Name: nano-differential scanning fluorimetry for screening in fragment-based lead discovery
Uploaded: 2021-05-16T15:00:00.000+00:00
Duration: 6 min 26 s
Description: Watch this Scientific Journal Video about Nano-Differential Scanning Fluorimetry for Screening in Fragment-based Lead Discovery at JoVE.com

&#34;This work benefited from access to the NKI Protein Facility, an Instruct-ERIC centre. Financial support has been provided by iNEXT, project number 653706, and iNEXT-Discovery, project number 871037, funded by the Horizon 2020 program of the European Commission&#34;.

NOTE: The proteins used in nano-DSF&#160;experiments should be pure (&#62;95%) and homogeneous as judged by sodium dodecylsulfate-polyacrylamide gel electrophoresis. Before performing the fragment screen, the stability of the proteins should be determined in various buffer conditions. A low ionic strength, low salt buffer that interferes minimally with the protein should be used so as not to affect its direct interaction with the fragments. The buffers typically used in this protocol for checking stability are shown in Supplemental Table S1. The concentration of the protein that needs to be used for this experiment as a stock solution is typically 0.2 mg mL-1. For screening the entire DSi-Poised library (768 compounds), a total of ~12 mL of protein of that concentration, a total of ~2.5 mg, is needed. The DSi-Poised library used in these experiments was supplied in 96-well format (Figure 1). The concentration of the fragments was adjusted to 100 mM in 20% v/v dimethylsulfoxide (DMSO). It should be noted that the mixing protocol described here results in low final DMSO concentrations of 0.4% v/v; although this is very unlikely to affect the stability of the protein, the effect of DMSO should be checked for each new protein.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62469/62469fig1v11.jpg" /> 
Figure 1: The types of plates used for these experiments. (A) U-bottom plate. (B) 96-well plate. (C) A close-up view of 96-well plate showing the subwell 1. <a href="https://www.jove.com/files/ftp_upload/62469/62469fig1v11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Plate preparation
<ol>
	<li>Take out a fragment plate from the -20 &#176;C/-80 &#176;C freezer, and let it thaw at room temperature, with gentle shaking on a benchtop shaker. Centrifuge the plate at 500 &#215; g for 30 s to collect any drops sticking to the side of the wells. 
	NOTE: As the fragment library is available dissolved in DMSO-d6 (melting point 19 &#176;C), it is essential to make sure that each compound is completely thawed and solubilized.</li>
	<li>Take an MRC 2-well crystallization plate (Figure 1A), and pipette 14.7&#160;&#181;L of the protein stock solution into each sub-well. To do this in a time-efficient manner, keep the protein in a reagent reservoir (Table of Materials), and use a multichannel pipette for dispensing (Figure 2A,B). 
	&#8203;NOTE: For the DSi-Poised library, depending on the format, not all the wells of the 96-well plate contain compounds. Typically, rows A, H and columns 1, 12 are filled with DMSO. Thus, rows A, H and columns 1, 12 are not to be filled with protein, as these wells will not contain any fragments at the end of the next step; only DMSO will be transferred there by the robot. Please note that the empty rows and columns do differ in some plates.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62469/62469fig2v11.jpg" /> 
Figure 2:&#160;Outline of the fragment screening procedure. (A) Using a multichannel pipette and reagent reservoir to dispense the protein. (B) Dispensing the protein in the 96-well plate. (C) Fragment dispensing by the dispenser robot. (D) Loading of the protein into the capillaries. (E) Drawer showing the capillary holder. (F) Close-up view of the capillary holder. <a href="https://www.jove.com/files/ftp_upload/62469/62469fig2v11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62469/62469fig3v11.jpg" /> 
Figure 3: Overview of the equipment used in these experiments. (A) Nanodispenser robot used for fragment dispensing. Plate positions are indicated. (B) Program interface for defining a new plate. (C) Interface of the dispensing program. (D) The dispensing program used to dispense the fragments. <a href="https://www.jove.com/files/ftp_upload/62469/62469fig3v11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Fragment nano-dispensing by the Mosquito robot
<ol>
	<li>Check the type of plate in which the fragments are supplied (Table of Materials).</li>
	<li>Check fragment and protein plate definitions on the Mosquito.
	<ol>
		<li>Switch on the nanodispenser (Figure 3A). Make sure that there are no obstacles around the moving components.</li>
		<li>Open the graphical user interface, click on the Setup tab, and under Deck Configuration, check whether the type of plate in which the compounds are supplied and the one in which the protein has been transferred in section 1 are already present in the list of the Available plates. If not, click Options|Plates and create a new plate definition by filling in the correct values for the Property type (Figure 3B). 
		NOTE: These values for the MRC 2-well plate and the 96-well plate used in this experiment are shown in Supplemental Table S2 and Supplemental Table S3, respectively.</li>
	</ol>
	</li>
	<li>The dispensing program
	<ol>
		<li>Under the Setup tab, specify the plate positions under Deck Configuration. 
		NOTE: The Mosquito used in this experiment has two plate positions on the deck. Position 1 is defined as the Source plate, and Position 2 is the Destination plate.</li>
		<li>Using the dropdown menu, choose Greiner U bottom plate for Position 1 and MRC 2-well plate for Position 2 (Figure 2C). Save the protocol.</li>
	</ol>
	</li>
	<li>Fragment dispensing
	<ol>
		<li>Place the fragment plate at Position 1 and the protein plate at Position 2 of the deck (Figure 3A).</li>
		<li>On the Protocol tab (Figure 3D), click on File and choose the protocol saved in section 2.3 for dispensing the fragments. Define the volume of the fragments to be dispensed; use 0.3 &#181;L. For a typical plate, where columns 1 and 12 do not contain fragments, define the Start location to column 2 and the End location to column 11. In some plates, if columns 2 or 11 are empty, use columns 3 and 10 as start and end values, respectively. 
		NOTE: Because of the Mosquito setup, only columns can be skipped, not rows; for the rows, the mosquito will pipette DMSO. Make sure that the Tip Changing option is selected as Always, so as not to cross-contaminate the fragment library.</li>
		<li>Click on Run to start the program. After dispensing, which takes ~2 min, is completed, remove the protein and the fragment plates from the robot, and seal them back with an adhesive sealing film. Briefly centrifuge the protein plate (500 &#215; g, 30 s) to collect any drops sticking to the sides of the wells before proceeding to the next step.</li>
	</ol>
	</li>
</ol>
3. Measurement of nano-DSF
NOTE: A detailed description about performing TSAs using the Prometheus NT.48 has been published previously20. Important points in the context of fragment screening are mentioned here.
<ol>
	<li>Plate inspection before the measurement
	<ol>
		<li>Visually inspect the wells of the protein plate for precipitation that might have occurred due to the addition of the fragments. If precipitation is observed in many wells, reduce the concentration of the fragments, and repeat the experiment. 
		NOTE: It is recommended to keep the protein plate at room temperature; the fragments tend to become insoluble at lower temperatures.</li>
	</ol>
	</li>
	<li>Preparing the Prometheus
	<ol>
