We would like to acknowledge all the previous members of the Bouchier-Hayes lab who contributed to the development of this technique. This work was funded in part by a Texas Children&#39;s Hospital Pediatric Pilot award to LBH. We thank Joya Chandra (MD Anderson, Houston, Texas) for permission to include data published in collaboration with her team. The development of the reagents described was supported by the Cytometry and Cell Sorting Core at Baylor College of Medicine with funding from the NIH (NIAID P30AI036211, NCI P30CA125123, and NCRR S10RR024574) and the assistance of Joel M. Sederstrom

1. Preparation of cells and culture dishes
NOTE: Perform Steps 1-3 in a tissue culture laminar flow hood. Wear gloves.
<ol>
	<li>If using glass bottom dishes, coat the glass with fibronectin. Skip to Step 1.2 if using plastic dishes.
	<ol>
		<li>Make a 0.1 mg/mL solution of fibronectin: dilute 1 mL of 1 mg/mL fibronectin solution in 9 mL of 1 X PBS.</li>
		<li>Cover the glass bottom of each well with 0.5-1 mL of fibronectin and incubate for 1-5 min at room temperature.</li>
		<li>Using a pipette, collect the fibronectin solution and store at 4 &#176;C. 
		NOTE: The fibronectin solution can be reused multiple times.</li>
		<li>Wash the glass once with 1-2 mL 1 X PBS, and aspirate off PBS.</li>
	</ol>
	</li>
	<li>Plate 1 x 105 cells per well of a 6-well plate in 2 mL of the appropriate cell culture media (see Table 1 for cell numbers to use for different sized wells). If using a cell line that stably expresses the caspase BiFC components (see Discussion), plate 2 x 105 cells and proceed to the treatment step (Step 3).</li>
	<li>Allow cells to adhere to the dish overnight at 37 &#176;C in a tissue culture incubator.</li>
</ol>
2. Transfection of cells to introduce the caspase BiFC components
<ol>
	<li>Transfect cells with the appropriate transfection reagent. 
	NOTE: This protocol outlines the instructions for Lipofectamine 2000, which has been optimized for minimal toxicity. This protocol has been successfully used in Hela cells, MCF7 cells and LN18 cells. If using additional transfection reagents follow the manufacturer&#39;s instructions and optimize as needed.
	<ol>
		<li>Add 12 &#181;L of the transfection reagent to 750 &#181;L of reduced serum media in a sterile tube.</li>
		<li>Incubate at room temperature for 5 min.</li>
		<li>Add 10 ng of a reporter plasmid (a plasmid encoding a fluorescent protein, e.g. red fluorescent protein [RFP], that will label the transfected cells) to a sterile 1.5 mL tube for each well to be transfected.</li>
		<li>Add 20 ng of each caspase BiFC plasmid (e.g., 20 ng of caspase-2 prodomain [C2 Pro]-VC and 20 ng of C2 Pro-VN) to each tube (Table 2).</li>
		<li>Bring each tube up to a total volume of 100 &#181;L by adding reduced serum media.</li>
		<li>Using a p200 pipette, add 100 &#181;L of transfection reagent solution from Step 2.1.2 to the plasmid mixture in a dropwise manner.</li>
		<li>Incubate the plasmid-transfection reagent mix for 20 min at room temperature.</li>
		<li>Aspirate the media from the cells using a pipette or with suction and then, using a p1000 pipette, pipette 800 &#181;L of serum free media gently down the side of the well.</li>
		<li>Using a p200 pipette, add 200 &#181;L of the DNA-lipid complex dropwise to each well.</li>
		<li>Incubate the cells in a tissue culture incubator at 37 &#176;C for 3 h.</li>
		<li>Taking care not to disrupt the monolayer, remove the serum free media containing the DNA-lipid complexes by aspiration using suction or with a pipette.</li>
