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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-Mercaptoethanol</td>
			<td>Fisher scientific</td>
			<td>O3446I-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Adenosine 5&#8242;-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>&#160;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

6.5K 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

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

Protein-protein interactions play an essential role in the function of a living organism. Once an interaction has been identified and validated it is necessary to characterize it at the structural and mechanistic level. Several biochemical and biophysical methods exist for such purpose. Among them, fluorescence anisotropy is a powerful technique particularly used when the fluorescence intensity of a fluorophore-labeled protein remains constant upon protein-protein interaction. In this technique, a fluorophore-labeled protein is excited with vertically polarized light of an appropriate wavelength that selectively excites a subset of the fluorophores according to their relative orientation with the incoming beam. The resulting emission also has a directionality whose relationship in the vertical and horizontal planes defines anisotropy (r) as follows: r=(IVV-IVH)/(IVV+2IVH), where IVV and IVH are the fluorescence intensities of the vertical and horizontal components, respectively. Fluorescence anisotropy is sensitive to the rotational diffusion of a fluorophore, namely the apparent molecular size of a fluorophore attached to a protein, which is altered upon protein-protein interaction. In the present text, the use of fluorescence anisotropy as a tool to study protein-protein interactions was exemplified to address the binding between the protein mutated in the Shwachman-Diamond Syndrome (SBDS) and the Elongation factor like-1 GTPase (EFL1). Conventionally, labeling of a protein with a fluorophore is carried out on the thiol groups (cysteine) or in the amino groups (the N-terminal amine or lysine) of the protein. However, SBDS possesses several cysteines and lysines that did not allow site directed labeling of it. As an alternative technique, the dye 4&#39;,5&#39;-bis(1,3,2 dithioarsolan-2-yl) fluorescein was used to specifically label a tetracysteine motif, Cys-Cys-Pro-Gly-Cys-Cys, genetically engineered in the C-terminus of the recombinant SBDS protein. Fitting of the experimental data provided quantitative and mechanistic information on the binding mode between these proteins.

Protein interactions are at the heart of a cell&#39;s function. Calorimetric and spectroscopic techniques are commonly used to characterize them. Here we describe fluorescence anisotropy as a tool to study the interaction between the protein mutated in the Shwachman-Diamond Syndrome (SBDS) and the Elongation factor-like 1 GTPase (EFL1).

Protein-protein interactions play an essential role in the function of a living organism. Once an interaction has been identified and validated it is ...

Biotinylated Cell-penetrating Peptides to Study Intracellular Protein-protein Interactions

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The modification presented includes the combination of this technique with cell-penetrating sequences. We propose to design cell-penetrating baits that can be incubated with living cells instead of cell lysates and therefore the interactions found will reflect those that occur within the intracellular context. Connexin43 (Cx43), a protein that forms gap junction channels and hemichannels is down-regulated in high-grade gliomas. The Cx43 region comprising amino acids 266-283 is responsible for the inhibition of the oncogenic activity of c-Src in glioma cells. Here we use TAT as the cell-penetrating sequence, biotin as the pull-down tag and the region of Cx43 comprised between amino acids 266-283 as the target to find intracellular interactions in the hard-to-transfect human glioma stem cells. One of the limitations of the proposed method is that the molecule used as bait could fail to fold properly and, consequently, the interactions found could not be associated with the effect. However, this method can be especially interesting for the interactions involved in signal transduction pathways because they are usually carried out by intrinsically disordered regions and, therefore, they do not require an ordered folding. In addition, one of the advantages of the proposed method is that the relevance of each residue on the interaction can be easily studied. This is a modular system; therefore, other cell-penetrating sequences, other tags, and other intracellular targets can be employed. Finally, the scope of this protocol is far beyond protein-protein interaction because this system can be applied to other bioactive cargoes such as RNA sequences, nanoparticles, viruses or any molecule that can be transduced with cell-penetrating sequences and fused to pull-down tags to study their intracellular mechanism of action.

This is a protocol to study intracellular protein-protein interactions based on the biotin-avidin pull-down system with the novelty of combining cell-penetrating sequences. The main advantage is that the target sequence is incubated with living cells instead of cell lysates and therefore the interactions will occur within the cellular context.

Here we present a protocol to study intracellular protein-protein interactions that is based on the widely used biotin-avidin pull-down system. The ...

Using Three-color Single-molecule FRET to Study the Correlation of Protein Interactions

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For many molecular machines in a cell proteins have to act together&#160;with interaction partners in a functional cycle to fulfill their task. The extension of two-color to multi-color smFRET makes it possible to simultaneously probe more than one interaction or conformational change. This not only adds a new dimension to smFRET experiments but it also offers the unique possibility to directly study the sequence of events and to detect correlated interactions when using an immobilized sample and a total internal reflection fluorescence microscope (TIRFM). Therefore, multi-color smFRET is a versatile tool for studying biomolecular complexes in a quantitative manner and in a previously unachievable detail.
Here, we demonstrate how to overcome the special challenges of multi-color smFRET experiments on proteins. We present detailed protocols for obtaining the data and for extracting kinetic information. This includes trace selection criteria, state separation, and the recovery of state trajectories from the noisy data using a 3D ensemble Hidden Markov Model (HMM). Compared to other methods, the kinetic information is not recovered from dwell time histograms but directly from the HMM. The maximum likelihood framework allows us to critically evaluate the kinetic model and to provide meaningful uncertainties for the rates.
By applying our method to the heat shock protein 90 (Hsp90), we are able to disentangle the nucleotide binding and the global conformational changes of the protein. This allows us to directly observe the cooperativity between the two nucleotide binding pockets of the Hsp90 dimer.

