Name: determining if dna stained with a cyanine dye can be digested with restriction enzymes
Uploaded: 2018-02-02T13:30:00.000+00:00
Duration: 6 min 58 s
Description: Watch this Scientific Journal Video about Determining if DNA Stained with a Cyanine Dye Can Be Digested with Restriction Enzymes at JoVE.com

This research was funded by the National Institute for General Medical Science (NIGMS) (5605100122001), a component of the National Institutes of Health (NIH), as well as University of Nebraska at Kearney (UNK) Summer Student Research Program (SSRP), and UNK Undergraduate Research Fellowship (URF).

1. Preparation of Dyes, Buffers and Agarose Gel
<ol>
	<li>Prepare the following solutions for staining DNA or the digestion of stained DNA.
	<ol>
		<li>Prepare 1x TE (Tris-HCl and ethylenediaminetetraacetic acid; EDTA) buffer using 10 mM Tris-HCl and 1 mM EDTA in a graduated cylinder or volumetric flask. Store at room temperature (18 - 25 &#176;C).</li>
		<li>Make aliquots of each dye: TOTO-1, YOYO-1, POPO-1, BOBO-1, TOTO-3, YOYO-3, POPO-3, BOBO-3 (Table 1). Pipette 4 &#181;L of 1 mM dye into a dark amber or black 1.5 mL tube and add 36 &#181;L of 1x TE, mix using the pipette; this concentration makes a 100 &#181;M dye solution (40 &#181;L total). Complete this step for each dye. Store at 4 &#176;C. 
		NOTE: Avoid exposing dye to light to prevent photobleaching or degradation of the dye.</li>
		<li>Prepare buffer 1 using 10 mM Bis-Tris-Propane-HCl, 10 mM MgCl2, and 100 &#181;g/mL bovine serum albumin (BSA). Prepare buffer 2 using 50 mM NaCl, 10 mM Tris-HCl, 10 mM MgCl2, and 100 &#181;g/mL BSA. Prepare buffer 3 using 50 mM Potassium Acetate, 20 mM Tris-acetate, 10 mM magnesium acetate, and 100 &#181;g/mL BSA. Each enzyme will require one of these specific buffers to function.</li>
	</ol>
	</li>
	<li>Prepare the following solutions for gel electrophoresis.
	<ol>
		<li>Make a 6x loading dye by combining 0.25% bromophenol blue, 0.25% xylene cyanol, 15% Ficoll in dH2O. Store at room temperature.</li>
		<li>To make a 0.7% agarose gel, weigh out 0.07 g of high gelling temperature (HGT) agarose in an Erlenmeyer flask, add 100 mL of 1x TAE buffer and place an inverted beaker on top of the flask.
		<ol>
			<li>Heat the solution on a hot plate until bubbles begin to form on the bottom of the flask and float to the top.</li>
			<li>Remove the solution from the hot plate, let the solution cool until the flask can be touched with a bare hand, and then pour the gel into a 24.5 cm x 21.8 cm gel mold with a gel comb to create wells.</li>
			<li>Remove bubbles near the comb or on the surface of the gel by popping them with a pipette tip and allow the gel to solidify for at least 15 minutes. This can be stored in a fridge (4 &#176;C) wrapped in plastic wrap for 1 week to prevent drying of the gel or contamination. For more information on how to run a gel refer to Lee et al.33. 
			NOTE: Use a gel comb with 30 individual wells. Each well should be 4.5 mm wide with a depth of 2 mm. The height of the well will depend on the amount of gel poured into the box.</li>
		</ol>
		</li>
		<li>Prepare 1x TAE (Tris - acetate - EDTA) buffer using 40 mM Tris-HCl, 20 mM acetic acid, and 1 mM EDTA. Store at room temperature (18 - 25 &#176;C).</li>
	</ol>
	</li>
</ol>
2. Preparation of Stained DNA
<ol>
	<li>Stain lambda DNA (300-800 ng) using different aliquots of dyes. Each dye (Table 1) will have six different concentrations tested, for a total of 48 reactions per restriction enzyme. The following process outlines a set up for 1 reaction (Figure 1).
	<ol>
		<li>Dilute lambda DNA using 1x TE to 100 ng/&#181;L. Store in 4 &#176;C. Make a solution with total volume of 300 &#181;L.</li>
		<li>Stain unmethylated lambda DNA. For each band expected from the digest, estimate 75 - 100 ng/band. Add appropriate amounts of dye to produce concentrations of 1.9 &#181;M, 3.8 &#181;M, 19.4 &#181;M, 32.2 &#181;M, 48.3 &#181;M, 63.5 &#181;M / 100 ng of DNA.</li>
		<li>Incubate overnight (15 h - 18 h) at 4 &#176;C.</li>
		<li>Leave one reaction unstained to act as a control25. 
		NOTE: Use unmethylated DNA for the digestion reaction. Methylation sensitive enzymes cannot cut methylated regions in DNA and will give an appearance of an incomplete digestion.</li>
	</ol>
	</li>
</ol>
3. Preparation of the Restriction Enzyme Assay
<ol>
	<li>Digest stained DNA with a restriction enzyme to determine if that enzyme can digest prestained DNA (Figure 1).
	<ol>
		<li>The following day, add 20 U of restriction enzyme, 3 &#181;L of the appropriate buffer ( Table 2), and distilled, autoclaved and filtered water for a total of 30 &#181;L. For each enzyme reaction, choose the buffer with the highest cutting efficiency, which can be found on the manufacturer&#39;s website.</li>
		<li>Place the reactions in a water bath at the temperature of optimal enzymatic activity, listed in Table 2 for the 6 enzymes used in Maschmann et al., for 2 - 4 h (Table 2)25.</li>
		<li>Stop the reaction by adding 2 &#181;L of 0.5 M EDTA pH 8.0.</li>
	</ol>
	</li>
</ol>
4. Electroeluting Stained DNA in an Agarose Gel
<ol>
	<li>Remove the sides from the 24.5 cm x 21.8 cm gel mold (Step 1.2.2) and place the gel into the gel electrophoresis box. Pour 2.5 L of 1x TAE buffer into the box. Carefully remove the gel comb from the gel, so the wells are accessible33.</li>
	<li>Pipette the restriction enzyme reactions onto a plastic film and combine each reaction with 6x loading dye. For each reaction solution, add 1 &#181;L of 6x loading dye per 5 - 6 &#181;L of DNA solution (1x). Then pipette the reactions (containing the loading dye, &#60;18 &#181;L) into the wells.</li>
	<li>Mix 1 &#181;L of 1 kb ladder, 9 &#181;L of 1x TE, and 2 &#181;L of 6x loading dye. Load solution into the wells that flank the other solutions.</li>
	<li>Cover the gel box with dark blue or black paper or fabric. Connect the gel box to a power supply, set the power supply to 30 V, and run it overnight (15 - 20 h). Turn off the lights while the gel is running, to prevent photocleavage of the labeled DNA.</li>
	<li>The next morning turn off the power supply. Take the gel out using the tray and transfer it to a container with 45 &#181;L ethidium bromide and 1 L of 1x TAE buffer. Store the container at room temperature in the dark or cover in foil to prevent exposure to light. Let gel stain for at least 45 min at room temperature. 
	CAUTION: Use ethidium bromide with gloves and wear safety glasses. When cleaning out the container containing ethidium bromide solution and throwing away the gel, consult your states disposal guidelines to determine how to deal with used ethidium bromide; additionally, other stains may be utilized instead of ethidium bromide.</li>
</ol>
5. Imaging the Agarose Gel
<ol>
	<li>Place the gel on top of a blue light transilluminator, which emits maximum light output between 400-500 nm. Take pictures with the camera attached to the station. 
	CAUTION: Make sure to use the cover for the illuminator and put on protective glasses prior to turning on the light source. 
	NOTE: This step may also be completed using any UV light source or standard gel scanner.</li>
	<li>View the pictures in ImageJ by dragging the file to the ImageJ status bar and the image will pop up.</li>
	<li>Determine the digestion rate for the experiment by comparing the bands on the gel to the expected in silico gel pattern. To determine the digestion rate, the number of experimental bands is divided by the expected number of DNA bands24,25. If the experimental digested DNA band pattern has the expected pattern, the digestion rate is 1. If the pattern has more fragments than expected, the digestion rate is greater than 1. If the pattern has less fragments than expected, the digestion rate is less than one.</li>
</ol>

