This work was funded by grants from the National Science Foundation (1929346) and the American Cancer Society (RSG-21-028-01). We would also like to thank members of the Balakrishnan laboratory for helpful discussions.

Details of the reagents and the equipment used in the study are listed in the Table of Materials.
1. Preparation of bait, analyte, buffers and drop-holder cleaning
<ol>
	<li>Setting up the instrument: Turn the instrument on at least 1 h before starting the experiment. This will allow the lamp to warm up.</li>
	<li>Preparation of stripping and cleaning buffers
	<ol>
		<li>Stripping buffer: Prepare 50 mL of 0.15 M Phosphoric acid [pH 2.0]).</li>
		<li>Cleaning buffer: Prepare 10 mL of 0.5 N HCl.</li>
	</ol>
	</li>
	<li>Preparation of BLI buffer
	<ol>
		<li>Prepare 50 mL of 1x BLI buffer (50 mM Tris pH 7.5, 1 mM EDTA, 100 mM NaCl, 0.1 mg/mL BSA, 1 mM DTT, 0.05% Tween 20). Store this buffer at 4 &#176;C. Before setting up this assay, bring the buffer to room temperature. 
		NOTE: It is highly recommended that fresh buffers be prepared for the assay, as aged buffers may undergo pH alterations or formation of aggregates and may have microbial contaminations, potentially compromising the reproducibility and accuracy of results. It is not recommended to use buffers that have been stored for more than two weeks.</li>
	</ol>
	</li>
	<li>Preparation of bait/substrate
	<ol>
		<li>Prepare 12.5 nM of 3&#39;Bio-ss-poly (dT)32 substrate (5&#39; TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TT 3&#39;Bio) using BLI buffer.</li>
	</ol>
	</li>
	<li>Preparation of protein concentration stocks
	<ol>
		<li>Prepare four concentrations of RPA (5 nM, 10 nM, 20 nM, 40 nM) using BLI buffer. The highest concentration of the protein (40 nM) is obtained by diluting the stock protein in BLI buffer. Subsequent dilutions are prepared by serial dilution of the 40 nM working solution. Ensure all protein stocks are kept on ice throughout the experiment to maintain stability.</li>
	</ol>
	</li>
	<li>Cleaning the drop holder
	<ol>
		<li>Add 4 &#181;L of distilled water and clean the drop holder with a lint-free wipe (repeat 3x). Next, repeat this step with 4&#181;L of 70% ethanol and clean with a lint-free wipe. This step ensures the removal of surface contaminants and dust particles.</li>
		<li>Add 4&#181;L of 0.5N HCl and leave it in the drop holder for 1 min. Clean it with a lint-free wipe and repeat this step once more. This ensures deep cleaning of the drop holder. Deep cleaning is especially useful to clean the drop holder of dried-up drops from previous use.</li>
		<li>1Repeat step 1 to wash the drop holder with distilled water and 70% ethanol and wipe it dry with a lint-free wipe. The drop holder is now ready to be used.</li>
	</ol>
	</li>
</ol>
2. Basic kinetics
<ol>
	<li>Setting up the instrument and the software
	<ol>
		<li>Open the software and select Basic Kinetics on the left panel (Supplementary Figure 1).</li>
		<li>Remove the rack of biosensors from the tray (obtained from commercial sources) and place a 96-well plate in the tray. Add 200 &#181;L of BLI buffer to the first well. Next, position the tray of biosensors back on top of the 96-well plate, ensuring each biosensor is inserted into its corresponding well. The first biosensor should now be submerged in the BLI buffer in the first well.</li>
		<li>On the software, click on Hydrate and turn the timer on for 10 min, allowing the single biosensor to hydrate for at least 10 min. The step hydrates the biosensor to make it ready for BLI assay.</li>
		<li>While the biosensor is being hydrated, pipette 250 &#181;L of BLI buffer into two 0.5 mL tubes. Label one as &#34;A&#34; (association) and the other one as &#34;D&#34; (dissociation).</li>
		<li>Fill in the details on the software (Name: Substrate Coating and description of the assay: RPA- 3&#39;Bio-ss-poly (dT)32: Basic Kinetics).</li>
		<li>Adjust the run setting for each step: Baseline (30 s), Association (120 s), Dissociation (120 s) (Table 1). Keep the shaker on for the tube and drop holder. The shaker speed is set at 2200 rpm (Range, 1000&#8211;2600 rpm), which prevents the mass transport effects caused by the limited diffusion near the biosensor surface. 
		NOTE: Optimizing the duration for each step (baseline, loading, associating, dissociating) when setting up a binding kinetics assay is recommended to improve data acquisition.</li>
	</ol>
	</li>
	<li>Establishing a baseline curve
	<ol>
		<li>Place tube A in the tube holder and attach the hydrated biosensor to the BLI system. Slide the tube holder under the biosensor (position A, Figure 2). 
		NOTE: Biosensors must not be completely dried at any time during the experiment. If left to dry, readings of the experiment can be significantly skewed.</li>
		<li>Hit Run and follow the instructions in the prompt message.</li>
		<li>After completing the initial baseline, open the lid and add 4 &#181;L of BLI buffer to the drop holder. Slide it to the right under the same biosensor (position B, Figure 2). Close the lid and monitor the BLI curve on the software (Figure 3, blue curve).</li>
		<li>After completion, open the lid to replace tube &#39;A&#39; with &#39;D&#39; and slide it under the biosensor (position A, Figure 2). Close the lid and proceed with the dissociation step. After completion, open the lid, detach the biosensor, place it back in the tray, and ensure the biosensor is dipped in BLI buffer.</li>
