We thank Dick Foley and Leigh Kimball for their key roles in filming and video production.

1. Peripheral blood mononuclear cell (PBMC) isolation
<ol>
<li>Obtain a blood sample &ndash; blood should be collected in syringes or blood tubes and anti-coagulated with heparin (1:50 ratio) to prevent clotting. Expect a yield of about 1&times;106 PBMC per mL of blood &ndash; about 40% of which will be CD4 positive (CD4+) T cells.</li>
<li>Aliquot the blood into 50 mL conical tubes, 25 mL per tube. If blood has separated, gently mix before aliquoting to distribute the plasma</li>
<li>Add PBS, bringing the total volume to 40 mL and underlay by drawing up 11 mL of Ficoll, inserting into blood tube, and carefully removing the pipet Aid from the pipette. The ficoll will slowly drain into the bottom of the tube. Once the level has equalized, slowly remove the pipette from the blood tube and discard.</li>
<li>Cap the blood tubes and move into the Aerosolve canisters. Firmly close the canisters and centrifuge at 900&times;g for 20 minutes &ndash; no brake. It is critical that no brake is applied at the end of this spin as the braking force will disturb the layer of cells.</li>
<li>After the spin, the PBMCs should form a distinct white layer between the yellow plasma (above) and transparent ficoll (below). Gently skim off white blood cell layer using a transfer pipette and transfer to new tube. Some plasma and Ficoll may be drawn up with the PBMCs, but avoid drawing up any of the red cell layer. Typically two blood tubes can be combined into one cell tube at this step to increase the pellet size.</li>
<li>Add PBS, bringing the total volume to 50 mL and centrifuge at 500&times;g for 15 minutes &ndash; low brake in aerosolve canisters.</li>
<li>Aspirate liquid using a Pasteur pipette, taking care not to disturb the pellet</li>
<li>Treat with hemolytic buffer by adding 5-6 mL to each tube and gently mixing to remove clumps. Incubate for no more than 5 minutes.</li>
<li>Add PBS, bringing the total volume to 50 mL and centrifuge at 230&times;g for 10 minutes &ndash; low brake. Aerosolve canisters are no longer needed for these slower spins.</li>
<li>Wash two times: Aspirate liquid using a Pasteur pipette, fill tube with PBS and centrifuge at 230&times;g for 10 minutes.</li>
<li>Prior to the final wash step, remove an aliquot of re-suspended cells, dilute with trypan blue, and count using a hemocytometer. This cell count will dictate the reagent volumes used in the T cell separation step.</li>
</ol>
2. CD4+ T cell separation*
<ol>
<li>Aspirate liquid and add running buffer to bring the total volume up to 40 uL per 10 million cells. Usually this requires 30 uL of buffer plus the residual pellet volume.</li>
<li>Transfer this volume to a 15 mL conical tube.</li>
<li>Add antibody cocktail from the CD4 isolation kit &ndash; 10 uL per 10 million cells, cap tube, and place on ice for 10 minutes</li>
<li>Remove tube from ice, remove cap, and add 30 uL of running buffer per 10 million cells</li>
<li>Add magnetic beads from the CD4 isolation kit &ndash; 20 uL per 10 million cells, cap tube, and place on ice for 15 minutes</li>
<li>Add ten volumes of running buffer to wash and centrifuge at 230&times;g for 10 minutes.</li>
<li>During the spin, set up the magnet and 5 mL polypropylene tube in your workspace</li>
<li>Aspirate liquid from cell pellet, add 2 mL running buffer and remove clumps using a transfer pipette</li>
<li>Using the same transfer pipette, transfer cells into the 5 mL tube and incubate in the magnet for 15 minutes</li>
<li>Carefully decant the CD4+ cell fraction into a marked 15 mL tube by inverting the magnet and emptying all but the last drop</li>
<li>Add another 2 mL of running buffer to the 5 mL tube and remove from magnet</li>
<li>Gently rinse the CD4- cells off the sides of tube and transfer to a marked 15 mL tube</li>