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The authors have nothing to disclose.

In order to digest fluorescently labeled DNA (Figure 1), a series of steps are required. First, DNA is stained with a fluorochrome overnight. DNA can be incubated with cyanine dimers for a shorter period of time; however, Carlsson et al. found that DNA created double bands for each DNA size due to incomplete staining20. To remedy this, the DNA can be stained overnight to prevent double bands from occurring. This is a critical step in the protocol. If the dye is not incubated for a long enough time, the dye-DNA may not be at equilibrium and will give a double-banding pattern. If this does occur, allow the DNA-dye mixture to equilibrate for a longer period of time.
Next, a restriction enzyme is added to the DNA-dye mixture and digested for 2 - 4 h at a specific temperature for each enzyme. The DNA must be unmethylated for methylation sensitive enzymes. If it is not, methylated DNA will inhibit the restriction enzyme from cutting DNA in methylated regions. Additionally, the reaction must be incubated in the correct buffer and temperature for the enzyme. Each enzyme has a list of buffers and the enzyme activity in each buffer. Choose the buffer with enzyme activity at 100%, if possible. As for temperature, make sure it is incubated in the correct temperature. If it is not, it may cause the enzyme to lose activity or denature if the temperature is too high. EDTA is utilized to stop the reaction. The company where the enzymes were purchased will have a chart with the optimal temperatures and buffer conditions for that enzyme. If the control does not give the correct number of bands at the right molecular weights, check to see if the enzyme is expired, increase the incubation time, or check to make sure the temperature is correct for that enzyme.
Once the reaction is completed, the samples can be loaded into a gel. First, a gel is loaded into the gel container and immersed in 1x TAE buffer. Next, the digested DNA is mixed with loading dye and the solution is loaded into the wells of the gel. The lid is slid over and the wires are attached to the gel box. A dark paper, that is taped together, is placed on top of the lid to cover the lid and drape over the sides. Another critical step is that the samples are protected from the light at all times, especially while the gel is running. The power supply should be turned on and the lights to the room turned off. In order to prevent photocleavage of DNA, samples are protected from light the whole time. If there is no dark paper, a dark fabric can be used as well. If the DNA does not move, make sure the wires are connected to the box and the run button has been pressed.
In the morning, take the gel out of the gel box and load the gel into a container that contains ethidium bromide. Ethidium bromide allows us to see the DNA bands under the blue light transilluminator (Figure 2). Other dyes can be used in place of ethidium bromide. A camera can be mounted above the blue light transilluminator to capture pictures of the gel. Depending on the setup, an imaging station can also be utilized.
This method can be utilized for any dye that intercalates or stains DNA, not just the TOTO series. Maschmann et al. modified a technique developed by Meng et al. to determine if fluorescently stained DNA could be digested with restriction enzymes24,25. This is the only technique utilized to determine if pre-stained DNA affects restriction enzymes. This method is limited to the restriction enzymes commercially available, unless a scientist has access to other restriction enzymes that are not listed commercially. Depending on DNA, the enzyme chosen should have a discernable DNA band pattern and enough bands (&#62;3) to determine if the DNA molecule is full length. Also, the DNA should be unmethylated if methylation sensitive enzymes are used. If DNA is methylated, then do not use methylation sensitive enzymes.
This method is an easy to use way to determine if restriction enzymes can cut DNA molecules at the recognition site stained with an intercalating dye without needing to design primers for a given cut site34. The future applications of this method would be to determine if fluorescently labeled molecules from an experiment were full length by using a restriction enzyme and determining the size of the DNA bands from the digest or to determine the methylation profiles of DNA molecules. Future experiments could also determine if nicking enzymes are affected by the cyanine dyes or other fluorescent dyes.

