Name: spatial and temporal control of t cell activation using a photoactivatable agonist
Uploaded: 2018-04-25T13:00:00.000+00:00
Duration: 7 min 48 s
Description: Watch this Scientific Journal Video about Spatial and Temporal Control of T Cell Activation Using a Photoactivatable Agonist at JoVE.com

We thank members of the Huse lab for advice and assistance. Supported by the US National Institutes of Health (R01-AI087644 to M.H. and P30-CA008748 to Memorial Sloan-Kettering Cancer Center).

1. Preparation of Stimulatory Glass Surfaces
<ol>
	<li>Coat eight-well chambered coverglass with biotinylated poly-L-lysine (Bio-PLL) diluted 1:500 in distilled, deionized water (ddH2O). Incubate for 30 min at room temperature (RT).</li>
	<li>Wash with H2O.</li>
	<li>Dry for 2 h at RT.</li>
	<li>Block Bio-PLL coated surfaces with blocking buffer (HEPES-buffered saline [10 mM HEPES pH 7.4, 150 mM NaCl], with 2% BSA) for 30 min at RT.
	<ol>
		<li>Dissolve 5 mg of poly-L-lysine hydrobromide in 1 mL of 10 mM NaPO4, pH 8.5.</li>
		<li>Add 125 &#956;mol (0.55 mg) of NHS-biotin (from a 100 mg/mL stock in DMSO) and check the pH of the reaction. If the pH is below 8.5, add 2 &#956;L of 4 N NaOH to raise it to between pH 8.5 and 9.5.</li>
		<li>Vortex for 30 min.</li>
		<li>Quench the reaction with 50 &#956;L of 100 mM glycine dissolved in 20 mM Tris pH 8.0.</li>
		<li>Spin at full power for 10 min and transfer the supernatant to a new tube. 
		NOTE: Bio-PLL quality is typically compared to previous preparations by coating serial two-fold dilutions of the material onto a 96-well ELISA plate and quantifying biotin content after incubation with alkaline phosphatase coupled streptavidin.</li>
	</ol>
	</li>
	<li>Remove blocking buffer. Do not allow wells to dry. Add streptavidin (100 &#181;g/mL in blocking buffer). Incubate for 1 h at 4 &#176;C.</li>
	<li>Wash in HBS. Fill and invert chamber slide wells 4 - 5 times, removing HBS from the wells. Do not allow wells to dry.</li>
	<li>Add the biotinylated pMHC ligands and adhesion molecules to the surface. 
	NOTE: MCC and OVA can be photocaged by adding ortho-nitrobenzyl based protecting groups (e.g., nitrophenylethyl (NPE) or nitroveratryloxycarbonyl (NVOC)) to the &#949;-amino group of key lysines (K12 in MCC, K7 in OVA). Photocaged MCC and OVA can be obtained from commercial vendors. After reconstitution in aqueous buffer, they are refolded into bacterially expressed I-Ek and H2-Kb using established approaches11,12.</li>
	<li>Validation of photoactivatable MCC or OVA peptide
	<ol>
		<li>Decage NVOC-protected peptides by UV irradiating with a handheld UV lamp (see Table of Materials) for 20 min at RT.</li>
		<li>Pulse 1.0 x 105 APCs with 1 &#181;M of nonstimulatory peptide (negative control, e.g. Hb), unirradiated photoactivatable peptide, irradiated photoactivatable peptide, and agonist peptide (positive control, e.g., MCC) for 1 h to overnight at 37 &#176;C.</li>
		<li>To stimulate T cells expressing the 5C.C7/2B4/AND TCR, use CH12 or CH27 B cells. To stimulate OT1 T cells, use RMA-s or EL4 thymoma cells.</li>
		<li>Mix the pulsed APCs with an equal number of T cells in 96-well round bottom plates at a final volume of 200 &#181;L. Incubate at 37 &#176;C for 12 - 24 h.</li>
		<li>Recover the supernatants from each well and analyze IL-2 by ELISA with streptavidin-horseradish peroxidase colorimetric detection. 
		NOTE: Confirmation of the caging status of all peptides by electrospray mass spectrometry prior to refolding into MHC complexes, is strongly recommended.</li>
	</ol>
	</li>
	<li>Perform the folding and purification of MHC so as to minimize exposure to light. For example, wrap folding reactions in aluminum foil and carry out gel filtration chromatography with the UV lamp off. 
	NOTE: It has been found that photoactivatable MCC- I-Ek and OVA- H2-Kb induce UV independent T cell activation at high density, possibly due to incomplete functional caging. Hence, the photoactivatable pMHC is diluted into a 10 - 30x excess of nonstimulatory pMHC (e.g. I-Ek containing the hemoglobin64-76 peptide (Hb)) during the immobilization step. This enhances the signal to noise ratio of the subsequent imaging experiment.</li>
	<li>To photoactivate CD4+ T cells, collectively use a mixture of biotinylated Hb-I-Ek (3 &#181;g/mL), biotinylated photoactivatable MCC- I-Ek (0.1 &#181;g/mL), and biotinylated antibody against the class I MHC H2-Kk (0.5 &#181;g/mL). The anti-H2-Kk antibody encourages 5C.C7/2B4/AND T cells, which express H2-Kk, to spread onto the glass surface without undergoing activation.</li>
	<li>For CD8+ T cells, use a mixture of biotinylated H2-Db bearing the peptide KAVYDFATL (1 &#181;g/mL), biotinylated photoactivatable OVA-H2-Kb (0.1 &#181;g/mL), and the extracellular domain of the adhesion molecule ICAM-1 (2 &#181;g/mL produced by insect cell culture13). The ICAM-1 encourages close contact formation by engaging the integrin LFA1 on the T cell surface.</li>
	<li>Apply all protein mixtures in blocking buffer, followed by incubation for 1 h at RT or at least 2 h at 4 &#176;C. 
	NOTE: The molecular density on surfaces of this kind to be ~8000 per &#181;m2 has been previously determined6. Given that photoactivatable pMHC represents ~1/30th of the biotinylated protein on the surface, its density will be ~267 molecules per &#181;m2, prior to decaging.</li>
	<li>Wash as in step 1.6 and leave in HBS until ready to use.</li>