		<li>Pipette 2 mL of pre-warmed (37 &#176;C) complete growth media gently down the side of each well.</li>
	</ol>
	</li>
	<li>Incubate the cells at 37 &#176;C in a tissue culture incubator for 24-48 h for optimal protein expression.</li>
</ol>
3. Induction of caspase activation
<ol>
	<li>Treat with a drug or stimulus that induces caspase activation approximately 24 h post-transfection.
	<ol>
		<li>Add the desired concentration of the drug to pre-warmed (37 &#176;C) imaging media (complete growth media supplemented with Hepes [20 mM, pH 7.2-7.5] and 2-mercaptoethanol [55 &#181;M]) and mix gently (Table 3).</li>
		<li>Aspirate the media from the cells using a pipette or with suction and then, using a pipette, pipette 2 mL of the solution from 3.1.1 gently down the side of the well.</li>
		<li>Add imaging media without the drug to one well as an untreated control.</li>
	</ol>
	</li>
	<li>Incubate the cells in a tissue culture incubator at 37 &#176;C or proceed to Step 4.3 for a time-lapse experiment.</li>
</ol>
4. Caspase BiFC data acquisition
<ol>
	<li>Caspase BiFC for single time point acquisition
	<ol>
		<li>Turn on the microscope and fluorescent light source following the manufacturer&#39;s instructions.</li>
		<li>Find the cells under the RFP filter and focus on the reporter gene fluorescence.</li>
		<li>Using a hand-tally counter, count all of the RFP-positive cells in the visual field. Record the number.</li>
		<li>While maintaining the same visual field, switch to the GFP filter (or YFP filter if available) and, using a hand-tally counter, count the number of red cells that are also green (Venus-positive). Switch back and forth between the two filters to ensure that only the red cells are evaluated for Venus-positivity. Record the number (Figure 3).</li>
		<li>Count at least 100 RFP-positive cells from each of three individual areas of the plate.</li>
		<li>In each area, calculate the percentage of Venus-positive transfected cells (i.e. red cells that are also green). Average the resulting percentages to get the standard deviation.</li>
	</ol>
	</li>
	<li>Three-dimensional imaging of caspase BiFC
	<ol>
		<li>Turn on the microscope following the manufacturer&#39;s instructions and launch the acquisition software.</li>
		<li>Select the 60x or 63x oil objective and place a drop of oil on the objective</li>
		<li>Place the culture dish on the microscope stage using the correct plate holder.</li>
		<li>Using either an epifluorescence light source and the microscope eye piece or the confocal lasers and the computer screen, navigate to a field of interest.</li>
		<li>Focus on the reporter fluorescence using the RFP laser or filter.</li>
		<li>If epifluorescence has been used up to this point, switch the light source to the confocal lasers and visualize the live image of the cells as acquired by the camera and displayed by the acquisition software on the computer screen.</li>
		<li>Using the joystick control and the focus wheel, fine tune the focus and position of the cells so that the cells are in the center of the view finder. Choose cells that are separated from each other and are not overlapping.</li>
		<li>Input the percentage laser power to 50% and the exposure time at 100 ms for both Venus and RFP as a starting point to determine the optimal settings to use for the experiment.</li>
		<li>Turn on the live capture and inspect the resulting image and the accompanying display histograms for both channels.</li>