<li>Add 2 mL T cell medium to each tube, remove aliquots for cell counting and centrifuge at 230&times;g for 10 minutes.</li>
<li>Dilute cell aliquots with trypan blue and count using a hemocytometer. These cell counts will dictate the number of wells used for the expansion culture step.</li>
</ol>
* Alternatively, MACS columns and beads (Miltenyi Biotec), the AutoMACS cell separator (Miltenyi Biotec), or Robosep cell separator (Stem Cell Technologies) can be used according to manufactures instructions in place of steps 2.2 &ndash; 2.11.
3. In vitro Expansion culture
<ol>
<li>Aspirate liquid from the CD4+ and CD4- cell pellets and, based on the cell counts, add culture media (typically 3 million CD4+ cells per mL and 10 million CD4- cells per mL works well for setting up the culture). The expansion culture requires 3-5 million CD4- cells per well and 2-3 million CD4+ cells per well in a total volume of 1 mL culture media in a 48 well culture plate. Typically, CD4+ cells are the limiting population</li>
<li>If desired, irradiate the CD4- cells (5000 Rads). Aliquot CD4- cells into the appropriate number of wells on a 48 well plate by adding 300-500 &mu;L to each. Place the plate in a 37&deg;C incubator for 1 hour. The adherent CD4- cells will serve as antigen presenting cells in the expansion culture.</li>
<li>Remove the plate from the incubator.</li>
<li>Wash the wells by adding 500 uL of fresh media and gently pipetting up and down 12-20 times using a transfer pipet and removing all of the liquid. This wash will leave behind a layer of adherent antigen presenting cells.</li>
<li>When the washing is complete, add just enough media to wet the bottom of each well (roughly 100 uL) &ndash; otherwise the adherent cells will dry out</li>
<li>Add 2-3 million CD4+ cells to each well. If necessary, add additional media to bring the total volume in each well up to 1 mL.</li>
<li>Add the desired peptide to each well. The typical concentration for stimulation is 10 ug/mL final.</li>
<li>Place plate in a 37&deg;C incubator for 7 days.</li>
<li>After 7 days, add 50 uL of Hemogen IL-2 (or equivalent, such as recombinant IL-2, 10 IU/mL) to each well.</li>
<li>On each subsequent day, monitor cell growth using a microscope. Feed wells that are not yet confluent by removing half of the media, adding fresh media and 50 uL of IL-2. Split confluent wells by resuspending the cells, moving half of the cells to an empty well, adding fresh media and 50 uL of IL-2. Return cells to incubator.</li>
<li>The expanding cells will be ready for tetramer after 13-15 days of culture.</li>
</ol>
4. Visualizing T cells by tetramer staining
<ol>
<li>Purchase or assemble tetramers to match the peptide/MHC combinations that match the stimulated CD4+ T cells (see discussion for sources of tetramer reagents). Tetramers should be ~0.5 mg/mL solution.</li>
<li>Remove plate from incubator</li>
<li>Carefully draw off half of the media from each well</li>
<li>Transfer an aliquot of cells from each well - typically 75 &mu;L or about 50,000 cells - into a 5 mL polystyrene FACS tube</li>
<li>Add tetramer - typically 1 &mu;L per 50 &mu;L total volume - and place in 37&deg;C incubator for 1-2 hours. It is also advisable to stain a second aliquot of cells from each well with a mismatched tetramer as a negative control.</li>
<li>Label cells with anti-CD3, anti-CD4, and anti-CD25 antibodies by adding 3-10 &mu;L of each antibody. Additional or alternative antibody markers can be used. However, do not use PE labeled antibodies since this channel must be reserved for the tetramers. At this time, also label single color or isotype controls using extra cells.</li>
<li>Incubate antibody labeled cells for 15-30 minutes on ice or at 4&deg;C.</li>
<li>Wash each tube with 0.5 to 2 mL running buffer and centrifuge at 230&times;g for 10 minutes.</li>
<li>Carefully decant liquid from FACS tubes. Store all tubes in a covered container (preferably on ice) for subsequent FACS analysis.</li>
</ol>
