The authors would like to thank Advanced Instrumentation Research Facility, JNU, for the CD spectrophotometer. V.J. and A.D. were supported by a fellowship from CSIR.

1. Working concentration of the reaction components
<ol>
	<li>Prepare the working concentrations of buffers for CD and other reaction components freshly (see Table 1) and keep them at 4 &#176;C before setting up the reactions. 
	NOTE: For the CD reactions described in this paper, the working concentrations of components are as follows: Sodium phosphate buffer (pH 7.0) 1 mM, ATP 2 mM, DNA 500 nM, Protein 1 &#181;M, MgCl2 10 mM, EDTA 50 mM, ADAADiN 5 &#181;M.</li>
</ol>
2. ATPase activity
<ol>
	<li>Before CD spectroscopy, establish the ATPase activity of the protein in the presence of the DNA molecules to ensure that the protein used in the CD spectroscopy is active and to identify the DNA molecules that are optimally effective in eliciting ATP hydrolysis.</li>
	<li>Measure the ATPase activity of the protein in the presence of different DNA molecules by an NADH-coupled oxidation assay consisting of the following two reactions.
	<ol>
		<li>Mix 0.1 &#181;M ADAAD, 2 mM ATP, 10 nM DNA, and 1x REG buffer in a 96-well plate to a final volume of 250 &#181;L. 
		NOTE: The pyruvate kinase enzyme uses the ADP and Pi to convert phosphoenolpyruvate to pyruvate, thus regenerating ATP. This ensures that ATP is always in a saturating concentration in the reaction. In the second reaction, the pyruvate formed by the action of pyruvate kinase is converted by lactate dehydrogenase to lactate. In this reaction, one NADH molecule is oxidized to NAD+. The consumption of NADH is measured by measuring the absorbance of the molecule at 340 nm.</li>
		<li>Incubate for 30 min at 37 &#176;C in an incubator.</li>
		<li>Measure the amount of NAD+ at 340 nm using a microplate reader.</li>
		<li>To measure the amount of NAD+, use the software provided along with the microplate reader.
		<ol>
			<li>Click on the NADH assay to measure the absorbance at 340 nm.</li>
			<li>Place the 96-well plate on the plate holder in the instrument. Click on the Read Plate button to record the absorbance. 
			NOTE: The concentration of NAD+ is calculated using the molar extinction coefficient of NADH as 6.3 mM&#8722;1 by using eq (1). 
			A = &#949;cl (1) 
			 
			Here, A = Absorbance 
			&#949; = Molar extinction coefficient 
			c = Molar concentration 
			l = Optical path length in cm</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Choosing and preparation of CD cuvettes
<ol>
	<li>Collect CD spectra in high-transparency quartz cuvettes. Use rectangular or cylindrical cuvettes. 
	NOTE: A CD quartz cuvette (nominal volume of 0.4 mL, path-length of 1 mm) was used for all the reactions described in this paper.</li>
	<li>Use a cuvette cleaning solution to clean the cuvette. Add 1% cuvette cleaning solution in water to make 400 &#181;L of the solution, pour it in the cuvette, and incubate it at 37 &#176;C for 1 h.</li>
	<li>Wash the cuvette with water several times to clean the cuvette. Take a scan of the water or buffer in the cuvette to check whether it is clean. 
	NOTE: The water or buffer must give a reading in the 0 to 1 mdeg range.</li>
</ol>
4. Preparation of proteins and DNA oligonucleotide
<ol>
	<li>Keep the volume of the protein below 50 &#181;L in the reaction to minimize the amounts of the buffer components that sometimes cause the formation of ambiguous peaks. Keep the protein on the ice throughout the experiment to avoid any degradation.</li>
	<li>Use PAGE-purified DNA oligonucleotides in the reactions. 
	NOTE: In the reactions described here, DNA was used both in native as well as heat-cooled forms (fast-cooled (FC) and slow-cooled (SC)). Fast cooling promotes intramolecular bonding in the DNA, yielding more secondary structures. In contrast, slow cooling promotes intermolecular bonding in the DNA, resulting in fewer secondary structures.</li>
	<li>For fast-cooling, heat DNA at 94 &#176;C for 3 min on the heating block and immediately cool it on ice. For slow-cooling, heat DNA at 94 &#176;C for 3 min and allow it to cool to room temperature at a rate of 1 &#176;C per minute.</li>
</ol>
5. Setting up control experiments to record the baseline spectra
<ol>
	<li>Keep the reaction volume at 300 &#181;L in all the reactions. Set up a total of 5 baseline reactions in 1.5 mL centrifuge tubes, one by one, as follows: i) Buffer + Water; ii) Buffer + MgCl2 + ATP + Water; iii) Buffer + MgCl2 + ATP + Protein + Water; iv) iii + EDTA or ADAADiN; v) Buffer + Protein +Water.</li>
</ol>
6. Setting up the experiments to record CD spectra
<ol>
	<li>Set up a total of 5 reactions, one by one, in 1.5 mL centrifuge tubes as follows: i) Buffer + DNA + Water; ii) Buffer + DNA + MgCl2 + ATP + Water; iii) Buffer + DNA + MgCl2 + ATP + Protein + Water; iv) iii + EDTA or ADAADiN; v) Buffer + DNA + Protein +Water.</li>
</ol>
7. Recording scan
<ol>
	<li>Turn on the gas and switch on the CD spectrometer.</li>
	<li>Switch on the lamp after 10-15 min. Switch on the water bath and set the holder temperature at 37 &#176;C.</li>
	<li>Open the CD spectrum software.
	<ol>
		<li>Set the temperature to 37 &#176;C.</li>
		<li>Set the wavelength range at 180 - 300 nm.</li>
		<li>Set the time per point to 0.5 s.</li>
		<li>Set the scan number to 5.</li>
		<li>Click on Pro-Data Viewer, make a new file, and rename it with details about the experiment and date.</li>
	</ol>
	</li>
	<li>Keep all the reaction components on ice to avoid any degradation. Make the baselines and reactions, one by one, in centrifuge tubes and mix them by pipetting. Transfer the reaction mix to the cuvette carefully, ensuring that there are no air bubbles.</li>
	<li>If performing a time-course experiment, incubate the reactions at 37 &#176;C for the required time and take the scan. Add EDTA to the buffer containing the DNA, ATP, Mg+2, and protein to stop ATP hydrolysis.</li>
	<li>Increase the concentration of EDTA and its incubation time to inhibit ATPase activity completely.</li>
	<li>Subtract the baselines from the corresponding reactions in the software (e.g., subtract reaction 1 from baseline 1). Smoothen the data either in the CD spectrum software or in the data plotting software. Plot the data in the data plotting software. 
	NOTE: Subtracting the baselines from the corresponding reactions will give the net CD spectra of only DNA.</li>
</ol>
8. Data analysis and interpretation
<ol>
	<li>Use the formula given by eq (2) to convert the values obtained in millidegrees to mean residue ellipticity. 
	<img alt="Equation 1" src="/files/ftp_upload/63147/63147eq01.jpg" />&#160; (2) 
	Here, S is the CD signal in millidegrees, c is the DNA concentration in mg/mL, mRw is the mean residue mass, and l is the path length in cm.</li>
	<li>Plot a graph against wavelength and mean residue ellipticity using the data plotting software and analyze the peaks.</li>
	<li>To plot the graph, select the mean residue ellipticity on the Y-axis and wavelength on the X-axis and plot a straight line graph. 
	NOTE: This graph will provide the characteristics peaks of different forms of DNA. The forms of DNA corresponding to the peaks can be identified using existing literature6.</li>
</ol>

The authors have no conflict of interest to declare.