		<li>Switch on the Prometheus instrument. On the touchscreen, press Open Drawer to access the capillary loading module of the instrument. Remove the magnetic strip from the loading module, and clean the mirror with ethanol to remove any dust particles.</li>
	</ol>
	</li>
	<li>Transfer from the protein-fragments plate to capillaries 
	NOTE: This step involves transferring the mixed protein/fragment sample from each well in the protein plate to the capillaries for use with the Prometheus. For this, although the Standard capillary type is typically used, it is possible to use the High Sensitivity capillaries for very low protein concentrations.
	<ol>
		<li>Place the protein plate and the capillaries next to the instrument to have easy access to the capillary loading module.</li>
		<li>Take one capillary, hold it at one end, and touch the solution in the protein plate with the far end of the capillary to transfer the sample by capillary action. Always wear gloves, and make sure not to touch the capillary in the middle, as impurities (e.g., dust particles) from the gloves would affect the measurement.</li>
		<li>Place the capillary in the designated position of the holder, making sure it is properly aligned and centered.</li>
		<li>Repeat steps 3.3.2 and 3.3.3, thereby filling up all positions in the loading module. Load all capillaries you will need for measurement. 
		NOTE: For a single run, a maximum of 48 capillaries can be loaded.</li>
		<li>At the end, place the magnetic strip on top of the capillaries to hold them in place, and press Close Drawer to start the experiment.</li>
	</ol>
	</li>
	<li>Perform the nano-DSF experiment
	<ol>
		<li>Fluorescence scan
		<ol>
			<li>Open the Prometheus application PR.ThermControl, and create a new project by clicking on Start New Session using the name ProteinName_ScreenName_PlateNumber. First, do a Discovery Scan to detect the fluorescence of the samples. 
			NOTE: The fluorescence levels should ideally be above 3,000 counts for each sample. Samples having fluorescence above the saturation limit of the instrument (20,000 counts) will not be measured.</li>
			<li>Alter the fluorescence signal of the samples by adjusting the Excitation Power of the laser (Figure 4).</li>
		</ol>
		</li>
		<li>Thermal denaturation
		<ol>
			<li>Click on the Melting Scan tab. To perform the experiment, set the Start Temperature to 20 &#176;C, the End Temperature to 95 &#176;C, and the Temperature Slope to 1 &#176;C min-1. Click on Start Measurement.</li>
		</ol>
		</li>
		<li>Annotating the experiment
		<ol>
			<li>After the experiments starts, click on the Annotation and Results tab. Annotate each capillary by giving it a unique plate- and well-number. 
			NOTE: For example, capillary 1B2 would correspond to Plate 1 and well B2. In this tab, extra columns can be added for the experiment, e.g., protein name, buffer, protein concentration. After the experiment is finished, the results are also displayed in this tab. This is a good time to pause at the end of the day; the experiments take place overnight, the results can be collected the next day.</li>
		</ol>
		</li>
		<li>Data visualization and export
		<ol>
			<li>After the experiment is finished, click on the Melting Scan tab to view the melting and scattering curves for the samples. To export the results in a spreadsheet, click on the Melting Scan tab, click on Export, and choose Export Processed Data from the drop-down menu.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62469/62469fig4v11.jpg" /> 
Figure 4: Adjusting the excitation power and fluorescence signal. (A) At 80% excitation power, the fluorescence signal for most of the samples is beyond the saturation limit. (B) The fluorescence signal is reduced to measurable levels by decreasing the excitation power to 60%. <a href="https://www.jove.com/files/ftp_upload/62469/62469fig4v11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Iteration
<ol>
	<li>Repeat steps 1-3 to perform 16 runs on the entire Enamine library of 768 compounds (16 &#215; 48 = 768).</li>
	<li>As the measurement of each plate takes ~1.5 h to complete, complete the annotation (3.4.3), and prepare the next protein-fragments plate by repeating steps in sections 1 and 2. 
	&#8203;NOTE: In a typical working day, 4-6 plates can be measured, depending on experience. The screening of the entire DSi-library can be finished in a total of 3-4 days.</li>
</ol>
5. Data analysis
<ol>
	<li>Examining the generated data
	<ol>
		<li>Once each run is completed, click on the Annotations and Results tab to display the results. For each sample, focus on two calculated values that are most important in this overview: 1) The scattering onset temperature (Onset #1 for Scattering), which indicates the temperature at the start of increased sample scattering events and is characteristic of aggregation; 2) the Tm (Inflection Point #1 for Ratio) value extracted from the ratio between fluorescence events at 330 and 350 nm-values that typically correspond to the maximal changes of tryptophan fluorescence upon changes in its environment.</li>
		<li>Be sure to check the scattering and the melting curves for each capillary in the Melting Scan tab to make sure that these values are reliable.</li>
	</ol>
	</li>
	<li>Export and inspection to a spreadsheet
	<ol>
		<li>Export the data from all different runs for inspection to a spreadsheet software, as described in 3.4.4, and to create overview tables and plots. Click on the various Sheets in the spreadsheet file to note the information in each sheet.
		<ol>
			<li>In the Overview sheet, note that each row corresponds to one experiment. Along an overview of parameters (e.g., start and end temperature), observe the calculated values for the Tm and the scattering onset (see 5.1 above for names and explanation, and Supplemental files 1 and 2 for an example). As the software calculates two or more Tm values (Inflection Point #1 and #2 for Ratio) for some samples, look at the melting transition curve to determine which of the Tm values is correct.</li>
			<li>In the Ratio sheet, note that each column corresponds to a sample, and each row to the data read at each temperature step. Observe the fluorescence count values corresponding to each temperature step, and use them to plot the melting curves for specific samples, plotting the temperature column against the fluorescence count column. Note that the data in the Ratio (first derivative), 330 nm, 330 nm (first derivative), 350 nm, 350 nm (first derivative) are used to calculate the ratio and its first derivative (which has a maximum at the maximum rate of change) in a similar format.</li>
			<li>Observe similar data for scattering in the Scattering sheet. Generate the scattering curve for each sample by plotting the temperature column against the scattering column. Look for the Sheet for the scattering first derivative.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Creating and validating a global overview for all fragments
	<ol>
		<li>After verifying the correct Tm values for the fragments, combine the columns Sample ID and Inflection point of 330/350 nm ratio (Tm) from all the runs in a single new result file, copying these columns from each run.</li>
		<li>Use the average Tm value of the native protein (typically calculated over ten runs) and subtract it from the Tm values of each sample, to get the &#916;Tm. Sort the results over &#916;Tm in descending order to identify the samples that result in the largest shift.</li>
		<li>Divide the fragments into bins depending on the &#916;Tm shift, and generate a frequency table for the whole library by plotting the &#916;Tm for each bin against the number of fragments (Figure 5). For convenience, show the axis representing the number of fragments in the log scale. Bin the sample with &#916;Tm &#177;1, and adapt the other bins to each run to adjust the number of outliers empirically.</li>
	</ol>
	</li>
</ol>