		<li>If the signal is low and the image is hard to make out, increase the percentage laser power and/or exposure time in increments until the signal in the image looks good.</li>
		<li>Ensure that the signal does not reach saturation. Inspect the display histogram to make sure a distinct peak is visible for each fluor. If the peak encompasses the entire histogram display, input lower laser power and exposure settings (Figure 2).</li>
		<li>Visualize control (untreated or non-transfected) cells under the same conditions to ensure that the signal is specific. 
		NOTE: Single color transfectants can also be used to control for crossover, but if the signals are spatially distinct (e.g. mitochondrial versus cytosolic) this is not necessary.</li>
		<li>Select or open the Z stack module in the microscope software.</li>
		<li>Using the focus knob, approximate the focal position corresponding to the center of the cell.</li>
		<li>Adjust the focus in one direction until the cell is no longer visible, select this as the top position.</li>
		<li>Adjust the focus in the opposite direction until the cell is again no longer visible and select this as the bottom position.</li>
		<li>Choose the optimal step size (usually approximately 20 &#181;m) and start the acquisition to instruct the camera to automatically take a single image in each channel at each of the 20 &#181;m steps between the top and bottom positions.</li>
		<li>Use 3D rendering software to reconstruct the 3D image (Figure 4, Movie 1).</li>
	</ol>
	</li>
	<li>Time-lapse imaging of caspase BiFC
	<ol>
		<li>At least 1 h prior to the experiment, turn on the microscope and set the temperature to 37 &#176;C.</li>
		<li>Select the 40x, 60x, or 63x oil objective and place a drop of oil on the objective.</li>
		<li>Place the culture dish on the microscope stage using the correct plate holder.</li>
		<li>If a CO2 source is available, place the CO2 delivery device (usually a plexiglass lid attached to the tube that delivers the CO2) on top of the plate holder. Set the CO2 level to 5% and turn on the CO2 controller from within the software. If a CO2 source is not available, include 20 mM Hepes to buffer the media.</li>
		<li>Navigate to a field of transfected cells and visualize the live image of the cells as acquired by the camera and displayed by the acquisition software on the computer screen using the RFP laser.</li>
		<li>Following Steps 4.2.8-4.2.12, input the settings for percentage laser power and exposure time for the RFP laser. Keep these values as low as possible while still being able to detect the fluorescent signal.</li>
		<li>Using a positive control sample (see Table 3 for examples), follow Steps 4.2.8-4.2.12 to input the settings for percentage laser power and exposure time for the YFP laser light. Keep these values as low as possible while still being able to detect Venus signal.</li>
		<li>For each well of the plate, choose a number of different positions that contain one or more cells expressing the reporter.</li>
		<li>Input the time interval between each frame of the time-lapse and total number of frames to be taken. Ensure there is time to acquire all the selected positions before the next frame is to be taken.</li>
		<li>Re-visit each position and correct and update the focus as needed.</li>
		<li>To correct for focal drift that can result from changes in temperature or external vibrations, turn on the focus drift correction system (if available).</li>
		<li>Begin the time-lapse and save the data.</li>
		<li>Analyze the data using available imaging software (Figure 5, Movie 2).</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

