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Mapping T cell epitopes by flow cytometry
Methods 29 (2003) 270–281 www.elsevier.com/locate/ymeth
Mapping T cell epitopes by flow cytometry Bodo Hoffmeister,a Felix Kiecker,a Lydia Tesfa,a Hans-Dieter Volk,a Louis J. Picker,b and Florian Kerna,* a
Department of Medical Immunology, Charit e, Humboldt Universit€at zu Berlin, Schumannstrasse 21, 10117 Berlin, Germany b Vaccine and Gene Therapy Institute, Oregon Health and Science University, Portland, OR 97201, USA Accepted 2 December 2002
Abstract Epitope mapping by flow cytometry is a very modern approach that not only identifies T-cell epitopes but simultaneously allows for detailed analysis of the responding T-cell subsets including lineage, activation marker expression, and other markers of interest. The most frequently used approach is based on the identification of intracellular cytokines in secretion-inhibited activated T cells following stimulation with peptides or peptide pools. A more recently developed assay analyzes T-cell proliferation by measuring the decrease in carboxyfluorescein diacetate succinimidyl ester staining in proliferated cells. This article includes information on peptide configuration, a section on the design and efficient application of peptide pools, and working laboratory protocols for both assays. 2003 Elsevier Science (USA). All rights reserved.
1. Introduction Epitopes are the portions of antigens that bind specifically with the binding site of an antibody or a receptor on a T lymphocyte. While antibodies (B-cell receptors) bind free spatial epitopes, T-cell receptors require their epitopes to be presented on major histocompatibility complex (MHC) molecules in a linear fashion. The T-cell receptor makes contact with portions of the peptide and portions of the MHC molecule so that, strictly speaking, a T-cell epitope comprises both the peptide and determinants of the presenting MHC molecule. A description of the details of antigen presentation by MHC molecules is clearly beyond the scope of this article but can be found elsewhere [1]. Although a very large number of protein sequences are known and stored in databases, for most proteins it is not known whether they are immunogenic (i.e., whether they bind one or more MHC alleles within a population and are recognizable by the T-cell receptor repertoire), and where the antigenic determinants are localized within the primary amino acid sequence. The identification of immunogenic proteins and determination of the T-cell epitopes contained in them often *
Corresponding author.
stand at the very beginning of vaccine development. While many vaccines use ‘‘bulk approaches’’ like whole recombinant proteins (hepatitis B), lysates of infectious agents (Bacillus Calmette–Guerin), complete toxoids (tetanus), there is increasing interest in more individualized peptide-based vaccinations. Possible applications include (but are not limited to) infectious diseases, tumor vaccines, and autoimmunity [2–5]. Whether or not a vaccination is peptide based in its final form, the exact knowledge of relevant epitopes is helpful in studying how a vaccine is applied most efficiently, since known epitopes may be packaged with adjuvants, applied within a longer stretch of amino acids or protein, coded by DNA using free plasmids or various vectors or even modified by amino acid substitutions to generate more immunogenic molecules. Peptide-based vaccinations may induce or modulate both CD4 and CD8 T-cell responses; however, depending on the form of the vaccine (e.g., peptide or protein plus adjuvant or DNA) CD4 or CD8 T cells may be the preferred responding population. The use of defined epitopes in vaccines allows determination of which T-cell subset (helper or cytotoxic/suppressor) is going to respond in most cases, the only exception being peptides that induce both CD4 and CD8 T-cell responses. A quite recent application of epitope identification is in the quantification of epitope-specific T-cell populations by
1046-2023/03/$ - see front matter 2003 Elsevier Science (USA). All rights reserved. doi:10.1016/S1046-2023(02)00349-3
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tetrameric MHC complexes (short: ‘‘tetramers’’) [6]. Such tetramers are composed of four recombinant MHC molecules, each of which is coupled to a biotin molecule and linked by binding to the four biotin binding sites of a fluorochrome-conjugated streptavidin molecule. The MHC molecules all contain the same peptide in their binding groove such that the tetrameric complexes can directly bind to T-cell receptors that recognize the peptide–MHC molecule combination. Such reagents may be used to directly examine T-cell responses at an antigenspecific level. Peptides alone, on the other hand, can be used for effectively stimulating T-cell responses in peripheral blood mononuclear cells (PBMC) or other cell suspensions. A very interesting application is the use of such peptides as ‘‘diagnostic stimulant,’’ i.e., to find out if infection with a certain agent has taken place (and has induced T cells) or not [7]. Whether using tetramers or peptides as a stimulant, T cell responses can be identified, analyzed, and monitored. Both approaches are widely used for monitoring T-cell responses in patients with human immunodeficiency virus (HIV), Epstein–Barr virus (EBV), or cytomegalovirus (CMV) infections. This article is dedicated exclusively to the mapping of T-cell epitopes by flow cytometry [8]. The method described is limited to epitopes within known protein sequences, since the stimulating agents for this approach are peptides that were synthesized according to a given primary protein sequence. The attraction of using a flow cytometer for mapping epitopes lies in the great versatility of this approach with respect to the number of parameters that may be acquired in a single measurement, i.e., cell lineage markers, activation markers on the cell surface and inside the cell [8]. The use of many parameters greatly enhances the resolution of the assay, i.e., identification of the T cell subset that responds to a given epitope including lineage and activation markers [9]. The approach we originally designed is based on the detection of rapid cytokine production in activated T cells in a short-term (as short as 6 h) ex vivo assay [10]. The biggest advantage of this approach apart from its unrivaled speed is that CD4 and CD8 T-cell responses can be measured in the same assay and even the same test tube [11]. However, proliferation may also be used as a readout parameter when using flow cytometry. Among the systems for measuring cell proliferation by flow cytometry that are available on the market, we favor the use of carboxyfluorescein diacetate succinimidyl ester (CFDA-SE) staining, because it is simple and reliable. The staining intensity of this protein dye is lost in regular increments as the intracellular protein content is passed on to the next generation by dividing cells. CFDA-SE is a nontoxic, fluorescein-related dye, which is able to permeate cells with the help of its two acetate side chains. Once inside cells, the acetate groups are removed by intracellular esterases and the resultant carboxyfluorescein exits at a much slower rate, allowing
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covalent binding to free amines of cytoplasmatic proteins, forming very stable amino bonds. Because CFSE also reacts with molecules that are rapidly degraded or transported out of the cell, a lot of the CFDA-SE that is initially taken up by cells and transformed into CFSE is lost during the first 24 h following labeling. The remaining portion that is bound to more stable intracellular components may be used for tracking cell proliferation. The use of CFDA-SE staining for the analysis of cell proliferation by flow cytometry was described in 2000 [12]. However, we have only recently started using it for the purpose of epitope identification and we therefore describe only our preliminary protocol. As regards proliferation assays, in general one fundamental difference between them and the detection of intracellular cytokines following short-term stimulation is that the latter does not require clonal expansion of responding cells for detection. On the other hand, such clonal expansion can amplify and thus enable detection of low-frequency responses that would otherwise be below the detection threshold of the cytokine-based assay. Our experience to date suggests that the CFDA-SE proliferation assay and the rapid cytokine induction assay give similar results in most donors; however, certain epitopes were identified only by one or the other approach (unpublished results). Because the detection of intracellular cytokines in secretion-inhibited cells is a standard method, we do not discuss the basic principles of this technology, which may be found elsewhere [13]. Instead, we focus our discussion on the design of the peptides and peptide pools that may be used for epitope mapping based on intracellular cytokine detection. There are several key issues to be discussed with respect to the peptides including length, overlap between consecutive peptides, and protective groups. These topics are more or less relevant depending on whether CD4 or CD8 epitopes, or both, are defined. Another important point is the intelligent design of peptide pools. When protein sequences are long, the number of overlapping peptides often exceeds the number of single assays that can be run due to a lack of material, which is generally a preparation of peripheral blood cells. As a result, more than one peptide has to be tested per tube. We discuss two intelligent and efficient setups for such pools that increase efficiency by saving material, work, and time. All the issues we discuss in detail apply primarily to the use of intracellular cytokine detection as a readout. To what degree they also apply to the proliferation assay using CFDA-SE has not yet been examined.