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There are no disclosures.

This protocol describes a medium-to-high throughput method for screening fragment libraries using some common robotics and measurement instruments. Screens like those described in this protocol can be routinely performed by the NKI Protein Facility in Amsterdam, for instance as an iNEXT-Discovery service, often even for free for users after proposal application and peer review. In such cases, the DSi-Poised library can be provided by the facility, but the use of other libraries can also be discussed in the context of each different user application and service agreement. The choice of instruments in this protocol represents practical solutions for many laboratories, but should not be considered as a gold standard. Label-free methods are recommended for measuring the thermal stability of the target protein for fragment screening, rather than methods that use environmentally sensitive labels for detecting unfolding in a reverse-transcription polymerase chain reaction thermocycler.
Label-free methods, such as the one presented here using the Prometheus instrument, have some advantages: they use low amounts of protein, often a couple of orders of magnitude less; they can be used to simultaneously measure scattering of the sample and thus aggregation; and the labels used for detecting unfolding in other approaches can interact differently with each fragment, resulting in measurement artifacts. This protocol has been described in the context of the Mosquito robot, which allows the pipetting of a very small volume of sample (0.3 &#181;L) that cannot be done manually. The Mosquito is a popular robot, present in many laboratories working on structural biology and drug discovery projects; however, the protocol can clearly use alternative approaches for low-volume pipetting.
Fragment libraries contain compounds dissolved in DMSO. One of the initial challenges is to find the optimal DMSO concentration at which the protein remains stable, and the compounds remain soluble. This involves performing the measurements at various DMSO concentrations to determine the optimal conditions for screening. The protein to fragment dilution used here results in DMSO concentrations of 0.2%; most proteins are fairly stable in these conditions. The amount of protein required for carrying out the screening for the 768-compound library is ~2-3 mg in total, as the measurements are typically carried out at low protein concentrations (0.2 mg mL-1). Working with such relatively low protein concentrations not only reduces protein production costs, but also reduces the chances of protein precipitation. The low protein concentration does not affect detection of fragment binding, as the concentration of the fragments in the experiment is ~2 mM, allowing also weak binders to be identified.
As the melting transition detection in these experiments is based on fluorescence intensity, a critical aspect is to determine the excitation power of the laser at which to carry out the measurements. The interaction of the compounds with the protein can (i) have no effect on its intrinsic fluorescence, (ii) result in quenching, or (iii) increase its intrinsic fluorescence. In addition to this, working with low protein concentrations means that the fluorescence count for the native protein would be low. The excitation power therefore must be adjusted in such a manner that most of the samples can be measured. The scattering profile of every run provides important information about the aggregation effects that might be triggered by the addition of any fragment. In addition, the effect of temperature on compound solubility can also be seen on the scattering profile.
Unexpectedly, for many compounds&#160;it was observed that the scattering actually decreased with increasing temperature (Figure 6). It is therefore important to look at both the melting transition curve and the accompanying scattering profile to decide about the reliability of each experiment, especially for those fragments that are considered candidates for more demanding measurements by X-ray crystallography or NMR spectroscopy, or even considered as hits for follow-up chemistry. One specific limitation of the method for fragment screening purposes is that many fragments in the DSi-Poised library have significant intrinsic fluorescence, sometimes even beyond the saturation limit of the detector, and therefore these cannot be properly screened for target binding even at low excitation power. Another point to note for this method is that it can only be used with proteins containing tryptophan residues.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62469/62469fig6v11.jpg" /> 
Figure 6: Effect of temperature on compound solubility. Melting transition curve and scattering profile of Hec1 with two different compounds. (A) The scattering profile shows that for this sample, the solubility is not affected by temperature. (B) The scattering profile shows that the solubility of this sample increases with increasing temperature. The melting transition curve in this case is therefore not reliable. <a href="https://www.jove.com/files/ftp_upload/62469/62469fig6v11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
An open question is what should be considered as a significant change in Tm, not from a mathematical perspective, but from the practical point of view: what change in Tm is important to consider as indicative of binding of a ligand to a protein? In these examples, shifts of less than 1 &#176;C are seen for 74% of the fragments for binding Hec1, 66% for Mps1, and 53% for Nsp5. Considering 1 &#176;C as &#39;significant change&#39; would hardly provide hits that are worthy to pursue by follow-up chemistry. In the overview graphs (Figure 5), bins of 1, 2, 5, or more than 5 degrees of Tm shift, either positive or negative, were considered. This requires modification according to each specific case to give a good overview and to allow informed decision-making, determining the next step. Notably, for some proteins, both stabilization and de-stabilization of the target were observed depending on the fragments considered. Both events are interesting, as both can be the result of fragment binding, and both can lead to a good follow-up molecule to manipulate protein behavior.
A final question remains, namely, &#34;what defines a useful hit?&#34;. In fact, the answer depends on the specific situation. For example, for Hec1, all fragments that stabilize the protein by more than 2 degrees or destabilize it by more than 5 were communicated to our chemistry collaborators, who designed new molecules based on these hits. For Nsp5, however, the most de-stabilizing hits were communicated to our NMR collaborators to confirm the nanoDSF-derived hits with NMR experiments. In other words, the screening results obtained from this protocol should be analyzed with caution and in a context-dependent manner, making informed decisions based on the specific question and surrounding methodology. In any case, the method described here is a complementary approach to existing methodologies such as X-ray and NMR-based screening, which can aim to confirm, prioritize, or give new ideas for chemistry campaigns.
Supplemental Table S1: List of buffers used for screening of proteins. Abbreviations: HEPES = 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid; DTT = dithiothreitol; MOBS = 4-(N-morpholino)butanesulfonic acid. <a href="https://www.jove.com/files/ftp_upload/62469/62469_Supp_Table_S1.xlsx" target="_blank">Please click here to download this Table.</a>
Supplemental Table S2: Properties for MRC 2-well plate for use with nanodispenser robot. <a href="https://www.jove.com/files/ftp_upload/62469/62469_Suppl_Table_S2.xlsx" target="_blank">Please click here to download this Table.</a>
Supplemental Table S3: Properties for 96-well V-bottom plate for use with nanodispenser robot. <a href="https://www.jove.com/files/ftp_upload/62469/62469_Suppl_Table_S3.xlsx" target="_blank">Please click here to download this Table.</a>
Supplemental Table S4: The buffer, protein concentration, and Tm of the proteins discussed in representative results. Abbreviations: DTT = dithiothreitol; Hec1 =&#160;Highly Expressed in Cancer 1 protein; Mps1 = monopolar spindle kinase 1; Nsp5 = SARS-CoV-2 3C-like protease. <a href="https://www.jove.com/files/ftp_upload/62469/62469_Suppl_Table_S4.xlsx" target="_blank">Please click here to download this Table.</a>
Supplemental File 1: Overview parameters-example data. <a href="https://www.jove.com/files/ftp_upload/62469/Supplementaty_file_1.xlsx" target="_blank">Please click here to download this File.</a>
Supplemental File 2: Tm and &#916;Tm values for 406 fragments-example data. <a href="https://www.jove.com/files/ftp_upload/62469/Supplementary_file_2.xlsx" target="_blank">Please click here to download this File.</a>