This protocol discusses the use of split fluorescent proteins to measure caspase induced proximity. Split Venus was chosen for this technique because it is very bright, highly photostable and the refolding is fast 13. Thus, the analysis of Venus refolding upon caspase induced proximity can provide close to real-time estimations of caspase protein interaction dynamics. Venus is split into two slightly overlapping fragments, the N terminus of Venus (Venus-N or VN) comprising amino acids 1-173 and th...

The caspase protease family are known for their critical roles in apoptosis and innate immunity 1. Because of their importance, determining when, where, and how efficiently specific caspases are activated can provide crucial insights into the mechanisms of caspase activation pathways. The imaging based protocol described here enables the visualization of the earliest steps in the caspase activation cascade. This technique takes advantage of the dynamic protein:protein interactions that drive caspase activation.
The caspases can be divided into two groups: the initiator caspases (caspase-1, -2, -4, -5, -8, -9, -10, and -12) and the executioner caspases (caspase-3, -6, and -7). The executioner caspases are present in the cell as preformed dimers and are activated by cleavage between the large and small subunit 2. When activated, they cleave numerous structural and regulatory proteins resulting in apoptosis 3. The initiator caspases are the first caspases to be activated in a pathway and generally trigger the activation of executioner caspases. In contrast to executioner caspases, initiator caspases are activated by dimerization 4,5. This dimerization is facilitated by recruitment of inactive monomers to specific large molecular weight complexes known as activation platforms. Assembly of activation platforms is governed by a series of specific protein:protein interactions. These are mediated by conserved protein interaction motifs present in the proform of the initiator caspase and include the death domain (DD), death effector domain (DED) and caspase recruitment domain (CARD) 6 (Figure 1A). Activation platforms generally include a receptor protein and an adaptor protein. The receptor is usually activated upon binding of a ligand, inducing a conformational change that allows for the oligomerization of numerous molecules. The receptor then either recruits the caspase directly or adaptor molecules that in turn bring the caspase to the complex. Thus, numerous caspase molecules come into close proximity permitting dimerization. This is known as the induced proximity model 7. Once dimerized, the caspase undergoes autoprocessing, which serves to stabilize the active enzyme 4,8. For example, assembly of the Apaf1 apoptosome is triggered by cytochrome c following its release from the mitochondria in a process called mitochondrial outer membrane permeabilization (MOMP). Apaf1 in turn recruits caspase-9 through an interaction that is mediated by a CARD present in both proteins 9. Similar protein interactions result in the assembly of the CD95 death inducing signaling complex (DISC) that leads to caspase-8 activation; the PIDDosome, which can activate caspase-2; and various inflammasome complexes that initiate caspase-1 activation 10,11,12. Thus, initiator caspases are recruited to specific activation platforms by a common mechanism resulting in induced proximity and dimerization, without which activation will not occur.
Caspase Bimolecular Fluorescence Complementation (BiFC) is an imaging-based assay that was developed to measure this first step in the activation of initiator caspases, allowing direct visualization of caspase induced proximity following activation platform assembly. This method takes advantage of the properties of the split fluorescent protein Venus. Venus is a brighter and more photostable version of yellow fluorescent protein (YFP) that can be separated into two non-fluorescent and slightly overlapping fragments: the N-terminus of Venus (Venus N or VN) and the C-terminus of Venus (Venus C or VC). These fragments retain the ability to refold and become fluorescent when in close proximity 13. Each Venus fragment is fused to the prodomain of the caspase, which is the minimal portion of the caspase that binds to the activation platform. This ensures that the caspases do not retain enzymatic activity and therefore concurrent analysis of downstream events associated with endogenous events is possible. The Venus fragments refold when the caspase prodomains are recruited to the activation platform and undergo induced proximity. (Figure 1B). The resulting Venus fluorescence can be used to accurately and specifically monitor the subcellular localization, the kinetics, and the efficiency of assembly of initiator caspase activation platforms in single cells. Imaging data can be acquired by confocal microscopy or by standard fluorescence microscopy and can be adapted to a number of different microscopy approaches including: time-lapse imaging to track caspase activation in real time; high resolution imaging for precise determinations of subcellular localization; and end point quantification of efficiency of activation.
This technique was first developed to investigate the activation of caspase-214. While the activation platform for caspase-2 is thought to be the PIDDosome, consisting of the receptor PIDD (p53-induced protein with a death domain) and the adaptor RAIDD (RIP-associated ICH-1/CAD-3 homologous protein with a death domain), PIDDosome-independent caspase-2 activation has been reported. This suggests that additional activation platforms for caspase-2 exist 15,16. Despite not knowing the full components of the caspase-2 activation platform, the caspase BiFC technique has allowed for successful interrogation of the caspase-2 signaling pathways at the molecular level 14,17. We have also successfully adapted this protocol for the inflammatory caspases (caspase-1, -4, -5 and -12) 18 and, in principle, the same approach should be sufficient to similarly analyze each of the remaining initiator caspases. This protocol could similarly be adapted to investigate other pathways were dimerization is a major activating signal. For example, STAT proteins are activated by dimerization following phosphorylation by Janus kinase (JAK) 19. Thus, the BiFC system could be used to visualize for STAT activation as well as many other pathways regulated by dynamic protein interactions. The following protocol provides step-by-step instructions for the introduction of the reporters into cells as well as methodologies for image acquisition and analysis.