	<li>Add 200,000 CD4+ or CD8+ T cells expressing the appropriate TCR into each well and allow cells to adhere at 37 &#176;C for 15 min. Once cells have attached to and spread on the surface, they are ready for photoactivation and imaging. 
	NOTE: Retroviral transduction of effector T cells with fluorescent imaging probes is described in detail elswhere6,7. Calcium signaling can also be studied using untransduced T cells loaded with the calcium sensitive dye Fluo-46. The ratiometric dye Fura-2 is not recommended because it requires excitation in the UV range, which also induces decaging of the photoactivatable pMHC.</li>
</ol>
2. Image Acquisition
<ol>
	<li>Use an inverted TIRF microscope outfitted with a UV compatible 150X objective lens for image acquisition. UV irradiate user-defined regions using a digital diaphragm system attached to a 100 W mercury lamp (HBO). Direct UV light from this lamp onto the sample using a 400 nm long pass mirror.</li>
	<li>Use image analysis software for localized photoactivation and time-lapse acquisition. In most experiments, monitor probes in both the green and red channels using 488 nm and 561 nm excitation lasers, respectively. Laser light is directed onto the sample using a dual-bandpass dichroic mirror that also transmits in the UV range (to enable decaging). Please see the Table of Materials and Figure 6 for additional information on microscope configuration.</li>
	<li>After mounting the chamber slide containing the T cells, adjust settings to obtain TIRF or epifluorescence illumination, as necessary. In live mode, select a field of cells that are expressing the fluorescent probe(s) of interest. Establish micron-scale regions for photoactivation beneath individual cells using software control.</li>
	<li>Begin time-lapse acquisition. Typically, 80 timepoints are acquired, with an interval of 5 s between each time point. This leaves more than enough time for sequential 488 nm and 563 nm exposures, in the case of dual color experiments.</li>
	<li>After 10 timepoints, photoactivate the selected regions by opening the digital diaphragm shutter for 1 - 1.5 s.</li>
	<li>After the time lapse is complete, select a new field of cells and repeat the process.</li>
</ol>
3. Data Analysis
NOTE: Localized photoactivation of immobilized ligands creates a well-defined, stationary stimulatory region that can be used for quantitative analysis of signaling responses and cytoskeletal remodeling events. Analysis protocols typically involve either quantifying fluorescence intensity within the irradiated region or using the irradiated region as a positional endpoint for distance measurements (e.g. for assessing polarization of the centrosome to the irradiated region). Both analysis protocols are described below. Various interactive image analysis programs can be used to make intensity and distance measurements, which can then be output for additional processing and analysis.
<ol>
	<li>Fluorescence intensity
	<ol>
		<li>Determine the fluorescence intensity (FI) of a background region outside of the cell (FIb). This will be used for background correction.
		<ol>
			<li>Draw a micron-sized square region outside of the cell and make a mask. To determine the fluorescence intensity, click on Analyze | Mask Statistics. Select Mean Fluorescence Intensity and export the values. 
			NOTE: Procedural details (e.g. 3.1.1.1) refer to Slidebook software (see Table of Materials). Implementation of this protocol using other software packages (e.g., FIJI) will be slightly different.</li>
		</ol>
		</li>
		<li>Determine the FI within the photoactivated region for each time point.
		<ol>
			<li>Select Mask to highlight the region that was photoactivated. To determine the fluorescence intensity, click on Analyze | Mask Statistics. Select Mean Fluorescence Intensity and export the values.</li>
		</ol>
		</li>
		<li>Subtract the background FI from the measured FI values within the region. Then, normalize the corrected FI measurements by dividing by the average, background corrected FI of the first 9 frames before photoactivation. &#916;F/F = ((FI-FIb)/mean(FI1-9)-FIb)). 
		NOTE: Graph &#916;F/F as a function of time.</li>
	</ol>
	</li>
	<li>Distance
	<ol>
		<li>Obtain the x and y coordinates of the center of the photoactivated region.
		<ol>
			<li>To determine the x and y coordinates of the center of the photoactivated region, select Mask to highlight the region that was photoactivated. Once the region of photoactivation is highlighted, select Analyze | Mask Statistics | Center of Area and export the values.</li>
		</ol>
		</li>
		<li>Determine the x and y coordinates for the fluorescent probe of interest for each time point. This is typically achieved through either manual or automated particle tracking.
		<ol>
			<li>To determine the x and y coordinates of the fluorescent probe of interest, select Manual Particle Tracking and click on the fluorescent probe of interest over time. Once all time points have been tracked, select Analyze | Mask Statistics | Center of Area and export the values.</li>
		</ol>
		</li>
		<li>Calculate the distance between the fluorescent protein of interest and the center of the photoactivated region for each time point using the equation: Distance = &#8730;((x2-x1)2+(y2-y1)2), in which x2 and y2 are the coordinates of the fluorescent protein of interest and x1 and y1 are the coordinates of the center of the photoactivated region.</li>
		<li>Graph the distance as a function of time.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