Drug discovery programs often start by screening chemical compounds for their ability to interact with and/or modify the function of drug targets, most often proteins. The so-called &#34;hits&#34; that are found in such screens, lay the basis for the discovery of novel leads and development candidates, and for most of the new drugs that are being licensed these days. The availability of high-throughput methods is therefore indispensable to screen an enormous number of available targets with a huge number of different compounds to quickly identify their tight binding or their ability to modulate a specific function of the target. After hits are identified, the most promising hit-target combinations are pushed into an extensive drug development pipeline using other, often expensive and time-consuming technologies, to understand &#34;structure-activity relationships&#34; (SAR).
Structural biology approaches, such as those offered by the EU-funded access programs&#160;&#34;Infrastructure for NMR, EM, and X-rays for Translational Research&#34; (iNEXT) and its current successor&#160;iNEXT-Discovery, are often used to study the interactions of numerous compounds in extreme detail while improving the affinity and pharmacological properties of the initial hits by typically several rounds of synthetic chemistry1. Lead compounds that emerge from these &#34;from hit to lead&#34; campaigns become development candidates and enter preclinical studies. The well-developed molecular screening methodology can be roughly categorized into two approaches, namely the ligand-based lead discovery (LBLD) and fragment-based lead discovery (FBLD). In LBLD campaigns, protein receptors are screened with either a few thousand hand-picked ligands (based on the structure of natural ligands or the structure of the target), or with many tens of thousands of compounds in drug-like ligand libraries that cover a large portion of chemical space.
Usually, the compounds are tested for their inhibitory activity in an activity assay, typically monitoring an enzymatic function. In FBLD campaigns2,3,4,5, however, some hundreds of compounds that are typically smaller than drugs (100-200 Dalton) are tested for their ability to bind the target directly, without the use of an activity assay. This binding could interfere with target activity and can be measured by many biophysical methods that report directly on the ability of fragments to bind to the target, or by structural methods such as X-ray crystallography6 and nuclear magnetic resonance spectroscopy7, and more recently, also cryo-electron microscopy. When fragments bind at different locations that are close to each other on the protein, the different, usually low-affinity binding fragments can be rationally combined chemically to create a small set of leads that can be studied in more detail. This frequently results in higher-affinity, more potent compounds, and this methodology has started to yield important molecules with clinical potential. The choice of an &#34;ideal&#34; fragment library that exploits chemical groups efficiently has been an active area of research for many years8,9,10.
While initial emphasis was on covering full chemical space, subsequent attention was focused on enabling the downstream chemical combination of fragment hits to produce lead compounds. Such research has led to the so-called &#34;poised&#34; libraries. These contain fragments with at least one functional group that allow rapid, cheap follow-up synthetic chemistry for efficient progress in studying SAR. One of the activities catalyzed by iNEXT was to update the poised library developed by researchers in the Diamond Light Source and Structural Genomics Consortium. This combined effort resulted in the DSi-Poised library11, which has also been validated within iNEXT12. Later, this library was aligned with the availability of compounds in the REAL Database of Enamine Ltd., a chemical research organization and producer of large collections of building blocks and compound libraries for screening. DSi-Poised is now available to anyone for purchase, but also available in many iNEXT-Discovery partner laboratories for supported fragment screening projects.
The high-end X-ray crystallography and NMR structural biology technologies both have their advantages and disadvantages for FBLD. Both require isolated target samples and yield the high-resolution atomic details that are required for FBLD. However, crystals are necessary for X-ray crystallography, and the fragments bind to cavities in the well-ordered protein regions that are not involved in building the three-dimensional crystal lattice. Solution NMR often yields different hits from X-ray crystallography, as it is unaffected by the crystal environment and is good in detecting binding also in partially ordered protein regions. However, while ligand-based NMR experiments are relatively fast, they still require a substantial amount of time and material and can routinely be done only for relatively small protein targets or domains. For the purpose of prioritizing compounds for crystallographic or NMR experiments, biophysical approaches have been used13,14,15.
As recent instrumentation and computational protocols allow efficient crystallographic screening for FBLD by determining structures and analyzing ~1,000 fragments very efficiently, this prioritization has become less essential in X-ray based research. For NMR however, it remains desirable to use cheaper and quicker experiments to prioritize library screening and save instrument time on highest-end equipment. At the same time, using a combination of essentially different technologies can provide independent confirmation of binding events, or even additional hits that are not picked up by employing only the crystallography or NMR method. Crystallographic and NMR techniques both require very expensive equipment and can often only be done in dedicated external facilities with help from local, highly skilled experts. In addition, the proper analysis of results also demands high expertise. While programs such as iNEXT and iNEXT-Discovery are democratizing access to such facilities16, it has been recognized that cheap, fast, and high-throughput FBLD screening by other methods can encourage drug-screening programs in a much wider range of laboratories. Such results can then be used as an indication to build collaborations with medicinal chemists, and to prioritize the most expensive screening experiments to the most promising compounds if NMR and crystallography facilities impose restrictions on the number of compounds that can be screened.
The TSA forms a quick, efficient, and relatively cheap and accessible biophysical method17 that can be used for FBLD screening. It has been used in multiple settings, from aiding to find stable protein conditions for crystallization trials18, to finding compounds that bind to specific targets in cells19. TSAs have also been used for measuring the dissociation constants for ligands binding target proteins, as ligand binding often leads to alterations in thermal stability. In all TSAs, the change in denaturation temperature of a protein (its stability) is measured as a function of a slow temperature increase. An efficient way to follow protein denaturation upon heating is by DSF or Thermofluor, which quantifies fluorescence quenching of a hydrophobic dye (typically Sypro Orange) upon interaction with exposed hydrophobic regions of protein that unfolds due to temperature increase.
Nano-DSF typically refers to measurement of the thermal stability of protein in the absence of external dyes. One of the first instruments that offered this possibility was the OPTIM1000 that measures a wide spectrum of light intensity as well as light scattering of a sample. This machine allowed the simultaneous measurement of protein unfolding (typically following tryptophan fluorescence) and protein aggregation (as formed nanoparticles result in an increase of light scattering at ~400 nm). Later, the Prometheus introduced the use of back-reflection for measuring aggregation and sensitive detection of the fluorescence signal, allowing screening of low protein concentrations with good sensitivity20. The following section describes how Prometheus was used to demonstrate a fragment screening protocol for detecting hits for different protein targets. A brief introduction about the expected protein quality and quantity is followed by a step-by-step protocol for preparing, performing, and analyzing the fragment screening experiments. Screening results for three proteins have been shown as example data obtained as part of iNEXT-Discovery collaborations.

