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...

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 ...

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

Research

JoVE Journal

Biochemistry

6.6K Views.  Jawaharlal Nehru University New Delhi. 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.

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;.

JoVE Encyclopedia of Experiments

Biological Techniques

Biomolecular Interaction Detection Techniques

Protein-protein interactions

EoE Sub Articles

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.