The purpose of this article is to introduce the CD spectroscopy technique as an approach to study the conformational changes occurring in the DNA in the presence of ATP-dependent chromatin remodeling proteins and to link these conformational changes to gene expression. CD spectroscopy provides a fast and easily accessible method to study the conformational changes in DNA.

A crucial point to be considered for this technique is the purity of the DNA and protein. It is advisable to ensure that both DNA and protein are &#62;95% pure. PAGE-purified oligonucleotides must be used in the assay, and the protein should be preferably affinity-purified to &#62;95% purity. The other critical parameter is that the cuvette should be clean such that the baseline reading does not exceed 1 mdeg. The buffers should be made using autoclaved water, and the baseline reading of the buffer should not exceed 1 mdeg. To study the conformation of the promoter, it is essential to identify the regions where the protein binds. Therefore, it is advisable to perform ChIP experiments using the protein of interest as this process helps to identify DNA sequences present in the promoter region of the effector gene bound by the protein. Once the region is identified, the ability of the sequence to adopt specific structures can be analyzed using available bioinformatics tools. This is important as the ChIP primers are usually 200 bp long and may have multiple conformations. Therefore, using bioinformatics tools to identify the structures would help shorten the length of the oligonucleotide to one structure.
Finally, if the protein of interest is an ATP-dependent chromatin remodeling protein, the ability of the oligonucleotides to act as an effector must be checked using ATPase assays. In both CD spectroscopy and ATPase assays, care should be taken to ensure that saturating concentrations of ligands are used in the reaction. If possible, the dissociation constant (Kd) for the protein-ligand interaction should be calculated before proceeding with CD spectroscopy. Numerous methods are available for calculating the binding parameter. Using ATPase assays, the Michaelis-Menten constant (KM) can be calculated by titrating increasing concentrations of DNA. The KM, in many cases, can be approximated to the binding constant. If the protein is fluorescent, binding constants can be calculated using fluorescence spectroscopy. If neither of these techniques is feasible, the electrophoresis mobility shift assay (EMSA) can be used.
The main problem with CD spectroscopy arises when the cuvettes are not cleaned or when the reagents are impure. If the baseline is too high, it is advisable to clean the cuvettes. Numerous cuvette cleaning solutions are available. Placing the cuvettes in a dilute acid solution for 16-24 h also helps clean the cuvette. It is advisable to purchase reagents that are &#62;95% pure and to use double-distilled and autoclaved water. Baseline drift is another potential problem. If doing a long-term experiment, it is advisable to periodically check the baseline. The peaks of DNA when studying protein-DNA interactions may not exactly correspond to the peaks/bands obtained with DNA alone. Protein peaks are usually observed in the Far UV range between 190 and 230 nm. Therefore, peaks below 250 nm might have interference from the protein peak and might not provide reliable information. DNA can adopt a variety of non-B conformations depending upon the sequence. The longer the DNA sequence, the higher the chance of multiple conformations co-existing within the DNA oligonucleotide. This can make analysis difficult. Hence, it is advisable to use shorter oligonucleotides corresponding to the potential structures predicted by the bioinformatic tools.
The other major drawback of CD spectroscopy is that it does not allow for atomic-level structure analysis, and the obtained spectrum is insufficient to identify the only viable structure. For example, both X-ray crystallography and protein NMR spectroscopy provide atomic resolution data, whereas CD spectroscopy provides less detailed structural information. However, CD spectroscopy is a rapid approach that does not necessitate vast quantities of proteins or considerable data processing. As a result, CD may be used to investigate a wide range of solvent variables such as temperature, pH, salinity, and the presence of various cofactors. It can also be used to monitor structural changes (due to complex formation, folding/unfolding, denaturation due to temperature, denaturants, and changes in amino acid sequence/mutation) in dynamic systems. By connecting it to the stop-flow apparatus, it can also be used to study the kinetics of protein/DNA-ligand interactions.
Well-characterized DNA conformers include A/B/Z DNA, triplex, hairpin, and G-quadruplexes. All these forms of DNA are associated with an open DNA conformation, i.e., unwound DNA that serves as the sink for negative supercoiling. Transcription is associated with negative supercoiling as the formation of an open complex is a prerequisite for the movement of RNA polymerase. Therefore, transcription of most genes involves increased negative supercoiling in the promoter region. Studies have shown that nucleosome unfolding leads to A-DNA conformation, which, however, is unstable39. One possibility is that the protein of interest, e.g., SMARCAL1, binds to such structures and stabilizes them, thus facilitating transcription, as seen in the case of DROSHA promoters. The guanine quadruplex is based on guanine tetrads bound by Hoogsteen hydrogen bonds. In silico analysis has confirmed that G4-forming sequences are notably enriched proximal to gene promoters and at transcription start sites. These G4 sequences can both activate and repress transcription.
In the case of the MYC promoter40, the formation of G-quadruplex acts as a repressor, while in the case of human vascular endothelial growth factor (VEGF), the G-quadruplex structure functions as a docking site for transcription factors41, thus activating the expression of this gene. In the case of SMARCAL1, the G-quadruplex structure was observed when ADAAD interacted with DGCR8 pair 7 promoter sequences. As the occupancy of SMARCAL1 and RNAPII increased on this primer pair, it is hypothesized that the formation of G-quadruplex, in this case, correlates with transcription activation of this gene. DNA can also be positively supercoiled, and the progress of RNA polymerase is known to generate positive supercoiling in front of it. This transcription-generated (+) supercoiling can disrupt or eliminate road-block proteins, destabilizing nucleosome structures to make the DNA more accessible to RNA polymerase. The X-DNA is a conformation of DNA that acts as sinks for positive supercoiling. The striking feature of an X-DNA is it can form in a sequence-specific manner on the promoter of a gene. In the case of SMARCAL1, ADAAD induced A-X and B-X transitions in an ATP-dependent manner in the DICER and DGCR8 promoters, respectively. Combined with in vivo data where increased SMARCAL1 and RNAPII occupancy was found on these promoters in the presence of doxorubicin-induced DNA damage, it can be hypothesized that X-DNA formation facilitates transcription by removing barriers/blocks. Triple DNA helices do not have a characteristic spectrum. The CD spectra of the DNA sequences present in the MYC promoter and the synthetic stem-loop DNA showed two positive peaks-one at 258 nm with a shoulder at 269 nm and a larger peak at 210 nm in the DNA. This type of spectrum can be obtained in the case of triplexes37. Triplexes are difficult to unwind and, therefore, are known to block transcription42. Hence, it is hypothesized that the formation of this structure in the c-MYC promoter by SMARCAL1 leads to repression of transcription.
It should be noted that ATP also binds to ATP-dependent chromatin remodeling proteins. The conserved arginine present in the motif VI of the helicase domain of these proteins interacts via electrostatic interactions with the &#947;-phosphate of the protein43. In the case of ADAAD, the Kd of protein-ATP interaction is (1.5 &#177; 0.1) &#215; 10-6 M38. The binding of ATP induces a conformational change in the protein such that the affinity of the DNA increases. The binding of DNA also induces a conformation change in the protein leading to an increased 10-fold affinity for ATP31. For example, in the case of ADAAD, bands/peaks are observed at -212 nm and -222 nm. ATP also gives bands at 197 nm, +210 nm, -222 nm, -247 nm, and -270 nm. These must be subtracted from the spectra of DNA + ADAAD + ATP to obtain the &#34;net&#34; conformation of the DNA in the presence of the ligands.
Thus, this paper shows the convenience of CD spectroscopy for the study of the conformational changes occurring in the DNA in the presence of ATP-dependent chromatin remodeling proteins. Correlating the changes in the DNA conformation with ChIP data can provide the investigators with information regarding how the DNA conformers activate/repress transcription.

