This work was supported by the Scientific Research Fund of Zhejiang Chinese Medicine University (2020ZG29), the Basic Public Welfare Research Project of Zhejiang Province (LGF19H150006, LTGY23B070001), the Project of Zhejiang Provincial Department of Education (Y202147028) and the Project of Experimental Technology of Zhejiang University Laboratory Department (SJS201712, SYB202130).

1. Cell culture
<ol>
	<li>Prepare a culture medium of Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS). Culture the human pancreatic cancer cell, PANC-1, in a 75 cm2 culture flask at a density of 1 x 106 cells/mL. Incubate the culture at 37 &#176;C with 5% CO2.</li>
	<li>Subculture the cells when cell density reaches 75%-80% under microscopic observation. Remove the medium, and rinse 2x with 3-5 mL of phosphate buffer saline (PBS; 0.01 M, pH 7.2-7.4).</li>
	<li>Discard the solution, add 1-2 mL of trypsin-EDTA solution, and let the cells detach until they become rounded and suspended in the medium under the microscope. Add fresh medium, aspirate, and disperse into 75 cm2 culture flasks with 15 mL culture medium. Use a sub-cultivation ratio of 1:2 to 1:4 (recommended). 
	&#8203;NOTE: All operations are performed in a biosafety cabinet to avoid cell contamination.</li>
</ol>
2. Culture medium collection
<ol>
	<li>Incubate the cells for 48 h at 37 &#176;C with 5% CO2. Collect cell supernatant (90 mL) in two 50 mL centrifuge tubes and centrifuge at 300 x g for 10 min at 4 &#176;C.</li>
	<li>Transfer the supernatant to new sterile tubes and centrifuge successively at 2,000 x g for 20 min, 10 000 x g for 30 min, and 12,000 x g for 70 min at 4 &#176;C.</li>
	<li>Remove the supernatant and rinse the pellet with 10 mL of PBS by pipetting gently. Centrifuge at 12,000 x g for 70 min at 4 &#176;C again and resuspend the pellet in 10 mL of PBS buffer to obtain a mixture of EVs.</li>
	<li>Perform nanoparticle tracking analysis (NTA) to quantify and size the EV mixture. Perform the NTA analysis according to the instrument operation reference manual and reference21.</li>
</ol>
3. Cell sorting
<ol>
	<li>Perform the standard startup procedure of the flow cell sorter. Turn ON the machine by pressing the Startup button. Click the Laser Power button on the laser control panel. Press the Change Tanks button in the smart sampler submenu to pressurize the fluidics. 
	NOTE: Ensure that the waste tank is empty, and the sheath tank is full before starting up.</li>
	<li>Use an ultrasonically cleaned 50 &#181;m jet-in-air nozzle and install it onto the nozzle assembly. Adjust the pressure to 80 psi for sheath fluid and 80.3 psi for sample flow on the pressure console.</li>
	<li>Press the Start Sheath Flow button to start the sheath stream flowing. Click the Bubble Button in the smart sampler submenu to automatically debubble for 10 min and then turn it off. 
	NOTE: For cell sorting, different sizes of air spray nozzles require different sheath fluid pressures to maintain liquid flow stability. In this method, a nozzle of 50 &#181;m was used, and only a sheath fluid pressure greater than or equal to 80 psi could generate stable side streams. The pressure difference between the sample flow and the sheath fluid determines the loading speed of sorting. Under the setting condition of the method (the pressure difference between the sample flow and the sheath fluid is 0.3-0.6 psi), the liquid flow was relatively stable, and the loading speed enabled the sorting efficiency to increase to 80% from 50% for pressure difference less than 0.3-0.6 psi.</li>
	<li>Align the stream and determine the laser spot. Turn ON the illumination chamber light to view the stream over the pinholes. Adjust the up-and-down, left-and-right, and front-and-back micrometers to center the stream on the pinhole and focus the stream.</li>
	<li>Select the Laser and Stream Intercept tab. Press the Green Arrow button continuously as prompted to complete the laser intercept and nozzle alignment calibration process.</li>
	<li>Access the fine laser alignment screen. Select 640-722/44 (H) channel as the X-axis parameter and 405-448/59 (H) channel as the Y-axis parameter. Press the Trigger Parameter Selector and choose the SSC Parameter of the 488 nm laser. Open all laser shutters. 
	NOTE: The X-axis and Y-axis parameters chosen depend on the laser configuration of the system. Other parameters are allowed if the system does not have a 640 nm or 405 nm laser. For best results, select lasers with the greatest spatial separation on the pinhole strip (not including the 355 nm laser).</li>
