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Immunology. Mar 2008; 123(3): 305–313.
PMCID: PMC2433336

Class II major histocompatibility complex tetramer staining: progress, problems, and prospects


The use of major histocompatibility complex (MHC) tetramers in the detection and analysis of antigen-specific T cells has become more widespread since its introduction 11 years ago. Early challenges in the application of tetramer staining to CD4+ T cells centred around difficulties in the expression of various class II MHC allelic variants and the detection of low-frequency T cells in mixed populations. As many of the technical obstacles to class II MHC tetramer staining have been overcome, the focus has returned to uncertainties concerning how oligomer valency and T-cell receptor/MHC affinity affect tetramer binding. Such issues have become more important with an increase in the number of studies relying on direct ex vivo analysis of antigen-specific CD4+ T cells. In this review we discuss which problems in class II MHC tetramer staining have been solved to date, and which matters remain to be considered.

Keywords: avidity, CD4, flow cytometry, multimer, oligomer, T cell


Analysis of antigen-specific T cells by flow cytometry using fluorescent oligomers of major histocompatibility complex (MHC)–peptide complexes, a technique known as ‘MHC tetramer staining’, was introduced in 1996 by Altman et al.1 Tetramer staining offered information that complemented many of the other immunological methods available and it quickly became a standard tool in the cellular immunologist’s toolbox. The procedure, little changed since the original report, allows identification of T cells of interest based on the binding specificity of their cell surface receptors for particular MHC–peptide complexes. Incorporation of MHC tetramer staining into multicolour flow cytometry protocols permits sophisticated phenotypic analysis and/or isolation of antigen-specific cells for further study. The method was originally described for human class I MHC tetramers and CD8+ T cells,1 with reports of application to murine2 and human3 class II MHC molecules and CD4+ T cells appearing not long after. However, broad application of this powerful technology to CD4+ T cells beyond a few well-defined test systems has been slow because of difficulty producing class II MHC-based reagents, a low frequency of CD4+ T cells of interest in many biological contexts, and complications caused by low T-cell receptor (TCR)–MHC avidity. There has been recent progress in overcoming each of these limitations, and several groups now routinely use class II MHC tetramer staining in analysis of human and murine T-cell populations (Table 1, Fig. 1). Recombinant class II MHC–peptide oligomers are available from the National Institutes of Health tetramer facility (http://www.niaid.nih.gov/reposit/tetramer/index.html) and from several commercial sources (Beckman-Coulter, Fullerton, CA;4 Proimmune, Oxford, UK), and it seems that class II MHC tetramer staining may finally be coming of age. Despite this progress, current questions remain about the ability of class II MHC tetramers to stain all relevant CD4+ T cells regardless of activation state or avidity. Several reviews of class II tetramer staining have appeared as the method has become more widely used.59 This review will focus on improvements in recombinant MHC–peptide production, detection of rare T cells, and developing an understanding of avidity modulation in T cells and its impact on MHC tetramer staining. In this review, we will use the term ‘MHC tetramer staining’ generally to refer to flow cytometry-based detection of T cells using MHC–peptide oligomers, including purposely-designed MHC dimers, pentamers and higher order oligomers, as well as biotin–streptavidin-based oligomers that may not have attained full tetrameric occupancy.

Table 1
Class II major histocompatibility complex (MHC) proteins used in MHC tetramer staining of antigen-specific CD4+ T cells
Figure 1
Publications reporting class II major histocompatibility complex (MHC) tetramer staining experiments. Open symbols, publications reporting use of class II MHC oligomers in T-cell staining experiments; closed symbols, publications reporting ex vivo class ...

Production of MHC–peptide complexes

A variety of expression systems have been used to prepare recombinant class II MHC–peptide complexes for use in tetramer studies. In general, recombinant expression systems are required both to subvert the antigen-processing pathways that normally load class II MHC proteins with endogenous peptides and to incorporate an oligomerization tag into the MHC molecule. Escherichia coli expression is the method of choice for production of class I MHC proteins,10 and can provide large quantities of highly purified protein. However, the folding conditions for class II MHC proteins need to be optimized individually for each species and allotype. At present, only four different MHC proteins have been produced by this method: I-E(k),11 human leucocyte antigen (HLA) -DR1,12,13 HLA-DR2a13,14 and HLA-DR4.15,16 The method involves the expression of insoluble (and presumably denatured) MHC subunits in E. coli inclusion bodies, followed by solubilization and folding in vitro in the presence of defined peptides. The set of class II MHC proteins suitable for folding in vitro has not been extended beyond the four noted above, despite considerable effort by several groups. An alternative system in which folded single-chain αβ–peptide constructs are expressed in E. coli1719 or yeast20 has been reported, but not widely adopted.