In recent years, light has emerged as an excellent tool for spatiotemporally controlled activation of cellular processes. Various methodologies have been developed, each with associated advantages and disadvantages. The system described here, which is based on the decaging of immobilized, extracellular ligands, is ideally suited for the analysis of rapid, subcellular, polarized signaling responses. This approach has been applied to examine IS formation in T cells as described above. Additionally, caged ligands for other receptors have been engineered, enabling us to apply the same strategy to assess inhibitory signaling in natural killing cells and nuclear translocation of the transcription factor NF-&#954;B in response to toll like receptor signaling14,15.
In this protocol, stimulatory ligands are immobilized to ensure that they remain as a stationary, polarized source of stimulation for the duration of the imaging experiment. This approach is well suited for studies of cell polarization. It should be noted however, that the TCR photoactivation approach is easily adapted for temporally controlled T cell activation on supported lipid bilayers. Obviously, one cannot achieve sustained, polarized stimulation in this context. This may be unnecessary, however, given the experimental objectives.
The system is also quite flexible with respect to the UV light source for decaging. Currently, a 100 W Hg lamp is used for focused illumination, however a pulsed nitrogen laser (pulse width 4 ns, pulse energy 120 mJ), was previously used6. At times, a handheld UV lamp (365 nm, 4 W) has been positioned over the imaging sample in order to decage larger regions of photostimulatable ligand. The specific features of the decaging system will in each case depend on the technical requirements of the experiment in question.
When implementing this system, it is important to confirm that the observed responses are specific for the photoactivation event, and not the result of spurious receptor activation or UV-induced photodamage. To rule out these artifacts, two specificity controls can be performed. First, fluorescence intensity in cells that are not directly irradiated during photoactivation experiments can be monitored. If the surface is properly caged, appreciable responses in these cells should not be observed (for example, Figure 5C). Second, experiments in which T cells are UV-irradiated on surfaces lacking photoactivatable ligands can be performed. This control is critical for identifying UV-induced effects that are independent of photoactivatable ligand (for example, see reference 6). During system optimization, it is also extremely helpful to have at hand a robust cellular imaging response (e.g., Grb2-GFP recruitment) that is both fast and localized. Assessing correlations between signaling events and photoactivation is much more straightforward if those events are spatiotemporally proximal to receptor stimulation.
Although the photoactivation approach offers exquisite spatiotemporal control over receptor signaling, it lacks the adaptability afforded by transfectable optogenetic systems16,17,18,19,20,21,22. Certain optogenetic proteins are also photoreversible, providing conditional control that is not available with the photoactivation approach described here. In optogenetic systems, however, constraining the diffusion of photoactivated proteins is technically challenging. This complicates the analysis of polarized signaling, particularly in a subcellular context.
One should also note that the engaging entity of this photoactivation system is a glass surface, which cannot recapitulate all aspects of a bona fide cell-cell interaction. To circumvent this issue while maintaining spatiotemporal control of signal initiation, investigators have used an optical trap system to bring an APC into contact with a T cell, allowing for the onset of cytoskeletal rearrangements and IS maturation to be visualized in real time23. Although this system enables the precise initiation and visualization of cytoskeletal dynamics using an actual APC, it is relatively low throughput and is not compatible with TIRF illumination, which adversely affects image quality.
In conclusion, the method described in this protocol provides spatiotemporal control of TCR-induced signaling within the T cell, ultimately allowing for the elucidation of the fast biochemical changes following TCR activation. This system provides us with a means to better understand how the signaling events following TCR activation promote T cell polarization, IS formation, and ultimately the delivery of effector responses.

T lymphocytes (T cells) play a central role in cellular immunity by recognizing antigenic peptides displayed in the context of cell surface MHC. Antigen recognition, which is mediated by the TCR, drives the differentiation of na&#239;ve T cells and promotes the delivery of cytolytic and communicative responses by effector populations. TCR engagement also induces dramatic changes in cellular architecture. Within minutes, the T cells gloms onto the side of the antigen-presenting cell (APC), forming a polarized interface known as the immunological synapse (IS)1,2. The IS potentiates T cell effector responses by enabling the directional release of cytokines or, in the case of cytotoxic T lymphocytes (CTLs), lytic proteins that destroy the APC.
TCR engagement by pMHC induces the rapid phosphorylation of multiple downstream adaptor molecules, including Linker for the Activation of T cells (LAT), which ultimately promotes robust remodeling of the synaptic cytoskeleton2. Cortical filamentous actin (F-actin) drives T cell spreading over the APC surface, and then resolves into an annular structure characterized by F-actin accumulation at the IS periphery and depletion from the center. F-actin ring formation is tightly coupled to the reorientation of the microtubule organizing center (MTOC, also called the centrosome in T cells) to a position just beneath the center of the interface. Both events occur within minutes of initial antigen recognition and establish the architectural context in which subsequent activation events and effector responses occur.
To study IS formation, various labs have developed approaches in which the APC is replaced by a glass surface that either contains immobilized TCR ligands or supports a lipid bilayer that itself contains the ligands3,4. T cells form IS-like contacts on these surfaces that can be imaged by total internal reflection fluorescence microscope (TIRF) or confocal microscopy, enabling high-resolution studies of early T cell activation and IS formation.
Although these approaches have allowed for excellent visualization of the fully assembled IS, much of the signaling following TCR:pMHC ligation occurs within seconds, complicating efforts to determine the sequence of events following TCR activation accurately. To circumvent this issue, a photoactivation approach has been developed, in which photoactivatable pMHC is used to achieve spatiotemporal control of TCR activation5,6,7. In this system, T cells are attached to glass surfaces containing photoactivatable pMHC that is non-stimulatory to the TCR until irradiated with ultraviolet (UV) light. UV irradiation of a micron sized region of the surface beneath the T cell removes the photocage creating a stimulatory zone that can be recognized by the T cell. Subsequent signaling events and cytoskeletal remodeling are then monitored using genetically encoded fluorescent reporters. Two photoactivatable versions of antigenic peptides, moth cytochrome c88-103 (MCC) and ovalbumin257-264 (OVA), which are presented in the context of the class II MHC I-Ek and the class I MHC H2-Kb, respectively, have been developed (Figure 1). This enables the analysis of both CD4+ T cells specific for MCC- I-Ek (expressing the 5C.C7, 2B4, or AND TCRs) and CD8+ T cells specific for OVA-H2-Kb (expressing the OT1 TCR).
Over the past decade, the TCR photoactivation and imaging approach has been utilized to establish the precise kinetics of early TCR signaling steps and also to identify the molecular pathways governing polarized cytoskeletal remodeling5,6,7,8,9,10. For example, the assay was instrumental in determining that centrosome reorientation toward the APC is mediated by a localized gradient of the lipid second messenger diacylglycerol centered at the IS. It is anticipated that this methodology will continue to be valuable for applications that demand high-resolution imaging analysis of T cell function.