2. Configuration of peptides used for epitope mapping The main function of MHC molecules is the presentation of peptides. Class I and class II MHC molecules
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differ in structure, function, and the peptides that may be presented. This is discussed in detail elsewhere [1]. It is interesting to note that class I MHC molecules have more stringent requirements regarding the peptides they bind. It is generally accepted that one important requirement is length (8–10 amino acids), and another one, the absence of protective chemical groups at the N and C termini. Theoretically all peptides found in the binding groove of class I MHC molecules should comply with these demands. In fact, peptides eluted from class I MHC molecules regularly exhibited a length of 8 to 11 amino acids with only few exceptions [1]. The maximum peptide length reported to bind a class I MHC molecule was 13 amino acids. How well a peptide fits into the binding groove is determined by the interactions of the amino acid side chains and the ‘‘back bones’’ of the two binding partners. So-called ‘‘anchors’’ are very important. These are positions in the peptide that can be occupied only by one particular amino acid or only a few, generally similar amino acids (e.g., aliphatic), for successful binding of peptide and MHC. Almost all MHC-presented peptides have at least two anchor positions. One of them is usually the C-terminal position, and with few excep-
tions, the second is position 2 (counting from the N terminus). [Refer to Rammensee et al. [1] for more details.] Although the binding requirements seem to be quite strict, it is clear that class I-restricted responses can be effectively elicited with peptides that are longer than 13 amino acids. The relative efficiency of 9- and 15-mer peptides in stimulating specific CD8+ T cells is shown in Fig. 1. Note that while the 9-amino-acid peptide generated a maximal response at considerably lower doses than the two 15-amino-acid peptides containing the respective 9-mer sequence, the 15-mers could stimulate near-maximal responses at a concentration of 1 lg/ml. Additional experiments (not shown) confirmed that the efficiency of stimulation is reduced as more amino acids are added, making 15-amino-acid peptides the longest that can reliably elicit CD8+ T-cell responses. Unlike class I MHC molecules, class II MHC molecules seem to tolerate much greater variability with respect to peptide length and protective groups at the C and N termini. It has been shown in crystal models that the class II MHC binding groove is ‘‘open’’ at both ends and thus peptides may stick out on both sides. As a result, the presence or absence of protective chemical groups does not seem to matter. In fact, class II
Fig. 1. PBMC from a CMV-seropositive donor were stimulated with one 9-amino-acid peptide and two different 15-amino-acid peptides all including the same known CD8 T-cell epitope. While the 9-amino-acid peptide produces a significant close-to-maximum stimulation at concentrations of 10 and 100 ng/ml, at least 1 lg/ml is required of the 15-amino-acid peptides to induce the same amount of interferon-c (IFN-c) positive events. Curves show the means from two determinations. Cells were stimulated according to the protocol described in the text.
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MHC-presented peptides have successfully been labeled with fluorochromes or other molecules this way, allowing for the tracking of cells that bind these peptides. Although the majority of these peptides are 13–18 amino acids long (binding takes place along a section of 9 amino acids) binding peptides between 9 and 24 amino acids in length were reported. Binding anchors seem less well defined than for class I MHC molecules [1]. Fig. 2 shows an example of dose–response curves obtained with three different 15-amino acid peptides that partly overlap and all induce a CD4 T-cell response. When using peptides for epitope mapping one should be satisfied that the set of peptides one uses can effectively stimulate the T-cell population in question. We found that in our assay peptides 15 amino acids in length can effectively stimulate both CD4 and CD8 T cells, allowing the search for CD4 and CD8 T-cell epitopes at the same time. Moreover the use of 15-mer peptides is to be preferred for practical and cost reasons: the number of peptides required to cover a complete protein sequence is smaller the longer the peptide used. Fewer peptides translate into lower cost and to the ability to complete epitope deconvolution on smaller samples of blood.
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The overlap between neighboring peptides is important with respect to the ability of a peptide scan to cover all possible epitopes along a given amino acid sequence. This overlap should not be shorter than the typical length of class I MHC-presented peptides (i.e., 9 amino acids) minus one (i.e., 8 amino acids). An overlap of 8 amino acids guarantees that all possible stretches 9 amino acids long are contained in the scan. For class II MHC-presented peptides this overlap is too short, since the typical epitope length exceeds 9 amino acids. Although most core epitopes are described to be 9 amino acids long there is evidence that flanking amino acids have a role in the recognition of class II MHC presented peptides by CD4 T cells. Fig. 3 shows our currently preferred setup for peptide scans: 15 amino acid peptides with 11 overlaps.
3. Intelligent design of peptide pools While the advantages and disadvantages associated with the use of different types of peptides (length, protective groups, etc.) were discussed in the previous section, here we discuss the design of peptide pools. The
Fig. 2. PBMC from an HLA-DR11-positive and CMV-seropositive donor were stimulated with three different but consecutive 15-amino-acid peptides covering an epitope-containing amino acid stretch of the CMV pp65 protein. While all three peptides are stimulating, their efficiency is quite variable. The middle peptide (EHPTFTSQYRIQGKL) best covers two possible theoretical sequences of HLA-DR11 presented epitopes contained in the concerned section of the pp65 sequence, including flanking amino acids at both the N and C termini (EHPTFTSQYRIQGKL or EHPTFTSQYRIQGKL, potential anchor amino acids in boldface type). This peptide is much more efficient than the other two peptides. Cells were stimulated according to the protocol described in the text.
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Fig. 3. Schematic representation of a peptide scan covering an amino acid sequence with peptides 15 amino acids in length and with four overlaps. Boxes represent amino acids.
objective of using peptide pools is to determine which peptides are stimulating without having to test every single peptide individually. This is a crucial issue as it essentially determines whether an analysis can be completed with a given amount of donor material. Whichever peptide length and overlap have been chosen, once the complete set of peptides has been obtained, one needs to find a way of testing them all on a certain (and generally limited) amount of cell suspension. We recommend using between 500,000 and 1,000,000 PBMC for each test, since a substantial percentage of cells are lost during the washing steps. To detect T cells that are antigen-specific but occur at low frequencies many T cells are needed to begin with. Assuming that 50% of T cells are lost during the procedure, starting with 500,000 PBMC would result in with 250,000 PBMC for flowcytometric analysis. Given a typical distribution in which T cells constitute about 70% of the PBMC and the CD8 subset about 35% of the T cells, a theoretical number of just over 60,000 CD8 T cells can be measured. If those T cells specific for a given peptide occur at a frequency of 0.01% (1/10,000), about 6 positive cells would be expected and it would be hard to argue this was a positive response. As a result, having more cells at the start is an advantage. In our experience 1 ml of blood generally yields between one and two million PBMC in healthy donors. Smaller numbers are obtained if the white blood cell counts (WBC) of donors (or patients) are low. We generally reckon that approximately 1 ml of blood is required for testing one peptide or peptide pool. The proteins we have chosen to analyze in the past were IE-1 and the pp65 proteins of human cytomegalovirus strain AD 169 (Swiss-Prot Accession Nos. P13202 and P06725, respectively). These protein sequences have total lengths of 491 and 561 amino acids, respectively, and peptide sets (15 amino acids length/11 amino acids overlap) include 120 and 138 peptides, respectively. Testing each peptide individually would require 120 106 or 138 106 PBMC. Because of this problem, we were obliged to group our peptides into
pools to save donor material, work, and money, just like many scientists in the past who performed epitope mapping using more conventional approaches. By far the most efficient way seemed to be a design in which peptide pools overlap in such a way that each individual peptide is contained in exactly two pools and the number of pools is minimum. The configuration that complies best with both demands is a square or rectangular matrix (Fig. 4A). For a given number of peptides, the next higher square number may be chosen (if the number is not a square number itself) and the square root of this number multiplied by 2 is the number of pools. For example for the IE-1 peptide set of 120 peptides, the closest higher square number is 121. So the ideal number of pools is 11 2 ¼ 22 (Fig. 4A). Some positions in such a square matrix can be left blank; sometimes it is a whole line (so that the resulting matrix in fact turns out rectangular). With such a matrix, the intersections of positive pools correspond to the stimulating peptides. These individual peptides, however, in this first step merely represent candidate stimulating peptides if two or more vertical and horizontal pools are positive. With two horizontal and two vertical pools positive, for example, there will be four intersections, even though only two peptides may in fact be stimulating. Such candidate peptides should be tested individually for confirmation. If PBMC are used, an aliquot of the cell suspension may be kept (unstimulated) in the incubator for this purpose for up to 48 h. Using whole blood by contrast, storage for 48 h may be problematic and produce artifacts. Because of the pool setup the number of tests to be run is reduced from 120 to 22 in the example plus a small number that depends on how many candidate peptides were identified. The ideal design of peptide pools, however, also depends on the number of epitopes that are eventually found. In a more recent project we have used a threedimensional design for the peptide pools, which in fact corresponds to a cube (Fig. 4B). So rather than having
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Fig. 4. Two- or three-dimensional pool setup. Using a two- (A), or three (B)-dimensional setup for mapping pools of patient/donor material, reagents and time may be saved. (A) Pools are represented by the (horizontal) lines and (vertical) columns of a table, each containing 10 or 11 peptides. Each number represents one peptide. The points of intersection between stimulating pools represent the stimulating peptides. (B) Pools are represented by horizontal, sagittal, or frontal sections of a cube, each containing 25 or 30 peptides. As in (A) the points of intersection between pools represent stimulating peptides. (C) Examples of positive responses to pools and the unstimulated control.