The first reported, and now most common, recombinant expression system for class II MHC proteins is the coexpression of α- and β-subunits in baculovirus-infected21 or stably transformed22 insect cells. As originally developed for HLA-DR1, this method relied on the lack of antigen processing and loading machinery in insect cells and the stability of the MHC protein in the absence of peptide.23 In this approach, peptide-free or ‘empty’ molecules could be isolated from the culture medium and loaded with defined peptide in vitro. Some other class II MHC proteins could be produced similarly, provided that they were either stable as empty molecules, or were able to bind adventitiously and be stabilized by low-affinity endogenous insect cell peptides.

Other class II MHC allele variants were found to assemble and load peptide inefficiently as soluble recombinant proteins. Insect cell expression of class II MHC molecules was extended to these less stable allotypes by fusion of peptide to the class II MHC β-subunit N terminus via a short linker.24 The covalently attached peptide had easy access to the peptide-binding site and was able to bind to and stabilize the class II MHC protein, in most cases not interfering with T-cell activation. This technique was originally developed for the murine MHC I-Ad, and subsequently extended to other class II MHC proteins from humans, mice and other species (Table 1) that could not be produced in the absence of an associating peptide. For recombinant proteins that incorporate covalent peptide, peptide loading is relatively independent of the host antigen processing machinery, and mammalian expression systems have been used as an alternative to insect cell expression.25 In some cases, coiled-coil ‘leucine zipper’ tails were added to the C termini of the subunits to promote assembly of the native αβ heterodimer26 for MHC allotypes in which covalent peptide alone was insufficient for stabilization of the structure. In many laboratories, such leucine zipper tails are included routinely in class II MHC production (Table 1). In addition to leucine zippers, immunoglobulin (Ig) domains have been used to promote heterodimerization of class II MHC α- and β-subunits.

A drawback of the covalent peptide method is that a separate custom expression vector is required for production of each different MHC–peptide complex. This complicates large-scale applications of tetramer staining, which often require a panel of MHC–peptide complexes comprising the same MHC but different candidate peptide antigens. To circumvent this limitation, the basic procedure can be modified so that the expression vector codes for a ‘stuffer’ peptide. This ‘stuffer’ peptide is first released through cleavage of a linker attaching it to the MHC, and is then replaced with antigenic peptides of interest in an in vitro peptide-exchange reaction.27 A favourite such stuffer peptide is the 81–104 region of the class II-associated invariant chain, known as CLIP (class II-associated invariant peptide).28,29 This is the region of the invariant chain that occupies the peptide-binding groove of nascent class II MHC proteins expressed in their native cells. HLA-DM, a catalytic peptide-exchange factor required in vivo for efficient exchange of CLIP, has also been used in vitro to promote CLIP exchange and loading of antigenic peptides of interest onto recombinant class II MHC molecules.14 In an interesting twist on this method, Day et al. utilize a hapten-labelled antigenic peptide (carrying an N-terminal dinitrophenyl group), which allows the peptide complexes of interest to be isolated using an anti-hapten antibody.30 For low-to-moderate affinity peptides and many MHC alleles, in vitro peptide-loading reactions may not go to completion; even for covalent peptide approaches, complete loading of the desired peptide may not be guaranteed.31 The hapten-labelling and tracking method represents an important advance, because quantitative peptide loading may be important for the detection of T cells carrying low-affinity TCR.