	<li>Dilute the rainbow fluorescent particles by adding one drop into 1 mL of double distilled water for a final concentration of 1 x 106 beads per mL. Open the door of the sample chamber and load a tube of the prepared rainbow fluorescent particles.</li>
	<li>Press the Start Sample button. Adjust up-and-down micrometers to optimize fluorescence intensity, making each signal as intense as possible. Press the Start Quality Control (QC) button on the touchscreen control panel to perform QC automatically.</li>
	<li>Perform drop delay. Access the sort setup screen and select the drop delay button. Press the IntelliSort Initialization button. The instrument automatically adjusts the frequency and amplitude of droplet oscillation to obtain the drop delay of 25 &#177; 5 and be able to visualize the break-off point of the drop clearly. Press the Maintain button.</li>
	<li>Select tube as the sort output type. Place a 15 mL tube holder in the sort chamber. Turn the plate voltage ON and set the plate voltage to approximately 4000 V.</li>
	<li>Turn on the test pattern by selecting the Stream Setup button. Adjust the charge phase slider and deflection slider to ensure that the image of the stream is lined up with the collection tube. Select the Check Mark button.</li>
	<li>Log in to the Summit workstation on the computer. Create a new protocol from the File main menu. Create a dot plot by selecting the Histogram tab in the Summit software control panel.</li>
	<li>Right-click the axes of the new dot plot in the workspace and choose FSC as the X-axis parameter and SSC as the Y-axis parameter. Present the FSC and SSC in logarithmic form.</li>
	<li>Select the Acquisition tab and locate the acquisition parameters panel. Enable all signals in the submenu of enabling parameters. Choose FSC as the trigger parameter and set the threshold of 0.01. Adjust the voltage and gain of FSC and SSC at 250 and 0.6 to ensure events within the FSC vs. SSC plot and populations are separated. 
	&#8203;NOTE: Threshold setting is equivalent to setting the particle size that can be detected by flow cytometry. The larger the threshold is, the larger the detected particles will be, and the small particles will be cut off by the threshold and cannot be detected. In this method, the threshold is set to 0.01, so all particles larger than electronic noise can be detected.</li>
</ol>
4. Isolation of sEV
<ol>
	<li>Dilute the fluorescent polystyrene microspheres to suspensions of 100 nm, 200 nm, and 300 nm. Add 1 &#181;L of microspheres to 1 mL of double distilled water.</li>
	<li>Load the microsphere suspension successively. Select Start under the acquisition menu of summit software and record 20,000 events. 
	NOTE: The number of events is optional. No less than 5,000 events are recommended to meet statistical significance.</li>
	<li>Right-click on the FSC vs. SSC dot plot to create rectangles and drag to resize and reposition the regions of electronic noise and 100 nm microsphere population. Right-click on the regions to rename them as R7 and R4, respectively. Repeat the above steps to frame the 200 nm and 300 nm microspheres with rectangles successively and rename them as R5 and R6, respectively.</li>
	<li>Finally, determine the 50-200 nm sorting region based on the electronic noise and the position of 200 nm particles defined as R8. 
	NOTE: The lower detection limit of 50 nm is placed just above the electronic noise of the flow cell sorter. The region between the noise and the 200 nm microsphere population can therefore be defined to be between 50-200 nm.</li>
	<li>Edit sort decisions in the sort logic and statistics panel. Double-click on the blank field of the left one (L1) stream. Select the region named R8 to create the sort logic. Choose the Abort mode of purity and select a droplet envelope of 1 drop for the L1 stream.</li>
	<li>Load 500 &#181;L sample of EVs mixture from step 2.3. Press the Start button under the acquisition menu in Summit to acquire data, and then press Start in the sort menu to collect 50-200 nm sEV into a 15 mL centrifuge tube. 
	NOTE: A sterile environment is necessary to minimize contamination during any procedure.</li>
	<li>Observe the presence of sEV by transmission electron microscopy (TEM) and verify the particle size of sEV using NTA. Perform western blot (WB) analysis to further confirm the expression of CD9, CD63, and CD81 markers18.</li>
</ol>

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The authors declare no conflicts of interest.

