequences. ...

Biochemistry

Technical Demonstration of Whole Genome Array Comparative Genomic Hybridization

Array comparative genomic hybridization (array CGH) is a method for detecting gains and losses of DNA segments or gene dosage in the genome 1. Recent advances in this technology have enabled high resolution comparison of whole genomes for the identification of genetic alterations in cancer and other genetic diseases 2. The Sub-Megabase Resolution Tiling-set array (or SMRT) array is comprised of a set of approximately thirty thousand overlapping bacterial artificial chromosome (BAC) clones that span the human genome in ~100 kilobase pair (kb) segments 2. These BAC targets are individually synthesized and spotted in duplicate on a single glass slide 2-4. Array CGH is based on the principle of competitive hybridization. Sample and reference DNA are differentially labeled with Cyanine-3 and Cyanine-5 fluorescent dyes, and co-hybridized to the array. After an incubation period the unbound samples are washed from the slide and the array is imaged. A freely available custom software package called SeeGH (www.flintbox.ca) is used to process the large volume of data collected - a single experiment generates 53,892 data points. SeeGH visualizes the log2 signal intensity ratio between the 2 samples at each BAC target which is vertically aligned with chromosomal position 5,6. The SMRT array can detect alterations as small as 50 kb in size 7. The SMRT array can detect a variety of DNA rearrangement events including DNA gains, losses, amplifications and homozygous deletions. A unique advantage of the SMRT array is that one can use DNA isolated from formalin fixed paraffin embedded samples. When combined with the low input requirements of unamplified DNA (25-100ng) this allows profiling of precious samples such as those produced by microdissection 7,8. This is attributed to the large size of each BAC hybridization target that allows the binding of sufficient labeled samples to produce signals for detection. Another advantage of this platform is the tolerance of tissue heterogeneity, decreasing the need for tedious tissue microdissection 8. This video protocol is a step-by-step tutorial from labeling the input DNA through to signal acquisition for the whole genome tiling path SMRT array.

This video is a technical demonstration of the hybridization protocol for whole genome tiling path array CGH, which scans the entire human genome using only 25-100 ng of DNA that can be isolated from a variety of sources, including archival formalin fixed material.

Array comparative genomic hybridization (array CGH) is a method for detecting gains and losses of DNA segments or gene dosage in the genome 1.  Recent ...

DNA Electrophoresis Using Thiazole Orange Instead of Ethidium Bromide or Alternative Dyes

DNA gel electrophoresis using agarose is a common tool in molecular biology laboratories, allowing separation of DNA fragments by size. After separation, DNA is visualized by staining. This article demonstrates how to use thiazole orange to stain DNA. Thiazole orange compares favorably to common staining methods, in that it is sensitive, inexpensive, excitable with UV or blue light (to prevent sample damage), and safer than ethidium bromide. Labs already equipped to run DNA electrophoresis experiments using ethidium bromide can generally switch dyes with no additional changes to existing protocols, using UV light for detection. Blue-light detection to avoid sample damage can additionally be achieved with a blue-light source and emission filter. Labs already equipped for blue-light detection can simply switch dyes with no additional changes to existing protocols.

Here, we present a protocol to use thiazole orange for the detection of DNA in gel electrophoresis experiments. The use of thiazole orange allows elimination of ethidium bromide, and fluorescence detection can be achieved with either UV or blue light.

DNA gel electrophoresis using agarose is a common tool in molecular biology laboratories, allowing separation of DNA fragments by size. After separation, ...

Combining QD-FRET and Microfluidics to Monitor DNA Nanocomplex Self-Assembly in Real-Time

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically functional inside the cell, nucleic acid-based therapeutics require an efficient intracellular delivery system. One widely adopted approach is to complex DNA with a gene carrier to form nanocomplexes via electrostatic self-assembly, facilitating cellular uptake of DNA while protecting it against degradation. The challenge lies in the rational design of efficient gene carriers, since premature dissociation or overly stable binding would be detrimental to the cellular uptake and therapeutic efficacy. Nanocomplexes synthesized by bulk mixing showed a diverse range of intracellular unpacking and trafficking behavior, which was attributed to the heterogeneity in size and stability of nanocomplexes. Such heterogeneity hinders the accurate assessment of the self-assembly kinetics and adds to the difficulty in correlating their physical properties to transfection efficiencies or bioactivities. We present a novel convergence of nanophotonics (i.e. QD-FRET) and microfluidics to characterize the real-time kinetics of the nanocomplex self-assembly under laminar flow. QD-FRET provides a highly sensitive indication of the onset of molecular interactions and quantitative measure throughout the synthesis process, whereas microfluidics offers a well-controlled microenvironment to spatially analyze the process with high temporal resolution (~milliseconds). For the model system of polymeric nanocomplexes, two distinct stages in the self-assembly process were captured by this analytic platform. The kinetic aspect of the self-assembly process obtained at the microscale would be particularly valuable for microreactor-based reactions which are relevant to many micro- and nano-scale applications. Further, nanocomplexes may be customized through proper design of microfludic devices, and the resulting QD-FRET polymeric DNA nanocomplexes could be readily applied for establishing structure-function relationships. 

We present a novel and powerful integration of nanophotonics (QD-FRET) and microfluidics to investigate the formation of polyelectrolyte polyplexes, which is expected to provide better control and synthesis of uniform and customizable polyplexes for future nucleic acid-based therapeutics.

Advances in genomics continue to fuel the development of therapeutics that can target pathogenesis at the cellular and molecular level. Typically ...

