Epitopes recognized by T cells are a collection of short peptide fragments derived from specific antigens or proteins. Immunological research to study T cell responses is hindered by the extreme degree of heterogeneity of epitope targets, which are usually derived from multiple antigens; within a given antigen, hundreds of different T cell epitopes can be recognized, differing from one individual to the next because T cell epitope recognition is restricted by the epitopes’ ability to bind to MHC molecules, which are extremely polymorphic in different individuals. Testing large pools encompassing hundreds of peptides is technically challenging because of logistical considerations regarding solvent-induced toxicity. To address this issue, we developed the MegaPool (MP) approach based on sequential lyophilization of large numbers of peptides that can be used in a variety of assays to measure T cell responses, including ELISPOT, intracellular cytokine staining, and activation-induced marker assays, and that has been validated in the study of infectious diseases, allergies, and autoimmunity. Here, we describe the procedures for generating and testing MPs, starting with peptide synthesis and lyophilization, as well as a step-by-step guide and recommendations for their handling and experimental usage. Overall, the MP approach is a powerful strategy for studying T cell responses and understanding the immune system's role in health and disease. © 2023 The Authors. Current Protocols published by Wiley Periodicals LLC.

Basic Protocol 1 : Generation of peptide pools (“MegaPools”)

Basic Protocol 2 : MegaPool testing and quantitation of antigen-specific T cell responses

T cell responses are key elements of the body's reaction to vaccination and infection and have likely evolved to provide a synergistic line of defense alongside antibody responses. T cells recognize epitopes, which are peptide fragments derived from the cellular processing of protein antigens, through their T cell receptors (TCRs), which are highly specialized proteins on the surface of T cells (Punt et al., 2018). T cells can be grouped into a series of subsets based on their function or protein/gene expression. There are several types of T cells, with the most prevalent being CD4+ T cells, also known as helper T cells, and CD8+ T cells, which are referred to as cytotoxic T cells or killer T cells.

Because of their key role in adaptive immunity, measuring T cell responses is an important component of vaccine and diagnostic evaluations, immunotherapy applications, and the general study of immunopathology. The most common and well-known methodologies to measure antigen-specific T cell responses are cytokine-based assays [e.g., interferon-gamma release assays, ELISPOT, and intracellular cytokine staining (ICS)] (Tian et al., 2018), tetramer/multimer staining assays (Klenerman et al., 2002), and activation-induced marker (AIM) assays, which are agnostic toward particular cytokine functionality (Dan et al., 2016; Poloni et al., 2023) (see Basic Protocol 2). The different approaches represent the fundamental tensions that exist between ease of performance, robustness, cost, and throughput on the one hand and physiological relevance, complexity, depth of analysis, and granularity of information on the other. A comprehensive discussion of the available techniques is beyond the scope of the present article, and some recent reviews have discussed the topic (Gondre-Lewis et al., 2023; Peters et al., 2020; Sidney et al., 2020).

In the following protocols, we will describe the generation (Basic Protocol 1) and use (Basic Protocol 2) of specialized reagents for probing and characterizing both CD4+ and CD8+ T cell responses. These reagents allow for the efficient elucidation of adaptive T cell immunity against any target of known sequence and in the context of infectious disease, cancer, allergy, and autoimmunity. With the capacity to probe large sequences, the reagents described below are of value for use in epitope discovery and are crucial for vaccine design and diagnostic purposes.

Human T Cell Responses Generally Recognize Multiple Epitopes Restricted by a Diverse Set of HLA Molecules

Measurement of human T cell responses has to contend with the large diversity of epitope targets recognized (Livingstone & Fathman, 1987; Peters et al., 2020). This diversity originates from two main interdependent sources: the diversity of the antigens recognized and the diversity of HLA molecules, which bind and present epitopes recognized by T cells. Importantly, this diversity is consequential because responses to different antigens and epitopes can be directly related to the preservation of T cell responses and the counteracting of immune escape or associated with differential disease outcomes (Dillon et al., 2015; Schulten, Westernberg, et al., 2018; Grifoni & Sette, 2022).