5. Flow Cytometer Acquisition and Analysis
<ol>
<li>Calibrate the Flow Cytometer using reference beads.</li>
<li>Verify instrument settings:<ol>
<li>Using unstained control cells or isotype controls, adjust the location of the viable lymphocyte gate (FSC vs. SSC). Remove auto-fluorescence from all channels by centering the population (by changing the gain settings) below the first decade on each axis.</li>
<li>Using single color control tubes, center events within the correct quadrants for each single color (by changing the compensation settings).</li>
<li>Do a fine adjustment of the instrument settings using a tube stained with an irrelevant tetramer. In particular settings for the PE channel may need adjustment because tetramers often stain brighter than the antibody control. The irrelevant tetramer should stain &lt;0.5% of the CD4+ cells.</li>
</ol></li>
<li>Acquire data for each tube<ol>
<li>Collect events for each sample tube and negative control tube</li>
<li>Acquire and save at least 20,000 events for analysys</li>
</ol></li>
<li>Analyze data<ol>
<li>Import data files into FACS analysis software (e.g CellQuest software, WinMDI, or FlowJo). Start your analysis using a tube stained using an irrelevant tetramer.</li>
<li>Plot FSC vs. SSC and draw a gate around the live lymphocytes (Figure 1, top left panel).</li>
<li>Plot CD4 vs. CD3 (gated to show live lymphocytes only) and draw gates around the CD3+ and CD4+ populations (Figure 1 top right panel).</li>
<li>Plot CD4 vs. Tetramer (gated to show CD3+ lymphocytes only) and set the quadrant boundaries so that the CD4+Tetramer+ population is less than 0.5% (Figure 1 bottom left panel).</li>
<li>Plot CD25 vs. Tetramer (gated to show CD4+ lymphocytes only) and set the quadrant boundaries so that the CD25+Tetramer+ population is less than 0.5% (Figure 1 bottom right panel).</li>
<li>Analyze all of the sample tubes without changing the gates.</li>
</ol></li>
</ol>
<img src="/files/ftp_upload/1167/JOVE%20FIGURE%201.png" alt="Figure 1" width="600" height="342" />
Figure 1. Representative negative control tetramer staining results. The upper left panel shows forward scatter verses side scatter and the lymphocyte size gate (R1). The upper right panel is gated for R1 and shows ant-CD4 versus anti-CD3 and the CD3+ (R2) and CD4+ (R3) gates. The lower left panel is gated for R2 and shows anti-CD4 versus tetramer. The lower right panel is gated for R3 and shows anti-CD25 versus tetramer. The quadrants for both tetramer plots were set to 0.5%.
6. Representative Results
<ol>
<li>The tetramer positive cells should be a distinct population among the expanded CD4+ cells (Figure 2).</li>
<li>Tetramer positive cells are usually CD25+ and often CD4 high.</li>
<li>In some cases, a MHC/peptide combination can give background staining (often caused by a peptide with poor solubility). Such background staining can be differentiated from a true positive because the &ldquo;false positive&rdquo; cells are not a single distinct population on either the CD4 vs. Tetramer or the CD25 vs. Tetramer plots (Figure 3).</li>
</ol>
<img src="/files/ftp_upload/1167/JOVE%20FIGURE%202.png" alt="" width="600" height="362" />
Figure 2. Representative positive tetramer staining results. The panel layout and gating strategy are identical to Figure 1. The tetramer positive cells appear as a distinct CD4 high population on the anti-CD4 versus tetramer panel and a distinct CD25+ population on the anti-CD25 versus tetramer panel.
<img src="/files/ftp_upload/1167/JOVE%20FIGURE%203.png" alt="Figure 3" width="600" height="336" />
Figure 3. Representative &ldquo;false positive&rdquo; tetramer staining results. The panel layout and gating strategy are identical to Figure 1. The anti-CD4 versus tetramer panel shows an indistinct &ldquo;mounded&rdquo; staining. The anti-CD25 versus tetramer also shows a &ldquo;mounded&rdquo; staining, with no clear trend toward being CD25+.