<table><tbody><tr><td>Nunc Lab-Tek Chambered Coverglass</td><td>Thermofischer Scientific</td><td>155361</td><td></td></tr><tr><td>Poly-L-lysine hydrobromide</td><td>Sigma-Aldrich</td><td>P2636</td><td>Will need to make Biotinylated Poly-L-Lysine</td></tr><tr><td>EZ-Link NHS-Biotin</td><td>Thermofischer Scientific</td><td>20217</td><td>Will need to make Biotinylated Poly-L-Lysine</td></tr><tr><td>Streptavidin</td><td>Thermofischer Scientific</td><td>434301</td><td></td></tr><tr><td>BirA-500: BirA biotin-protein ligase standard reaction kit</td><td>Avidity</td><td>BirA500</td><td>Will be used to biotinylate proteins</td></tr><tr><td>Biotinylated Hb I-E&lt;sup&gt;K&amp;nbsp;&lt;/sup&gt;</td><td></td><td></td><td>For protein folding, see reference 6. For biotinylation, use BirA kit</td></tr><tr><td>Biotinylated NPE-MCC I-E&lt;sup&gt;K&amp;nbsp;&lt;/sup&gt;</td><td>Anaspec</td><td></td><td>Custom NPE-MCC (H-ANERADLIAYL-K(Nvoc)-QATK-OH) can be purchased from Anaspec</td></tr><tr><td>Biotinylated &amp;alpha;H2-K&lt;sup&gt;k&lt;/sup&gt; antibody</td><td>BD Biosciences</td><td>553591</td><td></td></tr><tr><td>Biotinylated NPE-OVA H2-K&lt;sup&gt;b&lt;/sup&gt;</td><td>Anaspec</td><td></td><td>Custom NPE-OVA (H-SIINFE-K(Nvoc)-L-OH) can be purchased from Anaspec</td></tr><tr><td>Biotinylated KAVY H2-D&lt;sup&gt;b&lt;/sup&gt;&amp;nbsp;</td><td>Anaspec</td><td></td><td>Custom synthesized protein (KAVYDFATL) can be purchased from Anaspec</td></tr><tr><td>Biotinylated ICAM1&amp;nbsp;</td><td></td><td></td><td>For protein folding, see reference in protocol. For biotinylation, use BirA kit</td></tr><tr><td>Hand held UV lamp</td><td>UVP</td><td>UVGL-25</td><td>Lamp is held &amp;lt; 1 cm from the sample.&amp;nbsp; 30 s of 365 light is sufficient for detectable decaging, 20 min for quantitative decaging.</td></tr><tr><td>Olympus IX-81 OMAC TIRF system.</td><td>Olympus</td><td></td><td>Additional information about the imaging system can be found in &lt;strong class=&quot;xfig&quot;&gt;Figure 6&lt;/strong&gt;</td></tr><tr><td>Mosaic digital diaphragm</td><td>Andor</td><td></td><td></td></tr><tr><td>Slidebook software</td><td>Intelligent Imaging Innovations</td><td></td><td></td></tr></tbody></table>

spatial and temporal control of t cell activation using a photoactivatable agonist

T lymphocytes engage in rapid, polarized signaling, occurring within minutes following TCR activation. This induces formation of the immunological synapse, a stereotyped cell-cell junction that regulates T cell activation and directionally targets effector responses. To study these processes effectively, an imaging approach that is tailored to capturing fast, polarized responses is necessary. This protocol describes such a system, which is based on a photoactivatable peptide-major histocompatibility complex (pMHC) that is non-stimulatory until it is exposed to ultraviolet light. Targeted decaging of this reagent during videomicroscopy experiments enables precise spatiotemporal control of TCR activation and high-resolution monitoring of subsequent cellular responses by total internal reflection (TIRF) imaging. This approach is also compatible with genetic and pharmacological perturbation strategies. This allows for the assembly of well-defined molecular pathways that link TCR signaling to the formation of the polarized cytoskeletal structures that underlie the immunological synapse.