The genes encoding HLA molecules are polygenic and highly polymorphic. In humans, there are three main class I loci (A, B, and C) and several different class II loci (DRA, DRB1, DRB3/4/5, DPA, DPB, and DQA and DQB), resulting in the expression of four different types of HLA class II heterodimers. Because of heterozygosity and the high degree of HLA polymorphism, each individual can express up to 14 different HLA molecules (eight HLA class II and six HLA class I). Given that each HLA molecule has a different set of peptide specificities, HLA polygeny and polymorphism are a powerful force, amplifying the diversity of epitope repertoires recognized in humans (for review, see, e.g., Little & Parham, 1999; Madden, 1995; Parham, 1988; Sidney et al., 2008; Stevanovic, 2002).

At the same time, this large diversity poses unique challenges for approaches to measure T cell responses (McKinney et al., 2013; Nilsson et al., 2021; Sidney et al., 2020). Utilizing just a handful of epitopes and HLA molecules will most likely result in biased and incomplete coverage of the T cell responses and provide a skewed and incomplete assessment of responses. This situation is made more complex by the fact that different HLA variants are expressed at different frequencies in different ethnic groups and geographic regions (Gonzalez-Galarza et al., 2020). Perhaps unsurprisingly, epitopes presented by HLA molecules that are the most frequent in Caucasian populations are the ones that have been most highly characterized to date in the literature. This ethnic bias in coverage, a result of approaches utilizing few epitope specificities, has been repeatedly noted and is a serious limitation that needs to be addressed in developing approaches to study T cell responses in human populations (Sette et al., 2000; Sette et al., 2001; Sidney et al., 2020).

The MegaPool Approach

To address several of the concerns discussed above, including the need to probe for responses to a wide breadth of epitopes and in the context of diverse populations, our group developed the MegaPool (MP) approach (Carrasco Pro et al., 2015; Sidney et al., 2020). This approach, based on testing large pools of peptides and/or epitopes, provides an efficient means for comprehensive analysis of T cell responses in virtually any donor and has many advantages compared to other competing technologies (Table 1). To provide a comprehensive evaluation of diverse targets in a diverse population, it may be necessary to test peptide pools that can encompass several hundred different peptides. However, pooling such large numbers of peptides can result in a stock solution that is relatively diluted when each separate peptide is considered. Moreover, requiring the addition of a relatively large amount of solvent (such as dimethyl sulfoxide, or DMSO) into test cell cultures will contribute to cell toxicity if a sufficiently high amount of each peptide is required (de Abreu Costa et al., 2017; Verheijen et al., 2019). To overcome this limitation, we developed a sequential lyophilization approach that achieves much higher concentrations of each peptide in viable amounts of solvent, thus eliminating or reducing DMSO (or other solvent) toxicity (see Basic Protocol 1).

There are three main approaches to generate the MPs that we routinely utilize in our research. The first corresponds to the use of sets of overlapping peptides spanning the entire sequence of an antigen(s) of interest (Maecker et al., 2001). The second is based on assembling sets of predicted HLA binding epitopes (Peters et al., 2020). Finally, sets can be assembled to consider previously identified and experimentally defined epitopes (Dhanda et al., 2019). Each of these approaches is associated with distinctive advantages and disadvantages.

The use of overlapping peptides is the most comprehensive and unbiased approach, as it does not rely on any predictive algorithm or a priori knowledge of which epitopes are recognized in a given human population and is irrespective of known HLA phenotype. By selecting a 15-mer size overlapping by 10 amino acids, it further allows one to simultaneously assess CD4+ and CD8+ T cell responses in flow cytometry approaches by setting appropriate gating on the responding T cell populations. By definition, however, this approach requires the highest number of peptides; flow cytometry–based techniques, which are less high throughput and cost effective; and consequently, the highest number of cells required for testing if individual epitope identification is further desired.

MPs based on predictive approaches allow one to reduce the number of peptides tested. However, this approach requires careful consideration of HLA polymorphism to ensure good and appropriate population coverage. Furthermore, this approach requires separate pools for CD4+ and CD8+ T cell detection and will also detect a fraction of the total response, depending on the comprehensiveness of the HLA predictions and allele coverage. Additionally, there is the intrinsic limitation that predictive algorithms, while highly effective and accurate in most cases, are not 100% sensitive and specific, meaning that some epitopes may be missed.

Finally, MPs based on curated epitopes are effective because they include epitopes experimentally shown to be recognized by T cells. Epitope sequences can be retrieved by using web databases such as IEDB (https://www.iedb.org). The main limitation of this approach is that, by definition, it relies on the assumption that the epitope repertoire associated with a given indication has already been thoroughly defined in multiple HLA alleles and with accounting for diverse ethnic backgrounds representative of the general worldwide population.