<ol>
	<li>Novak, E. J., Liu, A. W., Nepom, G. T., Kwok, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MHC+Class+II+tetramers+permit+detection+and+characterization+of+peptide-specific+human+CD4++T+cells+proliferating+to+influenza+A+antigen.">MHC Class II tetramers permit detection and characterization of peptide-specific human CD4+ T cells proliferating to influenza A antigen.</a> J. Clin. Invest. 104, R63-R67 (1999).</li><li>Novak, E. J., Liu, A. W., Gebe, J. A., Falk, B., Nepom, G. T., Falk, B., Koelle, D. M., Kwok, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tetramer-guided+epitope+mapping:+rapid+identification+and+characterization+of+immunodominant+CD4++T+cell+epitopes+from+complex+antigens.">Tetramer-guided epitope mapping: rapid identification and characterization of immunodominant CD4+ T cell epitopes from complex antigens.</a> J. Immunol. 166, 6665-6670 (2001).</li></ol>

Understanding the role of CD4+ T cells in immunity is fundamentally important. However, antigen specific CD4+ T cells can be difficult to detect and isolate using traditional methods. In contrast, MHC class II tetramers allow the direct visualization of CD4+ T cells with the desired antigen specificity. This video demonstrates the isolation, purification, and in vitro amplification of CD4+ T cells and their subsequent visualization using tetramers. Class II Tetramers are fluorescent protein conjugates consisting of s...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Class 2 Laminar flow biosafety cabinet</td>
<td>equipment</td>
<td>Thermo Forma</td>
<td>1200</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>50 mL conical Tube</td>
<td>equipment</td>
<td>VWR</td>
<td>47747-182</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>1X PBS (Ca/Mg free)</td>
<td>Reagent</td>
<td>HyClone</td>
<td>SH30028.02</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Ficoll-paque PLUS</td>
<td>Reagent</td>
<td>GE Healthcare</td>
<td>17-1440-03</td>
<td>foil wrapped to protect from light</td>
</tr>
<tr>
<td>Pipet Aid XP</td>
<td>equipment</td>
<td>Drummond</td>
<td>4-000-101</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>10 mL pipet</td>
<td>equipment</td>
<td>Corning/Costar</td>
<td>CLS4492</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Aerosolve Canisters</td>
<td>equipment</td>
<td>Beckman</td>
<td>359481</td>
<td>with 50 mL inserts</td>
</tr>
<tr>
<td>Centrifuge</td>
<td>equipment</td>
<td>Beckman</td>
<td>GS-6R</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Transfer pipets </td>
<td>equipment</td>
<td>Samco</td>
<td>202-20S</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Pasteur pipets</td>
<td>equipment</td>
<td>VWR</td>
<td>14672-200</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Hemolytic Buffer</td>
<td>Reagent</td>
<td>N/A</td>
<td>N/A</td>
<td>8.3 g/L NH4Cl, 1.0 g/L NaHCO3, 0.04 g/L disodium EDTA</td>
</tr>
<tr>
<td>Trypan blue</td>
<td>Reagent</td>
<td>Sigma</td>
<td>T6146</td>
<td>0.2% in PBS</td>
</tr>
<tr>
<td>Hemocytometer</td>
<td>equipment</td>
<td>VWR </td>
<td>15170-208</td>
<td>&nbsp;</td>
<tr>
<td>15 mL conical tube</td>
<td>equipment</td>
<td>VWR</td>
<td>05-527-90</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Running Buffer</td>
<td>Reagent</td>
<td>N/A</td>
<td>N/A</td>
<td>PBS + 2mM EDTA + 5g/L BSA</td>
</tr>
<tr>
<td>MACS CD4+ T Cell Isolation Kit II</td>
<td>Reagent</td>
<td>Miltenyi Biotec</td>
<td>130-091-155</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>EasySep&reg; Magnet</td>
<td>equipment</td>
<td>Stem Cell Technologies</td>
<td>18000</td>
<td>		</td>
</tr>
<tr>
<td>5 mL polypropylene tube</td>
<td>equipment</td>
<td>Falcon</td>
<td>352063</td>
<td>		</td>
</tr>
<tr>
<td>RPMI 1640</td>
<td>Reagent</td>
<td> Invitrogen</td>
<td>22400-089</td>
<td>with 25 mM HEPES</td>
</tr>
<tr>
<td>Pooled human serum</td>
<td>Reagent</td>
<td>N/A</td>
<td>N/A</td>
<td>drawn from healthy donors, heat inactivated and filtered</td>
</tr>
<tr>
<td>Pen Strep</td>
<td>Reagent</td>
<td>Invitrogen</td>
<td>1570-063</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>500mL 0.22 micron bottle top filter</td>
<td>equipment</td>
<td>Nalgene</td>
<td>595-3320</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>48-well plate</td>
<td>equipment</td>
<td>Costar</td>
<td>3548</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>37&deg;C CO2 incubator</td>
<td>equipment</td>
<td>Sanyo Scientific</td>
<td>MCO-18AIC</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>20 mg/mL synthetic peptide</td>
<td>Reagent</td>
<td>N/A</td>
<td>N/A</td>
<td>Custom synthesis from any peptide vendor</td>
</tr>
<tr>
<td>5 mL FACS tube</td>
<td>equipment</td>
<td>Falcon</td>
<td>352008</td>
<td>Polystyrene</td>
</tr>
<tr>
<td>Peptide loaded class II tetramer (as described)</td>
<td>Reagent</td>
<td>N/A</td>
<td>N/A</td>
<td>Prepared by the BRI tetramer core</td>
</tr>
<tr>
<td>Anti-CD3</td>
<td>Reagent</td>
<td>BD pharmingen</td>
<td>347344</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Anti-CD4</td>
<td>Reagent</td>
<td>eBioscience</td>
<td>17-0049-73</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Anti-CD25</td>
<td>Reagent</td>
<td>eBioscience</td>
<td>11-0259-73</td>
<td>Or equivalent</td>
</tr>
<tr>
<td>Facs Calibur Flow Cytometer</td>
<td>equipment</td>
<td>BD Biosciences</td>
<td>342975</td>
<td>Or equivalent</td>
</tr>		</tbody>
		</table>
		</div>