The TOTO series (TOTO-1, YOYO-1, POPO-1, BOBO-1, TOTO-3, YOYO-3, POPO-3, and BOBO-3; Table 1) is utilized in a wide variety of experiments where the visualization of DNA is required1,2,3,4,5,6,7,8,9,10,11,12,13,14,15,16,17. The cyanine dimer family is widely used due to their quantum yield, sensitivity, and high affinity for DNA molecules18,19,20. Cyanine dimer dyes have great selectivity for double stranded DNA and when intercalated have a 100 to 1000 fold increase of fluorescence21. Pyridinium dyes (YOYO-1, TOTO-1, YOYO-3, and TOTO-3) have a shorter emission wavelength than their quinolium dye (BOBO-1, POPO-1, BOBO-3, and POPO-3) counterparts (Table 1)22. Also, the quantum yield for cyanine dimers intercalated into DNA is high (0.2 - 0.6)22. However, using an enzyme to determine the methylation profile of a DNA molecule2 or stretch23 of a DNA molecule already stained with fluorescent dye requires a method to determine what enzymes will digest stained DNA. Any type of dye that intercalates into DNA or any enzyme that gives a discernable pattern of the DNA substrate can be used for this method.
Meng et al. first determined the digestion rate of prestained DNA through gel electrophoresis using a variety of different dyes24. Maschmann et al. delved in deeper to look at the TOTO family of dyes. Both determined the digestion rate of stained DNA to see if DNA stained with a given dye could be digested with a restriction enzyme25. Other methods study binding effects of dyes intercalated with DNA using optical tweezers26 or NMR27. Either method requires specialized equipment; whereas, this method allows equipment that most molecular biology labs have to determine if a dye interferes with a restriction enzyme digestion.
Additionally, in other methods to measure the length of a given molecule, optical mapping has elongated unstained DNA molecules on a surface and digested DNA to determine the stretch and size of fragments. Intercalation of dye has been shown to increase the contour length of fluorescently stained DNA molecules and depending on the dye used, the contour lengths are different21. This method has been utilized in a variety of genomes1,3,4,6,13,28,29,30,31. However, if molecules were pre-stained, depending on the dye and enzyme, the enzyme may not be able to cut DNA stained with a given dye. Therefore, this method determines if DNA stained with a given dye can be digested with an enzyme. Additionally, depending on the concentration of the dye and the dye utilized, the mobility of the DNA bands in a gel will migrate more slowly than native DNA due to the partial unwinding of the DNA backbone to make room for the dye to insert between base pairs32.
However, sometimes these dyes can partially or completely inhibit the action of certain restriction enzymes7,24. This is thought to be due to a structural change in DNA caused by the attachment of the fluorescent dye, which may prevent the enzyme from recognizing its specific sequence. Understanding how these dyes affect restriction enzymes can help in experiments where the methylation profile or the stretch of stained DNA is required.
In our method, DNA was stained with a fluorochrome of interest and digested with a restriction enzyme. Then DNA was electrophoresed on a gel, imaged, and the restriction enzyme digestion rate was measured. The restriction enzymes were chosen based on the cut pattern on a gel. Too many bands caused overlap of DNA bands and too few bands did not give a complete picture of the DNA molecule. There is a sweet spot to be able to determine the profile of the digested DNA molecule; therefore, it will depend on the DNA used and the enzyme. An advantage of this method is its simplicity; it only requires equipment used in a restriction digestion and gel electrophoresis.

<table><tbody><tr><td>Unmethylated lambda DNA</td><td>NEB</td><td>N3013S</td><td></td></tr><tr><td>YOYO-1</td><td>ThermoFisher Scientific</td><td>Y3601</td><td></td></tr><tr><td>TOTO-1</td><td>ThermoFisher Scientific</td><td>T3600</td><td></td></tr><tr><td>POPO-1</td><td>ThermoFisher Scientific</td><td>P3580</td><td></td></tr><tr><td>BOBO-1</td><td>ThermoFisher Scientific</td><td>B3582</td><td></td></tr><tr><td>YOYO-3</td><td>ThermoFisher Scientific</td><td>Y3606</td><td></td></tr><tr><td>TOTO-3</td><td>ThermoFisher Scientific</td><td>T3604</td><td></td></tr><tr><td>POPO-3</td><td>ThermoFisher Scientific</td><td>P3584</td><td></td></tr><tr><td>BamHI</td><td>NEB</td><td>R0136S</td><td></td></tr><tr><td>PmlI</td><td>NEB</td><td>R0532S</td><td></td></tr><tr><td>ScaI</td><td>NEB</td><td>R3122S</td><td></td></tr><tr><td>EcoRI&amp;nbsp;</td><td>NEB</td><td>R0101S</td><td></td></tr><tr><td>HindIII</td><td>NEB</td><td>R0104S</td><td></td></tr><tr><td>SmaI</td><td>NEB</td><td>R0141S</td><td></td></tr><tr><td>Tris</td><td>Fisher</td><td>BP152-1</td><td>Irritant</td></tr><tr><td>EDTA</td><td>Fisher</td><td>S316-212</td><td>Toxic</td></tr><tr><td>HCl</td><td>Fisher</td><td>S25358</td><td>Corrosive and irritant</td></tr><tr><td>Buffer 1</td><td>NEB</td><td>B7201S</td><td>NEB 1.1</td></tr><tr><td>Buffer 2</td><td>NEB</td><td>B7202S</td><td>NEB 2.1</td></tr><tr><td>Buffer 3</td><td>NEB</td><td>B7204S</td><td>NEB Cutsmart</td></tr><tr><td>High gelling temperature agarose gel</td><td>Lonza</td><td>50041</td><td></td></tr><tr><td>Acetic Acid</td><td>Fisher</td><td>S25118</td><td>Hazardous</td></tr><tr><td>Blue light transilluminator</td><td>Dark Reader</td><td>DR196</td><td>Wear correct eyewear&amp;nbsp;</td></tr><tr><td>Ethidium bromide</td><td>Fisher</td><td>15585011</td><td>Carcinogen; other alternatives are Nancy 520, Gel red, etc.</td></tr><tr><td>Cannon EOS Rebel T3I camera</td><td>Cannon</td><td></td><td></td></tr><tr><td>Apogee Model H4</td><td>Biorad</td><td>1704510</td><td></td></tr><tr><td>Power Pac Basic Power Supply</td><td>Biorad</td><td>1645050&amp;nbsp;</td><td></td></tr><tr><td>Ficoll</td><td>Fisher Scientific</td><td>BP525-25</td><td></td></tr><tr><td>Bromophenol blue</td><td>Fisher Scientific</td><td>B-392</td><td></td></tr><tr><td>Xylene Cyanol</td><td>Eastmen</td><td>T1579</td><td></td></tr></tbody></table>

determining if dna stained with a cyanine dye can be digested with restriction enzymes