<table><tbody><tr><td>ClearVue Sheets</td><td>Molecular Dimensions</td><td></td><td>adhesive sealing film for protein plate</td></tr><tr><td>CORNING 6570 Aluminium Sealing Tape</td><td>CORNING</td><td></td><td>adhesive sealing film for fragment plate</td></tr><tr><td>DSi poised library</td><td>Enamine</td><td></td><td>Fragment library containing 768 compounds used in this study</td></tr><tr><td>Elisa Reagent Reservior</td><td>ThermoFisher Scientific</td><td>15075</td><td>Reagent reservior used for pipetting the protein</td></tr><tr><td>Greiner round (U) bottom plates</td><td></td><td>Cat. No.&amp;nbsp;650201</td><td>Fragments supplied in these plates</td></tr><tr><td>Mosquito type X1</td><td>sptlabtech</td><td>Part nr- 3019-0003</td><td>Nanolitre dispenser</td></tr><tr><td>MRC 2-well crystallization plate</td><td></td><td>MRC96T-PS</td><td></td></tr><tr><td>Pierce ELISA Reagent Reservoirs</td><td>Pierce</td><td></td><td></td></tr><tr><td>Prometheus High Sensitivity capillaries</td><td>Catalog PR-C006</td><td></td><td></td></tr><tr><td>Prometheus NT.48 nanoDSF</td><td>Nanotemper</td><td>Catalog nr PR001 (+ Aggregation Detection Optics, catalog nr PR-AGO)</td><td>nanoDSF and light back scattering</td></tr><tr><td>Prometheus Standard capillary type</td><td>Catalog PR-C002</td><td></td><td></td></tr><tr><td>TX-1000</td><td>Thermoscientific</td><td>Centrifuge for plates</td><td></td></tr></tbody></table>

nano-differential scanning fluorimetry for screening in fragment-based lead discovery

Thermal shift assays (TSAs) examine how the melting temperature (Tm) of a target protein changes in response to changes in its environment (e.g., buffer composition). The utility of TSA, and specifically of nano-Differential Scanning Fluorimetry (nano-DSF), has been established over the years, both for finding conditions that help stabilize a specific protein and for looking at ligand binding by monitoring changes in the apparent Tm. This paper presents an efficient screening of the Diamond-SGC-iNEXT Poised (DSi-Poised) fragment library (768 compounds) by the use of nano-DSF, monitoring Tm to identify potential fragment binding. The prerequisites regarding protein quality and concentration for performing nano-DSF experiments are briefly outlined followed by a step-by-step protocol that uses a nano-liter robotic dispenser commonly used in structural biology laboratories for preparing the required samples in 96-well plates. The protocol describes how the reagent mixtures are transferred to the capillaries needed for nano-DSF measurements. In addition, this paper provides protocols to measure thermal denaturation (monitoring intrinsic tryptophan fluorescence) and aggregation (monitoring light back-scattering) and the subsequent steps for data transfer and analysis. Finally, screening experiments with three different protein targets are discussed to illustrate the use of this procedure in the context of lead discovery campaigns. The overall principle of the method described can be easily transferred to other fragment libraries or adapted to other instruments.

A full screen of the DSi-Poised library (768 fragments) was performed on three proteins of medical interest, namely, the outer kinetochore Highly Expressed in Cancer 1 protein (Hec1, or Ndc80), the regulatory tetraricopeptide repeat (TPR) domain of the monopolar spindle kinase 1 (Mps1), and the SARS-CoV-2 3C-like protease, Nsp5, which cleaves off the C-terminus of the replicase polyprotein at 11 sites. The buffer conditions chosen for each protein, as well as the protein concentration and Tm of the proteins, are shown in Supplemental Table S4.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62469/62469fig5v12.jpg" /> 
Figure 5: Frequency distribution of the shift in melting temperature (&#916;Tm) for the three proteins, Hec1, Mps1, and Nsp5, presented in this study as representative results. Abbreviations: Hec1 =&#160;Highly Expressed in Cancer 1 protein; Mps1 = monopolar spindle kinase 1; Nsp5 = SARS-CoV-2 3C-like protease. <a href="https://www.jove.com/files/ftp_upload/62469/62469fig5v11large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The results of the three screenings using the protocol described above, displayed as the frequency distribution of the change in Tm versus the number of fragments, are shown in Figure 5. These plots have been generated as described in section 5.3 of the protocol, plotting the frequency of the observed change in Tm. The significance of the shift needs to be defined in a subjective manner for every different project, as described in the discussion section below. Negative values indicate a reduction in the melting temperature in the presence of a fragment, a positive value an increase in Tm. From such plots, it is easy to observe that for Nsp5, all the fragments have a destabilizing effect, whereas for Hec1 and Mps1, both stabilizing and destabilizing hits are observed. This can be expected and will be discussed.