The tetramer staining method relies on oligomerization of MHC–peptide complexes to overcome the inherently weak MHC–TCR interaction. Typically, the interactions of TCR with cognate MHC–peptide complexes are characterized by micromolar binding affinities and lifetimes in the order of seconds.32,33 This weak binding interaction precludes the use of washing and centrifugation steps common in flow cytometry practice. The conceptual advance of Altman et al.1 in the development of MHC tetramer staining was that oligomeric forms of MHC–peptide complexes could bind simultaneously to multiple TCR present on a T-cell surface, resulting in substantial increases in overall tetramer affinity34 and lifetime.35,36 The original and still most common method of oligomerization involves the introduction of biotin via a short linker attached to the membrane-proximal portion of the soluble MHC molecule. The MHC molecule is expressed with a biotinylation signal peptide sequence, of which an internal lysine can be modified to form biotinyl-lysine in a reaction catalysed by the bacterial biotin ligase BirA.37 Often, the reaction is carried out in vitro on purified MHC proteins, but the BirA enzyme can be coexpressed along with the bsp-tagged MHC in insect cells38 or E. coli (Soren Buus, personal communication), in which case the MHC proteins are biotinylated in vivo and can be used directly without the need for in vitro protein modification. In an alternative approach, biotin can be added by thiol-modification chemistry after the introduction of a cysteine residue at the MHC α or β C-terminus.39,40 Streptavidin is used to oligomerize the biotinylated MHC proteins for staining and flow cytometry. Although unmodified streptavidin has four potential biotin-binding sites, the fluorescent labelling approaches used by commercial suppliers can compromise a fraction of the biotin-binding sites.5 Even fully tetravalent streptavidin may not be able to engage a T cell using all four MHC–peptide complexes simultaneously because of steric constraints. Nonetheless, the exact stoichiometry does not appear to be crucial to the routine use of MHC tetramers as staining reagents, since much of the affinity enhancement can be seen already after dimerization.34 In some cases, class II MHC tetramers can stain antigen-specific T cells after being loaded with simple peptide mixtures, a technique that forms the basis of a tetramer-based epitope screening strategy.41 While streptavidin-mediated tetramerization of biotin-modified MHC proteins remains the most popular oligomerization strategy, other techniques for oligomerization of MHC molecules have been reported. These include an assortment of MHC oligomers of various valency assembled using peptide-based crosslinkers,42 and MHC–immunoglobulin dimers, which have been expressed in both insect and mammalian cells.25,43

Tetramer staining of low-frequency T cells

One of the issues complicating class II tetramer analysis of CD4+ T cells is their relatively low abundance in many sample populations of interest. With class I MHC tetramer staining, even from the initial report, it was possible to detect antigen-specific CD8+ T cells in samples of peripheral blood. In that instance, human immunodeficiency virus (HIV)-gag-specific and HIV-pol-specific, HLA-A2-restricted T cells present at ~ 0·1–1% of the total CD8+ T-cell pool were detected in peripheral blood mononuclear cell samples from HIV-infected individuals.1 Such direct staining of peripheral blood samples is now routine in the field of class I MHC tetramer staining. For class II MHC tetramer staining, direct detection of antigen-specific CD4+ T cells in samples of human peripheral blood has been more challenging, although there have been several examples published.3,4449 As early as 2000, Meyer et al. reported class II MHC tetramer staining of of Borrelia-reactive CD4+ T cells in patients with Lyme disease arthritis at frequencies up to 0·1%.3 More recent studies include detection of CD4+ T cells responding to infectious agents,45,46,48 autoantigens,47 allergens44 and tumour antigens,49,50 with frequencies generally in the range of 0·02–0·6% of total CD4+ T cells. However, resting CD4+ T-cell memory populations are typically present at much lower frequencies, beyond the detection limit of standard tetramer staining protocols, and require expansion or enrichment for detection by class II MHC tetramers (see below).

While most tetramer staining studies in humans have relied on the enumeration of T-cell populations present in circulating peripheral blood, higher frequencies might exist in a compartment more directly affected by the immune response of interest. In sampling both the synovial fluid and peripheral blood of patients with resistant Lyme disease arthritis, Meyer et al. found a nearly 33-fold increase in the abundance of OspA-specific CD4+ T cells at the primary site of inflammation.3 Interestingly, the synovial fluid population of CD4+ T cells exhibited a cell surface phenotype distinct from that of peripheral blood CD4+ cells and an increased sensitivity to antigenic stimulation. In another study, peripheral CD4+ memory populations from the lungs and parenchyma of mice infected with Sendai virus were isolated using MHC–immunoglobulin multimers (immunoglobulin dimers further oligomerized with Protein A). These populations were found to display acutely activated but distinct phenotypes.51 A later analysis of mice that were also infected with the Sendai virus used similar MHC–immunoglobulin multimers to show that a remarkable 13% of the activated CD4+ T cells in the lungs were specific for the antigen studied.25 In an analysis of autoreactive CD4+ T cells in experimental autoimmune encephalomyelitis, those cells which migrated to and participated in the central nervous system response to the virus exhibited different phenotypes from those exhibited by the cells active in the periphery.52 Such phenotypic differences among T-cell populations further underscore the careful selection of an appropriate T-cell sampling site.