Visualization of DNA for fluorescence microscopy utilizes a variety of dyes such as cyanine dyes. These dyes are utilized due to their high affinity and sensitivity for DNA. In order to determine if the DNA molecules are full length after the completion of the experiment, a method is required to determine if the stained molecules are full length by digesting DNA with restriction enzymes. However, stained DNA may inhibit the enzymes, so a method is needed to determine what enzymes one could use for fluorochrome stained DNA. In this method, DNA is stained with a cyanine dye overnight to allow the dye and DNA to equilibrate. Next, stained DNA is digested with a restriction enzyme, loaded into a gel and electrophoresed. The experimental DNA digest bands are compared to an in silico digest to determine the restriction enzyme activity. If there is the same number of bands as expected, then the reaction is complete. More bands than expected indicate partial digestion and less bands indicate incomplete digestion. The advantage of this method is its simplicity and it uses equipment that a scientist would need for a restriction enzyme assay and gel electrophoresis. A limitation of this method is that the enzymes available to most scientists are commercially available enzymes; however, any restriction enzymes could be used.

In order to determine if an intercalating dye will affect a restriction enzyme digesting DNA, the correct order of steps must be followed (Figure 1). Once DNA is stained and digested with a given enzyme, a picture of the gel can be taken to determine the number of fragments and their size (Figure 2). In order to determine the enzyme efficiency, the total number of expected visible bands divided by the number of visible bands. Enzyme efficiency equaled unity, if the number of bands expected and seen match. If there are more bands on the gel then expected, the value is greater than one and indicates partially digested reactions. If there are less bands then expected, then the value is less than one, which indicates incomplete digestion. In Figure 2a, lanes 3 and 4 show the mobility of the bands has decreased, causing the bands to shift towards the wells. In lanes 1 and 2, the amount of dye does not affect the mobility noticeably, so the bands match the control. In Figure 2a, lanes 5 and 6 show more bands than the control, so it indicates partial digestion. Which means, the dyes affected the cleavage of the DNA at the recognition site for that enzyme. In Figure 2b, all expected bands are seen for all dye concentrations, so the dye did not interfere with the digestion of DNA. Figure 2c tracks the mobility shifts with colored asterisks.
As dye concentration increases for dyes with -1, the mobility decreases. In order to determine the approximate size of the bands, a control is required to know where the native DNA molecules are expected compared to the stained DNA (Figure 2a). For dyes with -3, the mobility did not decrease to the extent that -1 dyes did, so it was easier to determine the fragment size (Figure 2b). The mobility difference is due to the type of dye utilized, as seen in Maschmann et al.25.The variance of these dyes is seen due to the structures of the dyes. The linker between the aromatic moiety for dyes -3 was longer than -1. The dyes may intercalate slightly differently due to the difference in the linker size, which may explain why the mobility shift is less for -3 dyes.
If there is no DNA, or the DNA bands are very faint in the gel, increasing the concentration of DNA is required. However, if the DNA concentration increases, then the amount of dye required will also increase to keep the correct concentration of dye to DNA ratio.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57141/57141fig1.jpg" /> 
Figure 1: A schematic of the steps required when digesting fluorochrome labeled DNA molecules. DNA and YOYO-1, or another dye from the TOTO series, is added to a black tube and incubated overnight (O/N; 15 - 20 h). A restriction enzyme is added to a tube; it is placed in an incubator, at the preferred temperature, for 2 - 4 h (Table 2). Load samples into an immersed 0.7% agarose gel made with 1x TAE buffer. Dark colored paper covers the gel box and the lights are turned off, while the gel is ran overnight to prevent photocleavage of the dye. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57141/57141fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57141/57141fig2.jpg" /> 