<table>
	<tbody>
		<tr>
			<td>6-well Uncoated No. 1.5 20 mm glass bottom dishes</td>
			<td>Mattek</td>
			<td>P06G-1.5-20-F</td>
			<td></td>
		</tr>
		<tr>
			<td>Human Plasma Fibronectin Purified Protein</td>
			<td>Millipore</td>
			<td>FC010-10MG</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS</td>
			<td>Sigma</td>
			<td>D8537-6x500ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine 2000 reagent</td>
			<td>Invitrogen</td>
			<td>11668019</td>
			<td></td>
		</tr>
		<tr>
			<td>OPTI MEM I</td>
			<td>Invitrogen</td>
			<td>31985088</td>
			<td></td>
		</tr>
		<tr>
			<td>C2-Pro VC plasmid</td>
			<td>Addgene</td>
			<td>49261</td>
			<td></td>
		</tr>
		<tr>
			<td>C2-Pro VN plasmid</td>
			<td>Addgene</td>
			<td>49262</td>
			<td></td>
		</tr>
		<tr>
			<td>Inflammatory caspase BiFC plasmids</td>
			<td></td>
			<td></td>
			<td>available by request from LBH</td>
		</tr>
		<tr>
			<td>HeLa cells stably expressing the C2-Pro BiFC components</td>
			<td></td>
			<td></td>
			<td>available by request from LBH</td>
		</tr>
		<tr>
			<td>DsRed mito plasmid</td>
			<td>Clontech</td>
			<td>632421</td>
			<td>similar plasmids that can be used as fluorescent reporters can be found on Addgene</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Invitrogen</td>
			<td>15630106</td>
			<td></td>
		</tr>
		<tr>
			<td>2 Mercaptoethanol 1000X</td>
			<td>Invitrogen</td>
			<td>21985023</td>
			<td></td>
		</tr>
		<tr>
			<td>q-VD-OPH</td>
			<td>Apex Bio</td>
			<td>A1901</td>
			<td></td>
		</tr>
	</tbody>
</table>

lighting up the pathways to caspase activation using bimolecular fluorescence complementation

The caspase family of proteases play essential roles in apoptosis and innate immunity. Among these, a subgroup known as initiator caspases are the first to be activated in these pathways. This group includes caspase-2, -8, and -9, as well as the inflammatory caspases, caspase-1, -4, and -5. The initiator caspases are all activated by dimerization following recruitment to specific multiprotein complexes called activation platforms. Caspase Bimolecular Fluorescence Complementation (BiFC) is an imaging-based approach where split fluorescent proteins fused to initiator caspases are used to visualize the recruitment of initiator caspases to their activation platforms and the resulting induced proximity. This fluorescence provides a readout of one of the earliest steps required for initiator caspase activation. Using a number of different microscopy-based approaches, this technique can provide quantitative data on the efficiency of caspase activation on a population level as well as the kinetics of caspase activation and the size and number of caspase activating complexes on a per cell basis.

An example of caspase-2 BiFC induced by DNA damage is shown in Figure 3. Camptothecin, a topoisomerase I inhibitor, was used to induce DNA damage and caspase-2 activation. The red fluorescent protein mCherry was used as a reporter to show that the cells express the BiFC probe and to help visualize the total number of cells. Venus fluorescence is shown in green and the large puncta represent caspase-2 induced proximity. These cells can be counted to determine ...

Watch this Scientific Journal Video about Lighting Up the Pathways to Caspase Activation Using Bimolecular Fluorescence Complementation at JoVE.com

Lighting Up the Pathways to Caspase Activation Using Bimolecular Fluorescence Complementation

This protocol describes caspase Bimolecular Fluorescence Complementation (BiFC); an imaging-based method that can be used to visualize induced proximity of initiator caspases, which is the first step in their activation.

The caspase family of proteases play essential roles in apoptosis and innate immunity. Among these, a subgroup known as initiator caspases are the first ...

lighting-up-pathways-to-caspase-activation-using-bimolecular

Universidad de Cartagena UDEC

Research

JoVE Journal

Biochemistry

9.0K Views.  Baylor College of Medicine. This protocol describes caspase Bimolecular Fluorescence Complementation (BiFC); an imaging-based method that can be used to visualize induced proximity of initiator caspases, which is the first step in their activation.