		<li>Remove the sample/buffer, clean the drop holder with a buffer before drying it with lint-free wipes, and remove tube &#39;D&#39; from the tube holder.</li>
	</ol>
	</li>
	<li>Substrate coating
	<ol>
		<li>Place tube A in the tube holder and attach the hydrated biosensor to the BLI system. Slide the tube holder under the biosensor and hit Run. </li>
		<li>After the initial baseline step, add 4 &#181;L of 12.5 nM substrate [3&#39;Bio-ss-poly (dT)32] to the drop holder and slide it under the biosensor. Close the lid and monitor the BLI curve on the software. Ensure that this response is higher than the baseline (Figure 3). 
		NOTE: Substrate concentration can be optimized by coating biosensors with different concentrations (only one concentration per biosensor) and observing the binding curves. A binding curve with a visibly higher signal compared to the baseline is enough to confirm substrate coating. Overloading the biosensor may lead to steric hindrance or crowding on the surface. Figure 3shows the curves for five concentrations of biotinylated 3&#39;Bio-ss-poly (dT)32 substrate. Additionally, after the bait loading step is completed, the biosensors can be dipped in streptavidin-blocking buffer (biocytin) for 30 s, followed by washing in BLI buffer. This will block the unbound streptavidin surface and prevent non-specific binding.</li>
		<li>After completing the association step, open the lid and replace tube &#39;A&#39; with tube &#39;D.&#39; Slide it under the biosensor and close the lid.</li>
		<li>After the dissociation step is completed, open the lid, detach the biosensor, and place it in the biosensor tray containing the BLI buffer. Ensure that the biosensor is completely dipped in the buffer and is adequately hydrated for the next steps. Clean the drop holder with assay buffer, dry it with lint-free wipes, and remove tube &#39;D&#39; from the tube holder. The biosensor is now coated with the 3&#39;Bio-ss-poly (dT)32 substrate.</li>
	</ol>
	</li>
	<li>Protein binding
	<ol>
		<li>Pipette 250 &#181;L of BLI buffer into ten 0.5 mL black microfuge tubes. Label them A1 to A5 and D1 to D5.</li>
		<li>In another 96-well plate, add 200 &#181;L of stripping buffer into the well, corresponding with the hydrated biosensor&#39;s position.</li>
		<li>Before starting the assay, clean the drop holder 3x with BLI buffer.</li>
		<li>Open a new file on the software and select Basic Kinetics on the left panel. Name the assay and add description as needed (Supplementary Figure 2). Adjust the run settings as required (Table 1).</li>
		<li>Place the tube labeled A1 in the tube holder and attach the 3&#39;Bio-ss-poly (dT)32-coated biosensor to the BLI system. Slide the tube under the biosensor and hit Run.</li>
		<li>After completing the initial baseline step, open the lid, add 4 &#181;L of the BLI buffer to the drop holder, and slide it under the biosensor. This will be the 0 nM protein concentration and serve as the reference curve.</li>
		<li>After completing the association step, replace tube A1 with D1 and slide it under the biosensor.</li>
		<li>Once the dissociation step is completed, remove the biosensor and place it back in its tray. Ensure that the biosensor is completely dipped in the buffer and adequately hydrated for the next steps. Remove tube D1, clean the drop holder with assay buffer, and dry it with a lint-free wipe.</li>
		<li>Next, the binding curve for 5 nM RPA stock will be obtained. Fill out sample details (Concentration: 5 nM, Molecular weight: 116 KDa], and press Calc. This will calculate the protein concentration in &#181;g/&#181;L.</li>
		<li>Attach the 3&#39;Bio-ss-poly (dT)32-coated biosensor and put tube A2 in the tube holder. Slide it under the biosensor and hit Run.</li>
		<li>After baseline, add 4 &#181;L of 5 nM RPA onto the drop holder and slide it under the biosensor. Close the lid and monitor the association curve. Protein binding to bait can be visualized by a curve running higher than the baseline curve. After the association step, replace A2 with D2, slide it under the biosensor, and complete the dissociation step.</li>
		<li>Remove the biosensor, dip it in the stripping buffer for 30 s, and then dip it in the BLI buffer for 3 min. 
		NOTE: The regeneration efficiency depends upon the immobilized bait and the disrupted analyte. Some immobilized biosensors can withstand ten or more regeneration cycles. We could strip biotinylated 3&#39;Bio-ss-poly (dT)32 substrate more than three times (Supplementary Figure 3) without observing a noticeable change in response.</li>
		<li>Repeat steps 2.4.9&#8211;2.4.12 for the other three concentrations of RPA 10 nM, 20 nM, and 40 nM. Make sure to strip the biosensor for each concentration. 
		NOTE: For BLI assays, experts recommend using four (excluding the reference curve) concentrations, spanning roughly from 0.1x to 10x the expected KD for accurate kinetic measurements. Here, the four concentrations were obtained by serial dilutions.</li>
	</ol>
	</li>
	<li>Analyzing data and calculating KD values
	<ol>
		<li>Before calculating the KD value, it is necessary to visually inspect the sensorgram for each concentration of RPA binding on the 3&#39;Bio-ss-poly (dT)32 substrate. Ensure that the BLI signal increases with increasing concentration of RPA until the immobilized DNA is saturated (Figure 4). 