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vertical and horizontal pools, we have slices of a cube that are either horizontal, frontal, or sagittal (to use anatomical descriptions). This design ensures that each intersection of a horizontal with a frontal and a sagittal pool points to precisely one single peptide and vice versa, that any positive peptide results in exactly three pools testing positive. With numbers of positive peptides ranging from 1 to 3 (as mostly the case with both CMV proteins we have tested) this design is highly efficient, as the number of pools (and thus the amount of material required) is further reduced and interpretation of the results including the selection of peptides to be confirmed is still manageable (Fig. 5). Thus, all IE-1 and pp65 peptides could be tested using just 5 þ 5 þ 5 ¼ 15 ð5 5 5 ¼ 125!Þ and 5 þ 5 þ 6 ¼ 16 ð5 5 6 ¼ 150!Þ peptide pools, respectively. This increase in efficiency is of advantage for the analysis of at-risk populations, for example, HIV patients or organ transplant recipients, since in most cases no more than 20 ml of citrated blood is required for a complete mapping. Increasing the number of peptides per pool and using a more complex setup of pools, however, is not always going to make epitope mapping more efficient. In donors responding to multiple epitopes in a given protein, using the complex ‘‘cube’’ approach, i.e., a threedimensional pool setup, will result in a large number of pools giving positive responses. In this situation, the number of tests required to confirm the candidate peptides may exceed by far the number of tests that would have been required for confirmation if only the square matrix, i.e., a two-dimensional setup, had been used. In such a scenario, however, candidate peptides can again be combined in a two-dimensional pool setup and individual candidate peptides can be confirmed in a third step.
4. Protocol for compiling peptide pools We generally recommend that peptides be dissolved in dimethyl sulfoxide (DMSO). Before compiling peptide pools from given peptide stock solutions one should work out how many peptides can be combined in one pool without the end concentration of the solvent becoming toxic in the assay. For peptides dissolved in pure DMSO the volume of peptide solution added to each test tube should not exceed 1% of the total test volume at any time. We recommend the following steps: • Work out number of peptides to be combined each pool. • Verify the concentrations of each of the peptide stock solutions to be combined in a given pool. • Calculate at least 1 lg/ml final concentration of each peptide per test (using PBMC! For whole blood higher concentrations may be required).
• Work out resulting volume of peptide solution to be added per test to achieve the desired concentration. • Reduce number of peptides per pool if the volume of peptide solution added exceeds 1% of the test volume. For solvents other than DMSO we cannot give detailed recommendations. The maximum amount of solvent added depends on its toxicity or its effect on the biological response. Tip. It is of advantage to compile the peptide pools in the same session as the peptides are first dissolved. For dissolving the peptides DMSO may be added to the freeze-dried peptides in fixed increments of volume until peptides are dissolved (all calculations must be done before starting). For each peptide a 10 mg/ml working solution may be prepared immediately. Once dissolved, each peptide solution should be placed on ice as soon as no more aliquoting from this solution is required (generally all aliquots required from a given peptide solution should be taken immediately). The peptide pools are then compiled from 10 mg/ml solutions (sufficient for very big peptide pools). Pools-to-be may be kept on ice in cryotubes and completed step by step as the individual peptides pertaining to each pool are dissolved and aliquoted. Because DMSO is not fluid on ice, the peptides are layered into the tube. Therefore, once completed, pools must be warmed slightly until fluid and mixed (vortex) prior to making aliquots (make one final aliquot per test!). DMSO may be added to completed pools to obtain identical volumes for all pools if desired. The aim should be aliquots that can be handled easily and quickly (rather 5 ll than one!). If the concentrations of the individual peptides in solvent are high enough, all peptides of a given scan can be combined in one single pool (‘‘total pool’’). Responses to such total pools reflect the sum of the responses to individual epitopes. They may be used as a positive control or as a screening prior to looking for individual epitopes [14,15].
5. Protocol for stimulation and intracellular cytokine detection We generally use heparinized or citrated blood and prepare PBMC by density gradient centrifugation using Ficoll–Paque. This protocol is for PBMC and alterations to this protocol may have to be made when using other materials such as, for example, bronchoalveolar lavage (BAL) fluid, whole blood, and joint fluid aspirates. Following gradient centrifugation cells are washed twice with sterile phosphate-buffered saline (PBS) and resuspended in ‘‘supplemented’’ RPMI-1640 medium
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Fig. 5. Example of a complete mapping procedure accommodating 120 peptides in a three-dimensional setup with five horizontal, five frontal, and five sagittal sections (5 5 5 ¼ 125). Candidate peptides for confirmation are chosen from the intersections of overlapping positive pools. Dot plots show results with all 15 pools, an IE-1 total pool (containing all peptides at once), and an unstimulated control sample. Percentages indicate IFN-c-positive events (highlighted dark). Tables on the right represent the frontal sections (I–V, see also Fig. 4). The points of intersection with the other pools (VII, VIII, and XI) are indicated by gray shading. Peptides 77 and 78 are identified as stimulating peptides.
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containing 2 mmol/L L -glutamine, 10% (v/v) heat-inactivated fetal calf serum (FCS), and 100 IU penicillin/ streptomycin. Place 100 ll of supplemented medium containing one test volume of peptide solution in sterile Falcon 2054 tubes (we generally use a test volume of 4 ll added to 96 ll of supplemented medium). Place 100 ll of supplemented medium containing a corresponding amount of DMSO alone in one tube which will serve as the unstimulated control. Notes. (1) The use of other tubes is possible. Using 15-ml conical polypropylene tubes largely prevents cell adhesion to the tube walls, so that the step using EDTA (see below) to detach cells may be left out. On the other hand, the advantage of using the Falcon 2054 tube is that the whole assay, including sample acquisition, may be performed in the same tube. (2) Placing the peptides or peptide pools in the tubes may be time-consuming, especially if many different peptides or peptide pools are used. Doing this first ensures that the cells in all tubes are stimulated for the same period once added quickly and more or less at the same time by a dispenser pipet. • Place 400 ll of cell suspension (containing 500,000 to 1,000,000 cells, i.e., 1.25 to 2:5 106 cells /ml) in each tube. • Place tubes in a rack and place the rack in the incubator (37 C, humidified CO2 atmosphere) at a 5 slant (tubes almost horizontal). Note. The final concentration of each peptide should be at least 1 lg/ml. The DMSO concentration should not exceed 1% (v/v) at any time. • After 2 h add 500 ll of supplemented medium containing 10 lg of brefeldin A (BFA) to each tube. BFA solution should be freshly prepared. • Return rack to the incubator and make sure that each tube is in the same slanting position as before (if a tube has turned, adherent cells may now be dry without medium covering them). Note. The final concentration of BFA should be 10 lg/ml. BFA stock solution of 5 mg/ml is prepared using DMSO and kept at )80 C. Two microliters of this stock solution added in 498 ll of supplemented medium will provide an addition of 10 lg of BFA to the assay. Tip. Label each tube and turn it so that all labels are either up or down when in slanting position. Prepare one batch of BFA in supplemented medium sufficient for all assay tubes shortly before adding the respective volume to each tube. • After an additional 4 h add 3 ml of ice-cold PBS to each tube. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid.
• Add 3 ml of PBS containing 2 mM EDTA. Make sure the area of the tube wall that may have adherent cells on it is covered with fluid (especially if you use tubes other than the above, 3 ml may not be sufficient). • Incubate all tubes for 10 min at 37 C (water bath). • Vortex at low speed for 30 s. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. • Add 1 ml of washing buffer ([PBS containing 0.5% (w/ v) bovine serum albumin (BSA) and 0.1% (w/v) sodium azide (NaN3 ).] • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. Note. Blotting tubes dry ensures that the remaining fluid is minimum and does not dilute the reagents added afterward to variable degrees. • Add 1 ml of BD lysing solution to each tube and incubate for 10 min at room temperature. Or add another fixing solution according to the manufacturerÕs instructions or use 1 ml of 4% paraformaldehyde in PBS solution and incubate tubes for 5 min at 37 C in a water bath. • Add at least 3 ml of washing buffer. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. • Add 0.5 ml of permeabilizing solution II to each tube and incubate for 10 min at room temperature. Or, use other permeabilizing solutions according to the respective manufacturerÕs instructions. Note. It is important that you follow the instructions of the manufacturer when using fixation and permeabilizing reagents. It is advisable also to use staining antibodies that were tested with these respective systems. It is probably safest to follow the manufacturerÕs recommendations on the combination of fixatives, permeabilizing reagents, and staining antibodies. • Add 3 ml of washing buffer. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Proceed to antibody staining according to manufacturerÕs instructions or your own protocols for intracellular staining. Tip. We prefer to stain in a total volume of 100 ll using a premixed ‘‘cocktail’’ of antibodies generally combining anti-IFN-c–FITC, anti-CD69-PE, anti-CD4PerCP, and anti-CD4 or anti-CD8-APC, or replacing anti-CD69-PE with anti-CD4 or anti-CD8-PE. Applying an antibody cocktail is a great way of saving time compared with adding all antibodies separately. Note. PerCP conjugates are available from BD only; all other antibodies can be obtained from a variety of companies of your choice.