Video: Lighting Up the Pathways to Caspase Activation Using Bimolecular Fluorescence Complementation

Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

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

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

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

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological functions. Such interactions can be highly dynamic, meaning the interacting subunits are constantly associated and dissociated at certain rates. While measuring the binding affinity using techniques such as quantitative pull-down reveals the strength of the interaction, studying the binding kinetics provides insights on how fast the interaction occurs and how long each complex can exist. Furthermore, measuring the kinetics of an interaction in the presence of an additional factor, such as a protein exchange factor or a drug, helps reveal the mechanism by which the interaction is regulated by the other factor, providing important knowledge for the advancement of biological and medical research. Here, we describe a protocol for measuring the binding kinetics of a protein complex that has a high intrinsic association rate and can be dissociated quickly by another protein. The method uses fluorescence resonance energy transfer to report the formation of the protein complex in vitro, and it enables monitoring the fast association and dissociation of the complex in real time on a stopped-flow fluorimeter. Using this assay, the association and dissociation rate constants of the protein complex are quantified.

Protein-protein interactions are critical for biological systems, and studies of the binding kinetics provide insights into the dynamics and function of protein complexes. We describe a method that quantifies the kinetic parameters of a protein complex using fluorescence resonance energy transfer and the stopped-flow technique. &#160;

Proteins are the primary operators of biological systems, and they usually interact with other macro- or small molecules to carry out their biological ...

High-Throughput Total Internal Reflection Fluorescence and Direct Stochastic Optical Reconstruction Microscopy Using a Photonic Chip

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced contrast due to optical sectioning. The conventional approach is to use high numerical aperture microscope TIRF objectives for both excitation and collection, severely limiting the field of view and throughput. We present a novel approach to generating TIRF excitation for imaging with optical waveguides, called chip-based nanoscopy. The aim of this protocol is to demonstrate how chip-based imaging is performed in an already built setup. The main advantage of chip-based nanoscopy is that the excitation and collection pathways are decoupled. Imaging can then be done with a low magnification lens, resulting in large field of view TIRF images, at the price of a small reduction in resolution. Liver sinusoidal endothelial cells (LSECs) were imaged using direct stochastic optical reconstruction microscopy (dSTORM), showing a resolution comparable to traditional super-resolution microscopes. In addition, we demonstrate the high-throughput capabilities by imaging a 500 &#181;m x 500 &#181;m region with a low magnification lens, providing a resolution of 76 nm. Through its compact character, chip-based imaging can be retrofitted into most common microscopes and can be combined with other on-chip optical techniques, such as on-chip sensing, spectroscopy, optical trapping, etc. The technique is thus ideally suited for high throughput 2D super-resolution imaging, but also offers great opportunities for multi-modal analysis.

Chip-based super-resolution optical microscopy is a novel approach to fluorescence microscopy and offers advantages in cost effectiveness and throughput. Here, the protocols for chip preparation and imaging are shown for TIRF microscopy and localization-based super-resolution microscopy.

Total internal reflection fluorescence (TIRF) is commonly used in single molecule localization based super-resolution microscopy as it gives enhanced ...

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Amyloid fibrils are associated with a number of neurodegenerative diseases such as Huntington&#39;s, Parkinson&#39;s, or Alzheimer&#39;s disease. These amyloid fibrils can sequester endogenous metastable proteins as well as components of the proteostasis network (PN) and thereby exacerbate protein misfolding in the cell. There are a limited number of tools available to assess the aggregation process of amyloid proteins within an animal. We present a protocol for fluorescence lifetime microscopy (FLIM) that allows monitoring as well as quantification of the amyloid fibrilization in specific cells, such as neurons, in a noninvasive manner and with the progression of aging and upon perturbation of the PN. FLIM is independent of the expression levels of the fluorophore and enables an analysis of the aggregation process without any further staining or bleaching. Fluorophores are quenched when they are in close vicinity of amyloid structures, which results in a decrease of the fluorescence lifetime. The quenching directly correlates with the aggregation of the amyloid protein. FLIM is a versatile technique that can be applied to compare the fibrilization process of different amyloid proteins, environmental stimuli, or genetic backgrounds in vivo in a non-invasive manner.