The photoactivation and imaging approach allows for observation and facile quantification of rapid, polarized signaling responses. To illustrate its capabilities, reproduced here is an experiment examining the spatiotemporal correlation between TCR-induced DAG accumulation and centrosome reorientation. 5C.C7 T cell blasts were retrovirally transduced with two fluorescent reporters: a DAG biosensor containing the tandem C1 domains from protein kinase C-&#952; linked to GFP (C1-GFP) and RFP-tubulin to monitor the centrosome. The T cells were then attached to chambered coverglass containing photoactivatable MCC- I-Ek. C1-GFP was imaged using TIRF microscopy and RFP-tubulin in epifluorescence mode. As shown in Figure 2, localized photoactivation of the surface beneath the T cell induces the accumulation of DAG in the irradiated zone. This is followed within seconds by the reorientation of the centrosome to the same region.
C1-GFP accumulation can be quantified by calculating the normalized fluorescence intensity, after background correction, at the center of the photoactivated region over time (Figure 3). Centrosome reorientation in response to photoactivation can be quantified by calculating the distance between the centrosome and the center of the photoactivated region as a function of time (Figure 4).
This method can be used to examine a wide array of rapid, polarized signaling responses, essentially anything that can be monitored using a fluorescent probe. For example, TCR photoactivation is utilized to monitor early signaling microcluster formation using fluorescently labeled Grb2 (Grb2-GFP) (Figure 5). Similar to the DAG accumulation and centrosome reorientation responses, Grb2-GFP polarization occurs shortly following photoactivation (Figure 5), and can be quantified using the normalized fluorescence intensity approach.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56655/56655fig1.jpg" /> 
Figure 1: Photoactivation system using photocaged MCC peptide. 
(A) Photocaged strategy for MCC peptide. A NPE group is attached to the lysine residue of the MCC peptide. UV irradiation results in the removal of NPE, restoring MCC to its native form. (B) When NPE is bound to MCC, the 5C.C7 TCR cannot bind to MCC. Removal of NPE results in recognition of the 5C.C7 TCR to the native MCC peptide. (C) The photocaged strategy has been adapted to work with CD8+ T cells, in which photocaged OVA peptide is used to activate the OT-1 TCR upon UV irradiation.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56655/56655fig2.jpg" /> 
Figure 2: Time-lapse montage of DAG accumulation and centrosome reorientation following photoactivation of TCR in 5C.C7 T cell. 
5C.C7 cell expressing a DAG biosensor, containing tandem C1-domains of PKC- fused to GFP (C1-GFP) and Tag-RFP Tubulin (RFP-Tub), a fluorescent reporter for the centrosome. The montages illustrate the polarized response of 5C.C7 T cells over time. The yellow oval and text indicate the time and location of photoactivation. Over time, C1-GFP accumulates at the photoactivated region, followed by centrosome reorientation towards the photoactivated region. Scale bar: 10 &#181;m.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56655/56655fig3.jpg" /> 
Figure 3: Quantification of DAG accumulation at the photoactivated region 
DAG, C1-GFP, accumulation at the photoactivated region (left) can be quantified by calculating the normalized fluorescence intensity, after background correction, at the photoactivated region (yellow circle with blue shading) over time. Data are normalized relative to the nine frames obtained just before photoactivation. C1-GFP enrichment at the photoactivated region can be calculated by the division of the mean fluorescence intensity in the irradiated region by the mean fluorescence intensity of the entire cell, following background correction. The equation: &#916;F/F = ((FI-FIb)/mean(FI1-9)-FIb)), in which FI is the mean fluorescence intensity, can be used to calculate the normalized fluorescence intensity. The normalized fluorescence intensity at the photoactivated region can then be graphed as a function of time (right). Purple line indicates the time of photoactivation. Scale bar: 10 &#181;m.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56655/56655fig4.jpg" /> 
Figure 4: Quantification of centrosome reorientation. 
Determine the coordinates of the centrosome (RFP-Tub) over time by manually tracking the centrosome path toward the photoactivated region (white arrow) at each time point (left). Determine the coordinates of the center of the photoactivated region (yellow circle). The distance between the centrosome and the center of the photoactivated region can be calculated at each time point using the equation for distance = &#8730;((x2-x1)2+(y2-y1)2), in which x2 and y2 are the x and y coordinates for the center of the area of the centrosome, and x1 and y1 are the x and y coordinates of the center of the photoactivated region. The distance of the centrosome from the center of the photoactivated region at each time point can then be graphed as a function of time (right). Purple line indicates the time of photoactivation. Scale bar: 10 &#181;m.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56655/56655fig5.jpg" /> 
Figure 5: Grb2-GFP microcluster formation. 
(A) Representative 5C.C7 cell expressing fluorescently labeled Grb2 (Grb2-GFP). The montages illustrate the polarized response of Grb2-GFP overtime. The yellow oval and text indicate the time and location of photoactivation. Over time, Grb2-GFP accumulates at the photoactivated region. (B) Quantification of Grb2-GFP fluorescence intensity at the photoactivated region over time. N = 15 cells. (C) Quantification of Grb2-GFP fluorescence intensity in a control region outside of the photoactivated zone. N = 15 cells.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56655/56655fig6.jpg" /> 
Figure 6: Microscope configuration for photoactivation and TIRF imaging. 
488 nm and 561 nm lasers are used for TIRF and epifluorescence imaging of green and red probes, respectively. UV light for photoactivation is taken from a mercury (Hg) lamp attached to a digital diaphragm system capable of illuminating small, user-defined regions. Light from the Hg lamp/diaphragm is reflected onto the sample by a longpass mirror positioned beneath the dichroic mirror that reflects the imaging lasers. The dichroic mirror must be designed to pass UV light. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56655/56655fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Spatial and Temporal Control of T Cell Activation Using a Photoactivatable Agonist at JoVE.com

Spatial and Temporal Control of T Cell Activation Using a Photoactivatable Agonist

This protocol describes an imaging-based method to activate T lymphocytes using photoactivatable peptide-MHC, enabling precise spatiotemporal control of T cell activation.