Here, we present a protocol to obtain three-color smFRET data and its analysis with a 3D ensemble Hidden Markov Model. With this approach, scientists can extract kinetic information from complex protein systems, including cooperativity or correlated interactions.

Single-molecule F&#246;rster resonance energy transfer (smFRET) has become a widely used biophysical technique to study the dynamics of biomolecules. For ...

Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

Lactate and pyruvate are key intermediates of intracellular energy metabolic pathways. Monitoring the lactate/pyruvate ratio in cells helps to determine whether there is an imbalance in age-related energy metabolism between mitochondrial oxidative phosphorylation and aerobic glycolysis. Here, we show the utilization of commercial colorimetric assay kits for lactate and pyruvate in the model organism C. elegans. Recently, the sensitivity and accuracy of the colorimetric/fluorimetric assay kits have been improved greatly by the research and development conducted by reagent manufacturers. The improved reagents have enabled the use of small-scale assays with a 96-well plate in C. elegans. In general, a fluorimetric assay is superior in sensitivity to a colorimetric assay; however, the colorimetric approach is more suitable for the use in common laboratories. Another important issue in these assays for quantitative determination is protein precipitation of homogenized C. elegans samples. In our protein precipitation method, common precipitants (e.g., trichloroacetic acid, perchloric acid and metaphosphoric acid) are used for sample preparation. A protein-free assay sample is prepared by directly adding cold precipitant (final concentration of 5%) during homogenization.

Small-Scale Colorimetric Assays of Intracellular Lactate and Pyruvate in the Nematode Caenorhabditis elegans

We describe a modified small-scale extraction and colorimetric assays of lactate and pyruvate in the nematode C. elegans. When utilizing commercial assay kits, the technical development of their sensitivity and accuracy is important. Protein precipitation in extraction is the most critical step for the quantitative determination of intracellular metabolites.

Lactate and pyruvate are key intermediates of intracellular energy metabolic pathways. Monitoring the lactate/pyruvate ratio in cells helps to determine ...

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 ...

A Fluorescence Fluctuation Spectroscopy Assay of Protein-Protein Interactions at Cell-Cell Contacts

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring cells. Of interest, only few assays are capable of specifically probing such interactions directly in living cells. Here, we present an assay to measure the binding of proteins expressed at the surfaces of neighboring cells, at cell-cell contacts. This assay consists of two steps: mixing of cells expressing the proteins of interest fused to different fluorescent proteins, followed by fluorescence fluctuation spectroscopy measurements at cell-cell contacts using a confocal laser scanning microscope. We demonstrate the feasibility of this assay in a biologically relevant context by measuring the interactions of the amyloid precursor-like protein 1 (APLP1) across cell-cell junctions. We provide detailed protocols on the data acquisition using fluorescence-based techniques (scanning fluorescence cross-correlation spectroscopy, cross-correlation number and brightness analysis) and the required instrument calibrations. Further, we discuss critical steps in the data analysis and how to identify and correct external, spurious signal variations, such as those due to photobleaching or cell movement.
In general, the presented assay is applicable to any homo- or heterotypic protein-protein interaction at cell-cell contacts, between cells of the same or different types and can be implemented on a commercial confocal laser scanning microscope. An important requirement is the stability of the system, which needs to be sufficient to probe diffusive dynamics of the proteins of interest over several minutes.

This protocol describes a fluorescence fluctuation spectroscopy-based approach to investigate interactions among proteins mediating cell-cell interactions, i.e. proteins localized in cell junctions, directly in living cells. We provide detailed guidelines on instrument calibration, data acquisition and analysis, including corrections to possible artefact sources.

A variety of biological processes involves cell-cell interactions, typically mediated by proteins that interact at the interface between neighboring ...

An Integrated Raman Spectroscopy and Mass Spectrometry Platform to Study Single-Cell Drug Uptake, Metabolism, and Effects

Cells are known to be inherently heterogeneous in their responses to drugs. Therefore, it is essential that single-cell heterogeneity is accounted for in drug discovery studies. This can be achieved by accurately measuring the plethora of cellular interactions between a cell and drug at the single-cell level (i.e., drug uptake, metabolism, and effect). This paper describes a single-cell Raman spectroscopy and mass spectrometry (MS) platform to monitor metabolic changes of cells in response to drugs. Using this platform, metabolic changes in response to the drug can be measured by Raman spectroscopy, while the drug and its metabolite can be quantified using mass spectrometry in the same cell. The results suggest that it is possible to access information about drug uptake, metabolism, and response at a single-cell level.