In many class II MHC tetramer staining protocols, T-cell samples are stimulated ex vivo with antigen to selectively induce proliferation of antigen-specific T cells. Such ex vivo stimulation can substantially increase the detection of low-frequency T cells above the background staining of non-specific cells. However, differential amplification of antigen-specific T cells complicates the determination of their frequency in the original population, which is often the goal of the measurement. A method has been reported53 in which labelling with carboxyfluorescein succinimidyl ester (CFSE) can be used to keep track of the number of divisions resulting from in vitro stimulation. However, the back-calculation of precursor frequency may underestimate the number of antigen-specific precursor cells, some of which might undergo apoptosis or fail to proliferate in response to stimulation.54,55 Another issue with ex vivo expansion of specific CD4+ populations is that it induces a uniform activated cellular functional state and so has the potential to change the T-cell phenotype.56

Recently, several groups have reported tetramer-based enrichment methods that can be used to preselect antigen-specific CD4+ T cells before tetramer staining,30,5658 extending a technique that was first applied to multimer-based sorting of CD8+ T cells.59,60 In the most common variation on this technique, the T cells of interest first are incubated with the MHC tetramer, and then the tetramer-positive population is enriched with magnetic beads before analysis by flow cytometry. In the most common approach, R-phycoerythrin (PE) is used both as the fluorescent label for the tetramer streptavidin core, and also as the capture tag for magnetic anti-PE coated beads. Control experiments established that the frequency of antigen-specific precursors in the original sample could be estimated using the frequency of tetramer-positive cells in the enriched population along with the total numbers of cells pre- and post-enrichment.57 Using magnetic bead enrichment combined with class II MHC tetramer staining, the frequency of CD4+ T cells in peripheral blood responding to particular MHC–peptide antigens was estimated at 1/100 000 or lower in memory cell populations responding to particular influenza,53,57 tuberculosis,46 melanoma,61 hepatitis,30 cytomegalovirus,57 HIV,57 or tumour56 antigens.


Several groups have observed an inability to stain certain antigen-specific CD4+ T cells despite using appropriate class II MHC tetramers6266 (C. Parra-Lopez, J.M. Calvo-Calle, S.S. Vollers, M.E. Patarroyo, E. Nardin and L.J. Stern, in preparation). The lack of staining is not the result of a low frequency of responding cells, because in some cases essentially all the cells in the population are specific for the MHC and peptide antigen tested. This holds true for human T-cell clones,66 murine T-cell hybridomas62,63 and splenocytes from TCR transgenic mice,65 as well as biological samples of mixed T-cell populations expected to contain a substantial population of specific CD4+ T cells64 (M. Calvo-Calle and L.J. Stern, unpublished observations). The MHC–peptide tetramers used were shown to carry the correct antigen, as the non-staining T cells either could be activated by immobilized versions of the class II tetramer65 (Parra Lopez, in preparation), or could be stained after artificial antibody-mediated TCR crosslinking.62 Why these apparently antigen-specific CD4+ T cells could not be stained by their respective class II MHC–peptide tetramers is not clear. Interestingly, this problem seems much less acute for CD8+ T cells, where it is more common to observe functionally inactive T cells that do stain than functionally active cells that do not.

It is possible that CD4+ T cells not stained by class II MHC tetramers might have MHC–TCR affinity that is too weak to support tetramer binding, even after the affinity enhancement resulting from oligomerization. For a set of murine CD4+ hybridomas, class II MHC tetramer staining intensity was shown to correlate with monomeric MHC–TCR affinity,2 although activation and avidity effects that are not apparent in the comparison of related hybridomas might modulate this simple relationship in examinations of mixed T-cell populations. In other studies, class II MHC tetramer binding was correlated with ‘functional avidity’, i.e. the peptide dose required to stimulate a functional response in an antigen-presenting cell assay.32,62 Recent work citing careful measurements of class I MHC staining of CD8+ T cells suggests a surprising sharp dependence on affinity, resulting from the effects of multivalency on overall tetramer association and dissociation kinetics.36,67 The first few affinity measurements for soluble purified MHC proteins and TCR from CD8+ and CD4+ T cells suggested that there might be a systematic difference between class I and class II MHC proteins,32 but as more examples have been collected,68 the difference has become less apparent. In this regard, it should be noted that there are few published biophysical measurements of MHC–TCR affinity for purified monomeric human class II MHC proteins binding to their respective TCR.69 Given the limited data available at present, it is impossible to say whether or not low MHC–TCR affinity per se is an important factor in restricting class II MHC tetramer staining to a subset of responding CD4+ T cells.