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5.1. Materials required (materials for PBMC preparation are not listed) DMSO-resistant cryotubes for peptide storage: no recommendation Falcon 2054 tubes (BD, Heidelberg, Germany) Racks that hold tubes in a fixed position (Nalgene, USA; for BD 2054 tubes) Dispenser bottles and dispenser pipets (recommended for wash buffer, EDTA buffer, cell suspension, etc., especially if many tubes are run in parallel): no recommendation
5.2. Reagents required
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CD4+, CD8+, or both T-cell subsets, (2) population percentages, (3) the degree of CD3/CD8 and CD3/CD4 receptor downregulation (look at the geometric mean of the corresponding fluorescence parameter), and (4) finally the ‘‘brightness’’ (IFN-c staining) of the responding population (i.e., amount of cytokine induced). Because the ‘‘brightness’’ clearly depends on the quality of cell permeabilization and the degree of antibody penetration, this criterion must be used with caution and is based on the assumption that within one experimental run, permeabilization and staining conditions were the same in all tubes. These features tend to be characteristic of specific peptides in certain donors. The combination of these criteria is most often sufficient to reliably identify a stimulating peptide. 5.4. Additional remarks
DMSO, silylation grade (Pierce, Germany) Ficoll–Paque (Pharmacia, Uppsala, Sweden) PBS (Gibco, Paisley, UK) RPMI-1640 medium (Biochrom, Berlin, Germany) L -Glutamine (Biochrom) Fetal calf serum (FCS; Biochrom) Penicillin/streptomycin (Biochrom) Brefeldin A (Sigma, Steinheim, Germany) EDTA (Sigma) (Sigma) BSA (Serva, Heidelberg, Germany) NaN3 (Serva) FACS lysing solution (BD, San Jose, CA, USA) Permeabilizing solution II (BD San Jose, CA, USA) Fluorescein isothiocyanate (FITC)-conjugated antiIFN-c Phycoerythrin (PE)-conjugated anti-CD69 PE-conjugated anti-CD4 Peridinin chlorophyll protein (PerCP)-conjugated anti-CD3 (BD, San Jose, CA, USA) Allophycocyanin (APC)-conjugated anti-CD8
We usually stain with a combination of anti-IFN-c– FITC, anti-CD4-PE, anti-CD3-PerCP, and anti-CD8APC to allow direct staining of potentially responding CD4 and CD8 T-cell subsets. The identified individual peptides may be confirmed using a combination of antiCD4 or anti-CD8-APC and anti-CD69-PE to improve gating of activated cells. One complete epitope mapping experiment in our hands usually comprises the initial peptide identification using the respective pools along with an unstimulated control and a complete mixture (containing all peptides in a set) and the subsequent confirmation of the identified peptides by individual testing, again in combination with an unstimulated control and the complete mixture. The complete mixture may serve as a positive control in donors/patients with known reactivity to the protein, as it contains all possible epitopes. Provided the background (i.e., the number of IFN-c producing cells within a given reference population in the unstimulated sample) is low, the sensitivity of CFC-based epitope mapping is an estimated 2 or even 1 positive cell per 10,000 cells.
5.3. Flow cytometric analysis and interpretation Data acquisition should ensure that no life gates are set that will exclude potentially reactive cells from later analysis. In particular, make sure that lymphocyte life gates (recommended!) are not too tight around the lymphocyte population. Do not use life gates on T cells based on CD3, CD4, or CD8 expression, since these surface molecules may be downregulated on activated cells and thus you may cut through a reactive T-cell population as you set these gates according to your normal experience. Otherwise, acquire as many events as possible to increase the quality of your analysis. Helpful criteria for correct identification of the stimulating peptides are (1) induction of responses in
6. Protocol for CFDA-SE-based proliferation assay for epitope mapping [For CFDA-SE based epitope mapping, we have so far used only the square matrix setup of peptide pools.] Following gradient centrifugation cells are washed twice with sterile phosphate-buffered saline and stained with CFDA-SE. 6.1. CFDA-SE staining of PBMC • Resuspend cells at a concentration of 107 /ml in PBS in a conical polypropylene tube of suitable size. • Add an equal volume of 5 lM CFDA-SE in PBS.
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• Mix gently for 3 min at room temperature. • Add a volume equal to the total volume of PBS containing 10% human AB serum (v/v) to quench unbound CFDA-SE or its deacetylated form, CFSE. 6.2. Proliferation assay • Resuspend CFDA-SE-labeled cells at 1 106 cells/ml in ‘‘supplemented’’ RPMI-1640 medium containing 2 mmol/litre L -glutamine, 10% (v/v) human AB serum (in our hands the use of AB serum is associated with less background proliferation than use of fetal calf serum or autologous serum), and 100 IU penicillin/ streptomycin. • Place 1 ml each of cell suspension (106 cells) in Falcon 2054 tubes. • Add peptides or peptide pools at high concentration observing that the final DMSO concentration must remain below 1% and the final peptide concentration should be at least 1 lg/ml). Run unstimulated samples (only the corresponding amount of DMSO if this was used to dissolve the peptides) in parallel as controls. • Incubate tubes upright in a rack in a standard incubator (37 C, humidified CO2 atmosphere). • After 48 h add interleukin (IL)-2 from IL-2 stock solution (105 IU/ml in PBS, keep at )70 C or below) to each tube to a final concentration of 25 IU/ml. • After a total incubation time of 96 to 144 h stop assay by adding 2 ml of ice-cold PBS to each tube. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant.
• Resuspend pellets in remaining fluid. • Add 3 ml of PBS containing 2 mM EDTA. • Incubate all tubes for 10 min at 37 C (water bath). • Vortex at low speed for 30 s. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Add 1 ml of washing buffer (PBS containing 0.5% (w/ v) BSA and 0.1% (w/v) NaN3 . • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. • Proceed to antibody staining. Remember to prepare extra tubes for instrument setup containing only CFSE-stained cells, only PE-stained cells, and only PerCP-stained cells (or other if different dyes are used). This makes it easier to set up PMTs and compensation. Note. CFDA-SE is a potent fluorescent dye that requires careful instrument setup when used in combination with PE and PerCP staining. 6.3. Flow cytometric analysis and interpretation Data acquisition should ensure that no life gates are set that will exclude potentially proliferated cells from later analysis. Do not use life gates on lymphocytes that are too small, allow for increase in size and granularity. Fig. 6 shows an example from a mapping procedure using CFDA-SE. It is sufficient to discriminate ‘‘response’’ from ‘‘no response’’ to identify a stimulating peptide. Working out the precise number of proliferated precursor cells is complicated (12) and unnecessary for identifying epitopes.
Fig. 6. Discrimination between ‘‘response’’ and ‘‘no response’’ using CFDA-SE. The plots show a stimulated (i.e., positive) sample and the respective unstimulated control.
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6.4. Materials required Materials for PBMC preparation are not listed. DMSO-resistant cryotubes for peptide storage: no recommendation Falcon 2054 tubes (BD, Heidelberg, Germany) Racks that hold tubes in a fixed position (Nalgene, USA, for BD 2054 tubes) Dispenser bottles and dispenser pipets (recommended for wash buffer, EDTA buffer, cell suspension, etc., especially if many tubes are run in parallel): no recommendation 6.5. Reagents required CFDA-SE (Molecular Probes, Leiden, The Netherlands) DMSO, silylation grade (Pierce, Germany) Ficoll–Paque (Pharmacia, Uppsala, Sweden) PBS (Gibco, Paisley, UK) RPMI-1640 medium (Biochrom) L -Glutamine (Biochrom) Human AB serum (Biochrom) Penicillin/streptomycin (Biochrom) EDTA (Sigma) BSA (Serva) NaN3 (Serva) PE-conjugated anti-CD4: no recommendation PerCP-conjugated anti-CD3 (BD, San Jose, CA, USA) APC-conjugated anti-CD8: no recommendation
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References [1] H.G. Rammensee, J. Bachmann, S. Stefanovic, MHC Ligands and Binding Motifs, Landes Bioscience, Georgetown, TX, 1997. [2] R.R. Singh, Curr. Opin. Rheumatol. 12 (2000) 399–406. [3] D.J. Diamond, J. York, J.Y. Sun, C.L. Wright, S.J. Forman, Blood 90 (1997) 1751–1767. [4] B. Min, K.L. Legge, L. Li, J.C. Caprio, C.D. Pack, R. Gregg, D. McGavin, D. Slauson, H. Zaghouani, Int. Rev. Immunol. 19 (2000) 247–264. [5] E. Wang, G.Q. Phan, F.M. Marincola, Expert Opin. Biol. Ther. 1 (2001) 277–290. [6] J.D. Altman, P.A.H. Moss, P.J.R. Goulder, D.H. Barouch, M.G. McHeyzer-Williams, J.I. Bell, A.J. McMichael, M.M. Davis, Science 274 (1996) 94–96. [7] H.M. Vordermeier, P.J. Cockle, A.O. Whelan, S. Rhodes, R.G. Hewinson, Clin. Infect. Dis. 30 (2000) S291–S298. [8] F. Kern, I.P. Surel, C. Brock, B. Freistedt, H. Radtke, A. Scheffold, R. Blasczyk, P. Reinke, J. Schneider-Mergener, A. Radbruch, P. Walden, H.D. Volk, Nat. Med. 4 (1998) 975–978. [9] F. Kern, E. Khatamzas, I. Surel, C. Frommel, P. Reinke, S.L. Waldrop, L.J. Picker, H.D. Volk, Eur. J. Immunol. 29 (1999) 2908–2915. [10] S.L. Waldrop, C.J. Pitcher, D.M. Peterson, V.C. Maino, L.J. Picker, J. Clin. Invest. 99 (1997) 1739–1750. [11] F. Kern, T. Bunde, N. Faulhaber, F. Kiecker, E. Khatamzas, I.M. Rudawski, A. Pruss, J.W. Gratama, R. Volkmer-Engert, R. Ewert, P. Reinke, H.D. Volk, L.J. Picker, J. Infect. Dis. 185 (2002) 1709–1716. [12] A.B. Lyons, J. Immunol. Methods 243 (2000) 147–154. [13] V.C. Maino, L.J. Picker, Cytometry 34 (1998) 207–215. [14] F. Kern, N. Faulhaber, C. Fr€ ommel, E. Khatamzas, S. Pr€ osch, C. Sch€ onemann, I. Kretzschmar, R. Volkmer-Engert, H.-D. Volk, P. Reinke, Eur. J. Immunol. 30 (2000) 1676–1682. [15] H.T. Maecker, H.S. Dunn, M.A. Suni, E. Khatamzas, C.J. Pitcher, T. Bunde, N. Persaud, W. Trigona, T.M. Fu, E. Sinclair, B.M. Bredt, J.M. McCune, V.C. Maino, F. Kern, L.J. Picker, J. Immunol. Methods 255 (2001) 27–40.