DNA Tension Probes to Map the Transient Piconewton Receptor Forces by Immune Cells

Mechanical forces transmitted at the junction between two neighboring cells and at the junction between cells and the extracellular matrix are critical for regulating many processes ranging from development to immunology. Therefore, developing the tools to study these forces at the molecular scale is critical. Our group developed a suite of molecular tension sensors to quantify and visualize the forces generated by cells and transmitted to specific ligands. The most sensitive class of molecular tension sensors are comprised of nucleic acid stem-loop hairpins. These sensors use fluorophore-quencher pairs to report on the mechanical extension and unfolding of DNA hairpins under force. One challenge with DNA hairpin tension sensors is that they are reversible with rapid hairpin refolding upon termination of the tension and thus transient forces are difficult to record. In this article, we describe the protocols for preparing DNA tension sensors that can be "locked" and prevented from refolding to enable "storing" of mechanical information. This allows for the recording of highly transient piconewton forces, which can be subsequently "erased" by the addition of complementary nucleic acids that remove the lock. This ability to toggle between real-time tension mapping and mechanical information storing reveals weak, short-lived, and less abundant forces, that are commonly employed by T cells as part of their immune functions.

This paper describes a detailed protocol for using DNA-based tension probes to image the receptor forces applied by immune cells. This approach can map receptor forces &gt;4.7pN in real-time and can integrate forces over time.

Mechanical forces transmitted at the junction between two neighboring cells and at the junction between cells and the extracellular matrix are critical ...

Bioengineering

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 ...

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 ...

Plasmid-derived DNA Strand Displacement Gates for Implementing Chemical Reaction Networks

DNA nanotechnology requires large amounts of highly pure DNA as an engineering material. Plasmid DNA could meet this need since it is replicated with high fidelity, is readily amplified through bacterial culture and can be stored indefinitely in the form of bacterial glycerol stocks. However, the double-stranded nature of plasmid DNA has so far hindered its efficient use for construction of DNA nanostructures or devices that typically contain single-stranded or branched domains. In recent work, it was found that nicked double stranded DNA (ndsDNA) strand displacement gates could be sourced from plasmid DNA. The following is a protocol that details how these ndsDNA gates can be efficiently encoded in plasmids and can be derived from the plasmids through a small number of enzymatic processing steps. Also given is a protocol for testing ndsDNA gates using fluorescence kinetics measurements. NdsDNA gates can be used to implement arbitrary chemical reaction networks (CRNs) and thus provide a pathway towards the use of the CRN formalism as a prescriptive molecular programming language. To demonstrate this technology, a multi-step reaction cascade with catalytic kinetics is constructed. Further it is shown that plasmid-derived components perform better than identical components assembled from synthetic DNA.

This protocol describes a method for deriving DNA strand displacement gates from plasmids and testing them using fluorescence kinetics measurements. Gates can be modularly composed into multi-component systems to approximate the behavior of formal chemical reaction networks (CRN), demonstrating a new use for CRNs as a molecular programming language.

DNA nanotechnology requires large amounts of highly pure DNA as an engineering material. Plasmid DNA could meet this need since it is replicated with high ...

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization

Single-molecule fluorescence in situ hybridization (smFISH) allows for counting the absolute number of mRNAs in individual cells. Here, we describe an application of smFISH to measure the rates of transcription and mRNA degradation in Escherichia coli. As smFISH is based on fixed cells, we perform smFISH at multiple time points during a time-course experiment, i.e., when cells are undergoing synchronized changes upon induction or repression of gene expression. At each time point, sub-regions of an mRNA are spectrally distinguished to probe transcription elongation and premature termination. The outcome of this protocol also allows for analyzing intracellular localization of mRNAs and heterogeneity in mRNA copy numbers among cells. Using this protocol many samples (~50) can be processed within 8 h, like the amount of time needed for just a few samples. We discuss how to apply this protocol to study the transcription and degradation kinetics of different mRNAs in bacterial cells.

Probing mRNA Kinetics in Space and Time in Escherichia coli using Two-Color Single-Molecule Fluorescence In Situ Hybridization

This protocol describes an application of single-molecule fluorescence in situ hybridization (smFISH) to measure the in vivo kinetics of mRNA synthesis and degradation.

Single-molecule fluorescence in situ hybridization (smFISH) allows for counting the absolute number of mRNAs in individual cells. Here, we describe an ...

Single-Molecule Fluorescence Visualization of DNA Polymerase Dynamics at G-Quadruplexes

The ability of proteins&#160;involved in eukaryotic DNA replication to&#160;overcome obstacles &#8212; such as protein and DNA &#39;roadblocks&#39;&#160;&#8212; is critical for ensuring faithful&#160;genome duplication.&#160;G-quadruplexes are higher-order nucleic acid structures that form in guanine-rich regions of DNA and have been shown to act as obstacles, interfering with genomic maintenance pathways. This study introduces a real-time, fluorescence microscopy-based method to observe DNA polymerase interactions with G-quadruplex structures. Short, primed DNA oligonucleotides containing a G-quadruplex were immobilized on functionalized glass coverslips within a microfluidic flow cell. Fluorescently labeled DNA polymerases were introduced, allowing their behavior and stoichiometry to be monitored over time. This approach enabled the observation of polymerase behavior as it was stalled by a G-quadruplex. Specifically, using fluorescently labeled yeast polymerase &#948;, it was found that upon encountering a G-quadruplex, the polymerase undergoes a continuous cycle of binding and unbinding. This single-molecule assay can be adapted to study interactions between various DNA-maintenance proteins and obstacles on the DNA substrate.

This protocol outlines a fluorescence microscopy-based single-molecule DNA replication assay, enabling real-time visualization of interactions between DNA polymerases and obstacles such as G-quadruplex structures.

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

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