Watch this Scientific Journal Video about Nano-Differential Scanning Fluorimetry for Screening in Fragment-based Lead Discovery at JoVE.com

Nano-Differential Scanning Fluorimetry for Screening in Fragment-based Lead Discovery

Monitoring changes in the melting temperature of a target protein (i.e., thermal shift assay, TSA) is an efficient method for screening fragment libraries of a few hundred compounds. We present a TSA protocol implementing robotics-assisted nano-Differential Scanning Fluorimetry (nano-DSF) for monitoring intrinsic tryptophan fluorescence and light back-scattering for fragment screening.

Thermal shift assays (TSAs) examine how the melting temperature (Tm) of a target protein changes in response to changes in its environment (e.g., buffer ...

nano-differential-scanning-fluorimetry-for-screening-fragment-based

Nanyang Technological University

Research

JoVE Journal

Biochemistry

4.7K Views.  the Netherlands Cancer Institute. Monitoring changes in the melting temperature of a target protein (i.e., thermal shift assay, TSA) is an efficient method for screening fragment libraries of a few hundred compounds. We present a TSA protocol implementing robotics-assisted nano-Differential Scanning Fluorimetry (nano-DSF) for monitoring intrinsic tryptophan fluorescence and light back-scattering for fragment screening. 

Video: Nano-Differential Scanning Fluorimetry for Screening in Fragment-based Lead Discovery

Preparation and Delivery of Protein Microcrystals in Lipidic Cubic Phase for Serial Femtosecond Crystallography

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural information, typically obtained by crystallography; however MPs are difficult to crystallize. Recent progress in MP structural determination has benefited greatly from the development of lipidic cubic phase (LCP) crystallization methods, which typically yield well-diffracting, but often small crystals that suffer from radiation damage during traditional crystallographic data collection at synchrotron sources. The development of new-generation X-ray free-electron laser (XFEL) sources that produce extremely bright femtosecond pulses has enabled room temperature data collection from microcrystals with no or negligible radiation damage. Our recent efforts in combining LCP technology with serial femtosecond crystallography (LCP-SFX) have resulted in high-resolution structures of several human G protein-coupled receptors, which represent a notoriously difficult target for structure determination. In the LCP-SFX technique, LCP is recruited as a matrix for both growth and delivery of MP microcrystals to the intersection of the injector stream with an XFEL beam for crystallographic data collection. It has been demonstrated that LCP-SFX can substantially improve the diffraction resolution when only sub-10 &#181;m crystals are available, or when the use of smaller crystals at room temperature can overcome various problems associated with larger cryocooled crystals, such as accumulation of defects, high mosaicity and cryocooling artifacts. Future advancements in X-ray sources and detector technologies should make serial crystallography highly attractive and practicable for implementation not only at XFELs, but also at more accessible synchrotron beamlines. Here we present detailed visual protocols for the preparation, characterization and delivery of microcrystals in LCP for serial crystallography experiments. These protocols include methods for conducting crystallization experiments in syringes, detecting and characterizing the crystal samples, optimizing crystal density, loading microcrystal laden LCP into the injector device and delivering the sample to the beam for data collection.

We describe procedures for the preparation and delivery of membrane protein microcrystals in lipidic cubic phase for serial crystallography at X-ray free-electron lasers and synchrotron sources. These protocols can also be applied for incorporation and delivery of soluble protein microcrystals, leading to substantially reduced sample consumption compared to liquid injection.

Membrane proteins (MPs) are essential components of cellular membranes and primary drug targets. Rational drug design relies on precise structural ...

Preparation and In Vivo Use of an Activity-based Probe for N-acylethanolamine Acid Amidase

Activity-based protein profiling (ABPP) is a method for the identification of an enzyme of interest in a complex proteome through the use of a chemical probe that targets the enzyme&#39;s active sites. A reporter tag introduced into the probe allows for the detection of the labeled enzyme by in-gel fluorescence scanning, protein blot, fluorescence microscopy, or liquid chromatography-mass spectrometry. Here, we describe the preparation and use of the compound ARN14686, a click chemistry activity-based probe (CC-ABP) that selectively recognizes the enzyme N-acylethanolamine acid amidase (NAAA). NAAA is a cysteine hydrolase that promotes inflammation by deactivating endogenous peroxisome proliferator-activated receptor (PPAR)-alpha agonists such as palmitoylethanolamide (PEA) and oleoylethanolamide (OEA). NAAA is synthesized as an inactive full-length proenzyme, which is activated by autoproteolysis in the acidic pH of the lysosome. Localization studies have shown that NAAA is predominantly expressed in macrophages and other monocyte-derived cells, as well as in B-lymphocytes. We provide examples of how ARN14686 can be used to detect and quantify active NAAA ex vivo in rodent tissues by protein blot and fluorescence microscopy.

Preparation and In Vivo Use of an Activity-based Probe for N-acylethanolamine Acid Amidase

Here, we describe the preparation and use of an activity-based probe (ARN14686, undec-10-ynyl-N-[(3S)-2-oxoazetidin-3-yl]carbamate) that allows for the detection and quantification of the active form of the proinflammatory enzyme N-acylethanolamine acid amidase (NAAA), both in vitro and ex vivo.

Activity-based protein profiling (ABPP) is a method for the identification of an enzyme of interest in a complex proteome through the use of a chemical ...