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Fluorescence lifetime imaging monitors, quantifies and distinguishes the aggregation tendencies of proteins in living, aging, and stressed C. elegans disease models.

Amyloid fibrils are associated with a number of neurodegenerative diseases such as Huntington&#39;s, Parkinson&#39;s, or Alzheimer&#39;s disease. These ...

Complementation of Splicing Activity by a Galectin-3 - U1 snRNP Complex on Beads

Classic depletion-reconstitution experiments indicate that galectin-3 is a required splicing factor in nuclear extracts. The mechanism of incorporation of galectin-3 into the splicing pathway is addressed in this paper. Sedimentation of HeLa cell nuclear extracts on 12%-32% glycerol gradients yields fractions enriched in an endogenous ~10S particle that contains galectin-3 and U1 snRNP. We now describe a protocol to deplete nuclear extracts of U1 snRNP with concomitant loss of splicing activity. Splicing activity in the U1-depleted extract can be reconstituted by the galectin-3 - U1 snRNP particle trapped on agarose beads covalently coupled with anti-galectin-3 antibodies. The results indicate that the galectin-3 - U1 snRNP - pre-mRNA ternary complex is a functional E complex leading to intermediates and products of the splicing reaction and that galectin-3 enters the splicing pathway through its association with U1 snRNP. The scheme of using complexes affinity- or immuno-selected on beads to reconstitute splicing activity in extracts depleted of a specific splicing factor may be generally applicable to other systems.

This article describes the experimental procedures for (a) depletion of U1 snRNP from nuclear extracts, with concomitant loss of splicing activity; and (b) reconstitution of splicing activity in the U1-depleted extract by galectin-3 - U1 snRNP particles bound to beads covalently coupled with anti-galectin-3 antibodies.

Classic depletion-reconstitution experiments indicate that galectin-3 is a required splicing factor in nuclear extracts. The mechanism of incorporation of ...

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One &#945;-Synuclein Monomer at a Time

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural dynamics and its contributions to biology. While the FRET-based ruler reports on inter-dye distances in the 3-10 nm range, other spectroscopic techniques, such as protein-induced fluorescence enhancement (PIFE), report on the proximity between a dye and a protein surface in the shorter 0-3 nm range. Regardless of the method of choice, its use in measuring freely-diffusing biomolecules one at a time retrieves histograms of the experimental parameter yielding separate centrally-distributed sub-populations of biomolecules, where each sub-population represents either a single conformation that stayed unchanged within milliseconds, or multiple conformations that interconvert much faster than milliseconds, and hence an averaged-out sub-population. In single-molecule FRET, where the reported parameter in histograms is the inter-dye FRET efficiency, an intrinsically disordered protein, such as the &#945;-Synuclein monomer in buffer, was previously reported as exhibiting a single averaged-out sub-population of multiple conformations interconverting rapidly. While these past findings depend on the 3-10 nm range of the FRET-based ruler, we sought to put this protein to the test using single-molecule PIFE, where we track the fluorescence lifetime of site-specific sCy3-labeled &#945;-Synuclein proteins one at a time. Interestingly, using this shorter range spectroscopic proximity sensor, sCy3-labeled &#945;-Synuclein exhibits several lifetime sub-populations with distinctly different mean lifetimes that interconvert in 10-100 ms. These results show that while &#945;-Synuclein might be disordered globally, it nonetheless attains stable local structures. In summary, in this work we highlight the advantage of using different spectroscopic proximity sensors that track local or global structural changes one biomolecule at a time.

Time-resolved single-molecule protein-induced fluorescence enhancement is a useful fluorescence spectroscopic proximity sensor sensitive to local structural changes in proteins. Here we show it can be used to uncover stable local conformations in &#945;-Synuclein, which is otherwise known as globularly unstructured and unstable when measured using the longer range FRET ruler.