T lymphocytes engage in rapid, polarized signaling, occurring within minutes following TCR activation. This induces formation of the immunological ...

spatial-temporal-control-t-cell-activation-using-photoactivatable

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

6.2K Views.  Memorial Sloan-Kettering Cancer Center. This protocol describes an imaging-based method to activate T lymphocytes using photoactivatable peptide-MHC, enabling precise spatiotemporal control of T cell activation.

Video: Spatial and Temporal Control of T Cell Activation Using a Photoactivatable Agonist

Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular functions and events1, 2. Their spatiotemporally dynamic nature has been revealed by visualization of their activity and localization in real time3. In order to gain deeper understanding of their roles in diverse cellular functions at the molecular level, the next step should be perturbation of protein activities at a precise subcellular location and timing. 
To achieve this goal, we have developed a method for light-induced, spatio-temporally controlled activation of small GTPases by combining two techniques: (1) rapamycin-induced FKBP-FRB heterodimerization and (2) a photo-caging method of rapamycin. With the use of rapamycin-mediated FKBP-FRB heterodimerization, we have developed a method for rapidly inducible activation or inactivation of small GTPases including Rac4, Cdc424, RhoA4 and Ras5, in which rapamycin induces translocation of FKBP-fused GTPases, or their activators, to the plasma membrane where FRB is anchored. For coupling with this heterodimerization system, we have also developed a photo-caging system of rapamycin analogs. A photo-caged compound is a small molecule whose activity is suppressed with a photocleavable protecting group known as a caging group. To suppress heterodimerization activity completely, we designed a caged rapamycin that is tethered to a macromolecule such that the resulting large complex cannot cross the plasma membrane, leading to virtually no background activity as a chemical dimerizer inside cells6. Figure 1 illustrates a scheme of our system. With the combination of these two systems, we locally recruited a Rac activator to the plasma membrane on a timescale of seconds and achieved light-induced Rac activation at the subcellular level6.

A method for spatio-temporal control of small GTPase activity by light is described. This method is based on rapamycin-induced FKBP-FRB heterodimerization and photo-caging systems. Optimization of light-irradiation enables the spatio-temporally controlled activation of small GTPases at the subcellular level.

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular ...

Bioengineering

An Optogenetic Method to Control and Analyze Gene Expression Patterns in Cell-to-cell Interactions

Cells should respond properly to temporally changing environments, which are influenced by various factors from surrounding cells. The Notch signaling pathway is one of such essential molecular machinery for cell-to-cell communications, which plays key roles in normal development of embryos. This pathway involves a cell-to-cell transfer of oscillatory information with ultradian rhythms, but despite the progress in molecular biology techniques, it has been challenging to elucidate the impact of multicellular interactions on oscillatory gene dynamics. Here, we present a protocol that permits optogenetic control and live monitoring of gene expression patterns in a precise temporal manner. This method successfully revealed that intracellular and intercellular periodic inputs of Notch signaling entrain intrinsic oscillations by frequency tuning and phase shifting at the single-cell resolution. This approach is applicable to the analysis of the dynamic features of various signaling pathways, providing a unique platform to test a functional significance of dynamic gene expression programs in multicellular systems.

Here, we present a protocol to analyze cell-to-cell transfer of oscillatory information by optogenetic control and live monitoring of gene expression. This approach provides a unique platform to test a functional significance of dynamic gene expression programs in multicellular systems.

Cells should respond properly to temporally changing environments, which are influenced by various factors from surrounding cells. The Notch signaling ...

Genetics

A TIRF Microscopy Technique for Real-time, Simultaneous Imaging of the TCR and its Associated Signaling Proteins

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex (pMHC) proteins expressed on the surface of antigen presenting cells (APCs). The TCR complex is composed of the ligand binding TCR&alpha;&beta; heterodimer that associates non-covalently with CD3 dimers (the &#949;&delta; and &#949;&gamma; heterodimers and the &zeta;&zeta; homodimer)1. Upon engagement of the receptor, the CD3 &zeta; chains are phosphorylated by the Src family kinase, Lck. This leads to the recruitment of the Syk family kinase, Zap70, which is then phosphorylated and activated by Lck. After that, Zap70 phosphorylates the adapter proteins LAT and SLP76, initiating the formation of the proximal signaling complex containing a large number of different signaling molecules2. 
The formation of this complex eventually results in calcium and Ras-dependent transcription factor activation and the consequent initiation of a complex series of gene expression programs that give rise to T cell differentiation2. TCR signals (and the resulting state of differentiation) are modulated by many other factors, including antigen potency and crosstalk with co-stimulatory/co-inhibitory, chemokine, and cytokine receptors 3-4. Studying the spatial and temporal organization of the proximal signaling complex under various stimulation conditions is, therefore, key to understanding the TCR signaling pathway as well as its regulation by other signaling pathways.
One very useful model system to study signaling initiated by the TCR at the plasma membrane in T cells is glass-supported lipid bilayers, as described previously5-6. They can be utilized to present antigenic pMHC complexes, adhesion, and co-stimulatory molecules to T cells-serving as artificial APCs. By imaging the T cells interacting with the lipid bilayer using total internal reflection fluorescence microscopy (TIRFM), we can restrict the excitation to within 100 nm of the space between the glass and the cell surface 7-8. This allows us to image primarily the signaling events occurring at the plasma membrane. As we are interested in imaging the recruitment of signaling proteins to the TCR complex, we describe a two-camera TIRF imaging system wherein the TCR, labeled with fluorescent Fab (fragment antigen binding) fragments of the H57 antibody (purified from hybridoma H57-597, ATCC, ATCC Number:HB-218) which is specific for TCR&beta;, and signaling proteins, tagged with GFP, may be imaged simultaneously and in real time. This strategy is necessary due to the highly dynamic nature of both the T cells and of the signaling events that are occurring at the TCR. This imaging modality has allowed researchers to image single ligands 9-11 as well as recruitment of signaling molecules to activated receptors and is an excellent system to study biochemistry in-situ12-16.