A Rapid and Quantitative Fluorimetric Method for Protein-Targeting Small Molecule Drug Screening

We demonstrate a new drug screening method for determining the binding affinity of small drug molecules to a target protein by forming fluorescent gold nanoclusters (Au NCs) within the drug-loaded protein, based on the differential fluorescence signal emitted by the Au NCs. Albumin proteins such as human serum albumin (HSA) and bovine serum albumin (BSA) are selected as the model proteins. Four small molecular drugs (e.g., ibuprofen, warfarin, phenytoin, and sulfanilamide) of different binding affinities to the albumin proteins are tested. It was found that the formation rate of fluorescent Au NCs inside the drug loaded albumin protein under denaturing conditions (i.e., 60 &#176;C or in the presence of urea) is slower than that formed in the pristine protein (without drugs). Moreover, the fluorescent intensity of the as-formed NCs is found to be inversely correlated to the binding affinities of these drugs to the albumin proteins. Particularly, the higher the drug-protein binding affinity, the slower the rate of Au NCs formation, and thus a lower fluorescence intensity of the resultant Au NCs is observed. The fluorescence intensity of the resultant Au NCs therefore provides a simple measure of the relative binding strength of different drugs tested. This method is also extendable to measure the specific drug-protein binding constant (KD) by simply varying the drug content preloaded in the protein at a fixed protein concentration. The measured results match well with the values obtained using other prestige but more complicated methods.

A protocol for small molecular drug screening based on in-situ synthesis of ultrasmall fluorescent gold nanoclusters (Au NCs) using drug-loaded protein as template is presented. This method is simple to determine the binding affinity of drugs to a target protein by a visible fluorescent signal emitted from the protein-templated Au NCs.

We demonstrate a new drug screening method for determining the binding affinity of small drug molecules to a target protein by forming fluorescent gold ...

Bioengineering

Natural Product Discovery with LC-MS/MS Diagnostic Fragmentation Filtering: Application for Microcystin Analysis

Natural products are often biosynthesized as mixtures of structurally similar compounds, rather than a single compound. Due to their common structural features, many compounds within the same class undergo similar MS/MS fragmentation and have several identical product ions and/or neutral losses. The purpose of diagnostic fragmentation filtering (DFF) is to efficiently detect all compounds of a given class in a complex extract by screening non-targeted LC-MS/MS datasets for MS/MS spectra that contain class specific product ions and/or neutral losses. This method is based on a DFF module implemented within the open-source MZmine platform that requires sample extracts be analyzed by data-dependent acquisition on a high-resolution mass spectrometer such as quadrupole Orbitrap or quadrupole time-of-flight mass analyzers. The main limitation of this approach is the analyst must first define which product ions and/or neutral losses are specific for the targeted class of natural products. DFF allows for the subsequent discovery of all related natural products within a complex sample, including new compounds. In this work, we demonstrate the effectiveness of DFF by screening extracts of Microcystis aeruginosa, a prominent harmful algal bloom causing cyanobacteria, for the production of microcystins.

Diagnostic fragmentation filtering, implemented into MZmine, is an elegant, post-acquisition approach to screen LC-MS/MS datasets for entire classes of both known and unknown natural products. This tool searches MS/MS spectra for product ions and/or neutral losses that the analyst has defined as being diagnostic for the entire class of compounds.

Natural products are often biosynthesized as mixtures of structurally similar compounds, rather than a single compound. Due to their common structural ...

Chemistry

Workflow and Tools for Crystallographic Fragment Screening at the Helmholtz-Zentrum Berlin

Fragment screening is a technique that helps to identify promising starting points for ligand design. Given that crystals of the target protein are available and display reproducibly high-resolution X-ray diffraction properties, crystallography is among the most preferred methods for fragment screening because of its sensitivity. Additionally, it is the only method providing detailed 3D information of the binding mode of the fragment, which is vital for subsequent rational compound evolution. The routine use of the method depends on the availability of suitable fragment libraries, dedicated means to handle large numbers of samples, state-of-the-art synchrotron beamlines for fast diffraction measurements and largely automated solutions for the analysis of the results.
Here, the complete practical workflow and the included tools on how to conduct crystallographic fragment screening (CFS) at the Helmholtz-Zentrum Berlin (HZB) are presented. Preceding this workflow, crystal soaking conditions as well as data collection strategies are optimized for reproducible crystallographic experiments. Then, typically in a one to two-day procedure, a 96-membered CFS-focused library provided as dried ready-to-use plates is employed to soak 192 crystals, which are then flash-cooled individually. The final diffraction experiments can be performed within one day at the robot-mounting supported beamlines BL14.1 and BL14.2 at the BESSY&#160; II electron storage ring operated by the HZB in Berlin-Adlershof (Germany). Processing of the crystallographic data, refinement of the protein structures, and hit identification is fast and largely automated using specialized software pipelines on dedicated servers, requiring little user input.
Using the CFS workflow at the HZB enables routine screening experiments. It increases the chances for successful identification of fragment hits as starting points to develop more potent binders, useful for pharmacological or biochemical applications.

Crystallographic fragment screening at the Helmholtz-Zentrum Berlin is performed using a workflow with dedicated compound libraries, crystal handling tools, fast data collection facilities&#160;and largely automated data analysis. The presented protocol intends to maximize the output of such experiments to provide promising starting points for downstream structure-based ligand design.

Fragment screening is a technique that helps to identify promising starting points for ligand design. Given that crystals of the target protein are ...

NMR-Based Fragment Screening in a Minimum Sample but Maximum Automation Mode

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest advantage of NMR-based fragment screening is its ability not only to detect binders over 7-8 orders of magnitude of affinity but also to monitor purity and chemical quality of the fragments and thus to produce high quality hits and minimal false positives or false negatives. A prerequisite within the FBS is to perform initial and periodic quality control of the fragment library, determining solubility and chemical integrity of the fragments in relevant buffers, and establishing multiple libraries to cover diverse scaffolds to accommodate various macromolecule target classes (proteins/RNA/DNA). Further, an extensive NMR-based screening protocol optimization with respect to sample quantities, speed of acquisition and analysis at the level of biological construct/fragment-space, in condition-space (buffer, additives, ions, pH, and temperature) and in ligand-space (ligand analogues, ligand concentration) is required. At least in academia, these screening efforts have so far been undertaken manually in a very limited fashion, leading to limited availability of screening infrastructure not only in the drug development process but also in the context of chemical probe development. In order to meet the requirements economically, advanced workflows are presented. They take advantage of the latest state-of-the-art advanced hardware, with which the liquid sample collection can be filled in a temperature-controlled fashion into the NMR-tubes in an automated manner. 1H/19F NMR ligand-based spectra are then collected at a given temperature. High-throughput sample changer (HT sample changer) can handle more than 500 samples in temperature-controlled blocks. This together with advanced software tools speeds up data acquisition and analysis. Further, application of screening routines on protein and RNA samples are described to make aware of the established protocols for a broad user base in biomacromolecular research.