Circular dichroism (CD) is a spectroscopic technique that relies on the inherent chirality of biological macromolecules that leads to differential absorption of right-handed and left-handed circularly polarized light. This differential absorption is known as circular dichroism. The technique, therefore, can be used to delineate the conformation of biological macromolecules, such as proteins and DNA, both of which contain chiral centers1,2.
Electromagnetic waves contain both electric and magnetic components. Both the electrical and the magnetic fields oscillate perpendicular to the direction of wave propagation. In the case of unpolarized light, these fields oscillate in many directions. When the light is circularly polarized, two electromagnetic fields are obtained at 90&#176; phase difference to each other. Chiral molecules show circular optical rotation (birefringence) such that they will absorb the right-handed circularly polarized light and the left-handed circularly polarized light to different extents3. The resulting electrical field will be traced as an ellipse, a function of the wavelength. The CD spectrum is, thus, recorded as ellipticity (q), and the data are presented as Mean Residue Ellipticity as a function of wavelength.
In the case of proteins, the C&#945; of amino acids (except glycine) is chiral, and this is exploited by CD spectroscopy to determine the secondary structure of this macromolecule4. The CD spectra of protein molecules are typically recorded in the Far UV range. &#945;-helical proteins have two negative bands at 222 nm and 208 nm and one positive peak at 193 nm4. Proteins with anti-parallel &#946;-sheet secondary structure show a negative peak at 218 nm and a positive peak at 195 nm4. Proteins with disordered structures show low ellipticity near 210 nm and a negative peak at 195 nm4. Thus, the well-defined peak/bands for different secondary structures make CD a convenient tool to elucidate the conformational changes occurring in the secondary structure of the proteins during denaturation as well as ligand binding.
Nucleic acids have three sources of chirality: the sugar molecule, the helicity of the secondary structure, and the long-range tertiary ordering of DNA in the environment5,6. The CD spectra of nucleic acids are typically recorded in the 190 to 300 nm range5,6. Each conformation of DNA, just like proteins, gives a characteristic spectrum, although the peaks/bands can vary by some degrees due to solvent conditions and differences in DNA sequences7. B-DNA, the most common form, is characterized by a positive peak around 260-280 nm and a negative peak around 245 nm6. The peaks/bands of B-form DNA are generally small because the base pairs are perpendicular to the double helix, conferring weak chirality to the molecule. A-DNA gives a dominant positive peak at 260 nm and a negative peak around 210 nm6. Z-DNA, the left-handed helix, gives a negative band at 290 nm and a positive peak around 260 nm6. This DNA also gives an extremely negative peak at 205 nm6.
In addition to these conformations, DNA can also form triplexes, quadruplexes, and hairpins, all of which can be distinguished by CD spectroscopy. The parallel G-quadruplex give a dominant positive band at 260 nm, while the anti-parallel G-quadruplex gives a negative band at 260 nm and a positive peak at 290 nm, making it easy to distinguish between the two forms of quadruplex structures6. Triplexes do not give a characteristic spectrum8. For example, the spectra of a 36 nucleotide-long DNA with the potential to form an intramolecular triple helix containing G.G.C and T.A.T base pairs in the presence of Na+ show a strong negative band at 240 nm and a broad positive peak. The broad positive peak shows contributions at 266, 273, and 286 nm. The same oligonucleotide in the presence of Na+ and Zn+ shows four negative bands (213, 238, 266, and 282 nm) and a positive peak at 258 nm. Thus, the spectra of triplex DNA can vary depending upon salt conditions8.
In addition to these conformations, CD spectra have enabled the identification of another form of DNA called X-DNA. X-DNA is formed when the DNA sequence contains alternate adenine and thymine residues. The CD spectra of X-DNA contain two negative peaks at 250 and 280 nm. Very little information is available about X-DNA, although it has been speculated to function as a sink for positive supercoiling6,9. Changes in CD spectra can also reveal details about ligand-protein interactions and, therefore, have been added to the arsenal of molecular methods for detecting drug-protein interactions10,11,12,13,14. CD spectra have also been used to monitor the changes in the secondary structure of proteins during the folding process15. Similarly, CD spectra can also be used for probing ligand-DNA interactions16,17.
CD spectroscopy, thus, is an easy, inexpensive method to distinguish between the different forms of DNA conformation, provided there is access to not-so-inexpensive equipment and software. The method is exceedingly sensitive and quick. It only requires a small amount of DNA, giving it an edge over the alternate technique of nuclear magnetic resonance (NMR) spectroscopy. Titrations with ligands and substrates are also easy to perform. The major constraint is that the DNA should be highly pure. It is advisable to use polyacrylamide gel electrophoresis (PAGE)-purified DNA.
The information obtained by CD spectra has been mainly used to deduce protein structural features and to identify distinct DNA conformers. In this study, CD spectra have been used to integrate the results obtained from an in vivo Chromatin Immunoprecipitation (ChIP) experiment to delineate whether the protein of interest/predicted transcription factor can bring about a conformational change in the promoter region of its effector genes. This collaboration aids in the progress of traditional CD spectroscopic techniques by predicting the mechanism of transcription regulation by the predicted transcription factor on and around the transcription start site (TSS) of a promoter.
Chromatin remodeling is a well-defined mechanism known to regulate DNA metabolic processes by making the tightly packed chromatin accessible to various regulatory factors such as transcription factors, components of DNA replication, or damage repair proteins. The ATP-dependent chromatin remodelers, also known as the SWI/SNF family of proteins, are key remodeler proteins present in eukaryotic cells18,19. Phylogenetic clustering has categorized the SWI/SNF family of proteins into 6 sub-groups20: Snf2-like, Swr1-like, SSO1653-like, Rad54-like, Rad5/16-like, and distant. SMARCAL1, the protein of interest in this study, belongs to the distant sub-group20. This protein has been used to investigate its mode of transcriptional regulation using CD spectroscopy.
Most of the members of the ATP-dependent chromatin remodeling proteins have been shown to either reposition or evict nucleosomes or mediate histone variant exchange in an ATP-dependent manner21,22. However, some members of this family have not been shown to remodel nucleosomes, e.g., SMARCAL1. Even though studies have shown that SMARCAL1 associates with polytene chromosomes, experimental evidence regarding its ability to remodel nucleosomes is lacking23. Therefore, it was postulated that SMARCAL1 may regulate transcription by altering the conformation of DNA24. CD spectroscopy provided an easy and accessible method to validate this hypothesis.
SMARCAL1 is an ATP-dependent chromatin remodeling protein that primarily functions as an annealing helicase25,26,27. It has been postulated to modulate transcription by remodeling the DNA conformation24. To test this hypothesis, the role of SMARCAL1 in regulating gene transcription during doxorubicin-induced DNA damage was studied. In these studies, SMARCAL1 was used for in vivo analysis and ADAAD for in vitro assays28,29. Previous studies have shown that ADAAD can recognize DNA in a structure-dependent but sequence-independent manner30,31. The protein binds optimally to DNA molecules possessing double-strand to single-strand transition regions, similar to stem-loop DNA, and hydrolyzes ATP 30,31.
In vivo experiments showed that SMARCAL1 regulates the expression of MYC, DROSHA, DGCR8, and DICER by binding to the promoter regions28,29. The region of interaction was identified by ChIP experiments28,29. The ChIP technique is used to analyze the interaction of a protein with its cognate DNA within the cell. Its goal is to determine whether specific proteins, such as transcription factors on promoters or other DNA binding sites, are bound to specific genomic areas. The protein bound to DNA is first cross-linked using formaldehyde. This is followed by isolation of the chromatin. The isolated chromatin is sheared to 500 bp fragments either by sonication or nuclease digestion, and the protein bound to DNA is immunoprecipitated using antibodies specific to the protein. The cross-linking is reversed, and the DNA is analyzed using either polymerase chain reaction (PCR) or quantitative real-time PCR.
The ChIP results led to the hypothesis that SMARCAL1 possibly mediates transcriptional regulation by inducing a conformational change in the promoter regions of these genes. QGRS mapper and Mfold software were used to identify the potential of these promoter regions to form secondary structures28,29. QGRS mapper is used for predicting G-quadruplexes32, while Mfold33 analyzes the ability of a sequence to form secondary structures such as stem-loops.
After secondary structure analysis, further in vitro experiments were performed with recombinant 6X His-tagged Active DNA-dependent ATPase A Domain (ADAAD), the bovine homolog of SMARCAL1, purified from Escherichia coli34. ATPase assays were performed using ADAAD to establish that the identified DNA sequences could act as effectors28,29. Finally, CD spectroscopy was performed to monitor the conformational changes induced in the DNA molecule by ADAAD28,29.
To prove that the ATPase activity of the protein was essential for inducing a conformational change in the DNA molecule, either ethylenediamine tetraacetic acid (EDTA) was added to chelate Mg+2 or Active DNA-dependent ATPase A Domain Inhibitor Neomycin (ADAADiN), a specific inhibitor of the SWI/SNF protein, was added35,36. This CD spectroscopic technique can be utilized with any purified protein that has been demonstrated by ChIP or any other relevant assay to bind to a predicted genomic region of a promoter.