		NOTE: For reliable results, ensure that at least four protein concentrations are bound successfully to the substrate, excluding the 0 nM analyte concentration. Refer to the discussion section to troubleshoot potential errors.</li>
		<li>In the table &#39;Run List,&#39; check Ref for 0 nM analyte concentration to serve as a reference curve. Check Analyze for all the concentrations of the analyte.</li>
		<li>Next, select both Start of association and Start of dissociation for step correction. This will align the beginning of association with the end of the baseline and the beginning of dissociation with the end of association, which is usually caused by different reflection patterns between the drop holder and tube if the buffer matches well.</li>
		<li>Select Global Fitting (1:1) for this assay. Global analysis calculates the KD values for an entire set of protein concentrations, while local analysis is for a single protein concentration. 
		NOTE: The software accompanying the instrument only allows for 1:1 fitting. This could be a potential caveat when the bait and analyte are expected to fit into other fitting models. Data from this software may be exported to other plotting software and fit into required fitting models.</li>
		<li>Next, select Analyze. This will calculate the KD, ka/kd values (Table 2). The ka and kd error within 10% of the value is generally considered acceptable. 
		NOTE: Ensure these values are within reasonable limits. If the error is less than 10%, it suggests that the fitting of data is reliable for studying interactions between biological molecules. Additionally, the calculated KD (equilibrium dissociation constant) using these ka and kd values should be sufficiently accurate for most practical applications18.</li>
		<li>To export the fitted data, right-click on the graph generated using the analyzed data. Choose Export Data, then select Text/Data. Click on File and Browse to save data in &#39;.dat&#39; format. This file can be opened in Microsoft Excel and used in other software for graph plotting. 
		NOTE: Global fitting considers all the binding curve data within the group, while local fitting focuses solely on the kinetic parameters for each individual analyte concentration.</li>
	</ol>
	</li>
</ol>
3. Setting up the instrument to run advanced kinetics
<ol>
	<li>Open the software and select Advanced Kinetics on the left panel (Supplementary Figure 1).</li>
	<li>Place a 96-well plate in the tray holding the biosensors. Add 200 &#181;L of BLI buffer in five wells. Place the tray of biosensors on the 96-well plate so that five biosensors dip into the well containing the buffer.</li>
	<li>On the software, click on Hydrate and turn the timer on for 10 min, allowing the single biosensor to hydrate for at least 10 min.</li>
	<li>While the biosensors are hydrating, pipette 250 &#181;L of BLI buffer into ten 0.5 mL black centrifuge tubes. Label them as A1 to A5 and D1 to D5.</li>
	<li>Fill in the details on the software (Name: Substrate Coating and description of the assay: RPA-3&#39;Bio-ss-poly (dT)32: Advanced Kinetics).</li>
	<li>Input the duration for each step (Supplementary Figure 4). Initial Baseline (30 s), Loading (120 s), Baseline (30 s), Association (120 s), Dissociation (120 s). Adjust the run settings as required (Table 1).
	<ol>
		<li>Fill in the details on the software (Concentration: 0 nM, Molecular Weight: 116 kDa) and press Calc.</li>
	</ol>
	</li>
	<li>Place tube A1 in the tube holder and attach the biosensor. Slide the tube holder under the biosensor and press Run&#160;to obtain the initial baseline.</li>
	<li>Add 4 &#181;L of 12.5 nM substrate into the drop holder and slide it under the biosensor. Close the lid. This will load the substrate onto the biosensor. Replace A1 with D1 in the tube holder and slide it under the biosensor. This will give the baseline curve.</li>
	<li>Clean the drop holder with a lint-free wipe and load 4 &#181;L of the BLI buffer. Slide it under the biosensor and close the lid. This will be the 0 nM protein concentration and will serve as the reference curve.</li>
	<li>After completing the association step, slide the D2 tube under the biosensor. Proceed with the dissociation step. Allow dissociation to complete.</li>
	<li>Detach the biosensor, remove D2 from the tube holder, and clean the drop holder with a lint-free wipe.</li>
	<li>Enter the details (Concentration: 5 nM, Molecular weight: 116 kDa) for the next run and press Calc.</li>
	<li>Attach the second biosensor to the BLItz system, place tube A2 in the tube holder, and slide it under the biosensor.</li>
	<li>After establishing the initial baseline, add 4 &#181;L of 12.5 nM substrate into the drop holder. Slide it under the biosensor and close the lid. Replace tube A2 with D3 and slide it under the biosensor. Next, add 4 &#181;L of 10 nM stock of RPA into the tube holder and slide it under the biosensor.</li>
	<li>After association, slide D4 under biosensor for dissociation.</li>
	<li>Repeat steps 3.1.12&#8211;3.1.15 for the subsequent three concentrations of RPA (10 nM, 20 nM, 40 nM). Binding curves for these concentrations are shown in Figure 5.</li>
	<li>Data analysis and KD value can be fitted in the same procedure as the BLI basic kinetics (step 2.5, Table 2).</li>
</ol>