This protocol presents an integrated Raman spectroscopy-mass spectrometry (MS) platform that is capable of achieving single-cell resolution. Raman spectroscopy can be used to study cellular response to drugs, while MS can be used for targeted and quantitative analysis of drug uptake and metabolism.

Cells are known to be inherently heterogeneous in their responses to drugs. Therefore, it is essential that single-cell heterogeneity is accounted for in ...

Use of Microscale Thermophoresis to Measure Protein-Lipid Interactions

The ability to determine the binding affinity of lipids to proteins is an essential part of understanding protein-lipid interactions in membrane trafficking, signal transduction and cytoskeletal remodeling. Classic tools for measuring such interactions include surface plasmon resonance (SPR) and isothermal titration calorimetry (ITC). While powerful tools, these approaches have setbacks. ITC requires large amounts of purified protein as well as lipids, which can be costly and difficult to produce. Furthermore, ITC as well as SPR are very time consuming, which could add significantly to the cost of performing these experiments. One way to bypass these restrictions is to use the relatively new technique of microscale thermophoresis (MST). MST is fast and cost effective using small amounts of sample to obtain a saturation curve for a given binding event. There currently are two types of MST systems available. One type of MST requires labeling with a fluorophore in the blue or red spectrum. The second system relies on the intrinsic fluorescence of aromatic amino acids in the UV range. Both systems detect the movement of molecules in response to localized induction of heat from an infrared laser. Each approach has its advantages and disadvantages. Label-free MST can use untagged native proteins; however, many analytes, including pharmaceuticals, fluoresce in the UV range, which can interfere with determination of accurate KD values. In comparison, labeled MST allows for a greater diversity of measurable pairwise interactions utilizing fluorescently labeled probes attached to ligands with measurable absorbances in the visible range as opposed to UV, limiting the potential for interfering signals from analytes.

Microscale thermophoresis obtains binding constants quickly at low material cost. Either labeled or label free microscale thermophoresis is commercially available; however, label free thermophoresis is not capable of the diversity of interaction measurements that can be performed using fluorescent labels. We provide a protocol for labeled thermophoresis measurements.

The ability to determine the binding affinity of lipids to proteins is an essential part of understanding protein-lipid interactions in membrane ...

In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans

Fast oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting (HRPF) method used to study protein structure, protein-ligand interactions, and protein-protein interactions. FPOP utilizes a KrF excimer laser at 248 nm for photolysis of hydrogen peroxide to generate hydroxyl radicals which in turn oxidatively modify solvent-accessible amino acid side chains. Recently, we expanded the use of FPOP of in vivo oxidative labeling in Caenorhabditis elegans (C. elegans), entitled IV-FPOP. The transparent nematodes have been used as model systems for many human diseases. Structural studies in C. elegans by IV-FPOP is feasible because of the animal&#8217;s ability to uptake hydrogen peroxide, their transparency to laser irradiation at 248 nm, and the irreversible nature of the modification. The assembly of a microfluidic flow system for IV-FPOP labeling, IV-FPOP parameters, protein extraction, and LC-MS/MS optimized parameters are described herein.

In Vivo Hydroxyl Radical Protein Footprinting for the Study of Protein Interactions in Caenorhabditis elegans

In vivo fast photochemical oxidation of proteins (IV-FPOP) is a hydroxyl radical protein footprinting technique that allows for mapping of protein structure in their native environment. This protocol describes the assembly and set-up of the IV-FPOP microfluidic flow system.

Fast oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting (HRPF) method used to study protein structure, protein-ligand interactions, ...

Optical Tweezers to Study RNA-Protein Interactions in Translation Regulation

RNA adopts diverse structural folds, which are essential for its functions and thereby can impact diverse processes in the cell. In addition, the structure and function of an RNA can be modulated by various trans-acting factors, such as proteins, metabolites or other RNAs. Frameshifting RNA molecules, for instance, are regulatory RNAs located in coding regions, which direct translating ribosomes into an alternative open reading frame, and thereby act as gene switches. They may also adopt different folds after binding to proteins or other trans-factors. To dissect the role of RNA-binding proteins in translation and how they modulate RNA structure and stability, it is crucial to study the interplay and mechanical features of these RNA-protein complexes simultaneously.&#160;This work illustrates how to employ single-molecule-fluorescence-coupled optical tweezers to explore the conformational and thermodynamic landscape of RNA-protein complexes at a high resolution. As an example, the interaction of the SARS-CoV-2 programmed ribosomal frameshifting element with the trans-acting factor short isoform of zinc-finger antiviral protein is elaborated. In addition, fluorescence-labeled ribosomes were monitored using the confocal unit, which would ultimately enable the study of translation elongation. The fluorescence coupled OT assay can be widely applied to explore diverse RNA-protein complexes or trans-acting factors regulating translation and could facilitate studies of RNA-based gene regulation.

This protocol presents a complete experimental workflow for studying RNA-protein interactions using optical tweezers. Several possible experimental setups are outlined including the combination of optical tweezers with confocal microscopy.