<table><tbody><tr><td>2-Mercaptoethanol</td><td>Fisher scientific</td><td>O3446I-100</td><td></td></tr><tr><td>Adenosine 5&amp;prime;-triphosphate disodium salt hydrate</td><td>Sigmaaldrich</td><td>A2383</td><td></td></tr><tr><td>CD Quartz Cuvette</td><td>STARNA</td><td>21-Q-1</td><td></td></tr><tr><td>Chirascan V100 CD spectrometer</td><td>Applied Photophysics</td><td>Not available</td><td></td></tr><tr><td>EDTA Disodium Salt Dihydrate</td><td>SRL</td><td>43272</td><td></td></tr><tr><td>Glutathione Sepharose 4B</td><td>GE Healthcare</td><td>17-0756-01</td><td>Glutathione affinity chromatography</td></tr><tr><td>Hellmanex III cleaning solution</td><td>Hellma</td><td>9-307-011-4-507</td><td></td></tr><tr><td>L-Lactic Dehydrogenase</td><td>Sigmaaldrich</td><td>&amp;nbsp;L2625</td><td></td></tr><tr><td>Magnesium Acetate Tetrahydrate</td><td>Fisher scientific</td><td>BP215-500</td><td></td></tr><tr><td>Magnesium Chloride Hexahydrate</td><td>Fisher scientific</td><td>M33-500</td><td></td></tr><tr><td>NADH disodium salt</td><td>Sigmaaldrich</td><td>10107735001</td><td></td></tr><tr><td>Phosphoenolpyruvate Monocyclohexylammonium Salt</td><td>SRL</td><td>40083</td><td></td></tr><tr><td>Potassium Acetate</td><td>Fisher scientific</td><td>P178-3</td><td></td></tr><tr><td>Pyruvate Kinase</td><td>Sigmaaldrich</td><td>P1506</td><td></td></tr><tr><td>Sodium Phosphate Dibasic Anhydrous</td><td>Fisher scientific</td><td>S374-500</td><td></td></tr><tr><td>Sodium Phosphate Monobasic Monohydrate</td><td>Fisher scientific</td><td>S369-500</td><td></td></tr><tr><td>Synergy HT microplate reader</td><td>BioTek</td><td>Not available</td><td></td></tr><tr><td>Tris Base</td><td>Fisher scientific</td><td>BP152-500</td><td></td></tr></tbody></table>

cd spectroscopy to study dna-protein interactions

Circular dichroism (CD) spectroscopy is a simple and convenient method to investigate the secondary structure and interactions of biomolecules. Recent advancements in CD spectroscopy have enabled the study of DNA-protein interactions and conformational dynamics of DNA in different microenvironments in detail for a better understanding of transcriptional regulation in vivo. The area around a potential transcription zone needs to be unwound for transcription to occur. This is a complex process requiring the coordination of histone modifications, binding of the transcription factor to DNA, and other chromatin remodeling activities. Using CD spectroscopy, it is possible to study conformational changes in the promoter region caused by regulatory proteins, such as ATP-dependent chromatin remodelers, to promote transcription. The conformational changes occurring in the protein can also be monitored. In addition, queries regarding the affinity of the protein towards its target DNA and sequence specificity can be addressed by incorporating mutations in the target DNA. In short, the unique understanding of this sensitive and inexpensive method can predict changes in chromatin dynamics, thereby improving the understanding of transcriptional regulation.

