Protocol Optimization

The detection of events as rare as 0.02% of total CD8+ T cells requires both the design of suitably controlled experiments and a well-maintained flow cytometer. If the number of antigen-specific events is expected to be low (< 0.05%), it is important to acquire a suitably large number of events (~500,000) within the live lymphocyte gate in order to collect sufficient events of your population of interest. The following sections set out important issues in detail.

Additionally, ProImmune’s Pro5® MHC Pentamer handbook is an essential reference for anyone carrying out antigen-specific T cell staining and analysis. We recommend that even experienced Pentamer users consult this handbook for practical tips and advice.


Cell Sample Setting the live lymphocyte gate
Titrating the Pentamer Using the correct anti-CD8 antibody
Incubation temperature and time Controls
Elimination of non-specific staining by exclusion of CD19+ B cells


Cell Sample

During their preparation, cells should be at room temperature (22°C) to maintain the metabolic activity and membrane lipid fluidity of the cells. Subsequently, unfixed cells should be kept cold (on ice) during antibody staining and analysis (note: Pentamer staining should be at room temperature). This will help to ensure that cell surface markers are not internalized following the interaction of the binding reagent (antibody or Pentamer) with the cell surface receptor. The cells must be in single cell suspension for the antibody labeling to work successfully.

Pentamers can also be used to stain cell samples that have previously been frozen for storage, although the number of viable cells will be reduced. Thaw the cells carefully and wash twice before proceeding with staining.

Cell preparation protocols can be downloaded from the protocols page. They are also available in the Pro5® MHC Pentamer handbook.

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Setting the Live Lymphocyte Gate

Light scatter is used to analyze specific subsets of cells, such as lymphocytes, within a heterogeneous population, and is commonly used to exclude dead cells, cell aggregates and cell debris from the fluorescence data. As cells pass through the laser beam of the flow cytometer, the light scatter is changed according to the size (forward scatter) and granularity (side scatter) of each cell. This allows large and very granular cells to be differentiated from the smaller, less granular lymphocytes in the blood. It can be difficult to identify the correct live lymphocyte population, but it is very important to do this correctly in order to minimize non-specific staining events seen in the density plot.

Forward scatter (FSC) is strongly influenced by cell size; the larger the cells, the further along the x-axis they appear. Side scatter (SSC) reflects granularity or internal cell structure; the larger and more granular a cell, the further up the y-axis it appears. Cell populations that are close together may be most easily distinguished using a density plot.

In the figure below, 3 populations of cells can be distinguished by their forward- and side-scatter profiles. Gating on these populations and looking at their CD8 and Pentamer expression easily distinguishes between them. Although cells in regions R1 and R3 are abundant, R2 is the correct population to gate upon. Looking at the profiles, R2 defines cells that have the expected percentage (20-30% of live cells) of CD8 positive (i.e. mature T cells) and a distinct population of Pentamer positive cells. A lower proportion of the R1 population are CD8 positive and do not stain cleanly for Pentamer. Judging by the FSC versus SSC properties of these cells they are likely to be dead cells and debris. The population of cells defined by R3 has no CD8+ cells within it, and these do not incorporate Pentamer staining. Because their FSC and SSC properties define them as large and granular, they are most probably granulocytes.

Example data plots showing that populations of cells can be distinguished by their forward and side scatter profiles

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Titrating the MHC Pentamer

The amount of Pentamer added to each staining condition should be titrated in order to find the optimum working dilution to use for each individual Pentamer and cell type. A suggested titration range would be doubling dilutions from 1 test quantity down to 1/16 test quantities.

Example data plots illustrating the importance of titrating Pentamer

The figure above illustrates the importance of titrating the Pentamer. The four plots are of live T cells from peripheral blood. For ease of viewing, Pentamer staining is shown against CD8 FITC. 1×106 peripheral blood mononuclear cells (PBMCs) were stained with either 1, 0.5, 0.25 or 0.1 test equivalents of B*07:02/CMV (TPRVTGGGAM) Pentamer for 10 min at 22°C. Following 2 washes, cells were incubated with an optimal dilution of anti-CD8-FITC for 20 minutes at 4°C. 100,000 live events were collected. When 1 test of Pentamer is used a clear population of antigen-specific T cells is seen (1.1% of total cells gated). The negative cells and the CD8-positive Pentamer-negative cells are well within the lower quadrant. The stain of the Pentamer is bright.

Comparing 0.5 test of Pentamer with 1 test shows that the mean fluorescence intensity has reduced (i.e. Pentamer staining is not as bright). With 0.25 test of Pentamer the frequency of cells detected is lower (1.0%). This is due to some events appearing in the lower quadrants of the plot and being indistinguishable from the negative population. When 0.1 test of Pentamer is used, the fluorescence intensity has significantly decreased, affecting the frequency of cells detected. Thus, by titration, any non-specific staining seen with high concentrations of MHC Pentamer can be reduced to an ideal level. Further titration below this level will reduce the signal intensity. The level of fluorescence intensity required for the application should also be taken into account.