Figure 2: Restriction digestion of DNA intercalated with a cyanine dimer dye. (a) Lambda DNA is stained with different concentrations of YOYO-1 overnight (lane C: digested lambda DNA with no dye (control), lane 1: 1.9 ng, lane 2: 3.8 ng, lane 3: 19.4 ng, lane 4: 32.2 ng, lane 5: 48.3 ng, lane 6: 63.5 ng of dye per 100 ng of DNA) and then digested with EcoRI at 37 &#176;C for 2 - 4 h. All reactions are electrophoresed (E = 0.85 V/cm) on a 0.7% high gelling temperature (HGT) agarose gel made with 1x TAE buffer (1x TAE; 40 mM Tris-HCl, 20 mM Acetic acid, 1 mM EDTA). Gels are stained with ethidium bromide and imaged with a blue light transilluminator coupled to a camera. Lane L is a 1 kb ladder (sizes, kb); lane &#955; is full length lambda DNA; lane E is the expected band pattern (sizes, kb). (b) Lambda DNA is stained with YOYO-3 (lane 1: 1.9 ng, lane 2: 3.8 ng, lane 3: 19.4 ng, lane 4: 32.2 ng, lane 5: 48.3 ng, lane 6: 63.5 ng of dye per 100 ng of DNA) and digested with HindIII at 37 &#176;C. (c) To see how the mobility shifted as the YOYO-1 concentration is increased, colored asterisks are located next to each band for each YOYO-1 concentration. For example, the band at 3.5 kb has a red asterisk next to each band in each YOYO-1 concentration. The unexpected bands in lane 5 and 6 do not have an asterisk next to the bands. The figure is modified from Maschmann et al.25. Copyright (2017) Nucleosides, Nucleotides, and Nucleic Acids. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/57141/57141fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Chemical Name</td>
			<td>Abbreviated Name</td>
			<td>Ex/Em (nm)</td>
		</tr>
		<tr>
			<td>[(1&#8211;1&#8242;-[1,3-propanediylbis[(dimethyliminio)-3,1-propanediyl] bis[4-[(3-methyl-2(3H)-benzothiazolylidene)methyl]]-, tetraiodide</td>
			<td>TOTO-1</td>
			<td>514/533</td>
		</tr>
		<tr>
			<td>1,1&#8242; - (4,4,8,8-tetramethyl-4,8-diazaundecamethylene)bis[4-[(3-methylbenzo-1,3-oxazol -2-yl)methylidene]-l,4-dihydroquinolinium] tetraiodide}</td>
			<td>YOYO-1</td>
			<td>491/509</td>
		</tr>
		<tr>
			<td>2,2&#8242;-[1,3-prop-anediylbis[(dimethyliminio)-3,1-propanediyl-1(4H)-pyridinyl-4-ylidenemethylid- yne]]bis[3-methyl]-, tetraiodide</td>
			<td>POPO-1</td>
			<td>434/456</td>
		</tr>
		<tr>
			<td>2,2&#8242;-[1,3-propanediylbis[(dimethylimi- nio)-3,1-propanediyl-1(4H)-pyridinyl-4&#8211;1,1&#8242; -[4,8-bis(dimethyliminio)undecane-1,11-diyl]bis{4-[3-(3-methyl-1,3-benzothiazol-2(3H)-ylidene)prop-1-en-1-yl]qui- nolinium}</td>
			<td>BOBO-1</td>
			<td>462/481</td>
		</tr>
		<tr>
			<td>2,2&#8242;-{[4,8-bis(dimethyliminio)undecane-1,11-diyl]bis[quinolin-1-yl-4- ylideneprop-1-en-1-yl-3-ylidene]}bis(3-methyl-1,3-benzothiazol-3-ium) tetraio- dide</td>
			<td>TOTO-3</td>
			<td>642/660</td>
		</tr>
		<tr>
			<td>1,1&#8242;-[1,3-propanediylbis[(dimethyliminio)-3,1-propanediyl]]bis[4- [3-(3-methyl-2(3H)-benzoxazolylidene)-1-propenyl]]-, tetraiodide</td>
			<td>YOYO-3</td>
			<td>612/631</td>
		</tr>
		<tr>
			<td>2,2&#8242; -[1,3-propanediylbis[(dimethyliminio)-3,1-propanediyl-1(4H)-pyridinyl-4- ylidene-1-pr-open-1-yl-3-ylidene]]bis[3-methyl]-, tetraiodide</td>
			<td>POPO-3</td>
			<td>534/570</td>
		</tr>
		<tr>
			<td>2,2&#39;-{propane-1,3-diylbis[(dimethylazaniumdiyl)propane-3,1-diylpyridin-1-yl-4-ylideneprop-1-en-1-yl-3-ylidene]}bis(3-methyl-1,3-benzothiazol-3-ium) tetraiodide</td>
			<td>BOBO-3</td>
			<td>570/602</td>
		</tr>
	</tbody>
</table>
Table 1: Abbreviated names of TOTO family of dyes with IUPAC names with emission (Em) and excitation (Ex) maximima22.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Enzyme</td>
			<td>Methylation Sensitive</td>
			<td>Buffer</td>
			<td>Water Bath Temperature</td>
		</tr>
		<tr>
			<td>BamHI</td>
			<td>No</td>
			<td>2</td>
			<td>37 &#176;C</td>
		</tr>
		<tr>
			<td>EcoRI</td>
			<td>Yes</td>
			<td>2</td>
			<td>37 &#176;C</td>
		</tr>
		<tr>
			<td>HindIII</td>
			<td>No</td>
			<td>2</td>
			<td>37 &#176;C</td>
		</tr>
		<tr>
			<td>PmlI</td>
			<td>Yes</td>
			<td>1</td>
			<td>37 &#176;C</td>
		</tr>
		<tr>
			<td>ScaI</td>
			<td>No</td>
			<td>2</td>
			<td>37 &#176;C</td>
		</tr>
		<tr>
			<td>SmaI</td>
			<td>Yes</td>
			<td>3</td>
			<td>25 &#176;C</td>
		</tr>
	</tbody>
</table>
Table 2: Listed in the table is a subset of possible enzymes that could be used in these experiments. Three enzymes are methylation sensitive and three are not methylation sensitive. The buffers used for the enzymes and incubation temperatures are listed for each enzyme.

Watch this Scientific Journal Video about Determining if DNA Stained with a Cyanine Dye Can Be Digested with Restriction Enzymes at JoVE.com

Determining if DNA Stained with a Cyanine Dye Can Be Digested with Restriction Enzymes

Staining DNA molecules for fluorescence microscopy allows a scientist to view them during an experiment. In the method presented here, DNA molecules are pre-stained with fluorescent dyes and digested with methylation and non-methylation sensitive restriction enzymes.

Visualization of DNA for fluorescence microscopy utilizes a variety of dyes such as cyanine dyes. These dyes are utilized due to their high affinity and ...

determining-if-dna-stained-with-cyanine-dye-can-be-digested-with

Nanyang Technological University

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Biochemistry

8.7K Views.  University of Nebraska - Kearney. Staining DNA molecules for fluorescence microscopy allows a scientist to view them during an experiment. In the method presented here, DNA molecules are pre-stained with fluorescent dyes and digested with methylation and non-methylation sensitive restriction enzymes.

Video: Determining if DNA Stained with a Cyanine Dye Can Be Digested with Restriction Enzymes

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, ...