ADAAD stabilizes a stem-loop like structure on the MYC promoter 
Previous experimental evidence showed that SMARCAL1 is a negative regulator of MYC29. Analysis of the 159 bp long promoter region of the MYC gene by QGRS mapper showed that the forward strand had the potential to form a G-quadruplex (Table 2). Mfold showed that both strands of the MYC DNA could form a stem-loop-like structure (Table 2). A 34 bp long DNA sequence containing the G-quadruplex (GECE) was synthesized. The Mfold structures of the forward and the reverse sequence of the GECE oligonucleotide are shown in Figure 1A,B.
ATPase assays using 6X His-ADAAD showed that fast-cooled GECE was a better effector than the native and the slow-cooled forms. Therefore, fast-cooled GECE was used to record the CD spectra in the absence and presence of ATP and ADAAD. The CD spectra showed that ADAAD induces two positive peaks-one at 258 nm with a shoulder at 269 nm and a larger peak at 210 nm in the DNA (Figure 1C). A dip towards the negative around 240 nm was also observed. This spectrum was similar to the one obtained when a synthetic stem-loop DNA, the optimal effector of ADAAD, was incubated with the protein and ATP (Figure 2A,B). Triplex DNA can give a similar spectrum37, leading to the hypothesis that the protein could be inducing such a structure in this case. ATP forms a coordination complex with Mg+2, and this cation is essential for ATP hydrolysis. The addition of EDTA chelates Mg+2, leading to the inhibition of ATP hydrolysis38. Therefore, EDTA was added to the reaction mix to understand whether ATP hydrolysis by ADAAD was important for conformational change. The addition of EDTA to the reaction abrogates this conformation. The CD spectra now have a negative 210 nm peak and a broad positive band with peaks at 230 and 250 nm (Figure 1C).
The importance of ATPase activity was also confirmed using an ATPase-dead mutant of ADAAD. The K241A mutation occurs in the conserved GKT box of motif I, and this mutant has been shown previously to lack the ability to hydrolyze ATP in the presence of DNA31. The mutant protein was expressed with a GST tag and purified using glutathione affinity chromatography. The conformational change induced in MYC DNA by this mutant was different from that induced by the wild-type ADAAD. The CD spectrum of the MYC DNA in the presence of the mutant protein possessed a positive 210 nm peak and a negative 260 nm peak (Figure 1D).
ADAAD induces A-form of conformation in DROSHA promoter 
The promoter regions of DROSHA, DGCR8, and DICER too were analyzed by QGRS mapper and Mfold software. Both QGRS and MFold showed that the promoter regions possess the potential to form G-quadruplex and stem-like structures (Table 2). The Mfold structures of the forward and reverse oligonucleotides are shown in Figure 3A,B. The ATPase activity showed that the native and heat-cooled DNA behaved similarly. Therefore, the slow-cooled form of these DNA sequences was used for CD studies. The CD spectra showed that ADAAD induces a negative peak at 210 nm and a positive peak at 260 nm in the DROSHA promoter (Figure 3C). This spectrum is a characteristic of A-DNA6.
ADAAD induces B-X transition and G-quadruplex formation in the DGCR8 promoter 
The Mfold structures of the forward and the reverse strands of the oligonucleotides used are shown in Figure 4A,B. A positive peak at 210 nm and a broad negative peak at 260 nm were observed for DGCR8 pair 1 (Figure 4C). This spectrum is characteristic of B-X transition6. The Mfold structures of the forward and the reverse strands of the oligonucleotides used are shown in Figure 4D,E. The CD spectra of DGCR8 pair 7 showed a strong positive peak at 210 and 270 nm and a negative peak at 250 nm (Figure 4F). This spectrum is characteristic of parallel G-quadruplex DNA structures6.
ADAAD induces A-X transition in DICER promoter 
The Mfold structures of the forward and reverse oligonucleotides are shown in Figure 5A,B. A positive peak at 210 nm and two negative peaks-one at 230 nm and the other at 260 nm peak-were observed for the DICER pair 1 (Figure 5C). These peaks are characteristic of A-X DNA transition6,9. All the CD spectra peaks and the forms of DNA having specific roles in the transcription process have been summarized in Table 3.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63147/63147fig01.jpg" /> 
Figure 1: ADAAD alters the conformation of GECE DNA. Mfold structures were predicted for the (A) forward strand and (B) reverse strand. (C) CD spectra of GECE alone (black), GECE incubated with ATP and ADAAD before (red) and after adding EDTA (blue). (D) CD spectra of GECE incubated with ATP and GST-tagged ADAAD before (black) and after adding EDTA (red) as well as CD spectra of GECE incubated with ATP and GST-tagged K241A mutant (blue). This figure has been modified from 29. Abbreviations: ADAAD = Active DNA-dependent ATPase A Domain; CD = circular dichroism. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63147/63147fig01large.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/63147/63147fig02.jpg" /> 
Figure 2: ADAAD alters the conformation of slDNA. (A) Mfold structure predicted for the stem-loop DNA. (B) CD spectra of slDNA alone (black), slDNA incubated with ATP and ADAAD (red). This figure has been modified from29. Abbreviations: ADAAD = Active DNA-dependent ATPase A Domain; slDNA = stem-loop DNA; CD = circular dichroism. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63147/63147fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63147/63147fig03.jpg" /> 
Figure 3: ADAAD alters the conformation of DROSHA pair 5 DNA. Mfold structure predicted for the (A) forward strand and (B) reverse strand. (C) CD spectra of DROSHA pair 5 DNA alone (black), DROSHA pair 5 DNA incubated with ATP and ADAAD (red). This figure has been modified from28. Abbreviations: ADAAD = Active DNA-dependent ATPase A Domain; CD = circular dichroism. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63147/63147fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63147/63147fig04.jpg" /> 
Figure 4: ADAAD alters the conformation of DGCR8 pair 1 and 7 DNA. Mfold structures predicted for the (A) forward strand and (B) reverse strand of DGCR8 pair 1 oligonucleotide. (C) CD spectra of DGCR8 pair 1 alone (black), DGCR8 pair 1 incubated with ATP and ADAAD (red). Mfold structures predicted for the (D) forward strand and (E) reverse strand of DGCR8 pair 7 oligonucleotide. (F) CD spectra of DGCR8 pair 7 alone (black), DGCR8 pair 7 incubated with ATP and ADAAD (red). This figure has been modified from28. Abbreviations: ADAAD = Active DNA-dependent ATPase A Domain; CD = circular dichroism. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63147/63147fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63147/63147fig05.jpg" /> 
Figure 5: ADAAD alters the conformation of DICER pair 1 DNA. Mfold structure predicted for the (A) forward strand and (B) reverse strand of DICER pair 1 oligonucleotide. (C) CD spectra of DICER pair 1 alone (black), DICER pair 1 incubated with ATP and ADAAD (red). This figure has been modified from28. Abbreviations: ADAAD = Active DNA-dependent ATPase A Domain; CD = circular dichroism. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63147/63147fig05large.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 colspan="2">5x REG buffer</td>
		</tr>
		<tr>
			<td>Component</td>
			<td>Working concentration</td>
		</tr>
		<tr>
			<td>Lactate dehydrogenase (LDH)</td>
			<td>50 units/mL</td>
		</tr>
		<tr>
			<td>Magnesium acetate (Mg(OAc)2)</td>
			<td>30 mM</td>
		</tr>
		<tr>
			<td>Phosphoenolpyruvate (PEP)</td>
			<td>6.8 mg/mL</td>
		</tr>
		<tr>
			<td>Pottasium acetate (KOAc)</td>
			<td>300 mM</td>
		</tr>
		<tr>
			<td>Pyruvate kinase (PK)</td>
			<td>50 units/mL</td>
		</tr>
		<tr>
			<td>Tris acetate (Tris-OAc)</td>
			<td>125 mM</td>
		</tr>
		<tr>
			<td>&#946;-mercaptoethanol (&#946;-ME)</td>
			<td>25 mM</td>
		</tr>
	</tbody>
</table>
Table 1: Buffer components.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Oligonucleotides</td>
			<td colspan="2">Forward sequence</td>
			<td>&#916;G (m-Fold) Kcal/mol</td>
			<td>G-score (QGRS mapper)</td>
			<td colspan="2">Reverse sequence</td>
			<td>&#916;G kcal/mol (Mfold prediction)</td>
			<td>G-score (QGRS mapper)</td>
		</tr>
		<tr>
			<td colspan="2">slDNA</td>
			<td colspan="2">GCGCAATTGCGCTCGA 
			CGATTTTTTAGCGCAA 
			TTGCGC</td>
			<td>-16.36</td>
			<td>-</td>
			<td colspan="2">-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td colspan="2">MYC GECE</td>
			<td colspan="2">CGCGCGTGGCGTGG 
			CGGTGGGCGCGCA 
			GTGCGTT</td>
			<td>-8.64</td>
			<td>19</td>
			<td colspan="2">AACGCACTGCGCGCC 
			CACCGCCACGCCA 
			CGCGCG</td>
			<td>-5.74</td>
			<td>-</td>
		</tr>
		<tr>
			<td colspan="2">DROSHA Pair 5</td>
			<td colspan="2">GGCCAGGCACGGTG 
			GCTTATGCCTGTAAT 
			CCCAGCCCTTTGGGA 
			GGCTGAGGCAGG</td>
			<td>-10.18</td>
			<td>19, 19</td>
			<td colspan="2">CCTGCCTCAGCCTCC 
			CAAAGGGCTGGGATT 
			ACAGGCATAAGCCAC 
			CGTGCCTGGCC</td>
			<td>-10.95</td>
			<td>-</td>
		</tr>
		<tr>
			<td colspan="2">DGCR8 Pair 1</td>
			<td colspan="2">GCCACCGTGCCCGG 
			CCGAACCACCGTGC 
			CCGGCCGAACCC</td>
			<td>-7.9</td>
			<td>-</td>
			<td colspan="2">GGGTTCGGCCGGG 
			CACGGTGGTTCGG 
			CCGGGCACGGTGGC</td>
			<td>-9.4</td>
			<td>21</td>
		</tr>
		<tr>
			<td colspan="2">DGCR8 Pair 7</td>
			<td colspan="2">TCATACTGCCGCTG 
			GGTTGGGCGGGAG 
			GCTGCAGCGGGAG</td>
			<td>-7.99</td>
			<td>33</td>
			<td colspan="2">TCCCGCTGCAGCCT 
			CCCGCCCAACCCAG 
			CGGCAGTATGAC</td>
			<td>-5.43</td>
			<td>-</td>
		</tr>
		<tr>
			<td colspan="2">DICER Pair 1</td>
			<td colspan="2">AAGTGCTCAGGAG 
			CCATGTGGGGCTG 
			GTGGCCCCAAACTG</td>
			<td>-8.82</td>
			<td>19</td>
			<td colspan="2">CAGTTTGGGGCCAC 
			CAGCCCCACATGGC 
			TCCTGAGCACTT</td>
			<td>-9.16</td>
			<td>-</td>
		</tr>
	</tbody>
</table>
Table 2:&#160;Oligonucleotide sequences. All the sequences are in the 5&#39;-3&#39; direction. Abbreviations: slDNA = stem-loop DNA.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Oligonucleotides</td>
			<td>CD Spectra Peaks in nm (After incubating with ATP and ADAAD)</td>
			<td>Form of DNA</td>
			<td>Role in Transcription</td>
		</tr>
		<tr>
			<td>slDNA</td>
			<td>+215, +250, +272</td>
			<td>Triplex</td>
			<td>Repression</td>
		</tr>
		<tr>
			<td>MYC GECE</td>
			<td>+210, +258, +269</td>
			<td>Triplex</td>
			<td>Repression</td>
		</tr>
		<tr>
			<td>DROSHA Pair 5</td>
			<td>-210, +260</td>
			<td>A-DNA</td>
			<td>Initiation/Activation</td>
		</tr>
		<tr>
			<td>DGCR8 Pair 1</td>
			<td>+210,-260</td>
			<td>B-X Transition</td>
			<td>Positive supercoiling/Activation</td>
		</tr>
		<tr>
			<td>DGCR8 Pair 7</td>
			<td>+210, -250, +270</td>
			<td>G-quadruplex</td>
			<td>Activation</td>
		</tr>
		<tr>
			<td>DICER Pair 1</td>
			<td>+210, -230, -257</td>
			<td>A-X Transition</td>
			<td>Positive supercoiling/Activation</td>
		</tr>
	</tbody>
</table>
Table 3:&#160;CD spectra peak corresponding to different forms of DNA with their role in transcription. Abbreviations: ADAAD = Active DNA-dependent ATPase A Domain; CD = circular dichroism.