This protocol outlines an optimized method to isolate and purify sEV with the specified particle size of 50-200 nm using a flow cell sorter, which was validated by NTA. The method solved the bottleneck problem of obtaining sEV with uniform particle size and high purity, avoiding interference from unrelated biological molecules wrapped in large-sized EVs22. Fast, high-throughput analyses are possible with flow cytometry, which can capture 100,000 particles per second and make 70,000 sorting decisio...

A cell releases extracellular vesicles (EVs) of varying sizes that result in signaling molecules and membrane inclusions, which are important for intercellular communication1. EVs of different sizes also play different biological roles, with 50-200 nm sEV being able to precisely distribute RNA, DNA, and proteins to the correct extracellular location. The sEV also helps determine their secretion mechanisms, involving not only the regulation of normal physiological processes such as immune surveillance, stem cell maintenance, blood coagulation, and tissue repair but also the pathology underlying several diseases such as tumor progression and metastasis2,3. Effective isolation and analysis of sEV are critical for identifying biomarkers and designing future drug interventions.
With continuous research on the clinical application of sEV, the isolation methods of sEV have put forward higher requirements. Due to the heterogeneity of sEV in size, source, and contents, as well as their similarity with other EVs in physicochemical and biochemical properties, there is no standard method for sEV isolation4,5. Currently, ultracentrifugation, size exclusion chromatography (SEC), polymer precipitation, and immunoaffinity capture are the most common sEV isolation methods6. Ultracentrifugation is still the gold standard for sEV isolation in research, despite being time-consuming, resulting in low purity with a wide size distribution of 40-500 nm and significant mechanical damage to the sEV after long-term centrifugation7,8,9. Polymer precipitation, which usually uses polyethylene glycol (PEG), suffers from unacceptable purity for subsequent functional analysis with concomitant precipitation of extracellular protein aggregates and polymer contamination10,11. Immunoaffinity capture-based methods require high-cost antibodies with varying specificity, as well as have problems with low processing volume and yields12,13,14. Particle size is one of the main indicators to evaluate the purity of isolated sEV. Although sEV purity remains an unattainable goal, SEC removes a considerable quantity of medium components, and the sEV particles extracted by the SEC method are mainly in the range of 50-200 nm15. The existing techniques have a few disadvantages, including but not limited to being time-consuming, low purity, low yield, poor reproducibility, low throughput of samples, and potential damage of sEV, which makes it incompatible with clinical utilization16. Thus, a rapid, inexpensive, and size-specified sEV isolation method applicable to diverse biofluids is an essential need in several research and clinical situations.
In flow cytometry, single particles are analyzed in a high-throughput, multiparametric manner, and subsets are sorted out17. Due to the heterogeneity of EVs, a single-particle flow cytometric measurement would be ideal, which has been used to investigate EVs following the paradigms of cell analysis, with light scatter and fluorescence labels being used to identify physiology-related features and protein components18,19,20. Nevertheless, conventional flow cytometry is challenged by the small size of sEV and low abundance of surface biomarkers. The sensitivity of flow cytometry could be improved by optimizing detection parameters to distinguish background noise and sEV regardless of using forward scatter (FSC), side scatter (SSC), or fluorescence threshold triggering parameter19.
With the present protocol, high-resolution flow cytometric sort settings were optimized using fluorescent beads as standard. By selecting the proper nozzle size, sheath fluid pressure, threshold triggering parameter, and voltages controlling scattered light intensities, we were able to isolate a specific subset of sEV from a complex mixture.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Centrifuge tube</td>
			<td>Beckman Coulter</td>
			<td>344058</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture flasks</td>
			<td>Corning&#160;</td>
			<td>430641</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s modified eagle medium</td>
			<td>Corning Cellgro</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum</td>
			<td>SUER</td>
			<td>SUER050QY</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow cell sorter</td>
			<td>Beckman Coulter</td>
			<td>Moflo Astrios EQ</td>
			<td></td>
		</tr>
		<tr>
			<td>Human pancreatic cancer cell, PANC-1</td>
			<td>NA</td>
			<td>NA</td>
			<td>PANC-1 cells were donated by Professor Weijun Yang, College of Life Sciences, Zhejiang University</td>
		</tr>
		<tr>
			<td>Laser particle size and zeta potential analyzer&#160;</td>
			<td>Malvern</td>
			<td>Zetasizer Nano ZS 90</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffer saline</td>
			<td>Gibco</td>
			<td>C20012500BT</td>
			<td></td>
		</tr>
		<tr>
			<td>Polystyrene fluorescent microspheres</td>
			<td>Beckman Coulter</td>
			<td>6602336</td>
			<td></td>
		</tr>
		<tr>
			<td>Transmission electron microscopy</td>
			<td>JEOL</td>
			<td>JEM-1200EX</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA solution</td>
			<td>Gibco</td>
			<td>1713949</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultra rainbow fluorescent particles</td>
			<td>Beckman Coulter</td>
			<td>B28479</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge</td>
			<td>Beckman Coulter</td>
			<td>Optima-L80XP</td>
			<td></td>
		</tr>
		<tr>
			<td>Ultracentrifuge rotor</td>
			<td>Beckman Coulter</td>
			<td>SW32TI</td>
			<td></td>
		</tr>
	</tbody>
</table>