In-Nucleus Hi-C in Drosophila Cells

The genome is organized into topologically associating domains (TADs) delimited by boundaries that isolate interactions between domains. In Drosophila, the mechanisms underlying TAD formation and boundaries are still under investigation. The application of the in-nucleus Hi-C method described here helped to dissect the function of architectural protein (AP)-binding sites at TAD boundaries isolating the Notch gene. Genetic modification of domain boundaries that cause loss of APs results in TAD fusion, transcriptional defects, and long-range topological alterations. These results provided evidence demonstrating the contribution of genetic elements to domain boundary formation and gene expression control in Drosophila. Here, the in-nucleus Hi-C method has been described in detail, which provides important checkpoints to assess the quality of the experiment along with the protocol. Also shown are the required numbers of sequencing reads and valid Hi-C pairs to analyze genomic interactions at different genomic scales. CRISPR/Cas9-mediated genetic editing of regulatory elements and high-resolution profiling of genomic interactions using this in-nucleus Hi-C protocol could be a powerful combination for the investigation of the structural function of genetic elements.

The genome is organized in the nuclear space into different structures that can be revealed through chromosome conformation capture technologies. The in-nucleus Hi-C method provides a genome-wide collection of chromatin interactions in Drosophila cell lines, which generates contact maps that can be explored at megabase resolution at restriction fragment level.

The genome is organized into topologically associating domains (TADs) delimited by boundaries that isolate interactions between domains. In Drosophila, ...

Genetics

Chromosome Conformation Capture (3C) for Undergraduate Research and Teaching

Getting an A with the 3Cs: Chromosome Conformation Capture for Undergraduates

Chromosome conformation capture (3C) is a powerful tool that has spawned a family of similar techniques (e.g., Hi-C, 4C, and 5C, referred to here as 3C techniques) that provide detailed information of the three-dimensional organization of chromatin. The 3C techniques have been used in a wide range of studies, from monitoring the changes in chromatin organization in cancer cells to identifying enhancer contacts made with gene promoters. While many of the studies using these techniques are asking big genome-wide questions with intricate sample types (i.e., single-cell analysis), what is often lost is that the 3C techniques are grounded in basic molecular biology methods that are applicable to a broad range of studies. By addressing tightly focused questions of chromatin organization, this cutting-edge technique can be used to enhance the undergraduate research and teaching lab experience. This paper presents a 3C protocol and provides adaptations and points of emphasis for implementation at primarily undergraduate institutions in undergraduate research and teaching experiences.

Author Spotlight: Getting an A with the 3Cs: Chromosome Conformation Capture for Undergraduates  

Here, we present an adaption of the chromosome conformation capture (3C) technique in detail with an emphasis on undergraduate involvement and learning.

Chromosome conformation capture (3C) is a powerful tool that has spawned a family of similar techniques (e.g., Hi-C, 4C, and 5C, referred to here as 3C ...

Single-Molecule Dwell-Time Analysis of Restriction Endonuclease-Mediated DNA Cleavage

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for hundreds of individual restriction endonuclease (REase) molecules in one experiment. This assay does not require protein labeling and can be carried out with a single imaging channel. In addition, the results of multiple individual experiments can be pooled to generate well-populated dwell-time distributions. Analysis of the resulting dwell-time distributions can help elucidate the DNA cleavage mechanism by revealing the presence of kinetic steps that cannot be directly observed. Example data collected using this assay with the well-studied REase, EcoRV - a dimeric Type IIP restriction endonuclease that cleaves the palindromic sequence GAT&#8595;ATC (where &#8595; is the cut site) - are in agreement with prior studies. These results suggest that there are at least three steps in the pathway to DNA cleavage that is initiated by introducing magnesium after EcoRV binds DNA in its absence, with an average rate of 0.17 s-1 for each step.

Using quantum-dot-labeled DNA and total internal reflection fluorescence microscopy, we can investigate the reaction mechanism of restriction endonucleases while using unlabeled protein. This single-molecule technique allows for massively multiplexed observation of individual protein-DNA interactions, and data can be pooled to generate well-populated dwell-time distributions.

This novel total internal reflection fluorescence microscopy-based assay facilitates the simultaneous measurement of the length of the catalytic cycle for ...

Hi-C: A Method to Study the Three-dimensional Architecture of Genomes.

The three-dimensional folding of chromosomes compartmentalizes the genome and and can bring distant functional elements, such as promoters and enhancers, into close spatial proximity 2-6. Deciphering the relationship between chromosome organization and genome activity will aid in understanding genomic processes, like transcription and replication. However, little is known about how chromosomes fold. Microscopy is unable to distinguish large numbers of loci simultaneously or at high resolution. To date, the detection of chromosomal interactions using chromosome conformation capture (3C) and its subsequent adaptations required the choice of a set of target loci, making genome-wide studies impossible 7-10.
We developed Hi-C, an extension of 3C that is capable of identifying long range interactions in an unbiased, genome-wide fashion. In Hi-C, cells are fixed with formaldehyde, causing interacting loci to be bound to one another by means of covalent DNA-protein cross-links. When the DNA is subsequently fragmented with a restriction enzyme, these loci remain linked. A biotinylated residue is incorporated as the 5' overhangs are filled in. Next, blunt-end ligation is performed under dilute conditions that favor ligation events between cross-linked DNA fragments. This results in a genome-wide library of ligation products, corresponding to pairs of fragments that were originally in close proximity to each other in the nucleus. Each ligation product is marked with biotin at the site of the junction. The library is sheared, and the junctions are pulled-down with streptavidin beads. The purified junctions can subsequently be analyzed using a high-throughput sequencer, resulting in a catalog of interacting fragments.
Direct analysis of the resulting contact matrix reveals numerous features of genomic organization, such as the presence of chromosome territories and the preferential association of small gene-rich chromosomes. Correlation analysis can be applied to the contact matrix, demonstrating that the human genome is segregated into two compartments: a less densely packed compartment containing open, accessible, and active chromatin and a more dense compartment containing closed, inaccessible, and inactive chromatin regions. Finally, ensemble analysis of the contact matrix, coupled with theoretical derivations and computational simulations, revealed that at the megabase scale Hi-C reveals features consistent with a fractal globule conformation.