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

Authors have no conflict of interest to declare.

The ability to analyze the binding kinetics of any protein to its substrate using BLI provides the means to isolate and characterize the specific factors (such as sequence, structure, or length of a DNA) governing protein-DNA interactions within the cell19. The Octet N1 system, which relies on the principles of biolayer interferometry, allows for quantitative measurement of protein-protein and protein-nucleic acids interactions. Further, interactions between lipids, antibodies, and small molecules...

Cellular proteins play a pivotal role in orchestrating the complex biological processes that occur within living organisms. The optimal functioning of these pathways is dependent on the interplay between proteins and other biomolecules inside the cell, including interactions with partner proteins and nucleic acids1. Thus, comprehending the intricacies of cellular processes necessitates a deep understanding of the dynamics of protein-nucleic acid interactions.
Traditionally, protein-nucleic acid interactions have been studied using electrophoretic mobility gel shift assays (EMSAs)2. In this assay, proteins are incubated with synthetic oligonucleotides (either DNA/RNA, containing specific lengths or sequences) for a short period, and the reaction is then electrophoresed on a native polyacrylamide (PAGE) gel3,4,5. To enable visualization of the protein-oligonucleotide interaction, the oligonucleotides are typically radioactively labeled with 32P or tagged with fluorescent molecules. If the protein interacts and binds the oligonucleotide, then the binding of the protein slows down the mobility of the nucleic acid within the gel6,7. Thus, this method is also called the gel shift or gel retardation assay. Though this method has been extensively used, there are a few limitations to consider while using this method to obtain KD values, including low resolution from either weak or dynamic binding, the requirement for a significantly high concentration of proteins, and more effort. Additionally, EMSAs are not real-time assays and, therefore, cannot accurately measure binding kinetics7,8.
Innovative techniques such as surface plasmon resonance (SPR) and biolayer interferometry (BLI) have emerged to overcome these limitations9,10,11,12. Both methods measure association/dissociation rate constants and affinity constants between molecules in a label-free manner. Since the protein is not required to be tagged, these techniques eliminate the risk of altering the properties of the protein or blocking the binding site. In SPR, polarized light interacts with a sensor (a metal conducting film, typically gold), generating an electron charge density wave called plasmon. This interaction diminishes the reflected beam&#39;s intensity and a detector measures the change in the specific angle, known as the resonance angle13. To study ligand (nucleic acid) - analyte (protein) interactions, the ligand is immobilized in one flow cell on the sensor chip, and the analyte is injected into the flow cell containing the immobilized ligand. On binding the analyte to the ligand, there is a detectable change in the refractive index near the sensor surface, thus allowing for the measurement of molecular interactions14,15,16.
Bio-Layer Interferometry (BLI) measures the pattern of light interference as light passes through an optical fiber with a biolayer, composed of a ligand (bait) and its binding partner (analyte), at the bottom surface of biosensor&#39;s tip. Light is transmitted through the tip and reflected at both ends of the biolayer due to the biolayer&#39;s properties. The biolayer thickness is proportional to the number of bound molecules that influence the pattern of the reflected light. By comparing the change of the curve of relative intensity vs. wavelength caused by interference between the reflected light from the reference interface and from the biolayer/buffer interface, the thickness change of the biolayer can be determined. When more molecule binds, a greater shift occurs, making BLI a powerful tool for studying biomolecular interactions in real time. The change in the interference pattern is measured and represented on a sensorgram as a spectral shift17,18 (Figure 1A). The nature of the binding interaction can be accurately determined by using appropriate controls, such as a setup lacking the ligand-varying concentrations of the binding partner19,20,21. Compared to SPR, BLI is cost-effective and user-friendly, rendering it accessible to a broad range of researchers. Additionally, samples used in BLI remain intact if there is no degradation or aggregation. This allows them to be possibly recovered and reused, thereby minimizing waste. The BLI instrument operates without the use of microfluidics, thereby eliminating the disadvantages of the fluidics system, such as the need for maintenance/care, clogging, or use of degassed buffers. It also minimizes the risk of contaminating the instrument due to unfiltered or crude protein samples.
In a BLI experiment, the biolayer is established by immobilizing the bait molecule on the biosensor. The biolayer of biosensors can interact with various tags, making it possible to study the interactions between molecules (including nucleic acids, proteins, antibodies, viruses, small molecules, etc.)22. Various capturing strategies, including biotin/streptavidin, 6X-His-tag/Ni-NTA, FLAG/anti-FLAG, GST/anti-GST, and antibody/anti-Fc, can be employed to establish this immobilization. Maintaining the structure and activity of the immobilized ligand is crucial when choosing the biosensor. The changes in the interference pattern are influenced by the amount of bound molecule as well as the matrix23. Protein-nucleic acid interactions are studied using biotinylated oligonucleotide probes that can be immobilized on streptavidin-coated biosensors. Protein samples expected to interact with the immobilized bait (oligonucleotide) are in contact with the oligonucleotide for a specific period in a binding buffer to measure association and subsequently switched to a blank binding buffer to measure dissociation. A schematic representation of analyte binding and corresponding changes in the binding curve is shown in Figure 1B. Additionally, protein-nucleic interactions can also be studied by a reversed immobilization route, in which the protein (bait) is captured on the biosensor and interacts with the nucleic acid (analyte).