Fragment-based screening by NMR is a robust method to rapidly identify small molecule binders to biomacromolecules (DNA, RNA, or proteins). Protocols describing automation-based sample preparation, NMR experiments &#38; acquisition conditions, and analysis workflows are presented. The technique allows for optimal exploitation of both 1H and 19F NMR-active nuclei for detection.

Fragment-based screening (FBS) is a well-validated and accepted concept within the drug discovery process both in academia and industry. The greatest ...

Achieving Efficient Fragment Screening at XChem Facility at Diamond Light Source

In fragment-based drug discovery, hundreds or often thousands of compounds smaller than ~300 Da are tested against the protein of interest to identify chemical entities that can be developed into potent drug candidates. Since the compounds are small, interactions are weak, and the screening method must therefore be highly sensitive; moreover, structural information tends to be crucial for elaborating these hits into lead-like compounds. Therefore, protein crystallography has always been a gold-standard technique, yet historically too challenging to find widespread use as a primary screen.
Initial XChem experiments were demonstrated in 2014 and then trialed with academic and industrial collaborators to validate the process. Since then, a large research effort and significant beamtime have streamlined sample preparation, developed a fragment library with rapid follow-up possibilities, automated and improved the capability of I04-1 beamline for unattended data collection, and implemented new tools for data management, analysis and hit identification.
XChem is now a facility for large-scale crystallographic fragment screening, supporting the entire crystals-to-deposition process, and accessible to academic and industrial users worldwide. The peer-reviewed academic user program has been actively developed since 2016, to accommodate projects from as broad a scientific scope as possible, including well-validated as well as exploratory projects. Academic access is allocated through biannual calls for peer-reviewed proposals, and proprietary work is arranged by Diamond's Industrial Liaison group. This workflow has already been routinely applied to over a hundred targets from diverse therapeutic areas, and effectively identifies weak binders (1%-30% hit rate), which both serve as high-quality starting points for compound design and provide extensive structural information on binding sites. The resilience of the process was demonstrated by continued screening of SARS-CoV-2 targets during the COVID-19 pandemic, including a 3-week turn-around for the main protease.

This paper describes the complete XChem process for crystal-based fragment screening, starting from applying for access and all subsequent steps to data dissemination.

In fragment-based drug discovery, hundreds or often thousands of compounds smaller than ~300 Da are tested against the protein of interest to identify ...

Quantitative-Irreversible Tethering: Screening Electrophilic Ligands Efficiently

Covalent Fragment Screening Using the Quantitative Irreversible Tethering Assay

Compounds that form covalent bonds with specific target proteins offer a variety of advantages&#160;as chemical probes and therapeutic agents. Most commonly, mildly reactive, electrophilic small molecules are employed to form covalent bonds with select cysteine side chains in specific proteins. Electrophile-first approaches of ligand discovery, whereby a library of electrophilic small molecules are screened against a protein target, have become popular as they avoid the need for time-consuming downstream installation of an electrophilic warhead. Such screening is complicated, however, as electrophilic ligands can exhibit a wide range of different rates of spontaneous reaction with cysteines. Quantitative-irreversible tethering (qIT) offers a fluorescence-based method for hit identification and development that normalizes data for these differences in intrinsic compound reactivity. Rates of reaction of individual compounds with a target protein are determined and compared to compound reactivity with the unstructured tripeptide glutathione (this being a proxy for spontaneous compound reaction), enabling the identification of compounds that preferentially react with the protein of interest. This methodology has been successfully applied to identify selective covalent fragments against several drug targets, including SARS-CoV-2 main protease, cyclin-dependent kinase 2, and RAP27A. Here, we demonstrate the application of qIT to a target protein to generate a quantitative and robust data set, allowing prioritization of hit ligands for future development.

The quantitative irreversible tethering (qIT) assay is a fluorescence-based method of identifying covalent fragments that selectively engage with a target protein. Here, a detailed protocol is outlined to describe the qIT assay, with example results included.

Compounds that form covalent bonds with specific target proteins offer a variety of advantages&#160;as chemical probes and therapeutic agents. Most ...

Determination of Protein-ligand Interactions Using Differential Scanning Fluorimetry

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

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

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

Biology

High-Sensitivity Detection of Compound-Protein Binding Using Fluorescence

Detection of CD40 Protein-Umbelliferone Interaction via Differential Scanning Fluorescence

The investigation of interactions between different molecules is a crucial aspect of understanding disease pathogenesis and screening for drug targets. Umbelliferone, an active ingredient in Tibetan medicine Vicatia thibetica, exhibits an immunomodulatory effect with an unknown mechanism. The CD40 protein is a key target in the immune response. Therefore, this study employs the principle of differential scanning fluorescence technology to analyze the interactions between CD40 protein and umbelliferone using fluorescent enzyme markers. Initially, the stability of the protein fluorescent orange dye was experimentally verified, and the optimal dilution ratio of 1:500 was determined. Subsequently, it was observed that the temperature melting (Tm) value of CD40 protein tended to decrease with an increase in concentration. Interestingly, the interaction between CD40 protein and umbelliferone was found to enhance the thermal stability of CD40 protein. This study represents the first attempt to detect the binding potential of small molecule compounds and proteins using fluorescence microplates and fluorescent dyes. The technique is characterized by high sensitivity and accuracy, promising advancements in the fields of protein stability, protein structure, and protein-ligand interactions, thus facilitating further research and exploration.

Author Spotlight: Streamlining PCR Methods for Enhanced Accessibility and Efficiency

This protocol describes a unique method for detecting the binding between compounds and protein molecules, offering the advantages of minimal protein sample loss and high data accuracy.

The investigation of interactions between different molecules is a crucial aspect of understanding disease pathogenesis and screening for drug targets. ...

Nano-Differential Scanning Fluorimetry for Screening in Fragment-based Lead Discovery

Summary

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Determination of Protein-ligand Interactions Using Differential Scanning Fluorimetry

Detection of CD40 Protein-Umbelliferone Interaction via Differential Scanning Fluorescence