visualizing antigen specific cd4+ t cells using mhc class ii tetramers

Major histocompatibility complex (MHC) class II tetramers allow the direct visualization of antigen specific CD4+ T cells by flow cytometry.  This method relies on the highly specific interaction between peptide loaded MHC and the corresponding T-cell receptor.  While the affinity of a single MHC/peptide molecule is low, cross-linking MHC/peptide complexes with streptavidin increases the avidity of the interaction, enabling their use as staining reagents.  Because of the relatively low frequencies of CD4+ T cells (~1 in 300,000 for a single specificity) this assay utilizes an in vitro amplification step to increase its threshold of detection.  Mononuclear cells are purified from peripheral blood by Ficoll underlay.  CD4+ cells are then separated by negative selection using biotinylated antibody cocktail and anti-biotin labeled magnetic beads.  Using adherent cells from the CD4- cell fraction as antigen presenting cells, CD4+ T cells are expanded in media by adding an antigenic peptide and IL-2.  The expanded cells are stained with the corresponding class II tetramer by incubating at 37 C for one hour and subsequently stained using surface antibodies such as anti-CD4, anti-CD3, and anti-CD25.  After labeling, the cells can be directly analyzed by flow cytometry.  The tetramer positive cells typically form a distinct population among the expanded CD4+ cells.  Tetramer positive cells are usually CD25+ and often CD4 high.  Because the level of background tetramer staining can vary, positive staining results should always be compared to the staining of the same cells with an irrelevant tetramer.  Multiple variations of this basic assay are possible.  Tetramer positive cells may be sorted for further phenotypic analysis, inclusion in ELISPOT or proliferation assays, or other secondary assays.  Several groups have also demonstrated co-staining using tetramers and either anti-cytokine or anti-FoxP3 antibodies.  

Watch this Scientific Journal Video about Visualizing Antigen Specific CD4+ T Cells using MHC Class II Tetramers at JoVE.com

Visualizing Antigen Specific CD4+ T Cells using MHC Class II Tetramers

This procedure demonstrates the purification and in vitro expansion of antigen specific CD4+ T cells from whole peripheral blood and their visualization using MHC class II tetramers. Tetramers permit the direct visualization of T cells with a single antigen specificity and defined MHC class II restriction.

Major histocompatibility complex (MHC) class II tetramers allow the direct visualization of antigen specific CD4+ T cells by flow cytometry.  This method ...

visualizing-antigen-specific-cd4-t-cells-using-mhc-class-ii-tetramers

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JoVE Journal

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21.6K Views.  Benaroya Research Institute. This procedure demonstrates the purification and in vitro expansion of antigen specific CD4+ T cells from whole peripheral blood and their visualization using MHC class II tetramers. Tetramers permit the direct visualization of T cells with a single antigen specificity and defined MHC class II restriction. 