Currently, multiple instruments are available commercially that operate on the principle of biolayer interferometry. A straightforward and cost-effective instrument is known as the Octet N1 system, which features a single channel for data throughput. It requires manual operation, consumes minimal sample volume (4 &#181;L), and performs sample analysis at ambient temperatures9,23. This single-channel instrument can detect binding efficiencies of proteins larger than 10 kDa and measures affinities in the micromolar (&#181;M) to nanomolar (nM) range11,12. Some instruments with 2, 8, 16, and 96 channels for automated data reading are also available and compatible with both 96 or 384 well formats19,20. Some of these instruments can also operate between 4 &#176;C to 40 &#176;C, detect biomolecules with molecular weight as low as 150 Da, and measure affinities within the millimolar to picomolar range19,21. While the described system is cost-effective, the pricier instruments with multiple channels offer high-throughput automatic processing. These advanced systems are commonly employed in characterizing and developing biological drug molecules9,23.
The current protocol describes the steps involved in measuring the binding parameters of replication protein A (RPA) to single-stranded DNA (ss-DNA) using the single-channel, manual biolayer interferometry system. RPA is a heterotrimeric, ssDNA-binding protein complex that plays a critical role in almost all aspects of DNA metabolism, including DNA replication, repair, and recombination24. Due to its high affinity to ssDNA (measured to be in the sub-nanomolar range), it can rapidly bind to ssDNA generated during various DNA transactions, protect it from nucleolytic degradation, and prevent impromptu binding of other downstream proteins. RPA also plays a critical role in preventing the formation of non-canonical secondary and tertiary structures, such as G-quadruplexes25. Human RPA is made up of three subunits: RPA70 (70 kDa), RPA32 (32 kDa), and RPA14 (14 kDa), also called RPA 1, RPA 2, and RPA 3, respectively24,26,27. These subunits house six oligonucleotide/oligosaccharide binding folds (OB-folds) labeled A to F. Among these, the DNA binding domains (DBDs) are on the RPA1 (DBD-A, DBD-B, DBD-C) and RPA2 (DBD-D) subunits28. It was first thought that depending on the DNA&#39;s length, RPA exhibits different binding modes where different DBDs were engaged to specific DNA lengths29. Data from recent structural and single molecule studies have helped refine these models to suggest that the binding of different DBDs of RPA is more dynamic than initially suggested30,31,32,33,34,35,36. This feature of the dynamic binding property is essential to the function of RPA because RPA should be able to bind tightly to the DNA substrate during certain DNA transactions; however, it should also be able to displace from the substrate to hand over the substrate to the next protein during the biological process. Using different biochemical techniques, the KD for RPA has been determined to be about 0.4 nM for a ss-(polydT)30 substrate, and 80 nM and 200 nM for ss-(polydA)257 and dG-(polydG)602 substrates respectively, as measured by fluorescence polarization anisotropy (FPA)37,38. RPA also shows about a 50-fold higher preference for binding to pyrimidine rich sequences compared to purines. Though different biochemical techniques have determined differing KD measurements for RPA, all measurements have been in the nanomolar range, suggesting RPA&#39;s high binding affinity for ssDNA. Employing total internal reflection fluorescence microscopy (TIRFM), it was discovered that based on the length of the DNA, RPA associated with at least two binding modes: one that was defined at fast dissociating (Kd, 680 pM) and a slow dissociating (Kd, 60 pM) complex33,39. Given its pivotal involvement in nearly all DNA metabolic pathways, there has been a keen interest in creating inhibitors that can hinder the interaction between RPA and single-stranded DNA (ssDNA). These chemical inhibitors are vital in disrupting the DNA damage response, rendering cancer cells more susceptible to DNA-damaging agents employed in clinical therapies. Utilizing the BLI assay allows for a precise quantitative evaluation of inhibitor efficacy in modulating RPA&#39;s binding function40,41.
BLI setup allows for distinct settings: basic and advanced kinetics. In basic kinetics mode, the assay comprises three primary stages: baseline establishment, association, and dissociation. However, before performing this assay, the biosensors need to be coated separately, using either the instrument or manually on the laboratory bench. Conversely, the advanced kinetics mode extends beyond the fundamental three steps, allowing for the incorporation of additional steps. These supplementary steps can serve various purposes, such as conditioning the biosensor to the buffer alterations or facilitating the detection of subsequent protein interactions. Advanced kinetics also allows stripping the biosensor of the analyte by a good regeneration method for potential reuse, provided the bait and the biosensor remain intact. Generally, advanced kinetics is used over basic kinetics when multiple steps are involved in the assay, or the assay needs to be performed in a more complex format.
This protocol outlines both methods using the same bait [3&#39;Bio-ss-poly (dT)32] and analyte (RPA). A streptavidin-coated biosensor will be used to immobilize the bait, and KD measurements for the analyte will be obtained. This protocol describes a complete approach to perform a BLI assay using a single-channel instrument biolayer interferometry, covering biosensor-bait preparation, buffer considerations, and a step-by-step outline of the procedure.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5 mL Micro Centrifuge Tubes</td>
			<td>Globe Scientific</td>
			<td>111554A</td>
			<td></td>
		</tr>
		<tr>
			<td>96 Well Standard Black Microplate</td>
			<td>Dot Scientific</td>
			<td>4ti-0223</td>
			<td></td>
		</tr>
		<tr>
			<td>Biotinylated poly dT Oligonucleotide</td>
			<td>IDT</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Sigma-Aldrich</td>
			<td>A2153-10G</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithiothreitol (DTT)</td>
			<td>Dot Scientific</td>
			<td>DSD11000-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid (EDTA)</td>
			<td>Dot Scientific</td>
			<td>DSE57020-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric Acid&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>A144-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimtec Science Kimwipes&#160;</td>
			<td>Kimtech</td>
			<td>34120</td>
			<td></td>
		</tr>
		<tr>
			<td>Octet N1 Software</td>
			<td>Sartorius&#160;</td>
			<td>1.4.0.13</td>
			<td></td>
		</tr>
		<tr>
			<td>Octet SA Biosensor&#160;</td>
			<td>Sartorius&#160;</td>
			<td>18-5019</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS pH 7.2 (10x)</td>
			<td>Gibco</td>
			<td>1666711</td>
			<td></td>
		</tr>
		<tr>
			<td>Personal Assay Octet N1 System</td>
			<td>Sartorius&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphoric Acid&#160;</td>
			<td>Ward&#39;s Science</td>
			<td>470302-024</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride (NaCl)</td>
			<td>Dot Scientific</td>
			<td>DSS23020-5000</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris Base</td>
			<td>Dot Scientific</td>
			<td>DST60040-5000</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween20&#160;</td>
			<td>Bio-Rad</td>
			<td>170-6531</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