The compartmentalization of proteins either within the plasma membrane or into intracellular locations is one regulatory mechanism that can greatly influence signaling outcomes; hence, to understand signaling it is important to study the spatial and temporal behavior of the proteins involved. We describe here a TIRF microscopy based system to study signal transduction in T cells, but is broadly applicable.

Signaling is initiated through the T Cell Receptor (TCR) when it is engaged by antigenic peptide fragments bound by Major Histocompatibility Complex ...

Real-time Live Imaging of T-cell Signaling Complex Formation

Protection against infectious diseases is mediated by the immune system 1,2. T lymphocytes are the master coordinators of the immune system, regulating the activation and responses of multiple immune cells 3,4. T-cell activation is dependent on the recognition of specific antigens displayed by antigen presenting cells (APCs). The T-cell antigen receptor (TCR) is specific to each T-cell clone and determines antigen specificity 5. The binding of the TCR to the antigen induces the phosphorylation of components of the TCR complex. In order to promote T-cell activation, this signal must be transduced from the membrane to the cytoplasm and the nucleus, initiating various crucial responses such as recruitment of signaling proteins to the TCR;APC site (the immune synapse), their molecular activation, cytoskeletal rearrangement, elevation of intracellular calcium concentration, and changes in gene expression 6,7. The correct initiation and termination of activating signals is crucial for appropriate T-cell responses. The activity of signaling proteins is dependent on the formation and termination of protein-protein interactions, post translational modifications such as protein phosphorylation, formation of protein complexes, protein ubiquitylation and the recruitment of proteins to various cellular sites 8. Understanding the inner workings of the T-cell activation process is crucial for both immunological research and clinical applications.
Various assays have been developed in order to investigate protein-protein interactions; however, biochemical assays, such as the widely used co-immunoprecipitation method, do not allow protein location to be discerned, thus precluding the observation of valuable insights into the dynamics of cellular mechanisms. Additionally, these bulk assays usually combine proteins from many different cells that might be at different stages of the investigated cellular process. This can have a detrimental effect on temporal resolution. The use of real-time imaging of live cells allows both the spatial tracking of proteins and the ability to temporally distinguish between signaling events, thus shedding light on the dynamics of the process 9,10. We present a method of real-time imaging of signaling-complex formation during T-cell activation. Primary T-cells or T-cell lines, such as Jurkat, are transfected with plasmids encoding for proteins of interest fused to monomeric fluorescent proteins, preventing non-physiological oligomerization 11. Live T cells are dropped over a coverslip pre-coated with T-cell activating antibody 8,9, which binds to the CD3/TCR complex, inducing T-cell activation while overcoming the need for specific activating antigens. Activated cells are constantly imaged with the use of confocal microscopy. Imaging data are analyzed to yield quantitative results, such as the colocalization coefficient of the signaling proteins.

We describe a live-cell imaging method that provides insight into protein dynamics during the T-cell activation process. We demonstrate the combined usage of the T-cell spreading assay, confocal microscopy and imaging analysis to yield quantitative results to follow signaling complex formation throughout T-cell activation.

Protection against infectious diseases is mediated by the immune system 1,2. T lymphocytes are the master coordinators of the immune system, regulating ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Biology

Imaging CD4 T Cell Interstitial Migration in the Inflamed Dermis

The ability of CD4 T cells to carry out effector functions is dependent upon the rapid and efficient migration of these cells in inflamed peripheral tissues through an as-yet undefined mechanism. The application of multiphoton microscopy to the study of the immune system provides a tool to measure the dynamics of immune responses within intact tissues. Here we present a protocol for non-invasive intravital multiphoton imaging of CD4 T cells in the inflamed mouse ear dermis. Use of a custom imaging platform and a venous catheter allows for the visualization of CD4 T cell dynamics in the dermal interstitium, with the ability to interrogate these cells in real-time via the addition of blocking antibodies to key molecular components involved in motility. This system provides advantages over both in vitro models and surgically invasive imaging procedures. Understanding the pathways used by CD4 T cells for motility may ultimately provide insight into the basic function of CD4 T cells as well as the pathogenesis of both autoimmune diseases and pathology from chronic infections.

The mechanisms that govern the interstitial motility of CD4 effector T cells at sites of inflammation are relatively unknown. We present a non-invasive approach to visualize and manipulate in vitro-primed CD4 T cells in the inflamed ear dermis, allowing for study of the dynamic behavior of these cells in situ.