Video: Visualizing Antigen Specific CD4+ T Cells using MHC Class II Tetramers

CD4+ T-Lymphocyte Capture Using a Disposable Microfluidic Chip for HIV

Isolation of CD4+ T cells from Mouse Lymph Nodes Using Miltenyi MACS Purification

Isolation of cells from the primary source is a necessary step in many more complex protocols. Miltenyi offers kits to isolate cells from several organisms including humans, non-human primates, rat and, as we describe here, mice. Magnetic bead-based cell separation allows for either positive selection (or cell depletion) as well as negative selection. Here, we demonstrate negative selection of untouched or na ve CD4+ helper T cells. Using this standard protocol we typically purify cells that are &#8805; 96% pure CD4+/CD3+. This protocol is used in conjunction with the protocol Dissection and 2-Photon Imaging of Peripheral Lymph Nodes in Mice published in issue 7 of JoVE, for purification of T cells and other cell types to adoptively transfer for imaging purposes. Although we did not demonstrate FACS analysis in this protocol video, it is highly recommended to check the overall purity of isolated cells using the appropriate antibodies via FACS. In addition, we demonstrate the non-sterile method of T cell isolation. If sterile cells are needed for your particular end-user application, be sure to do all of the demonstrated procedures in the tissue culture hood under standard sterile conditions. Thank you for watching and good luck with your own experiments!

Isolation of lymphocytes using the Miltenyi MACs kit is a reliable way to purify cells from whole lymphoid tissue homogenates. Cells purified using the Miltenyi system are typically &#8805; 96% pure. Here, we demonstrate the steps taken to isolate CD4+ T cells, one of the many kits offered by Miltenyi.

Isolation of cells from the primary source is a necessary step in many more complex protocols. Miltenyi offers kits to isolate cells from several ...

A Technique to Simultaneously Visualize Virus-Specific CD8+ T Cells and Virus-Infected Cells In situ

The numbers and locations of virus-specific CD8+ T cells relative to the numbers and locations of their infected cell targets is thought to be critical in determining outcomes that range from clearance to chronic persistent infections. We describe here a method for assessing the spatial and quantitative relationships between immune effector (E) virus-specific CD8+ T cells and infected targets (T) that combines in situ tetramer (IST) staining to detect virus-specific CD8+ T cells and in situ hybridization (ISH) to detect viral-RNA+ cells in the tissues where the battle between immune defenses and infection takes place. The combination of IST staining and ISH, abbreviated ISTH, enables visualization and mapping of the locations of immune effector cells and targets, and facile determination of E:T ratios.  These parameters in turn can then be used to determine the relationships between spatial proximity, and the timing and magnitude of the immune response that predict outcomes in early infection.

A Technique to Simultaneously Visualize Virus-Specific CD8+ T Cells and Virus-Infected Cells In situ

A technique combining in situ tetramer staining and in situ hybridization (ISTH) enables visualization, mapping and analysis of the spatial proximity of virus-specific CD8+ T cells to their virus-infected targets, and determination of the quantitative relationships between these immune effectors and targets to infection outcomes.

The numbers and locations of virus-specific CD8+ T cells relative to the numbers and locations of their infected cell targets is thought to be critical in ...