In BLI, white light is reflected from the interface of biolayer/buffer and the internal reference interface to the spectrometer. The resulting interference pattern is recorded, and the spectral shift is measured over a period of time and depicted as binding curves response in nm. A representative figure showing the biosensor tip with and without analyte binding and the corresponding spectral wavelength shift (nm) is shown in Figure 1. The binding curves correspond to the baselines, bait load...

Analyzing DNA-Protein Interactions with Streptavidin-Based Biolayer Interferometry

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 ...

analyzing-dna-protein-interactions-with-streptavidin-based-biolayer

Research

JoVE Journal

Biochemistry

1.1K Views.  Indiana University Indianapolis. Biomolecuulinteracties, waaronder eiwit-eiwit en eiwit-DNA interacties, zijn essentieel in biochemisch onderzoek. In deze studie gebruiken we biolaaginterferometrie, of BLI, een efficiënte techniek om deze interacties te analyseren. We richten ons op replicatie-eiwit A en de bindingsaffiniteit ervan voor een enkelstrengs DNA om de principes en mogelijkheden van BLI te demonstreren.Biolaaginterferometrie is een gemakkelijke en kosteneffectieve methode ...

Video: Analyse van DNA-eiwitinteracties met op streptavidine gebaseerde biolaaginterferometrie

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

Analyzing Supercomplexes of the Mitochondrial Electron Transport Chain with Native Electrophoresis, In-gel Assays, and Electroelution

The mitochondrial electron transport chain (ETC) transduces the energy derived from the breakdown of various fuels into the bioenergetic currency of the cell, ATP. The ETC is composed of 5 massive protein complexes, which also assemble into supercomplexes called respirasomes (C-I, C-III, and C-IV) and synthasomes (C-V) that increase the efficiency of electron transport and ATP production. Various methods have been used for over 50 years to measure ETC function, but these protocols do not provide information on the assembly of individual complexes and supercomplexes. This protocol describes the technique of native gel polyacrylamide gel electrophoresis (PAGE), a method that was modified more than 20 years ago to study ETC complex structure. Native electrophoresis permits the separation of ETC complexes into their active forms, and these complexes can then be studied using immunoblotting, in-gel assays (IGA), and purification by electroelution. By combining the results of native gel PAGE with those of other mitochondrial assays, it is possible to obtain a completer picture of ETC activity, its dynamic assembly and disassembly, and how this regulates mitochondrial structure and function. This work will also discuss limitations of these techniques. In summary, the technique of native PAGE, followed by immunoblotting, IGA, and electroelution, presented below, is a powerful way to investigate the functionality and composition of mitochondrial ETC supercomplexes.

This protocol describes the separation of functional mitochondrial electron transport chain complexes (Cx) I-V and supercomplexes thereof using native electrophoresis to reveal information about their assembly and structure. The native gel can be subjected to immunoblotting, in-gel assays, and purification by electroelution to further characterize individual complexes.