The ability of CD4 T cells to carry out effector functions is dependent upon the rapid and efficient migration of these cells in inflamed peripheral ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Neuroscience

Use of Single Chain MHC Technology to Investigate Co-agonism in Human CD8+ T Cell Activation

Non-stimulatory self peptide MHC (pMHC) complexes do not induce T cell activation and effector functions, but can enhance T cell responses to agonist pMHC, through a process termed co-agonism. This protocol describes an experimental system to investigate co-agonism during human CD8+ T cell activation by expressing human MHC class I molecules presenting pre-determined peptides as single polypeptides (single chain MHC) in a xenogeneic cell line. We expressed single chain MHCs under conditions where low levels of agonist single chain p-MHC complexes and high levels of non-stimulatory single chain p-MHC complexes were expressed. Use of this experimental system allowed us to compare CD8+ T cell responses to agonist pMHC in the presence or absence of non-stimulatory pMHC. The protocol describes cell line transfection with single chain MHC constructs, generation of stable cell lines, culture of hepatitis B virus-specific human CD8+ T cells and T cell activation experiments simultaneously quantifying cytokine production and degranulation. The presented methods can be used for research on different aspects of CD8+ T cell activation in human T cell systems with known peptide MHC specificity.

This protocol describes the use of single chain MHC class I complexes to investigate molecular interactions in human CD8+ T cell activation: generation of engineered antigen presenting cells expressing single chain constructs, culture of human CD8+ T cell clone and T cell activation experiments.

Non-stimulatory self peptide MHC (pMHC) complexes do not induce T cell activation and effector functions, but can enhance T cell responses to agonist ...

Interrogating Individual Autoreactive Germinal Centers by Photoactivation in a Mixed Chimeric Model of Autoimmunity

Autoimmune diseases present a significant health burden. Fundamental questions regarding the development and progression of autoimmune disease remain unanswered. One requirement for advancements in our understanding of the underlying disease mechanisms and cellular dynamics is the precise coupling of the microanatomical location of cell subsets with downstream molecular or functional analyses; a goal that has traditionally been difficult to achieve. The development of stable photoactivatable biological fluorophores and their integration into reporter strains has recently enabled precise microanatomical labeling and tracking of cellular subsets in murine models. Here, we describe how the ability to analyze autoreactive lymphocytes from single germinal centers may help to provide novel insights into autoimmunity, using the combination of a novel chimeric model of autoimmunity with a photoactivatable reporter as an example. We demonstrate a procedure for generating mixed chimeras with spontaneous autoreactive germinal centers populated by lymphocytes carrying a photoactivatable green fluorescent protein reporter. Using in vivo labeling strategies, single germinal centers can be visualized in explanted lymphoid tissues and their cellular constituents photoactivated by two-photon microscopy. Photoactivated lymphocytes from single germinal centers can then be analyzed or sorted flow cytometrically, as single cells or in bulk, and may be subjected to additional downstream molecular and functional analyses. This approach may directly be applied to provide renewed insights in the field of autoimmunity, but the procedure for generating bone marrow chimeras and the photoactivation procedure may additionally find broad application in studies of infectious diseases and tumor metastases.

This protocol describes the generation of mixed murine bone marrow chimeras with spontaneous autoimmune germinal centers, in which autoreactive lymphocytes carry a photoactivatable green fluorescent protein (PA-GFP) reporter. This provides the ability to link cellular location in tissues with downstream molecular and functional analyses.

Autoimmune diseases present a significant health burden. Fundamental questions regarding the development and progression of autoimmune disease remain ...

Optogenetic Control of Enzyme Activity Using Light-Regulated Allosteric Switch

Spatiotemporal Control of Protein Activity through Optogenetic Allosteric Regulation

Optogenetics offers the potential for mimicking complex spatiotemporal control of enzyme activity down to a subcellular resolution. However, most optogenetic approaches often face significant challenges in integrating multiple capabilities in a single tool applicable to a wide range of target proteins. Achieving precise control over ON/OFF kinetics, ensuring minimum leakiness in the dark, and demonstrating efficient performance in mammalian cells with subcellular precision are some of the most common challenges faced in this field. A promising solution lies in the application of rationally designed light-sensitive domains to allosterically control protein activity. Using that strategy, we generated an optogenetic method combining all the desired features. The approach involves the incorporation of the Light-regulated allosteric switch module (LightR) in the target protein to regulate enzyme activity using blue (465 nm) light. The LightR domain is generated by linking two Vivid (VVD) photoreceptor domains, creating a light-sensitive clamp that can be incorporated into a small flexible loop within the catalytic domain of an enzyme. In its dark state, LightR clamp is open, thus distorting the enzyme&#39;s catalytic domain and inactivating it. Upon exposure to blue light, the LightR domain closes and restores the catalytic domain&#39;s structure and enzyme activity. In this manuscript, we discuss design strategies&#160;to generate a light-regulated protein kinase and demonstrate its control by blue light, reversibility, kinetics, and precise regulation at the subcellular level, enabling tight spatiotemporal precision. Utilizing Src tyrosine kinase as a model, we showcase a protocol for effectively regulating LightR-Src kinase activity. We also demonstrate LightR applicability across different enzyme classes, expanding the utility of the tool system in addressing mechanistic questions of signaling pathways in different diseases.

This protocol describes the application of an engineered blue-light-activated allosteric switch (LightR) domain for reversible spatiotemporal control of protein activity. Utilizing Src tyrosine kinase as a model, this study offers an elaborate protocol for developing and characterizing light-regulated Src (LightR-Src). It demonstrates the versatility of this approach across enzyme classes.