Watch this Scientific Journal Video about CD Spectroscopy to Study DNA-Protein Interactions at JoVE.com

CD Spectroscopy to Study DNA-Protein Interactions

The interaction of an ATP-dependent chromatin remodeler with a DNA ligand is described using CD spectroscopy. The induced conformational changes on a gene promoter analyzed by the peaks generated can be used to understand the mechanism of transcriptional regulation.

Circular dichroism (CD) spectroscopy is a simple and convenient method to investigate the secondary structure and interactions of biomolecules. Recent ...

cd-spectroscopy-to-study-dna-protein-interactions

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Biochemistry

6.6K Views.  Jawaharlal Nehru University New Delhi. Het protocol stelt ons in staat om de veranderingen in de tweede herstructurering van DNA geïnduceerd door ATP-afhankelijke chromatineremodelleringseiwitten te visualiseren. De verandering in de DNA-structuur kan worden gecorreleerd aan transcriptieregulatie. Dit is een eenvoudige en zeer gevoelige techniek die een zeer kleine hoeveelheid zuiver DNA gebruikt om de structurele veranderingen in het oligonucleotide vast te leggen.Deze methode kan inzicht geven in transcriptieregulatie ge...

Video: CD Spectroscopy to Study DNA-Protein Interactions

Measuring Interactions of Globular and Filamentous Proteins by Nuclear Magnetic Resonance Spectroscopy (NMR) and Microscale Thermophoresis (MST)

Filamentous proteins such as vimentin provide organization within cells by providing a structural scaffold with sites that bind proteins containing plakin repeats. Here, a protocol for detecting and measuring such interactions is described using the globular plakin repeat domain of envoplakin and the helical coil of vimentin. This provides a basis for determining whether a protein binds vimentin (or similar filamentous proteins) and for measurement of the affinity of the interaction. The globular protein of interest is labeled with 15N and titrated with vimentin protein in solution. A two-dimensional NMR spectrum is acquired to detect interactions by observing changes in peak shape or chemical shifts, and to elucidate effects of solution conditions including salt levels, which influence vimentin quaternary structure. If the protein of interest binds the filamentous ligand, the binding interaction is quantified by MST using the purified proteins. The approach is a straightforward way for determining whether a protein of interest binds a filament, and for assessing how alterations, such as mutations or solution conditions, affect the interaction.

Measuring Interactions of Globular and Filamentous Proteins by Nuclear Magnetic Resonance Spectroscopy NMR and Microscale Thermophoresis MST

Here, we present a protocol for the production and purification of proteins that are labeled with stable isotopes, and subsequent characterization of protein-protein interactions using Nuclear Magnetic Resonance (NMR) spectroscopy and MicroScale Thermophoresis (MST) experiments.

Filamentous proteins such as vimentin provide organization within cells by providing a structural scaffold with sites that bind proteins containing plakin ...

DNA Curtains Shed Light on Complex Molecular Systems During Homologous Recombination

Homologous recombination (HR) is important for the repair of double-stranded DNA breaks (DSBs) and stalled replication forks in all organisms. Defects in HR are closely associated with a loss of genome integrity and oncogenic transformation in human cells. HR involves coordinated actions of a complex set of proteins, many of which remain poorly understood. The key aspect of the research described here is a technology called "DNA curtains", a technique which allows for the assembly of aligned DNA molecules on the surface of a microfluidic sample chamber. They can then be visualized by total internal reflection fluorescence microscopy (TIRFM). DNA curtains was pioneered by our laboratory and allows for direct access to spatiotemporal information at millisecond time scales and nanometer scale resolution, which cannot be easily revealed through other methodologies. A major advantage of DNA curtains is that it simplifies the collection of statistically relevant data from single molecule experiments. This research continues to yield new insights into how cells regulate and preserve genome integrity.

DNA curtains present a novel method for visualizing hundreds or even thousands of DNA-binding proteins in real-time as they interact with DNA molecules aligned on the surface of a microfluidic sample chamber.

Homologous recombination (HR) is important for the repair of double-stranded DNA breaks (DSBs) and stalled replication forks in all organisms. Defects in ...

Genetics

Deciphering Molecular Mechanism of Histone Assembly by DNA Curtain Technique

Chromatin is a higher-order structure that packages eukaryotic DNA. Chromatin undergoes dynamic alterations according to the cell cycle phase and in response to environmental stimuli. These changes are essential for genomic integrity, epigenetic regulation, and DNA metabolic reactions such as replication, transcription, and repair. Chromatin assembly is crucial for chromatin dynamics and is catalyzed by histone chaperones. Despite extensive studies, the mechanisms by which histone chaperones enable chromatin assembly remains elusive. Moreover, the global features of nucleosomes organized by histone chaperones are poorly understood. To address these problems, this work describes a unique single-molecule imaging technique named DNA curtain, which facilitates the investigation of the molecular details of nucleosome assembly by histone chaperones. DNA curtain is a hybrid technique that combines lipid fluidity, microfluidics, and total internal reflection fluorescence microscopy (TIRFM) to provide a universal platform for real-time imaging of diverse protein-DNA interactions.Using DNA curtain, the histone chaperone function of Abo1, the Schizosaccharomyces pombe bromodomain-containing AAA+ ATPase, is investigated, and the molecular mechanism underlying histone assembly of Abo1 is revealed. DNA curtain provides a unique approach for studying chromatin dynamics.

DNA curtain, a high-throughput single-molecule imaging technique, provides a platform for real-time visualization of diverse protein-DNA interactions. The present protocol utilizes the DNA curtain technique to investigate the biological role and molecular mechanism of Abo1, a Schizosaccharomyces pombe bromodomain-containing AAA+ ATPase.

Chromatin is a higher-order structure that packages eukaryotic DNA. Chromatin undergoes dynamic alterations according to the cell cycle phase and in ...