optimization of flow cytometric sorting parameters for high-throughput isolation and purification of small extracellular vesicles

Small extracellular vesicles (sEV) can be released from all cell types and carry protein, DNA, and RNA. Signaling molecules serve as indicators of the physiological and pathological state of a cell. However, there is no standard method for sEV isolation, which prevents downstream biomarker identification and drug intervention studies. In this article, we provide a detailed protocol for the isolation and purification of 50-200 nm sEV by a flow cell sorter. For this, a 50 &#181;m nozzle and 80 psi sheath fluid pressure were selected to obtain a good sorting rate and stable side stream. Standard sized polystyrene microspheres were used to locate populations of 100, 200, and 300 nm particles. With additional optimization of the voltage, gain, and forward scatter (FSC) triggering threshold, the sEV signal could be separated from the background noise. These optimizations provide a panel of critical sort settings that enables one to obtain a representative population of sEV using FSC vs. side scatter (SSC) only. The flow cytometry-based isolation method not only allows for high-throughput analysis but also allows for synchronous classification or proteome analysis of sEV based on the biomarker expression, opening numerous downstream research applications.

The flow chart diagram for the experimental protocol is shown in Figure 1. In this method, standard sized polystyrene microspheres were used as reference standards for particle size distribution. Under the specific instrumental parameter condition, the particle signal could be clearly distinguished from the background noise in the FSC vs. SSC plot using the logarithmic form. Gating strategies are shown in Figure 2. R4, R5, and R6 refer to the positions of 100 nm...

Watch this Scientific Journal Video about Optimization of Flow Cytometric Sorting Parameters for High-Throughput Isolation and Purification of Small Extracellular Vesicles at JoVE.com

Optimization of Flow Cytometric Sorting Parameters for High-Throughput Isolation and Purification of Small Extracellular Vesicles

This protocol provides a rapid and size-specific isolation method for small extracellular vesicles by optimizing the size of the air spray nozzle, sheath fluid pressure, sample flow pressure, voltage, gain, and triggering threshold parameters.

Small extracellular vesicles (sEV) can be released from all cell types and carry protein, DNA, and RNA. Signaling molecules serve as indicators of the ...

optimization-flow-cytometric-sorting-parameters-for-high-throughput

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Biochemistry

2.1K Views.  Zhejiang University School of Medicine. This protocol provides a rapid and size-specific isolation method for small extracellular vesicles by optimizing the size of the air spray nozzle, sheath fluid pressure, sample flow pressure, voltage, gain, and triggering threshold parameters.

Video: Optimization of Flow Cytometric Sorting Parameters for High-Throughput Isolation and Purification of Small Extracellular Vesicles

Method for Identifying Small Molecule Inhibitors of the Protein-protein Interaction Between HCN1 and TRIP8b

Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels are expressed ubiquitously throughout the brain, where they function to regulate the excitability of neurons. The subcellular distribution of these channels in pyramidal neurons of hippocampal area CA1 is regulated by tetratricopeptide repeat-containing Rab8b interacting protein (TRIP8b), an auxiliary subunit. Genetic knockout of HCN pore forming subunits or TRIP8b, both lead to an increase in antidepressant-like behavior, suggesting that limiting the function of HCN channels may be useful as a treatment for Major Depressive Disorder (MDD). Despite significant therapeutic interest, HCN channels are also expressed in the heart, where they regulate rhythmicity. To circumvent off-target issues associated with blocking cardiac HCN channels, our lab has recently proposed targeting the protein-protein interaction between HCN and TRIP8b in order to specifically disrupt HCN channel function in the brain. TRIP8b binds to HCN pore forming subunits at two distinct interaction sites, although here the focus is on the interaction between the tetratricopeptide repeat (TPR) domains of TRIP8b and the C terminal tail of HCN1. In this protocol, an expanded description of a method for purifying TRIP8b and executing a high throughput screen to identify small molecule inhibitors of the interaction between HCN and TRIP8b, is described. The method for high throughput screening utilizes a Fluorescence Polarization (FP) -based assay to monitor the binding of a large TRIP8b fragment to a fluorophore-tagged eleven amino acid peptide corresponding to the HCN1 C terminal tail. This method allows &#39;hit&#39; compounds to be identified based on the change in the polarization of emitted light. Validation assays are then performed to ensure that &#39;hit&#39; compounds are not artifactual.

The interaction between HCN channels and their auxiliary subunit has been identified as a therapeutic target in Major Depressive Disorder. Here, a fluorescence polarization-based method for identifying small molecule inhibitors of this protein-protein interaction, is presented.

Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels are expressed ubiquitously throughout the brain, where they function to regulate the ...

Extracellular Protein Microarray Technology for High Throughput Detection of Low Affinity Receptor-Ligand Interactions

Secreted factors, membrane-tethered receptors, and their interacting partners are main regulators of cellular communication and initiation of signaling cascades during homeostasis and disease, and as such represent prime therapeutic targets. Despite their relevance, these interaction networks remain significantly underrepresented in current databases; therefore, most extracellular proteins have no documented binding partner. This discrepancy is primarily due to the challenges associated with the study of the extracellular proteins, including expression of functional proteins, and the weak, low affinity, protein interactions often established between cell surface receptors. The purpose of this method is to describe the printing of a library of extracellular proteins in a microarray format for screening of protein-protein interactions. To enable detection of weak interactions, a method based on multimerization of the query protein under study is described. Coupled to this microbead-based multimerization approach for increased multivalency, the protein microarray allows robust detection of transient protein-protein interactions in high throughput. This method offers a rapid and low sample consuming-approach for identification of new interactions applicable to any extracellular protein. Protein microarray printing and screening protocol are described. This technology will be useful for investigators seeking a robust method for discovery of protein interactions in the extracellular space.

Here, we present a protocol to screen extracellular protein microarrays for identification of novel receptor-ligand interactions in high throughput. We also describe a method to enhance detection of transient protein-protein interactions by using protein-microbead complexes.

Secreted factors, membrane-tethered receptors, and their interacting partners are main regulators of cellular communication and initiation of signaling ...

Isolation of Tissue Extracellular Vesicles from the Liver

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in cell-to-cell communication by shuttling bioactive molecules such as RNA or protein from one cell to another. Most studies of EVs have been performed in cell culture models or in EVs isolated from body fluids. There is emerging interest in the isolation of EVs from tissues to study their contribution to physiological processes and how they are altered in disease. The isolation of EVs with sufficient yield from tissues is technically challenging because of the need for tissue dissociation without cellular damage. This method describes a procedure for the isolation of EVs from mouse liver tissue. The method involves a two-step process starting with in situ collagenase digestion followed by differential ultra-centrifugation. Tissue perfusion using collagenase provides an advantage over mechanical cutting or homogenization of liver tissue due to its increased yield of obtained EVs. The use of this two-step process to isolate EVs from the liver will be useful for the study of tissue EVs.

This is a protocol to isolate tissue extracellular vesicles (EVs) from the liver. The protocol describes a two-step process involving collagenase perfusion followed by differential ultracentrifugation to isolate liver tissue EVs.

Extracellular vesicles (EVs) can be released from many different cell types and detected in most, if not all, body fluids. EVs can participate in ...