Mapping T cell epitopes by flow cytometry Bodo Hoffmeister,a Felix Kiecker,a Lydia Tesfa,a Hans-Dieter Volk,a Louis J. Picker,b and Florian Kerna,* a
Department of Medical Immunology, Charit e, Humboldt Universit€at zu Berlin, Schumannstrasse 21, 10117 Berlin, Germany b Vaccine and Gene Therapy Institute, Oregon Health and Science University, Portland, OR 97201, USA Accepted 2 December 2002
Abstract Epitope mapping by flow cytometry is a very modern approach that not only identifies T-cell epitopes but simultaneously allows for detailed analysis of the responding T-cell subsets including lineage, activation marker expression, and other markers of interest. The most frequently used approach is based on the identification of intracellular cytokines in secretion-inhibited activated T cells following stimulation with peptides or peptide pools. A more recently developed assay analyzes T-cell proliferation by measuring the decrease in carboxyfluorescein diacetate succinimidyl ester staining in proliferated cells. This article includes information on peptide configuration, a section on the design and efficient application of peptide pools, and working laboratory protocols for both assays. 2003 Elsevier Science (USA). All rights reserved.
1. Introduction Epitopes are the portions of antigens that bind specifically with the binding site of an antibody or a receptor on a T lymphocyte. While antibodies (B-cell receptors) bind free spatial epitopes, T-cell receptors require their epitopes to be presented on major histocompatibility complex (MHC) molecules in a linear fashion. The T-cell receptor makes contact with portions of the peptide and portions of the MHC molecule so that, strictly speaking, a T-cell epitope comprises both the peptide and determinants of the presenting MHC molecule. A description of the details of antigen presentation by MHC molecules is clearly beyond the scope of this article but can be found elsewhere [1]. Although a very large number of protein sequences are known and stored in databases, for most proteins it is not known whether they are immunogenic (i.e., whether they bind one or more MHC alleles within a population and are recognizable by the T-cell receptor repertoire), and where the antigenic determinants are localized within the primary amino acid sequence. The identification of immunogenic proteins and determination of the T-cell epitopes contained in them often *
Corresponding author.
stand at the very beginning of vaccine development. While many vaccines use ‘‘bulk approaches’’ like whole recombinant proteins (hepatitis B), lysates of infectious agents (Bacillus Calmette–Guerin), complete toxoids (tetanus), there is increasing interest in more individualized peptide-based vaccinations. Possible applications include (but are not limited to) infectious diseases, tumor vaccines, and autoimmunity [2–5]. Whether or not a vaccination is peptide based in its final form, the exact knowledge of relevant epitopes is helpful in studying how a vaccine is applied most efficiently, since known epitopes may be packaged with adjuvants, applied within a longer stretch of amino acids or protein, coded by DNA using free plasmids or various vectors or even modified by amino acid substitutions to generate more immunogenic molecules. Peptide-based vaccinations may induce or modulate both CD4 and CD8 T-cell responses; however, depending on the form of the vaccine (e.g., peptide or protein plus adjuvant or DNA) CD4 or CD8 T cells may be the preferred responding population. The use of defined epitopes in vaccines allows determination of which T-cell subset (helper or cytotoxic/suppressor) is going to respond in most cases, the only exception being peptides that induce both CD4 and CD8 T-cell responses. A quite recent application of epitope identification is in the quantification of epitope-specific T-cell populations by
1046-2023/03/$ - see front matter 2003 Elsevier Science (USA). All rights reserved. doi:10.1016/S1046-2023(02)00349-3
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tetrameric MHC complexes (short: ‘‘tetramers’’) [6]. Such tetramers are composed of four recombinant MHC molecules, each of which is coupled to a biotin molecule and linked by binding to the four biotin binding sites of a fluorochrome-conjugated streptavidin molecule. The MHC molecules all contain the same peptide in their binding groove such that the tetrameric complexes can directly bind to T-cell receptors that recognize the peptide–MHC molecule combination. Such reagents may be used to directly examine T-cell responses at an antigenspecific level. Peptides alone, on the other hand, can be used for effectively stimulating T-cell responses in peripheral blood mononuclear cells (PBMC) or other cell suspensions. A very interesting application is the use of such peptides as ‘‘diagnostic stimulant,’’ i.e., to find out if infection with a certain agent has taken place (and has induced T cells) or not [7]. Whether using tetramers or peptides as a stimulant, T cell responses can be identified, analyzed, and monitored. Both approaches are widely used for monitoring T-cell responses in patients with human immunodeficiency virus (HIV), Epstein–Barr virus (EBV), or cytomegalovirus (CMV) infections. This article is dedicated exclusively to the mapping of T-cell epitopes by flow cytometry [8]. The method described is limited to epitopes within known protein sequences, since the stimulating agents for this approach are peptides that were synthesized according to a given primary protein sequence. The attraction of using a flow cytometer for mapping epitopes lies in the great versatility of this approach with respect to the number of parameters that may be acquired in a single measurement, i.e., cell lineage markers, activation markers on the cell surface and inside the cell [8]. The use of many parameters greatly enhances the resolution of the assay, i.e., identification of the T cell subset that responds to a given epitope including lineage and activation markers [9]. The approach we originally designed is based on the detection of rapid cytokine production in activated T cells in a short-term (as short as 6 h) ex vivo assay [10]. The biggest advantage of this approach apart from its unrivaled speed is that CD4 and CD8 T-cell responses can be measured in the same assay and even the same test tube [11]. However, proliferation may also be used as a readout parameter when using flow cytometry. Among the systems for measuring cell proliferation by flow cytometry that are available on the market, we favor the use of carboxyfluorescein diacetate succinimidyl ester (CFDA-SE) staining, because it is simple and reliable. The staining intensity of this protein dye is lost in regular increments as the intracellular protein content is passed on to the next generation by dividing cells. CFDA-SE is a nontoxic, fluorescein-related dye, which is able to permeate cells with the help of its two acetate side chains. Once inside cells, the acetate groups are removed by intracellular esterases and the resultant carboxyfluorescein exits at a much slower rate, allowing
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covalent binding to free amines of cytoplasmatic proteins, forming very stable amino bonds. Because CFSE also reacts with molecules that are rapidly degraded or transported out of the cell, a lot of the CFDA-SE that is initially taken up by cells and transformed into CFSE is lost during the first 24 h following labeling. The remaining portion that is bound to more stable intracellular components may be used for tracking cell proliferation. The use of CFDA-SE staining for the analysis of cell proliferation by flow cytometry was described in 2000 [12]. However, we have only recently started using it for the purpose of epitope identification and we therefore describe only our preliminary protocol. As regards proliferation assays, in general one fundamental difference between them and the detection of intracellular cytokines following short-term stimulation is that the latter does not require clonal expansion of responding cells for detection. On the other hand, such clonal expansion can amplify and thus enable detection of low-frequency responses that would otherwise be below the detection threshold of the cytokine-based assay. Our experience to date suggests that the CFDA-SE proliferation assay and the rapid cytokine induction assay give similar results in most donors; however, certain epitopes were identified only by one or the other approach (unpublished results). Because the detection of intracellular cytokines in secretion-inhibited cells is a standard method, we do not discuss the basic principles of this technology, which may be found elsewhere [13]. Instead, we focus our discussion on the design of the peptides and peptide pools that may be used for epitope mapping based on intracellular cytokine detection. There are several key issues to be discussed with respect to the peptides including length, overlap between consecutive peptides, and protective groups. These topics are more or less relevant depending on whether CD4 or CD8 epitopes, or both, are defined. Another important point is the intelligent design of peptide pools. When protein sequences are long, the number of overlapping peptides often exceeds the number of single assays that can be run due to a lack of material, which is generally a preparation of peripheral blood cells. As a result, more than one peptide has to be tested per tube. We discuss two intelligent and efficient setups for such pools that increase efficiency by saving material, work, and time. All the issues we discuss in detail apply primarily to the use of intracellular cytokine detection as a readout. To what degree they also apply to the proliferation assay using CFDA-SE has not yet been examined.