Measuring Ubiquitylated, SUMOylated, and ADP-Ribosylated DNA-Protein Crosslinks

Quantitative Detection of DNA-Protein Crosslinks and Their Post-Translational Modifications

DNA-protein crosslinks (DPCs) are frequent, ubiquitous, and deleterious DNA lesions, which arise from endogenous DNA damage, enzyme (topoisomerases, methyltransferases, etc.) malfunctioning, or exogenous agents such as chemotherapeutics and crosslinking agents. Once DPCs are induced, several types of post-translational modifications (PTMs) are promptly conjugated to them as early response mechanisms. It has been shown that DPCs can be modified by ubiquitin, small ubiquitin-like modifier (SUMO), and poly-ADP-ribose, which prime the substrates to signal their respective designated repair enzymes and, in some cases, coordinate the repair in sequential manners. As PTMs transpire quickly and are highly reversible, it has been challenging to isolate and detect PTM-conjugated DPCs that usually remain at low levels. Presented here is an immunoassay to purify and quantitatively detect ubiquitylated, SUMOylated, and ADP-ribosylated DPCs (drug-induced topoisomerase DPCs and aldehyde-induced non-specific DPCs) in vivo. This assay is derived from the RADAR (rapid approach to DNA adduct recovery) assay that is used for the isolation of genomic DNA containing DPCs by ethanol precipitation. Following normalization and nuclease digestion, PTMs of DPCs, including ubiquitylation, SUMOylation, and ADP-ribosylation, are detected by immunoblotting using their corresponding antibodies. This robust assay can be utilized to identify and characterize novel molecular mechanisms that repair enzymatic and non-enzymatic DPCs&#160;and has the potential to discover small molecule inhibitors targeting specific factors that regulate PTMs to repair DPCs.

Author Spotlight: Quantitative Detection of DNA Protein Crosslinks and Their Post-Translational Modifications  

The present protocol highlights a modified method to detect and quantify DNA-protein crosslinks (DPCs) and their post-translational modifications (PTMs), including ubiquitylation, SUMOylation, and ADP-ribosylation induced by topoisomerase inhibitors and by formaldehyde, thereby allowing the study of the formation and repair of DPCs and their PTMs.

DNA-protein crosslinks (DPCs) are frequent, ubiquitous, and deleterious DNA lesions, which arise from endogenous DNA damage, enzyme (topoisomerases, ...

Quantitative Analysis of RPA-ssDNA Binding Kinetics Using Label-Free Methods

Analyzing DNA-Protein Interactions with Streptavidin-Based Biolayer Interferometry

Protein-DNA interactions underpin essential cellular processes. Understanding these interactions is critical for elucidating the molecular mechanisms of various pathways. Key factors such as the structure, sequence, and length of a DNA molecule can significantly influence protein binding. Bio-layer interferometry (BLI) is a label-free technique that measures binding kinetics between molecules, offering a straightforward and precise approach to quantitatively study protein-DNA interactions. A major advantage of BLI over traditional gel-based methods is its ability to provide real-time data on binding kinetics, enabling accurate measurement of the equilibrium dissociation constant (KD) for dynamic protein-DNA interactions. This article presents a basic protocol for determining the KD value of the interaction between a DNA replication protein, replication protein A (RPA), and a single-stranded DNA (ssDNA) substrate. RPA binds ssDNA with high affinity but must also be easily displaced to facilitate subsequent protein interactions within biological pathways. In the described BLI assay, biotinylated ssDNA is immobilized on a streptavidin-coated biosensor. The binding kinetics (association and dissociation) of RPA to the biosensor-bound DNA are then measured. The resulting data are analyzed to derive precise values for the association rate constant (ka), dissociation rate constant (kd), and equilibrium binding constant (KD) using system software.

This article describes a protocol for studying DNA-protein interactions using a streptavidin-based biolayer interferometry (BLI) system. It outlines the essential steps and considerations for utilizing either basic or advanced binding kinetics to determine the equilibrium binding affinity (KD) of the interaction.

Protein-DNA interactions underpin essential cellular processes. Understanding these interactions is critical for elucidating the molecular mechanisms of ...

Combining Single-molecule Manipulation and Imaging for the Study of Protein-DNA Interactions

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method offers the opportunity of investigating interactions occurring in solution (thus avoiding problems due to closeby surfaces as in other single molecule methods), controlling the DNA extension and tracking interaction dynamics as a function of both mechanical parameters and DNA sequence. The methods for establishing successful optical trapping and nanometer localization of single molecules are illustrated. We illustrate the experimental conditions allowing the study of interaction of lactose repressor (lacI), labeled with Atto532, with a DNA molecule containing specific target sequences (operators) for LacI binding. The method allows the observation of specific interactions at the operators, as well as one-dimensional diffusion of the protein during the process of target search. The method is broadly applicable to the study of protein-DNA interactions but also to molecular motors, where control of the tension applied to the partner track polymer (for example actin or microtubules) is desirable.

Here we describe the instrumentation and methods for detecting single fluorescently-labeled protein molecules interacting with a single DNA molecule suspended between two optically trapped microspheres.

The paper describes the combination of optical tweezers and single molecule fluorescence detection for the study of protein-DNA interaction. The method ...

Biology

Determination of Protein-ligand Interactions Using Differential Scanning Fluorimetry

A wide range of methods are currently available for determining the dissociation constant between a protein and interacting small molecules. However, most of these require access to specialist equipment, and often require a degree of expertise to effectively establish reliable experiments and analyze data. Differential scanning fluorimetry (DSF) is being increasingly used as a robust method for initial screening of proteins for interacting small molecules, either for identifying physiological partners or for hit discovery. This technique has the advantage that it requires only a PCR machine suitable for quantitative PCR, and so suitable instrumentation is available in most institutions; an excellent range of protocols are already available; and there are strong precedents in the literature for multiple uses of the method. Past work has proposed several means of calculating dissociation constants from DSF data, but these are mathematically demanding. Here, we demonstrate a method for estimating dissociation constants from a moderate amount of DSF experimental data. These data can typically be collected and analyzed within a single day. We demonstrate how different models can be used to fit data collected from simple binding events, and where cooperative binding or independent binding sites are present. Finally, we present an example of data analysis in a case where standard models do not apply. These methods are illustrated with data collected on commercially available control proteins, and two proteins from our research program. Overall, our method provides a straightforward way for researchers to rapidly gain further insight into protein-ligand interactions using DSF.

Differential scanning fluorimetry is a widely used method for screening libraries of small molecules for interactions with proteins. Here, we present a straightforward method to extend these analyses to provide an estimate of the dissociation constant between a small molecule and its protein partner.

A wide range of methods are currently available for determining the dissociation constant between a protein and interacting small molecules. However, most ...

Advanced Confocal Microscopy Techniques to Study Protein-protein Interactions and Kinetics at DNA Lesions

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological approach to analyzing protein kinetics at DNA lesions over time or protein-protein interactions on locally microirradiated chromatin. We also show how to recognize individual phases of the cell cycle using the Fucci cellular system to study cell-cycle-dependent protein kinetics at DNA lesions. A methodological description of the use of two UV lasers (355 nm and 405 nm) to induce different types of DNA damage is also presented. Only the cells microirradiated by the 405-nm diode laser proceeded through mitosis normally and were devoid of cyclobutane pyrimidine dimers (CPDs). We also show how microirradiated cells can be fixed at a given time point to perform immunodetection of the endogenous proteins of interest. For the DNA repair studies, we additionally describe the use of biophysical methods including FRAP (Fluorescence Recovery After Photobleaching) and FLIM (Fluorescence Lifetime Imaging Microscopy) in cells with spontaneously occurring DNA damage foci. We also show an application of FLIM-FRET (Fluorescence Resonance Energy Transfer) in experimental studies of protein-protein interactions.