Isolation of Valvular Endothelial Cells

Heart valves are solely responsible for maintaining unidirectional blood flow through the cardiovascular system. These thin, fibrous tissues are subjected to significant mechanical stresses as they open and close several billion times over a lifespan. The incredible endurance of these tissues is due to the resident valvular endothelial (VEC) and interstitial cells (VIC) that constantly repair and remodel in response to local mechanical and biological signals. Only recently have we begun to understand the unique behaviors of these cells, for which in vitro experimentation has played a key role. Particularly challenging is the isolation and culture of VEC. Special care must be used from the moment the tissue is removed from the host through final plating. Here we present protocols for direct isolation, side specific isolation, culture, and verification of pure populations of VEC. We use enzymatic digestion followed by a gentle swab scraping technique to dislodge only surface cells. These cells are then collected into a tube and centrifuged into a pellet. The pellet is then resuspended and plated into culture flasks pre-coated with collagen I matrix. VEC phenotype is confirmed by contact inhibited growth and the expression of endothelial specific markers such as PECAM1 (CD31), Von Willebrand Factor (vWF), and negative expression of alpha-smooth muscle actin (&alpha;-SMA). The functional characteristics of VEC are associated with high levels of acetylated LDL. Unlike vascular endothelial cells, VEC have the unique capacity to transform into mesenchyme, which normally occurs during embryonic valve formation1. This can also occur during significantly prolonged post confluent in vitro culture, so care should be made to passage at or near confluence. After VEC isolation, pure populations of VIC can then be easily acquired.

We provide a method for isolating and culturing pure populations of heart valve endothelial cells (VEC). VEC can be isolated from either side of the cusp or leaflet and immediately following, underlying interstitial cell (VIC) isolation is straightforward.

Heart valves are solely responsible for maintaining unidirectional blood flow through the cardiovascular system.  These thin, fibrous tissues are ...

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the Arabidopsis leaf is a complex organ made up of many different cell types and several structures. At the same time, the growing leaf contains cells at different stages of development, with the cells furthest from the petiole being the first to stop expanding and undergo senescence1. Different cells within the leaf are therefore dividing, elongating or differentiating; active, stressed or dead; and/or responding to stimuli in sub-sets of their cellular type at any one time. This makes genomic study of the leaf challenging: for example when analyzing expression data from whole leaves, signals from genetic networks operating in distinct cellular response zones or cell types will be confounded, resulting in an inaccurate profile being generated.
To address this, several methods have been described which enable studies of cell specific gene expression. These include laser-capture microdissection (LCM)2 or GFP expressing plants used for protoplast generation and subsequent fluorescence activated cell sorting (FACS)3,4, the recently described INTACT system for nuclear precipitation5 and immunoprecipitation of polysomes6.
FACS has been successfully used for a number of studies, including showing that the cell identity and distance from the root tip had a significant effect on the expression profiles of a large number of genes3,7. FACS of GFP lines have also been used to demonstrate cell-specific transcriptional regulation during root nitrogen responses and lateral root development8, salt stress9 auxin distribution in the root10 and to create a gene expression map of the Arabidopsis shoot apical meristem11. Although FACS has previously been used to sort Arabidopsis leaf derived protoplasts based on autofluorescence12,13, so far the use of FACS on Arabidopsis lines expressing GFP in the leaves has been very limited4. In the following protocol we describe a method for obtaining Arabidopsis leaf protoplasts that are compatible with FACS while minimizing the impact of the protoplast generation regime. We demonstrate the method using the KC464 Arabidopsis line, which express GFP in the adaxial epidermis14, the KC274 line, which express GFP in the vascular tissue14 and the TP382 Arabidopsis line, which express a double GFP construct linked to a nuclear localization signal in the guard cells (data not shown; Figure 2). We are currently using this method to study both cell-type specific expression during development and stress, as well as heterogeneous cell populations at various stages of senescence.

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

A method for producing Arabidopsis leaf protoplasts that are compatible with fluorescence activated cell sorting (FACS), allowing for studies of specific cell populations. This method is compatible with any Arabidopsis line that expresses GFP in a subset of cells.

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the ...

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

A High Throughput MHC II Binding Assay for Quantitative Analysis of Peptide Epitopes

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or design1,2. Here, a peptide-MHC II binding assay is scaled to 384-well format. The scaled down protocol reduces reagent costs by 75% and is higher throughput than previously described 96-well protocols1,3-5. Specifically, the experimental design permits robust and reproducible analysis of up to 15 peptides against one MHC II allele per 384-well ELISA plate. Using a single liquid handling robot, this method allows one researcher to analyze approximately ninety test peptides in triplicate over a range of eight concentrations and four MHC II allele types in less than 48 hr. Others working in the fields of protein deimmunization or vaccine design and development may find the protocol to be useful in facilitating their own work. In particular, the step-by-step instructions and the visual format of JoVE should allow other users to quickly and easily establish this methodology in their own labs.