2. Configuration of peptides used for epitope mapping The main function of MHC molecules is the presentation of peptides. Class I and class II MHC molecules
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differ in structure, function, and the peptides that may be presented. This is discussed in detail elsewhere [1]. It is interesting to note that class I MHC molecules have more stringent requirements regarding the peptides they bind. It is generally accepted that one important requirement is length (8–10 amino acids), and another one, the absence of protective chemical groups at the N and C termini. Theoretically all peptides found in the binding groove of class I MHC molecules should comply with these demands. In fact, peptides eluted from class I MHC molecules regularly exhibited a length of 8 to 11 amino acids with only few exceptions [1]. The maximum peptide length reported to bind a class I MHC molecule was 13 amino acids. How well a peptide fits into the binding groove is determined by the interactions of the amino acid side chains and the ‘‘back bones’’ of the two binding partners. So-called ‘‘anchors’’ are very important. These are positions in the peptide that can be occupied only by one particular amino acid or only a few, generally similar amino acids (e.g., aliphatic), for successful binding of peptide and MHC. Almost all MHC-presented peptides have at least two anchor positions. One of them is usually the C-terminal position, and with few excep-
tions, the second is position 2 (counting from the N terminus). [Refer to Rammensee et al. [1] for more details.] Although the binding requirements seem to be quite strict, it is clear that class I-restricted responses can be effectively elicited with peptides that are longer than 13 amino acids. The relative efficiency of 9- and 15-mer peptides in stimulating specific CD8+ T cells is shown in Fig. 1. Note that while the 9-amino-acid peptide generated a maximal response at considerably lower doses than the two 15-amino-acid peptides containing the respective 9-mer sequence, the 15-mers could stimulate near-maximal responses at a concentration of 1 lg/ml. Additional experiments (not shown) confirmed that the efficiency of stimulation is reduced as more amino acids are added, making 15-amino-acid peptides the longest that can reliably elicit CD8+ T-cell responses. Unlike class I MHC molecules, class II MHC molecules seem to tolerate much greater variability with respect to peptide length and protective groups at the C and N termini. It has been shown in crystal models that the class II MHC binding groove is ‘‘open’’ at both ends and thus peptides may stick out on both sides. As a result, the presence or absence of protective chemical groups does not seem to matter. In fact, class II
Fig. 1. PBMC from a CMV-seropositive donor were stimulated with one 9-amino-acid peptide and two different 15-amino-acid peptides all including the same known CD8 T-cell epitope. While the 9-amino-acid peptide produces a significant close-to-maximum stimulation at concentrations of 10 and 100 ng/ml, at least 1 lg/ml is required of the 15-amino-acid peptides to induce the same amount of interferon-c (IFN-c) positive events. Curves show the means from two determinations. Cells were stimulated according to the protocol described in the text.
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MHC-presented peptides have successfully been labeled with fluorochromes or other molecules this way, allowing for the tracking of cells that bind these peptides. Although the majority of these peptides are 13–18 amino acids long (binding takes place along a section of 9 amino acids) binding peptides between 9 and 24 amino acids in length were reported. Binding anchors seem less well defined than for class I MHC molecules [1]. Fig. 2 shows an example of dose–response curves obtained with three different 15-amino acid peptides that partly overlap and all induce a CD4 T-cell response. When using peptides for epitope mapping one should be satisfied that the set of peptides one uses can effectively stimulate the T-cell population in question. We found that in our assay peptides 15 amino acids in length can effectively stimulate both CD4 and CD8 T cells, allowing the search for CD4 and CD8 T-cell epitopes at the same time. Moreover the use of 15-mer peptides is to be preferred for practical and cost reasons: the number of peptides required to cover a complete protein sequence is smaller the longer the peptide used. Fewer peptides translate into lower cost and to the ability to complete epitope deconvolution on smaller samples of blood.
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The overlap between neighboring peptides is important with respect to the ability of a peptide scan to cover all possible epitopes along a given amino acid sequence. This overlap should not be shorter than the typical length of class I MHC-presented peptides (i.e., 9 amino acids) minus one (i.e., 8 amino acids). An overlap of 8 amino acids guarantees that all possible stretches 9 amino acids long are contained in the scan. For class II MHC-presented peptides this overlap is too short, since the typical epitope length exceeds 9 amino acids. Although most core epitopes are described to be 9 amino acids long there is evidence that flanking amino acids have a role in the recognition of class II MHC presented peptides by CD4 T cells. Fig. 3 shows our currently preferred setup for peptide scans: 15 amino acid peptides with 11 overlaps.
3. Intelligent design of peptide pools While the advantages and disadvantages associated with the use of different types of peptides (length, protective groups, etc.) were discussed in the previous section, here we discuss the design of peptide pools. The
Fig. 2. PBMC from an HLA-DR11-positive and CMV-seropositive donor were stimulated with three different but consecutive 15-amino-acid peptides covering an epitope-containing amino acid stretch of the CMV pp65 protein. While all three peptides are stimulating, their efficiency is quite variable. The middle peptide (EHPTFTSQYRIQGKL) best covers two possible theoretical sequences of HLA-DR11 presented epitopes contained in the concerned section of the pp65 sequence, including flanking amino acids at both the N and C termini (EHPTFTSQYRIQGKL or EHPTFTSQYRIQGKL, potential anchor amino acids in boldface type). This peptide is much more efficient than the other two peptides. Cells were stimulated according to the protocol described in the text.
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Fig. 3. Schematic representation of a peptide scan covering an amino acid sequence with peptides 15 amino acids in length and with four overlaps. Boxes represent amino acids.
objective of using peptide pools is to determine which peptides are stimulating without having to test every single peptide individually. This is a crucial issue as it essentially determines whether an analysis can be completed with a given amount of donor material. Whichever peptide length and overlap have been chosen, once the complete set of peptides has been obtained, one needs to find a way of testing them all on a certain (and generally limited) amount of cell suspension. We recommend using between 500,000 and 1,000,000 PBMC for each test, since a substantial percentage of cells are lost during the washing steps. To detect T cells that are antigen-specific but occur at low frequencies many T cells are needed to begin with. Assuming that 50% of T cells are lost during the procedure, starting with 500,000 PBMC would result in with 250,000 PBMC for flowcytometric analysis. Given a typical distribution in which T cells constitute about 70% of the PBMC and the CD8 subset about 35% of the T cells, a theoretical number of just over 60,000 CD8 T cells can be measured. If those T cells specific for a given peptide occur at a frequency of 0.01% (1/10,000), about 6 positive cells would be expected and it would be hard to argue this was a positive response. As a result, having more cells at the start is an advantage. In our experience 1 ml of blood generally yields between one and two million PBMC in healthy donors. Smaller numbers are obtained if the white blood cell counts (WBC) of donors (or patients) are low. We generally reckon that approximately 1 ml of blood is required for testing one peptide or peptide pool. The proteins we have chosen to analyze in the past were IE-1 and the pp65 proteins of human cytomegalovirus strain AD 169 (Swiss-Prot Accession Nos. P13202 and P06725, respectively). These protein sequences have total lengths of 491 and 561 amino acids, respectively, and peptide sets (15 amino acids length/11 amino acids overlap) include 120 and 138 peptides, respectively. Testing each peptide individually would require 120 106 or 138 106 PBMC. Because of this problem, we were obliged to group our peptides into
pools to save donor material, work, and money, just like many scientists in the past who performed epitope mapping using more conventional approaches. By far the most efficient way seemed to be a design in which peptide pools overlap in such a way that each individual peptide is contained in exactly two pools and the number of pools is minimum. The configuration that complies best with both demands is a square or rectangular matrix (Fig. 4A). For a given number of peptides, the next higher square number may be chosen (if the number is not a square number itself) and the square root of this number multiplied by 2 is the number of pools. For example for the IE-1 peptide set of 120 peptides, the closest higher square number is 121. So the ideal number of pools is 11 2 ¼ 22 (Fig. 4A). Some positions in such a square matrix can be left blank; sometimes it is a whole line (so that the resulting matrix in fact turns out rectangular). With such a matrix, the intersections of positive pools correspond to the stimulating peptides. These individual peptides, however, in this first step merely represent candidate stimulating peptides if two or more vertical and horizontal pools are positive. With two horizontal and two vertical pools positive, for example, there will be four intersections, even though only two peptides may in fact be stimulating. Such candidate peptides should be tested individually for confirmation. If PBMC are used, an aliquot of the cell suspension may be kept (unstimulated) in the incubator for this purpose for up to 48 h. Using whole blood by contrast, storage for 48 h may be problematic and produce artifacts. Because of the pool setup the number of tests to be run is reduced from 120 to 22 in the example plus a small number that depends on how many candidate peptides were identified. The ideal design of peptide pools, however, also depends on the number of epitopes that are eventually found. In a more recent project we have used a threedimensional design for the peptide pools, which in fact corresponds to a cube (Fig. 4B). So rather than having
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Fig. 4. Two- or three-dimensional pool setup. Using a two- (A), or three (B)-dimensional setup for mapping pools of patient/donor material, reagents and time may be saved. (A) Pools are represented by the (horizontal) lines and (vertical) columns of a table, each containing 10 or 11 peptides. Each number represents one peptide. The points of intersection between stimulating pools represent the stimulating peptides. (B) Pools are represented by horizontal, sagittal, or frontal sections of a cube, each containing 25 or 30 peptides. As in (A) the points of intersection between pools represent stimulating peptides. (C) Examples of positive responses to pools and the unstimulated control.