The Hi-C method allows unbiased, genome-wide identification of chromatin interactions (1). Hi-C couples proximity ligation and massively parallel sequencing. The resulting data can be used to study genomic architecture at multiple scales: initial results identified features such as chromosome territories, segregation of open and closed chromatin, and chromatin structure at the megabase scale.

The three-dimensional folding of chromosomes compartmentalizes the genome and and can bring distant functional elements, such as promoters and enhancers, ...

Biology

DNA-based Fish Species Identification Protocol

We have developed a fast, simple, and accurate DNA-based screening method to identify the fish species present in fresh and processed seafood samples. This versatile method employs PCR amplification of genomic DNA extracted from fish samples, followed by restriction fragment length polymorphism (RFLP) analysis to generate fragment patterns that can be resolved on the Agilent 2100 Bioanalyzer and matched to the correct species using RFLP pattern matching software.
The fish identification method uses a simple, reliable, spin column- based protocol to isolate DNA from fish samples. The samples are treated with proteinase K to release the nucleic acids into solution. DNA is then isolated by suspending the sample in binding buffer and loading onto a micro- spin cup containing a silica- based fiber matrix. The nucleic acids in the sample bind to the fiber matrix. The immobilized nucleic acids are washed to remove contaminants, and total DNA is recovered in a final volume of 100 &mu;l. The isolated DNA is ready for PCR amplification with the provided primers that bind to sequences found in all fish genomes. The PCR products are then digested with three different restriction enzymes and resolved on the Agilent 2100 Bioanalyzer. The fragment lengths produced in the digestion reactions can be used to determine the species of fish from which the DNA sample was prepared, using the RFLP pattern matching software containing a database of experimentally- derived RFLP patterns from commercially relevant fish species.

This publication describes how to use the Agilent Fish Species Identification System to identify the species of a fish by extracting DNA and performing PCR and RFLP analysis.

We have developed a fast, simple, and accurate DNA-based screening method to identify the fish species present in fresh and processed seafood samples. ...

Studying TRF1/2 Interactions with Telomeric DNA Using Single-Molecule Assay

Analyzing Telomeric Protein-DNA Interactions Using Single-Molecule Magnetic Tweezers

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper function depends on tightly regulated processes of replication, elongation, and damage response. The shelterin complex, especially Telomere Repeat-binding Factor 1 (TRF1) and TRF2, plays a pivotal role in telomere protection and has emerged as a potential anti-cancer target for drug discovery. These proteins bind to the repetitive telomeric DNA motif TTAGGG, facilitating the formation of protective structures and recruitment of other telomeric proteins. Structural methods and advanced imaging techniques have provided insights into telomeric protein-DNA interactions, but probing the dynamic processes requires single-molecule approaches. Tools like magnetic tweezers, optical tweezers, and atomic force microscopy (AFM) have been employed to study telomeric protein-DNA interactions, revealing important details such as TRF2-dependent DNA distortion and telomerase catalysis. However, the preparation of single-molecule constructs with telomeric repetitive motifs continues to be a challenging task, potentially limiting the breadth of studies utilizing single-molecule mechanical methods. To address this, we developed a method to study interactions using full-length human telomeric DNA with magnetic tweezers. This protocol describes how to express and purify TRF2, prepare telomeric DNA, set up single-molecule mechanical assays, and analyze data. This detailed guide will benefit researchers in telomere biology and telomere-targeted drug discovery.

Author Spotlight: Advanced Single-Molecule Techniques for Investigating Telomeric Protein-DNA Interactions

This protocol demonstrates using single-molecule magnetic tweezers to study interactions between telomeric DNA-binding proteins (Telomere Repeat-binding Factor 1 [TRF1] and TRF2) and long telomeres extracted from human cells. It describes the preparatory steps for telomeres and telomeric repeat-binding factors, the execution of single-molecule experiments, and the data collection and analysis methods.

Telomeres, the protective structures at the ends of chromosomes, are crucial for maintaining cellular longevity and genome stability. Their proper ...

CMG-Mediated DNA Unwinding Observed by ssDNA-RPA Tract Growth

Single-Molecule Real-Time Visualization of DNA Unwinding by CMG Helicase

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a dynamic complex of proteins termed the replisome. At the heart of the replisome is the&#160;CDC45-MCM2-7-GINS (CMG) helicase, which separates the two strands of the DNA double helix such that DNA polymerases can copy each strand. During genome duplication, replisomes must overcome a plethora of obstacles and challenges. Each of these threatens genome stability, as failure to replicate DNA completely and accurately can lead to mutations, diseases, or cell death. Therefore, it is of great interest to understand how CMG functions in the replisome during both normal replication and replication stress. Here, we describe a total internal reflection fluorescence (TIRF) microscopy assay using recombinant purified proteins, which allows for real-time visualization of surface-tethered stretched DNA molecules by individual CMG complexes. This assay provides a powerful platform to investigate CMG behavior at the single-molecule level, allowing helicase dynamics to be directly observed with real-time control over reaction conditions.

Author Spotlight: Unraveling the Dynamics of Eukaryotic DNA Replication Through Single-Molecule Visualization

This protocol demonstrates performing a single-molecule assay for live visualization of DNA unwinding by CMG helicase. It describes (1) preparing a DNA substrate, (2) purifying fluorescently labeled Drosophila melanogaster CMG helicase, (3) preparing a microfluidic flow cell for total internal reflection fluorescence (TIRF) microscopy, and (4) the single-molecule DNA unwinding assay.

Faithful genome duplication is essential for preserving the genetic stability of dividing cells. DNA replication is carried out during the S phase by a ...

DNA Isolation and Restriction Enzyme Analysis

The revelation of DNA as the hereditary molecule in all organisms has led to enormous scientific and medical breakthroughs and significantly enhanced our understanding of ourselves and other organisms. DNA isolation and profiling have been the fundamental first steps for many of the advancements in the past century; from identification of gene function, to revolutions of agriculture and forensics.