The mitochondrial electron transport chain (ETC) transduces the energy derived from the breakdown of various fuels into the bioenergetic currency of the ...

Measuring Biomolecular DSC Profiles with Thermolabile Ligands to Rapidly Characterize Folding and Binding Interactions

Differential scanning calorimetry (DSC) is a powerful technique for quantifying thermodynamic parameters governing biomolecular folding and binding interactions. This information is critical in the design of new pharmaceutical compounds. However, many pharmaceutically relevant ligands are chemically unstable at the high temperatures used in DSC analyses. Thus, measuring binding interactions is challenging because the concentrations of ligands and thermally-converted products are constantly changing within the calorimeter cell. Here, we present a protocol using thermolabile ligands and DSC for rapidly obtaining thermodynamic and kinetic information on the folding, binding, and ligand conversion processes. We have applied our method to the DNA aptamer MN4 that binds to the thermolabile ligand cocaine. Using a new global fitting analysis that accounts for thermolabile ligand conversion, the complete set of folding and binding parameters are obtained from a pair of DSC experiments. In addition, we show that the rate constant for thermolabile ligand conversion may be obtained with only one supplementary DSC dataset. The guidelines for identifying and analyzing data from several more complicated scenarios are presented, including irreversible aggregation of the biomolecule, slow folding, slow binding, and rapid depletion of the thermolabile ligand.

We present a protocol for rapid characterization of biomolecular folding and binding interactions with thermolabile ligands using differential scanning calorimetry.

Differential scanning calorimetry (DSC) is a powerful technique for quantifying thermodynamic parameters governing biomolecular folding and binding ...

Capturing the Interaction Kinetics of an Ion Channel Protein with Small Molecules by the Bio-layer Interferometry Assay

The bio-layer interferometry (BLI) assay is a valuable tool for measuring protein-protein and protein-small molecule interactions. Here, we first describe the application of this novel label-free technique to study the interaction of human EAG1 (hEAG1) channel proteins with the small molecule PIP2. hEAG1 channel has been recognized as potential therapeutic target because of its aberrant overexpression in cancers and a few gain-of-function mutations involved in some types of neurological diseases. We purified&#160;hEAG1 channel proteins from a mammalian stable expression system and measured the interaction with PIP2 by BLI. The successful measurement of the kinetics of binding between hEAG1 protein and PIP2 demonstrates that the BLI assay is a potential high-throughput approach used for novel small-molecule ligand screening in ion channel pharmacology.

The protocol here describes the interactions of purified hEAG1 ion channel protein with the small molecule lipid ligand phosphatidylinositol 4, 5-bisphosphate (PIP2). The measurement demonstrates that BLI could be a potential method for novel small-molecule ion channel ligand screening.

The bio-layer interferometry (BLI) assay is a valuable tool for measuring protein-protein and protein-small molecule interactions. Here, we first describe ...

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this work, the pre-endosomal anthrax toxin is assembled and transitioned into the endosomal complex. First, the N-terminal domain of a cysteine mutant lethal factor (LFN) is attached to a biolayer interferometry (BLI) biosensor through disulfide coupling in an optimal orientation, allowing protective antigen (PA) prepore to bind (Kd 1 nM). The optimally oriented LFN-PAprepore complex then binds to soluble capillary morphogenic gene-2 (CMG2) cell surface receptor (Kd 170 pM), resulting in a representative anthrax pre-endosomal complex, stable at pH 7.5. This assembled complex is then subjected to acidification (pH 5.0) representative of the late endosome environment to transition the PAprepore into the membrane inserted pore state. This PApore state results in a weakened binding between the CMG2 receptor and the LFN-PApore and a substantial dissociation of CMG2 from the transition pore. The thio-attachment of LFN to the biosensor surface is easily reversed by dithiothreitol. Reduction on the BLI biosensor surface releases the LFN-PAprepore-CMG2 ternary complex or the acid transitioned LFN-PApore complexes into microliter volumes. Released complexes are then visualized and identified using electron microscopy and mass spectrometry. These experiments demonstrate how to monitor the kinetic assembly/disassembly of specific protein complexes using label-free BLI methodologies and evaluate the structure and identity of these BLI assembled complexes by electron microscopy and mass spectrometry, respectively, using easy-to-replicate sequential procedures.

Here we present a protocol to monitor the assembly and disassembly of the anthrax toxin using biolayer interferometry (BLI). Following assembly/disassembly on the biosensor surface, the large protein complexes are released from the surface for visualization and identification of components of the complexes using electron microscopy and mass spectrometry, respectively.

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this ...