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vertical and horizontal pools, we have slices of a cube that are either horizontal, frontal, or sagittal (to use anatomical descriptions). This design ensures that each intersection of a horizontal with a frontal and a sagittal pool points to precisely one single peptide and vice versa, that any positive peptide results in exactly three pools testing positive. With numbers of positive peptides ranging from 1 to 3 (as mostly the case with both CMV proteins we have tested) this design is highly efficient, as the number of pools (and thus the amount of material required) is further reduced and interpretation of the results including the selection of peptides to be confirmed is still manageable (Fig. 5). Thus, all IE-1 and pp65 peptides could be tested using just 5 þ 5 þ 5 ¼ 15 ð5 5 5 ¼ 125!Þ and 5 þ 5 þ 6 ¼ 16 ð5 5 6 ¼ 150!Þ peptide pools, respectively. This increase in efficiency is of advantage for the analysis of at-risk populations, for example, HIV patients or organ transplant recipients, since in most cases no more than 20 ml of citrated blood is required for a complete mapping. Increasing the number of peptides per pool and using a more complex setup of pools, however, is not always going to make epitope mapping more efficient. In donors responding to multiple epitopes in a given protein, using the complex ‘‘cube’’ approach, i.e., a threedimensional pool setup, will result in a large number of pools giving positive responses. In this situation, the number of tests required to confirm the candidate peptides may exceed by far the number of tests that would have been required for confirmation if only the square matrix, i.e., a two-dimensional setup, had been used. In such a scenario, however, candidate peptides can again be combined in a two-dimensional pool setup and individual candidate peptides can be confirmed in a third step.
4. Protocol for compiling peptide pools We generally recommend that peptides be dissolved in dimethyl sulfoxide (DMSO). Before compiling peptide pools from given peptide stock solutions one should work out how many peptides can be combined in one pool without the end concentration of the solvent becoming toxic in the assay. For peptides dissolved in pure DMSO the volume of peptide solution added to each test tube should not exceed 1% of the total test volume at any time. We recommend the following steps: • Work out number of peptides to be combined each pool. • Verify the concentrations of each of the peptide stock solutions to be combined in a given pool. • Calculate at least 1 lg/ml final concentration of each peptide per test (using PBMC! For whole blood higher concentrations may be required).
• Work out resulting volume of peptide solution to be added per test to achieve the desired concentration. • Reduce number of peptides per pool if the volume of peptide solution added exceeds 1% of the test volume. For solvents other than DMSO we cannot give detailed recommendations. The maximum amount of solvent added depends on its toxicity or its effect on the biological response. Tip. It is of advantage to compile the peptide pools in the same session as the peptides are first dissolved. For dissolving the peptides DMSO may be added to the freeze-dried peptides in fixed increments of volume until peptides are dissolved (all calculations must be done before starting). For each peptide a 10 mg/ml working solution may be prepared immediately. Once dissolved, each peptide solution should be placed on ice as soon as no more aliquoting from this solution is required (generally all aliquots required from a given peptide solution should be taken immediately). The peptide pools are then compiled from 10 mg/ml solutions (sufficient for very big peptide pools). Pools-to-be may be kept on ice in cryotubes and completed step by step as the individual peptides pertaining to each pool are dissolved and aliquoted. Because DMSO is not fluid on ice, the peptides are layered into the tube. Therefore, once completed, pools must be warmed slightly until fluid and mixed (vortex) prior to making aliquots (make one final aliquot per test!). DMSO may be added to completed pools to obtain identical volumes for all pools if desired. The aim should be aliquots that can be handled easily and quickly (rather 5 ll than one!). If the concentrations of the individual peptides in solvent are high enough, all peptides of a given scan can be combined in one single pool (‘‘total pool’’). Responses to such total pools reflect the sum of the responses to individual epitopes. They may be used as a positive control or as a screening prior to looking for individual epitopes [14,15].
5. Protocol for stimulation and intracellular cytokine detection We generally use heparinized or citrated blood and prepare PBMC by density gradient centrifugation using Ficoll–Paque. This protocol is for PBMC and alterations to this protocol may have to be made when using other materials such as, for example, bronchoalveolar lavage (BAL) fluid, whole blood, and joint fluid aspirates. Following gradient centrifugation cells are washed twice with sterile phosphate-buffered saline (PBS) and resuspended in ‘‘supplemented’’ RPMI-1640 medium
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Fig. 5. Example of a complete mapping procedure accommodating 120 peptides in a three-dimensional setup with five horizontal, five frontal, and five sagittal sections (5 5 5 ¼ 125). Candidate peptides for confirmation are chosen from the intersections of overlapping positive pools. Dot plots show results with all 15 pools, an IE-1 total pool (containing all peptides at once), and an unstimulated control sample. Percentages indicate IFN-c-positive events (highlighted dark). Tables on the right represent the frontal sections (I–V, see also Fig. 4). The points of intersection with the other pools (VII, VIII, and XI) are indicated by gray shading. Peptides 77 and 78 are identified as stimulating peptides.
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containing 2 mmol/L L -glutamine, 10% (v/v) heat-inactivated fetal calf serum (FCS), and 100 IU penicillin/ streptomycin. Place 100 ll of supplemented medium containing one test volume of peptide solution in sterile Falcon 2054 tubes (we generally use a test volume of 4 ll added to 96 ll of supplemented medium). Place 100 ll of supplemented medium containing a corresponding amount of DMSO alone in one tube which will serve as the unstimulated control. Notes. (1) The use of other tubes is possible. Using 15-ml conical polypropylene tubes largely prevents cell adhesion to the tube walls, so that the step using EDTA (see below) to detach cells may be left out. On the other hand, the advantage of using the Falcon 2054 tube is that the whole assay, including sample acquisition, may be performed in the same tube. (2) Placing the peptides or peptide pools in the tubes may be time-consuming, especially if many different peptides or peptide pools are used. Doing this first ensures that the cells in all tubes are stimulated for the same period once added quickly and more or less at the same time by a dispenser pipet. • Place 400 ll of cell suspension (containing 500,000 to 1,000,000 cells, i.e., 1.25 to 2:5 106 cells /ml) in each tube. • Place tubes in a rack and place the rack in the incubator (37 C, humidified CO2 atmosphere) at a 5 slant (tubes almost horizontal). Note. The final concentration of each peptide should be at least 1 lg/ml. The DMSO concentration should not exceed 1% (v/v) at any time. • After 2 h add 500 ll of supplemented medium containing 10 lg of brefeldin A (BFA) to each tube. BFA solution should be freshly prepared. • Return rack to the incubator and make sure that each tube is in the same slanting position as before (if a tube has turned, adherent cells may now be dry without medium covering them). Note. The final concentration of BFA should be 10 lg/ml. BFA stock solution of 5 mg/ml is prepared using DMSO and kept at )80 C. Two microliters of this stock solution added in 498 ll of supplemented medium will provide an addition of 10 lg of BFA to the assay. Tip. Label each tube and turn it so that all labels are either up or down when in slanting position. Prepare one batch of BFA in supplemented medium sufficient for all assay tubes shortly before adding the respective volume to each tube. • After an additional 4 h add 3 ml of ice-cold PBS to each tube. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid.