<h4>Isolating DNA from Cells</h4>

DNA isolation is a fairly simple procedure that requires few but essential steps: cell harvesting, lysis, protein degradation, and finally DNA precipitation. Generally, DNA can be harvested from small amounts of tissue, such as a single leaf of a plant, hair follicles of animals, or epithelial cells that can be easily sloughed off within the mouth with a simple saline solution. Since DNA is compartmentalized inside the nucleus or mitochondria within eukaryotic cells, scientists first break apart the cell and nuclear membrane to expose the DNA, which is done with the lysis step. However, once DNA is exposed, it can be degraded by heavy metal ions and extremes of pH. Therefore, a detergent-containing buffer solution is used to not only dissolve the cell membrane, but also to protect DNA from degrading elements in the solution. Moreover, DNA is an unusually long molecule that is condensed inside the nucleus by packing tightly around special proteins called histones. Therefore, an enzyme called Proteinase K, which degrades peptide bonds of proteins, is added to separate the DNA from histones and other proteins. Next, scientists separate DNA from the rest of the cellular extract by using its chemical properties. Due to the negatively charged structure of nucleic acids, Na+ ions in a sodium chloride solution bind to and integrate into entangled DNA strands. Once the DNA is clumped, it can be observed by the naked eye. DNA is insoluble in alcohol and readily precipitates at low temperatures, thus scientists add cold alcohol to precipitate DNA. Finally, the precipitated DNA can be dissolved and stored in deionized water or a mild buffer, free of any enzymes that can degrade nucleic acids, until it is used.

<h4>Quantifying DNA</h4>

Reactions that use DNA often require high purity and precise amounts of DNA. Therefore, following DNA isolation, scientists use a spectrophotometer to quantify the amount of DNA in their sample, as well as the purity of the DNA in the sample. A spectrophotometer measures the intensity of a light beam passed through a sample as a function of its wavelength. Nucleic acids absorb light at 260 nm, whereas proteins absorb light at 280 nm. By measuring the intensity of a light beam at 260 nm, scientists can determine the DNA concentration. Furthermore, by assessing the light intensity at both 260 nm and 280 nm, the purity of the DNA content can be determined. In a sample of pure DNA, the 260/280 nm ratio should approach the value of 2.

<h4>Restriction Enzymes</h4>

For a number of analyses, such as cloning, sequencing, and mapping DNA fragments and genomes, the isolated DNA needs to be cut at specific sequences using restriction enzymes. These enzymes are produced by bacteria as mechanisms of innate immunity against intruding viral DNA and function as molecular scissors. Hence, after restriction enzymes identify a viral DNA, they start to break it apart by cutting it into segments at certain nucleotide sequences they recognize. These nucleotide sequences are often six to twelve nucleotides long and are generally palindromic, which means that they have the same nucleotide sequence in the 3&#8217;-5&#8217; and 5&#8217;-3&#8217; directions. For instance, the restriction enzyme EcoRI cuts DNA at the sequence 3&#8217;...GAATTC&#8230;-5&#8217;, which reads the same in the 5&#8217;-3&#8217; direction. Like EcoRI, there are hundreds of enzymes that correspond to many distinct palindromic sequences. Once DNA is cut by these enzymes, scientists can load the DNA onto a gel in an electrophoresis chamber and pass electrical current through the gel. DNA carries negative charges, which causes DNA fragments to migrate towards the anode. However, the pores of the gel slow down larger DNA fragments relative to smaller DNA sequences, thus the pieces separate according to their size. After the completion of the electrophoresis, DNA fragments can be visualized with the aid of dyes that specifically bind to DNA molecules.

Cutting DNA with restriction enzymes and separating DNA fragments with electrophoresis allows researchers to identify the sizes of unknown fragments relative to a standard ladder. These fragments of interest can then be further isolated and ligated into a plasmid vector for genetic cloning. Genetic engineering and laboratory analyses have been improved immensely with these methods. With DNA profiling, scientists are able to sequence genomes to further understand the workings of organisms and are also able to identify and characterize certain genes1.

DNA isolation and profiling are also practical for events directly impacting everyday life. For example, forensics scientists routinely analyze DNA samples to provide evidence for criminal and civil cases2. Similarly, healthcare workers extract and analyze DNA to check for paternity and genetic diseases. Recently, DNA ancestry analysis kits became popular by allowing users determine their genetic lineage, find relatives, and even discover their genetic disposition to various health conditions3. Finally, with advancements in biotechnology and genetic engineering, personalized medicine is gaining traction among clinical researchers to develop treatments by using an individual&#8217;s genetic data, paving the way to effectively treat patients without detrimental side effects4.

<h4>References</h4>

<ol class="references">
	<li>Chambers, GK, et al. DNA fingerprinting in zoology: past, present, future. Investig Genet. 2014, Vol. 5, 3 doi: 10.1186/2041-2223-5-3.</li>
	<li>Roewer, L. DNA fingerprinting in forensics: past, present, future. Investig Genet. . 2013, Vol. 4, 22 doi: 10.1186/2041-2223-4-22.</li>
	<li>Kirkpatrick, BE and Rashkin, MD. Ancestry Testing and the Practice of Genetic Counseling. J Gen Counseling. 2016, Vol. 26, 1 (6-20).</li>
	<li>Hamburg, MA and Collins, FS. The Path to Personalized Medicine. N Engl J Med . 2010; , Vol. 363, 301-304.</li>
</ol>

The revelation of DNA as the hereditary molecule in all organisms has led to enormous scientific and medical breakthroughs and significantly enhanced our ...

Education

JoVE Lab Manual

Restriction Enzyme Digests

Restriction enzymes or endonucleases recognize and cut DNA at a specific sequence. These enzymes occur naturally in bacteria as a defense against bacteriophages - viruses that infect bacteria. Bacterial restriction enzymes cut the invading bacteriophage DNA while leaving the bacterial genomic DNA unharmed due to addition of methyl groups.
This video explains the basic principles of restriction enzymes including: how restriction enzymes are named and the types of recognition sites and overhangs that exist. Also provided is a step-by-step generalized procedure for how to set up a restriction digest including the necessary components, the order in which the mixture should be assembled, and the typical incubation temperature and time. The importance of inactivating restriction enzymes to prevent star activity is mentioned. Tips for performing multiple enzymes digests and using controls in digestion reactions are also provided.