Imaging of Extracellular Vesicles by Atomic Force Microscopy

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane proteins and other molecules, and diverse luminal content inherited from a parent cell, which includes RNAs, proteins, and DNAs. The characterization of the hydrodynamic sizes of EVs, which depends on the size of the vesicle and its coronal layer formed by surface decorations, has become routine. For exosomes, the smallest of EVs, the relative difference between the hydrodynamic and vesicles sizes is significant. The characterization of vesicles sizes by the cryogenic transmission electron microscopy (cryo-TEM) imaging, a gold standard technique, remains a challenge due to the cost of the instrument, the expertise required to perform the sample preparation, imaging and data analysis, and a small number of particles often observed in images. A widely available and accessible alternative is the atomic force microscopy (AFM), which can produce versatile data on three-dimensional geometry, size, and other biophysical properties of extracellular vesicles. The developed protocol guides the users in utilizing this analytical tool and outlines the workflow for the analysis of EVs by the AFM, which includes the sample preparation for imaging EVs in hydrated or desiccated form, the electrostatic immobilization of vesicles on a substrate, data acquisition, its analysis, and interpretation. The representative results demonstrate that the fixation of EVs on the modified mica surface is predictable, customizable, and allows the user to obtain sizing results for a large number of vesicles. The vesicle sizing based on the AFM data was found to be consistent with the cryo-TEM imaging.&#160;

A step-by-step procedure is described for label-free immobilization of exosomes and extracellular vesicles from liquid samples and their imaging by atomic force microscopy (AFM). The AFM images are used to estimate the size of the vesicles in the solution and characterize other biophysical properties.&#160;

Exosomes and other extracellular vesicles (EVs) are molecular complexes consisting of a lipid membrane vesicle, its surface decoration by membrane ...

A Semi-High-Throughput Adaptation of the NADH-Coupled ATPase Assay for Screening Small Molecule Inhibitors

ATPase enzymes utilize the free energy stored in adenosine triphosphate to catalyze a wide variety of endergonic biochemical processes in vivo that would not occur spontaneously. These proteins are crucial for essentially all aspects of cellular life, including metabolism, cell division, responses to environmental changes and movement. The protocol presented here describes a nicotinamide adenine dinucleotide (NADH)-coupled ATPase assay that has been adapted to semi-high throughput screening of small molecule ATPase inhibitors. The assay has been applied to cardiac and skeletal muscle myosin II's, two actin-based molecular motor ATPases, as a proof of principle. The hydrolysis of ATP is coupled to the oxidation of NADH by enzymatic reactions in the assay. First, the ADP generated by the ATPase is regenerated to ATP by pyruvate kinase (PK). PK catalyzes the transition of phosphoenolpyruvate (PEP) to pyruvate in parallel. Subsequently, pyruvate is reduced to lactate by lactate dehydrogenase (LDH), which catalyzes the oxidation of NADH in parallel. Thus, the decrease in ATP concentration is directly correlated to the decrease in NADH concentration, which is followed by change to the intrinsic fluorescence of NADH. As long as PEP is available in the reaction system, the ADP concentration remains very low, avoiding inhibition of the ATPase enzyme by its own product. Moreover, the ATP concentration remains nearly constant, yielding linear time courses. The fluorescence is monitored continuously, which allows for easy estimation of the quality of data and helps to filter out potential artifacts (e.g., arising from compound precipitation or thermal changes).

A nicotinamide adenine dinucleotide (NADH)-coupled ATPase assay has been adapted to semihigh throughput screening of small molecule myosin inhibitors. This kinetic assay is run in a 384-well microplate format with total reaction volumes of only 20 &#181;L per well. The platform should be applicable to virtually any ADP producing enzyme.

ATPase enzymes utilize the free energy stored in adenosine triphosphate to catalyze a wide variety of endergonic biochemical processes in vivo that would ...

High-Throughput Screening for Protein Crystallization

High-Throughput Screening to Obtain Crystal Hits for Protein Crystallography

X-ray crystallography is the most commonly employed technique to discern macromolecular structures, but the crucial step of crystallizing a protein into an ordered lattice amenable to diffraction remains challenging. The crystallization of biomolecules is largely experimentally defined, and this process can be labor-intensive and prohibitive to researchers at resource-limited institutions. At the National High-Throughput Crystallization (HTX) Center, highly reproducible methods have been implemented to facilitate crystal growth, including an automated high-throughput 1,536-well microbatch-under-oil plate setup designed to sample a wide breadth of crystallization parameters. Plates are monitored using state-of-the-art imaging modalities over the course of 6 weeks to provide insight into crystal growth, as well as to accurately distinguish valuable crystal hits. Furthermore, the implementation of a trained artificial intelligence scoring algorithm for identifying crystal hits, coupled with an open-source, user-friendly interface for viewing experimental images, streamlines the process of analyzing crystal growth images. Here, the key procedures and instrumentation are described for the preparation of the cocktails and crystallization plates, imaging the plates, and identifying hits in a way that ensures reproducibility and increases the likelihood of successful crystallization.