• Add 3 ml of PBS containing 2 mM EDTA. Make sure the area of the tube wall that may have adherent cells on it is covered with fluid (especially if you use tubes other than the above, 3 ml may not be sufficient). • Incubate all tubes for 10 min at 37 C (water bath). • Vortex at low speed for 30 s. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. • Add 1 ml of washing buffer ([PBS containing 0.5% (w/ v) bovine serum albumin (BSA) and 0.1% (w/v) sodium azide (NaN3 ).] • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. Note. Blotting tubes dry ensures that the remaining fluid is minimum and does not dilute the reagents added afterward to variable degrees. • Add 1 ml of BD lysing solution to each tube and incubate for 10 min at room temperature. Or add another fixing solution according to the manufacturerÕs instructions or use 1 ml of 4% paraformaldehyde in PBS solution and incubate tubes for 5 min at 37 C in a water bath. • Add at least 3 ml of washing buffer. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. • Add 0.5 ml of permeabilizing solution II to each tube and incubate for 10 min at room temperature. Or, use other permeabilizing solutions according to the respective manufacturerÕs instructions. Note. It is important that you follow the instructions of the manufacturer when using fixation and permeabilizing reagents. It is advisable also to use staining antibodies that were tested with these respective systems. It is probably safest to follow the manufacturerÕs recommendations on the combination of fixatives, permeabilizing reagents, and staining antibodies. • Add 3 ml of washing buffer. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Proceed to antibody staining according to manufacturerÕs instructions or your own protocols for intracellular staining. Tip. We prefer to stain in a total volume of 100 ll using a premixed ‘‘cocktail’’ of antibodies generally combining anti-IFN-c–FITC, anti-CD69-PE, anti-CD4PerCP, and anti-CD4 or anti-CD8-APC, or replacing anti-CD69-PE with anti-CD4 or anti-CD8-PE. Applying an antibody cocktail is a great way of saving time compared with adding all antibodies separately. Note. PerCP conjugates are available from BD only; all other antibodies can be obtained from a variety of companies of your choice.
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5.1. Materials required (materials for PBMC preparation are not listed) DMSO-resistant cryotubes for peptide storage: no recommendation Falcon 2054 tubes (BD, Heidelberg, Germany) Racks that hold tubes in a fixed position (Nalgene, USA; for BD 2054 tubes) Dispenser bottles and dispenser pipets (recommended for wash buffer, EDTA buffer, cell suspension, etc., especially if many tubes are run in parallel): no recommendation
5.2. Reagents required
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CD4+, CD8+, or both T-cell subsets, (2) population percentages, (3) the degree of CD3/CD8 and CD3/CD4 receptor downregulation (look at the geometric mean of the corresponding fluorescence parameter), and (4) finally the ‘‘brightness’’ (IFN-c staining) of the responding population (i.e., amount of cytokine induced). Because the ‘‘brightness’’ clearly depends on the quality of cell permeabilization and the degree of antibody penetration, this criterion must be used with caution and is based on the assumption that within one experimental run, permeabilization and staining conditions were the same in all tubes. These features tend to be characteristic of specific peptides in certain donors. The combination of these criteria is most often sufficient to reliably identify a stimulating peptide. 5.4. Additional remarks
DMSO, silylation grade (Pierce, Germany) Ficoll–Paque (Pharmacia, Uppsala, Sweden) PBS (Gibco, Paisley, UK) RPMI-1640 medium (Biochrom, Berlin, Germany) L -Glutamine (Biochrom) Fetal calf serum (FCS; Biochrom) Penicillin/streptomycin (Biochrom) Brefeldin A (Sigma, Steinheim, Germany) EDTA (Sigma) (Sigma) BSA (Serva, Heidelberg, Germany) NaN3 (Serva) FACS lysing solution (BD, San Jose, CA, USA) Permeabilizing solution II (BD San Jose, CA, USA) Fluorescein isothiocyanate (FITC)-conjugated antiIFN-c Phycoerythrin (PE)-conjugated anti-CD69 PE-conjugated anti-CD4 Peridinin chlorophyll protein (PerCP)-conjugated anti-CD3 (BD, San Jose, CA, USA) Allophycocyanin (APC)-conjugated anti-CD8
We usually stain with a combination of anti-IFN-c– FITC, anti-CD4-PE, anti-CD3-PerCP, and anti-CD8APC to allow direct staining of potentially responding CD4 and CD8 T-cell subsets. The identified individual peptides may be confirmed using a combination of antiCD4 or anti-CD8-APC and anti-CD69-PE to improve gating of activated cells. One complete epitope mapping experiment in our hands usually comprises the initial peptide identification using the respective pools along with an unstimulated control and a complete mixture (containing all peptides in a set) and the subsequent confirmation of the identified peptides by individual testing, again in combination with an unstimulated control and the complete mixture. The complete mixture may serve as a positive control in donors/patients with known reactivity to the protein, as it contains all possible epitopes. Provided the background (i.e., the number of IFN-c producing cells within a given reference population in the unstimulated sample) is low, the sensitivity of CFC-based epitope mapping is an estimated 2 or even 1 positive cell per 10,000 cells.
5.3. Flow cytometric analysis and interpretation Data acquisition should ensure that no life gates are set that will exclude potentially reactive cells from later analysis. In particular, make sure that lymphocyte life gates (recommended!) are not too tight around the lymphocyte population. Do not use life gates on T cells based on CD3, CD4, or CD8 expression, since these surface molecules may be downregulated on activated cells and thus you may cut through a reactive T-cell population as you set these gates according to your normal experience. Otherwise, acquire as many events as possible to increase the quality of your analysis. Helpful criteria for correct identification of the stimulating peptides are (1) induction of responses in
6. Protocol for CFDA-SE-based proliferation assay for epitope mapping [For CFDA-SE based epitope mapping, we have so far used only the square matrix setup of peptide pools.] Following gradient centrifugation cells are washed twice with sterile phosphate-buffered saline and stained with CFDA-SE. 6.1. CFDA-SE staining of PBMC • Resuspend cells at a concentration of 107 /ml in PBS in a conical polypropylene tube of suitable size. • Add an equal volume of 5 lM CFDA-SE in PBS.
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• Mix gently for 3 min at room temperature. • Add a volume equal to the total volume of PBS containing 10% human AB serum (v/v) to quench unbound CFDA-SE or its deacetylated form, CFSE. 6.2. Proliferation assay • Resuspend CFDA-SE-labeled cells at 1 106 cells/ml in ‘‘supplemented’’ RPMI-1640 medium containing 2 mmol/litre L -glutamine, 10% (v/v) human AB serum (in our hands the use of AB serum is associated with less background proliferation than use of fetal calf serum or autologous serum), and 100 IU penicillin/ streptomycin. • Place 1 ml each of cell suspension (106 cells) in Falcon 2054 tubes. • Add peptides or peptide pools at high concentration observing that the final DMSO concentration must remain below 1% and the final peptide concentration should be at least 1 lg/ml). Run unstimulated samples (only the corresponding amount of DMSO if this was used to dissolve the peptides) in parallel as controls. • Incubate tubes upright in a rack in a standard incubator (37 C, humidified CO2 atmosphere). • After 48 h add interleukin (IL)-2 from IL-2 stock solution (105 IU/ml in PBS, keep at )70 C or below) to each tube to a final concentration of 25 IU/ml. • After a total incubation time of 96 to 144 h stop assay by adding 2 ml of ice-cold PBS to each tube. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant.
• Resuspend pellets in remaining fluid. • Add 3 ml of PBS containing 2 mM EDTA. • Incubate all tubes for 10 min at 37 C (water bath). • Vortex at low speed for 30 s. • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Add 1 ml of washing buffer (PBS containing 0.5% (w/ v) BSA and 0.1% (w/v) NaN3 . • Centrifuge (430g, 8 min, 4 C) and decant or aspirate supernatant. • Resuspend pellets in remaining fluid. • Proceed to antibody staining. Remember to prepare extra tubes for instrument setup containing only CFSE-stained cells, only PE-stained cells, and only PerCP-stained cells (or other if different dyes are used). This makes it easier to set up PMTs and compensation. Note. CFDA-SE is a potent fluorescent dye that requires careful instrument setup when used in combination with PE and PerCP staining. 6.3. Flow cytometric analysis and interpretation Data acquisition should ensure that no life gates are set that will exclude potentially proliferated cells from later analysis. Do not use life gates on lymphocytes that are too small, allow for increase in size and granularity. Fig. 6 shows an example from a mapping procedure using CFDA-SE. It is sufficient to discriminate ‘‘response’’ from ‘‘no response’’ to identify a stimulating peptide. Working out the precise number of proliferated precursor cells is complicated (12) and unnecessary for identifying epitopes.
Fig. 6. Discrimination between ‘‘response’’ and ‘‘no response’’ using CFDA-SE. The plots show a stimulated (i.e., positive) sample and the respective unstimulated control.
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6.4. Materials required Materials for PBMC preparation are not listed. DMSO-resistant cryotubes for peptide storage: no recommendation Falcon 2054 tubes (BD, Heidelberg, Germany) Racks that hold tubes in a fixed position (Nalgene, USA, for BD 2054 tubes) Dispenser bottles and dispenser pipets (recommended for wash buffer, EDTA buffer, cell suspension, etc., especially if many tubes are run in parallel): no recommendation 6.5. Reagents required CFDA-SE (Molecular Probes, Leiden, The Netherlands) DMSO, silylation grade (Pierce, Germany) Ficoll–Paque (Pharmacia, Uppsala, Sweden) PBS (Gibco, Paisley, UK) RPMI-1640 medium (Biochrom) L -Glutamine (Biochrom) Human AB serum (Biochrom) Penicillin/streptomycin (Biochrom) EDTA (Sigma) BSA (Serva) NaN3 (Serva) PE-conjugated anti-CD4: no recommendation PerCP-conjugated anti-CD3 (BD, San Jose, CA, USA) APC-conjugated anti-CD8: no recommendation
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