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or design. &#160;Here, a peptide-MHC II binding assay scaled to 384-well plates is described. This cost effective format should prove useful in the fields of protein deimmunization and vaccine design and development.

Biochemical assays with recombinant human MHC II molecules can provide rapid, quantitative insights into immunogenic epitope identification, deletion, or ...

Detection of Intracellular Gene Expression in Live Cells of Murine, Human and Porcine Origin Using Fluorescence-labeled Nanoparticles

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, rat, human, pig and others. The verification of iPS clones mainly relies on the detection of the endogenous expression of different pluripotency genes. These genes mostly represent transcription factors which are located in the cell nucleus. Traditionally, the proof of their endogenous expression is supplied by immunohistochemical staining after fixation of the cells. This approach requires replicate cultures of each clone at this early stage to preserve validated clones for further experiments. The present protocol describes an approach with gene-specific nanoparticles which allows the evaluation of intracellular gene expression directly in live cells by fluorescence. The nanoparticles consist of a central gold particle coupled to a capture strand carrying a sequence complementary to the target mRNA as well as a quenched reporter strand. These nanoparticles are actively endocytosed and the target mRNA displaces the reporter strand which then start to fluoresce. Therefore, specific target gene expression can be detected directly under the microscope. In addition, the emitted fluorescence allows the identification, isolation and enrichment of cells expressing a specific gene by flow cytometry. This method can be applied directly to live cells in culture without any manipulation of the target cells.

This manuscript describes a novel technique which allows the detection of intracellular gene expression after endocytosis of fluorescence-labeled nanoparticles directly in live cells. The method does not require manipulation of the cells and is not restricted with respect to the target gene or species.

The reprogramming of somatic cells to induced pluripotent stem cells (iPS) has successfully been performed in different mammalian species including mouse, ...

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here protocols are presented for preparing embryos and adult C. elegans animals for short- and long-term time-lapse microscopy and methods for tracking and quantification of developmental processes. The methods presented are all based on C. elegans strains available from the Caenorhabditis Genetics Center and on open-source software that can be easily implemented in any laboratory independently of the microscopy system used. A reconstruction of a 3D cell-shape model using the modelling software IMOD, manual tracking of fluorescently-labeled subcellular structures using the multi-purpose image analysis program Endrov, and an analysis of cortical contractile flow using PIVlab (Time-Resolved Digital Particle Image Velocimetry Tool for MATLAB) are shown. It is discussed how these methods can also be deployed to quantitatively capture other developmental processes in different models, e.g., cell tracking and lineage tracing, tracking of vesicle flow.

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Here it is shown how to track and quantify developmental processes in C. elegans. The methods presented are based on open-source tools that can be easily implemented. It is demonstrated how to reconstruct 3D cell-shape models, how to manually track subcellular structures, and how to analyze cortical contractile flow.

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here ...

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper (cTfh) Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the lymph node microenvironment in B-cell non-Hodgkin&#39;s lymphoma is analyzed. Subsets of circulating T-follicular helper cells (cTfh), the circulating memory compartment of Tfh cells in the blood, are also perturbed in disease and therefore represent potential novel predictive biomarkers. Peripheral blood-based testing is advantageous because it is relatively non-invasive and allows simple serial monitoring.This article describes a method for isolating CD4+ T-cells from human blood, and further analysis by flow-cytometry to enumerate cTfh cells and the proportions of their various subsets (cTfhPD-1-/+/hi, cTfh1,2,17 and cTfh1/17). The level of these subsets was then compared between normal subjects and patients with lymphoma. We found that the method was robust enough to obtain reliable results from routinely collected patient material. The technique we describe for the analysis can be easily adapted to cell sorting and downstream applications such as RT-PCR.

Isolation of CD4+ T-cells and Analysis of Circulating T-follicular Helper cTfh Cell Subsets from Peripheral Blood Using 6-color Flow Cytometry

Here, a protocol for the isolation and characterization of CD4+ T-cell subsets from human peripheral blood is described. Purified CD4+ T-cells are analyzed by flow cytometry to determine proportions of T-follicular helper cell subsets.

Aberrant T-follicular helper (Tfh) cell activity is detectable in autoimmune conditions and their presence is associated with clinical outcomes when the ...