Author Spotlight: High-Throughput Screening to Obtain Crystal Hits for Protein Crystallography  

This protocol details high-throughput crystallization screening, ranging from the 1,536 microassay plate preparation to the end of a 6 week experimental time window. Details are included about the sample setup, the imaging obtained, and how users can perform analyses using an artificial intelligence-enabled graphical user interface to quickly and efficiently identify macromolecular crystallization conditions.

X-ray crystallography is the most commonly employed technique to discern macromolecular structures, but the crucial step of crystallizing a protein into ...

Isolating Small Extracellular Vesicles and Extracting Their Peptidome

Identification of Peptides of Small Extracellular Vesicles from Bone Marrow-Derived Macrophages

Small extracellular vesicles (sEVs) are typically secreted by the exocytosis of multivesicular bodies (MVBs). These nanovesicles with a diameter of &lt;200 nm are present in various body fluids. These sEVs regulate various biological processes such as gene transcription and translation, cell proliferation and survival, immunity and inflammation through their cargos, such as proteins, DNA, RNA, and metabolites. Currently, various techniques have been developed for sEVs isolation. Among them, the ultracentrifugation-based method is considered the gold standard and is widely used for sEVs isolation. The peptides are naturally biomacromolecules with less than 50 amino acids in length. These peptides participate in a variety of biological processes with biological activity, such as hormones, neurotransmitters, and cell growth factors. The peptidome is intended to systematically analyze endogenous peptides in specific biological samples by liquid chromatography-tandem mass spectrometry (LC-MS/MS). Here, we introduced a protocol to isolate sEVs by differential ultracentrifugation and extracted peptidome for identification by LC-MS/MS. This method identified hundreds of sEVs-derived peptides from bone marrow-derived macrophages.

Author Spotlight: Peptidome Extraction from Small Extracellular Vesicles Isolated from Bone Marrow-Derived Macrophages 

This protocol describes a procedure to isolate small extracellular vesicles from macrophages by differential ultracentrifugation and extract the peptidome for identification by mass spectrometry.

Small extracellular vesicles (sEVs) are typically secreted by the exocytosis of multivesicular bodies (MVBs). These nanovesicles with a diameter of ...

Strategies for Expressing and Purifying Solute Carrier Transporters

High-Throughput Expression and Purification of Human Solute Carriers for Structural and Biochemical Studies

Solute carriers (SLCs) are membrane transporters that import and export a range of endogenous and exogenous substrates, including ions, nutrients, metabolites, neurotransmitters, and pharmaceuticals. Despite having emerged as attractive therapeutic targets and markers of disease, this group of proteins is still relatively underdrugged by current pharmaceuticals. Drug discovery projects for these transporters are impeded by limited structural, functional, and physiological knowledge, ultimately due to the difficulties in the expression and purification of this class of membrane-embedded proteins. Here, we demonstrate methods to obtain high-purity, milligram quantities of human SLC transporter proteins using&#160;codon-optimized gene sequences. In conjunction with a systematic exploration of construct design and high-throughput expression, these protocols ensure the preservation of the structural integrity and biochemical activity of the target proteins. We also highlight critical steps in the eukaryotic cell expression, affinity purification, and size-exclusion chromatography of these proteins. Ultimately, this workflow yields pure, functionally active, and stable protein preparations suitable for high-resolution structure determination, transport studies, small-molecule engagement assays, and high-throughput in vitro screening.

Author Spotlight: Expression and Purification of Human Solute Carrier Transporters Using Codon-Optimized Genes 

Structural and biochemical studies of human membrane transporters require milligram quantities of stable, intact, and homogeneous protein. Here we describe scalable methods to screen, express, and purify human solute carrier transporters using codon-optimized genes.

Solute carriers (SLCs) are membrane transporters that import and export a range of endogenous and exogenous substrates, including ions, nutrients, ...