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CD8 Antibody

Your anti-CD8 antibody co-stain should be carefully selected since some clones interfere with MHC multimer binding to the T cell receptor. We recommend using clone LT8 with human Pentamers and clone KT15 with mouse Pentamers, which may be purchased from us with your order. The staining sequence you carry out may also need to be optimized. For example, adding Pentamer first and incubating for 10 minutes, then washing the cells before addition of anti-CD8 co-stain is preferred to the addition of both reagents simultaneously.

It is important to cross-titrate the Pentamer against the anti-CD8 antibody. If the concentration of both reagents is too high, background staining will interfere with data analysis. If the quantity of reagent used is too little, the fluorescence intensity signal may be too low to analyze results accurately. Cross-titration of the regents enables the reduction of background staining, whilst maintaining an optimum fluorescence intensity level.

The figure below illustrates the importance of cross titration of Pentamer against anti-CD8 monoclonal antibody. 1×106 PBMCs were stained with either 1.0, 0.25 or 0.1 test equivalents of R-PE labeled B*07:02/CMV (TPRVTGGGAM) Pentamer for 10 minutes at 22°C. Following 2 washes, cells were incubated with either 1.0, 0.5, or 0.1 ul of anti-CD8-FITC (clone LT8) for 20 minutes at 4°C. From this example, the Pentamer should be used at 1 test quantity (10ul) and the anti-CD8 antibody at either 1 or 0.5 ul per sample, depending on the desired degree of separation of positive from negative cells.

Example data plots showing the importance of cross titration of Pentamer against anti-CD8 monoclonal antibody

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Incubation temperature and time

Staining at 22°C (room temperature) for 10 minutes is recommended in the first instance. Incubation at 4°C for 40 minutes or at 37°C for 5 to 10 minutes may be tested in order to optimize the signal to noise ratio. The higher the incubation temperature, the shorter the incubation time required.
In the figure below, the upper central density plot (22°C for 10 minutes) shows a higher mean fluorescence compared with the first plot of 4°C for 10 minutes. When the temperature is increased to 37°C, a higher level of non-specific staining is seen (1.3%) and base line negative staining is raised. These events interfere with the positive events under investigation and affect the data significantly. Although the number of CD8 positive and Pentamer positive cells have increased (0.9%), some of these events are likely to be Pentamer negative. Fewer cells can be visualized due to clumping that occurs at the higher temperature. Similar characteristics are seen in the lower plots when a longer staining time (45 minutes) is tried.

Example data for testing incubation temperature and time

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Positive control: MHC Pentamers should be tested against a specific T cell line or clone, if available. Use T cells that have not been stimulated recently as this has been shown to cause down-regulation of T cell receptors. Cells should not be used for a minimum of 3-4 days after stimulation. If possible, wait 10 days after stimulation for best results. If a T cell line is not available, it is possible to use PBMCs from a known positive donor. In this situation the frequency of positive cells will be much lower and more cells will be required per stain (at least 1×106). Ideally, collect functional data using a technique such as ELISPOT, to indicate the frequency of positive cells that should be expected.

Negative control: Cells obtained from an unexposed (seronegative) individual may be used. To control for non-specific staining it is also advisable to stain the cell sample with either the A*02:01 Human Negative Control Pro5® Pentamer, or a mismatched Pro5® Pentamer (irrelevant MHC allele and/or irrelevant peptide.

The ProImmune HLA-A*02:01 negative control Pentamer product consists of multimeric HLA-peptide complexes assembled with irrelevant peptide antigen which is known to have no T cell response. Any staining achieved by using the negative control Pentamer is genuinely non-specific, so this reagent is ideal for use when studying a low frequency of antigen-specific T cells.

Negative Pentamer Product Sheet

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Elimination of non-specific staining by exclusion of CD19+ B cells

Background (non-specific) staining is sometimes observed when performing flow cytometric analysis of murine splenocyte and blood samples, and of human PBMCs, particularly in the CD8-negative, Pentamer-positive quadrant. This problem is largely caused by non-specific staining of B cells, and can be reduced or eliminated by co-staining with anti-CD19 antibody and gating on the CD19-negative cells when performing analysis. Alternatively, anti-CD19 magnetic beads could be used to remove B cells prior to staining.

The figure below demonstrates the effect of gating out B cells from murine splenocytes. 1 x 106 naive C57BL/6 (a) – (c) or SIINFEKL-immunized C57BL/6 (d) – (f) splenocytes were incubated with 1 test unlabeled H-2Kb/OVA (SIINFEKL) Pro5® MHC Pentamer for 10 minutes at room temperature (22°C). Cells were washed, then further incubated with 1 test Pro5® R-PE Fluorotag, 1 test anti-mouse CD8-FITC (clone KT15) and 1 test anti-mouse CD19-PECy5 (clone 6D5) for 30 minutes at 4°C. In (a) & (d), the live splenocyte population shows considerable non-specific staining in the CD8-negative, Pentamer-positive quadrant; (b) & (e) A region R2 is set upon the CD19-negative cells; (c) & (f) The live splenocyte population is re-gated to show only events within R2 (which excludes B cells), and the non-specific staining has now been eliminated. This illustrates that a substantial majority of the non-specific staining can be attributed to B cells.

Example data showing elimination of non-specific staining by exclusion of CD19+ B cells

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