Laser microirradiation is a useful tool for studies of DNA repair in living cells. A methodological approach for the use of UVA lasers to induce various DNA lesions is shown. We have optimized a method for local microirradiation that maintains the normal cell cycle; thus, irradiated cells proceed through mitosis.

Local microirradiation with lasers represents a useful tool for studies of DNA-repair-related processes in live cells. Here, we describe a methodological ...

Nuclear Magnetic Resonance to Study Atomic Level Protein-Protein Interactions

Turn on the airflow with the eject command &#34;ej&#34;. This will bring the sample up from the magnet. Now, place the sample within a spinner on top of the magnet in the opening. Insert with the command &#34;ij&#34;. Wait until the sample settles inside the magnet before proceeding.
Create a new data set using the &#34;edc&#34; command, and load Standard Proton NMR parameters by selecting the experiment &#34;ZGPR&#34;. Fill in the name, experiment number, and processed data folder number fields.
Select the solvent in the &#34;Set Solvent&#34; field and click on &#34;Execute &#39;getprosol&#39;&#34; to read standard probehead head and solvent-dependent parameters. Lock the sample to the deuterated solvent using the &#34;Lock&#34; command, and wait until it is finished sweeping and achieves lock.
Correct the resonance frequency of the magnet by tuning the sample using the automatic tuning command &#34;atma&#34;. Monitor the wobble curve until the automatic tuning is complete.
Shim the magnetic field using &#34;TOPSHIM&#34;. Shimming makes adjustment to the magnetic field to achieve uniformity around the sample. It is good practice to store the same values with the command &#34;wsh&#34; and read them using &#34;rsh&#34; before topshim if using the same or similar samples.
Now, adjust the receiver gain with the &#34;rga&#34; command to achieve the maximum signal-to-noise ratio. Place the center of the spectrum on the water resonance offset and set the 90-degree proton pulse at high power using &#34;calibo1p1&#34;.
Collect the proton spectrum using the Zero Go &#34;zg&#34; command and process with &#34;efp&#34;. This includes exponential multiplication, the free induction decay incorporating line broadening, Fourier transformation of FID, and &#34;pk&#34; to apply phase correction.
Apply the automatic phase correction &#34;apk&#34; and the automatic baseline correction &#34;absn&#34; using the polynomial without integration option. Create a new data set for the SOFAST HMBC experiment by selecting &#34;SFHMQC3GPPH&#34; in experiment.
Copy the optimized P1 and O1 from the proton spectrum and populate P1-dependent pulses by using the command &#34;getprosol 1H p1 plw1&#34; where P1 is the optimized P1 value and plw1 is the power level for P1. Now, optimize the CNST54 constant to set the offset for amide chemical shift. Also, optimize CNST55 to define the bandwidth in order to encompass the spectral regions of interest, allowing the receiver gain to be optimized.
To select these parameters, extract the first FID from the two-dimensional spectrum, and look for the observed signal to define them. In addition, vary the relaxation delay, number of scans, and dummy scans to obtain acceptable signal sensitivity with the command &#34;gs&#34;, which enables go and scan to monitor data quality in real-time. Finally, record the spectra using Zero Go &#34;zg&#34;.

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Biological Techniques

Biomolecular Interaction Detection Techniques

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Differential Scanning Calorimetry to Study DNA Aptamer-Thermolabile Ligand Interaction

Degas the buffer, biomolecule, and ligand solutions in a table top degasser prior to loading into the differential scanning calorimeter, or DSC. Unscrew the pressure handle from the DSC. Then, run silicone tubing from the working buffer, and attach it to the front flange of the reference capillary.
Create a bridge between the reference and sample capillaries by connecting the rear reference flange to the front sample flange. Next, attach a piece of silicone tubing to the rear sample flange that runs to a waste flask with a vacuum line attached. Turn on the vacuum line to flush the DSC with 200 milliliters of working buffer.
Begin by attaching roughly 3- to 5-centimeter sections of silicone tubing to the reference capillary flanges. Next, insert a 1-milliliter pipette tip into the silicone tubing of the rear flange. Draw 0.8 milliliters of working buffer with a pipette, and insert the pipette tip with buffer into the silicone tubing of the front reference flange.
Gently press the pipette plunger down to pass the working buffer through the front silicone tubing into the reference capillary and up into the attached pipette tip of the rear flange. Press the pipette plunger down until the working buffer level reaches just above the front silicone tubing. Then, release the pipette plunger until the working buffer level reaches just above the rear silicone tubing. Continue passing the working buffer back and forth in the reference capillary to purge the volume of bubbles.
Next, cap the rear pipette tip with a forefinger and gently pull up on the rear pipette tip and front pipette to remove them from the reference flanges with the silicone tubing attached. Load the sample capillary with working buffer as before.
Place a black plastic cap on the rear reference and sample flanges, leaving the front flanges uncovered. Attach the pressure handle to the DSC and then, open the DSC software and pressurize the instrument by clicking the red &#34;Up&#34; arrow at the top of the interface once the power reading has stabilized. The DSC power is indicated in a box at the top right of the interface, along with the instrument temperature and pressure reading.
Equilibrate the DSC with working buffer by performing a forward and reverse scan. In the &#34;Experimental Method&#34; tab on the left side of the screen, ensure that the &#34;Scanning&#34; option is selected to run the DSC in temperature-scanning mode.
In the &#34;Temperature parameters&#34; inset under the &#34;Experimental Method&#34; tab, click the button for heating. Enter 1 and 100 degrees Celsius for the lower and upper experimental temperatures, 1 degree Celsius per minute for the scan rate, and 60 seconds for the equilibration period. Click the &#34;Add Series&#34; button under the input field for the equilibration period. Enter 2 into the &#34;Steps to add&#34; field in the pop-up window and check the &#34;Alternate Heating/Cooling&#34; box. Click &#34;OK.&#34;
The added scans appear in the lower portion of the interface. Check that the parameters for each scan are as desired. Start the experiment by clicking the green &#34;Play&#34; button at the top of the interface. Navigate to the desired window and input a file name for saving the experiment in the pop-up window. View the experiment progress by clicking the &#34;Data&#34; tab to the right of the &#34;Experiment Method&#34; tab.
Run reference experiments for baseline subtraction of the sample data by reloading the DSC with working buffer in both capillaries. Collect multiple forward and reverse scans over a suitable temperature range at 1 degree Celsius per minute with an upper equilibration time of 120 seconds.
Delete the previous buffer equilibration scans from the lower portion of the interface by highlighting each individually and clicking the red &#34;X&#34; to the middle right of the interface.
Add the new scans by clicking the &#34;Add Series&#34; button, entering 20 in the field for &#34;Steps to add&#34; and checking the &#34;Alternate Heating/Cooling&#34; box. Then, click &#34;OK&#34; and run the experiment by clicking the green &#34;Play&#34; button as before.
Repeat these steps with working buffer containing the desired concentration of ligand in both capillaries to obtain the reference experiments for the ligand. Collect two separate experiments to be used in acquiring the rate constant for thermolabile ligand conversion.
For the free biomolecule data set, ensured that the reference capillary contains the working buffer, while the sample capillary contains the free biomolecule at the desired concentration in working buffer. Next, run the sample experiments using the same DSC-loading procedure and experimental parameters used in the reference scans.
For the ligand-bound experiments, ensure that the ligand is in the working buffer in the reference capillary and the biomolecule plus ligand are in the working buffer in the sample capillary. Flush the system between additions of different ligands as before.
Perform one additional experiment with the biomolecule bound to the thermolabile ligand where the high-temperature equilibration period is increased to 600 seconds, and all other experimental parameters are the same as before. Proceed to data processing and analysis as described in the text protocol.

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