Enhancing EV Isolation for Precise Biofluid Analysis

Chemical Affinity-Based Isolation of Extracellular Vesicles from Biofluids for Proteomics and Phosphoproteomics Analysis

Extracellular vesicles (EVs) from biofluids have recently gained significant attention in the field of liquid biopsy. Released by almost every type of cell, they provide a real-time snapshot of host cells and contain a wealth of molecular information, including proteins, in particular those with post-translational modifications (PTMs) such as phosphorylation, as the main player of cellular functions and disease onset and progression. However, the isolation of EVs from biofluids remains challenging due to low yields and impurities from current EV isolation methods, making the downstream analysis of EV cargo, such as EV phosphoproteins, difficult. Here, we describe a rapid and effective EV isolation method based on functionalized magnetic beads for EV isolation from biofluids such as human urine and downstream proteomics and phosphoproteomics analysis following EV isolation. The protocol enabled a high recovery yield of urinary EVs and sensitive profiles of EV proteome and phosphoproteome. Furthermore, the versatility of this protocol and relevant technical considerations are also addressed here.

Author Spotlight: Advancing EVtrap for High-Throughput Proteomics in Disease Biomarker Discovery 

The present protocol provides detailed descriptions for the efficient isolation of urinary extracellular vesicles utilizing functionalized magnetic beads. Moreover, it encompasses subsequent analyses, including western blotting, proteomics, and phosphoproteomics.

Extracellular vesicles (EVs) from biofluids have recently gained significant attention in the field of liquid biopsy. Released by almost every type of ...

Investigating Vitamin A Receptor Kinetics and Function for Retinoid Homeostasis

Methodology for Studying Interactions of Vitamin A Membrane Receptors and Opsin Protein with their Ligands in Generating the Retinylidene Protein

Distribution of dietary vitamin A/all-trans retinol (ROL) throughout the body is critical for maintaining retinoid function in peripheral tissues and generating the retinylidene protein for visual function. RBP4-ROL is the complex of ROL with retinol-binding protein 4 (RBP4), which is present in the blood. Two membrane receptors, Retinol Binding Protein 4 Receptor 2 (RBPR2) in the liver and STimulated by Retinoic Acid 6 Retinol (STRA6) in the eye, bind circulatory RBP4 and this mechanism is critical for internalizing ROL into cells. Establishing methods to investigate receptor-ligand kinetics is essential in understanding the physiological function of vitamin A receptors for retinoid homeostasis. Using Surface Plasmon Resonance (SPR) assays, we can analyze the binding affinities and kinetic parameters of vitamin A membrane receptors with its physiological ligand RBP4. 
These methodologies can reveal new structural and biochemical information of RBP4-binding motifs in RBPR2 and STRA6, which are critical for understanding pathological states of vitamin A deficiency. In the eye, internalized ROL is metabolized to 11-cis retinal, the visual chromophore that binds to opsin in photoreceptors to form the retinylidene protein, rhodopsin. The absorbance of light causes the cis-to-trans isomerization of 11-cis retinal, inducing conformational changes in rhodopsin and the subsequent activation of the phototransduction cascade. Decreased concentrations of serum and ocular ROL can impact retinylidene protein formation, which in turn can cause rhodopsin mislocalization, apoprotein opsin accumulation, night blindness, and photoreceptor outer segment degeneration, leading to Retinitis Pigmentosa or Leber Congenital Amaurosis. 
Therefore, spectrophotometric methodologies to quantify the G protein-coupled receptor opsin-11-cis retinal complex in the retina are critical for understanding mechanisms of retinal cell degeneration in the above-mentioned pathological states. With these comprehensive methodologies, investigators will be able to better assess dietary vitamin A supply in maintaining systemic and ocular retinoid homeostasis, which is critical for generating and maintaining retinylidene protein concentrations in photoreceptors, which is critical for sustaining visual function in humans.

Author Spotlight: Unraveling Vitamin A Transport Mechanisms &#8212; Linking Liver Receptors to Vision Health Through RBPR2 and RBP4 Interactions

Here, we describe two quantitative methods for studying the protein-ligand interactions of vitamin A membrane receptors and photoreceptor opsin with their respective physiological ligands.

Distribution of dietary vitamin A/all-trans retinol (ROL) throughout the body is critical for maintaining retinoid function in peripheral tissues and ...

Methods Article

Optimization of Flow Cytometric Sorting Parameters for High-Throughput Isolation and Purification of Small Extracellular Vesicles