Using spectroscopic rulers to track multiple conformations of single biomolecules and their dynamics have revolutionized the understanding of structural ...

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

A Pipeline to Investigate the Structures and Signaling Pathways of Sphingosine 1-Phosphate Receptors

Lysophospholipids (LPLs) are bioactive lipids that include sphingosine 1-phosphate (S1P), lysophosphatidic acid, etc. S1P, a metabolic product of sphingolipids in the cell membrane, is one of the best-characterized LPLs that regulates a variety of cellular physiological responses via signaling pathways mediated by sphingosine 1-phosphate receptors (S1PRs). This implicated that the S1P-S1PRs signaling system is a remarkable potential therapeutic target for disorders, including multiple sclerosis (MS), autoimmune disorders, cancer, inflammation, and even COVID-19. S1PRs, a small subset of the class A G-protein coupled receptor (GPCR) family, are composed of five subtypes: S1PR1, S1PR2, S1PR3, S1PR4, and S1PR5. The lack of detailed structural information, however, impedes the drug discovery targeting S1PRs. Here, we applied the cryo-electron microscopy method to solve the structure of the S1P-S1PRs complex, and elucidated the mechanism of activation, selective drug recognition, and G-protein coupling by using cell-based functional assays. Other lysophospholipid receptors (LPLRs) and GPCRs can also be studied using this strategy.

S1P exerts its diverse physiological effects through the S1P receptors (S1PRs) subfamily. Here, a pipeline is described to expound on the structures and function of S1PRs.

Lysophospholipids (LPLs) are bioactive lipids that include sphingosine 1-phosphate (S1P), lysophosphatidic acid, etc. S1P, a metabolic product of ...

Measuring Caspase Activity Using a Fluorometric Assay or Flow Cytometry

The activation of cysteine proteases, known as caspases, remains an important process in multiple forms of cell death. Caspases are critical initiators and executioners of apoptosis, the most studied form of programmed cell death. Apoptosis occurs during developmental processes and is a necessary event in tissue homeostasis. Pyroptosis is another form of cell death that utilizes caspases and is a critical process in activating the immune system through the activation of the inflammasome, which results in the release of members of the interleukin-1 (IL-1) family. To assess caspase activity, target substrates can be assessed. However, sensitivity can be an issue when examining single cells or low-level activity. We demonstrate how a fluorogenic substrate can be used with a population-based assay or single-cell assay by flow cytometry. With proper controls, different amino acid sequences can be used to identify which caspases are active. Using these assays, the simultaneous loss of the inhibitors of apoptosis proteins upon tumor necrosis factor (TNF) stimulation has been identified, which primarily induces apoptosis in macrophages rather than other forms of cell death.

The present protocol describes two methods to measure caspase activity through a fluorogenic substrate using flow cytometry or a spectrofluorometer.

The activation of cysteine proteases, known as caspases, remains an important process in multiple forms of cell death. Caspases are critical initiators ...

Lighting Up the Pathways to Caspase Activation Using Bimolecular Fluorescence Complementation

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Fluorescence Anisotropy as a Tool to Study Protein-protein Interactions

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Using In Vitro Fluorescence Resonance Energy Transfer to Study the Dynamics Of Protein Complexes at a Millisecond Time Scale

High-Throughput Total Internal Reflection Fluorescence and Direct Stochastic Optical Reconstruction Microscopy Using a Photonic Chip

Characterization of Amyloid Structures in Aging C. Elegans Using Fluorescence Lifetime Imaging

Complementation of Splicing Activity by a Galectin-3 - U1 snRNP Complex on Beads

Utilizing Time-Resolved Protein-Induced Fluorescence Enhancement to Identify Stable Local Conformations One α-Synuclein Monomer at a Time

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

A Pipeline to Investigate the Structures and Signaling Pathways of Sphingosine 1-Phosphate Receptors

Measuring Caspase Activity Using a Fluorometric Assay or Flow Cytometry