We have argued elsewhere that multivalent avidity effects might limit CD4+ T cell staining by class II MHC tetramers.40 To bind an MHC tetramer tightly, multiple TCR on the cell surface must simultaneously engage multiple MHC from a particular tetramer. It is possible that such TCR clustering might be more difficult in some cells than others. Lower avidity in general may be an underlying molecular explanation for the requirement for many CD4+ T cells to stain at elevated temperatures (usually 25 or 37°),40,57 in contrast to CD8+ T cells, which can usually be stained with class I MHC tetramers at 4°. In some cases, the inability of a CD4+ T-cell sample to be stained by a cognate class II MHC tetramer has been overcome by increasing the valency of the overall interaction. This was accomplished either by preclustering the TCR with antibody,62 or by using very highly multivalent MHC reagents, such as synthetic lipid vesicles coated with MHC–peptide complexes65 or fixed staphylococcal A particles coated with MHC–immunoglobulin.6 The MHC-coated semiconductor nanoparticles or ‘quantum dots,’ having approximately 10 MHC per particle, have been developed for class I MHC proteins,70 and might eventually prove useful as highly multivalent class II MHC reagents.

Another possible explanation for differential avidity of CD4+ and CD8+ T cells is the extent to which the CD4 and CD8 cofactors participate in tetramer binding. For CD8+ T cells and class I MHC oligomers, low-affinity TCR require the participation of CD8 for tetramer staining to be observed.36,67,71 However, CD4 is not believed to contribute substantially to class II MHC tetramer binding.2,72 Interestingly, class I MHC tetramers that have been engineered with a reduced CD8 affinity have some of the characteristics of CD4+ T-cell tetramers: they only bind high-affinity clones, and the staining of low-avidity clones is improved at higher temperatures.67 Similarly, a lack of productive coreceptor participation in class II MHC tetramer staining of CD4+ T cells might limit detection to a high-affinity subset. In principle, the differential role of CD8 and CD4 in binding tetramers could be the result of differences in intrinsic monomeric MHC–coreceptor affinity26,7375 or to differences in constraints on coreceptors colocalizing with TCR on the T-cell membrane.76,77 As with questions of differential MHC–TCR affinity in CD4+ and CD8+ T cells, resolution of the issue of differential coreceptor affinity for MHC class I and II proteins may have to wait for experimental characterization of additional examples.


Recent improvements in class II MHC tetramer staining technology have provided solutions to the problems of class II MHC production and low frequency of CD4+ T cells. For a large number of class II MHC proteins of interest, well-defined class II MHC–peptide complexes can be made using standard methods and then oligomerized with streptavidin. Leucine zipper26 and covalent peptide24 approaches have extended recombinant protein production to different species and many allelic variants. A peptide-exchange procedure has been developed for in vitro cleavage and replacement of covalent peptide,30 and it promises to relieve the tedium of cloning new expression vectors for each peptide of interest. Magnetic bead enrichment techniques have been developed to allow phenotypic analysis of very-low-frequency T cells by flow cytometry without substantial phenotypic modification of the responding T-cell population.30,56,57 In many instances, antigen-specific CD4+ T cells can be directly detected in peripheral blood without amplification or enrichment.3,4449,78,79 Expansion of the technology to include more complementary techniques pioneered for class I MHC tetramers appears to be possible, including live cell sorting,3,49,80 combined tetramer and cytokine secretion analysis,81 and in situ staining applications.46,82 Employing such techniques in conjunction with tetramer staining may offer additional, useful information on cell phenotype, location of immune response, and effector function.

In any class II MHC tetramer staining experiment in which mixed populations of CD4+ T cells are analysed, there is the possibility that not all of the relevant antigen-specific T cells will be detected. What fraction of the total population is represented by these non-staining cells, and how they differ from the antigen-specific cells that do stain, is not known at this time. Low functional avidity for antigen seems to be a characteristic of these cells,62 but whether this is the result of low MHC–TCR affinity, reduced TCR clustering, coreceptor effects, or other phenomena, is not known. A few studies of memory T-cell populations have found similar frequencies of bead-enriched class II MHC tetramer-positive cells and interferon-γ-secreting cells,30,57 suggesting that in these cases the non-staining cells are either few or non-functional. In other studies, for example those directed at avidity modulation, autoimmune responses, T-cell cross-reactivity, or functional regulation, the non-staining CD4+ T-cell populations might be of interest. With this caveat, class II tetramer staining is clearly ready to join other more established T-cell assays in the arsenal of the cellular immunologist.


The authors acknowledge helpful discussions with John D. Altman, Mauricio Calvo-Calle, William W. Kwok and Alexander Sigalov. This work was supported by National Institutes of Health grant NIH U19-057319.


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