Site-Directed Mutagenesis for In Vitro and In Vivo Experiments Exemplified with RNA Interactions in Escherichia Coli

Site-directed mutagenesis is a technique used to introduce specific mutations in DNA to investigate the interaction between small non-coding ribonucleic acid (sRNA) molecules and target messenger RNAs (mRNAs). In addition, site-directed mutagenesis is used to map specific protein binding sites to RNA. A 2-step and 3-step PCR based introduction of mutations is described. The approach is relevant to all protein-RNA and RNA-RNA interaction studies. In short, the technique relies on designing primers with the desired mutation(s), and through 2 or 3 steps of PCR synthesizing a PCR product with the mutation. The PCR product is then used for cloning. Here, we describe how to perform site-directed mutagenesis with both the 2- and 3-step approach to introduce mutations to the sRNA, McaS, and the mRNA, csgD, to investigate RNA-RNA and RNA-protein interactions. We apply this technique to investigate RNA interactions; however, the technique is applicable to all mutagenesis studies (e.g., DNA-protein interactions, amino-acid substitution/deletion/addition). It is possible to introduce any kind of mutation except for non-natural bases but the technique is only applicable if a PCR product can be used for downstream application (e.g., cloning and template for further PCR).

Site-Directed Mutagenesis for In Vitro and In Vivo Experiments Exemplified with RNA Interactions in Escherichia Coli

Site-directed mutagenesis is a technique used to introduce specific mutations in deoxyribonucleic acid (DNA). This protocol describes how to do site-directed mutagenesis with a 2-step and 3-step polymerase chain reaction (PCR) based approach, which is applicable to any DNA fragment of interest.

Site-directed mutagenesis is a technique used to introduce specific mutations in DNA to investigate the interaction between small non-coding ribonucleic ...

Application of Biolayer Interferometry (BLI) for Studying Protein-Protein Interactions in Transcription

A transcription factor&#160;(TF) is a protein that regulates gene expression by interacting with the RNA polymerase, another TF, and/or template DNA. GrgA is a novel transcription activator found specifically in the obligate intracellular bacterial pathogen Chlamydia. Protein pulldown assays using affinity beads have revealed that GrgA binds two &#963; factors, namely &#963;66 and &#963;28, which recognize different sets of promoters for genes whose products are differentially required at developmental stages. We have used BLI to confirm and further characterize the interactions. BLI demonstrates several advantages over pulldown: 1) It reveals real-time association and dissociation between binding partners, 2) It generates quantitative kinetic parameters, and 3) It can detect bindings that pulldown assays often fail to detect. These characteristics have enabled us to deduce the physiological roles of GrgA in gene expression regulation in Chlamydia, and possible detailed interaction mechanisms. We envision that this relatively affordable technology can be extremely useful for studying transcription and other biological processes.

Application of Biolayer Interferometry BLI for Studying Protein-Protein Interactions in Transcription

Interactions of transcription factors (TFs) with the RNA polymerase are usually studied using pulldown assays. We apply a Biolayer Interferometry (BLI) technology to characterize the interaction of GrgA with the chlamydial RNA polymerase. Compared to pulldown assays, BLI detects real-time association and dissociation, offers higher sensitivity, and is highly quantitative.

A transcription factor&#160;(TF) is a protein that regulates gene expression by interacting with the RNA polymerase, another TF, and/or template DNA. GrgA ...

Interactions with and Membrane Permeabilization of Brain Mitochondria by Amyloid Fibrils

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary mechanism of amyloid aggregate-induced toxicity in neurodegenerative diseases. However, most reports describing the mechanisms of membrane disruption are based on phospholipid model systems, and studies directly targeting events occurring at the level of biological membranes are rare. Described here is a model for studying the mechanisms of amyloid toxicity at the membrane level. For mitochondrial isolation, density gradient medium is used to obtain preparations with minimal myelin contamination. After mitochondrial membrane integrity confirmation, the interaction of amyloid fibrils arising from &#945;-synuclein, bovine insulin, and hen egg white lysozyme (HEWL) with rat brain mitochondria, as an in vitro biological model, is investigated. The results demonstrate that treatment of brain mitochondria with fibrillar assemblies can cause different degrees of membrane permeabilization and ROS content enhancement. This indicates structure-dependent interactions between amyloid fibrils and mitochondrial membrane. It is suggested that biophysical properties of amyloid fibrils and their specific binding to mitochondrial membranes may provide explanations for some of these observations.

Provided here is a protocol for investigating the interactions between native form, prefibrillar, and mature amyloid fibrils of different peptides and proteins with mitochondria isolated from different tissues and various areas of the brain.

A growing body of evidence indicates that membrane permeabilization, including internal membranes such as mitochondria, is a common feature and primary ...

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.

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 ...

Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An important first step in any membrane-dependent reaction is binding of protein on the phospholipid membrane. An approach to studying protein interaction with lipid membranes has been developed using fluorescently labeled proteins and flow cytometry. This method allows the study of protein-membrane interactions using live cells and natural or artificial phospholipid vesicles. The advantage of this method is the simplicity and availability of reagents and equipment. In this method, proteins are labeled using fluorescent dyes. However, both self-made and commercially available, fluorescently labeled proteins can be used. After conjugation with a fluorescent dye, the proteins are incubated with a source of the phospholipid membrane (microvesicles or cells), and the samples are analyzed by flow cytometry. The obtained data can be used to calculate the kinetic constants and equilibrium Kd. In addition, it is possible to estimate the approximate number of protein binding sites on the phospholipid membrane using special calibration beads.

Here, we describe a set of methods for characterizing the interaction of proteins with membranes of cells or microvesicles.

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An ...

Analyzing DNA-Protein Interactions with Streptavidin-Based Biolayer Interferometry

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