In general, MPs are envisioned for determining antigen-specific T cell responses for any particular indication, such as, but not limited to, allergies, autoimmunity, and bacterial or viral infectious diseases. However, MPs designed with overlapping peptides spanning an individual antigen or a combination of several could be initially used to screen for donor responsiveness, further introduced into an epitope screening pipeline, and sequentially deconvoluted to map CD4+ T cell reactivity for individual peptides. This multi-step approach has been recently employed for genome-wide screening and epitope identification for SARS-CoV-2, common cold coronaviruses, and Bordetella pertussis (da Silva Antunes et al., 2023; Tarke, Sidney, Kidd, et al., 2021; Tarke et al., 2023).

In the following sections, we will give a brief account of the main MPs we have defined for probing and characterizing both CD4+ and CD8+ T cell responses for several different indications (Table 2). These pools exemplify each of the considerations above and demonstrate how they can be efficient tools for characterizing immune responses in a wide range of different immunological contexts.

Allergens

MPs have been used extensively to characterize allergen specific T cell responses. The first use of MPs was associated with the characterization of timothy grass responses, addressing both known and novel allergens (Hinz et al., 2016; Oseroff et al., 2010; Schulten et al., 2013), and was later expanded to a broad collection of allergens (Oseroff, Sidney, Vita, et al., 2012). Further studies investigated Japanese cedar (Oseroff et al., 2016), cockroach (Birrueta et al., 2019; da Silva Antunes, Sutherland, et al., 2021; Dillon et al., 2015; Oseroff, Sidney, Tripple, et al., 2012), dust mite (Hinz et al., 2015; Seumois et al., 2020), murine (da Silva Antunes, Pham, et al., 2018; Grifoni, da Silva Antunes, et al., 2019; Schulten, Westernberg, et al., 2018), and cow milk (Lewis et al., 2023) allergens.

Some of the salient and original applications of MPs include the characterization of T cell responses to antigens identified by immunoproteomic approaches (Oseroff et al., 2017; Schulten et al., 2013) and epitopes differentially recognized in different clinical manifestations (asthma versus rhinitis) (Dillon et al., 2015) or different levels of sensitization (Schulten et al., 2019) and the detection of T cell responses in non-allergic subjects. Interestingly, by probing these reagents, allergic and non-allergic subjects were both found to mount T cell responses but were associated with different phenotypes (da Silva Antunes, Pham, et al., 2018; Grifoni, da Silva Antunes, et al., 2019; Hinz et al., 2016; Schulten, Westernberg, et al., 2018; Yu, Westernberg, et al., 2021). Moreover, specific mouse allergen–derived epitopes were identified by coupling mass spectrometry and bioinformatic approaches (da Silva Antunes, Pham, et al., 2018). Dust mite peptide pools were used to define the transcriptional profiles associated with allergic asthma (Seumois et al., 2020), and timothy grass–specific and cockroach-specific MPs were used to characterize T cell reactivity in the context of allergen-specific immunotherapy (Rudman Spergel et al., 2021; Schulten et al., 2014; Schulten et al., 2016; Schulten, Tripple, et al., 2018). Lastly, timothy grass and mouse allergen MPs were used to study the modulation of T cell responses associated with allergen exposure in non-allergic subjects (Hinz et al., 2016; Yu, Westernberg, et al., 2021) and, most recently, to reveal recognition of allergen-specific responses by gamma delta T cells (Yu, Wang, Garrigan, Sutherland, et al., 2022).

Autoimmunity

MPs have also been utilized in the study of autoimmunity, specifically in the study of neurodegenerative and cardiovascular diseases. In the case of Parkinson's disease (PD), MPs derived from alpha-synuclein (Lindestam Arlehamn et al., 2020; Singhania, Pham, et al., 2021; Sulzer et al., 2017), tau (Lindestam Arlehamn et al., 2019), and other neuronal-associated antigens (Dhanwani et al., 2020) have been studied. Similar studies have been conducted in the context of Alzheimer's disease (AD). The results thus far indicate an autoimmune component in PD, possibly associated with early disease stages (Lindestam Arlehamn et al., 2020), with enhanced reactivity against alpha-synuclein (Sulzer et al., 2017) and possibly other antigens. No increased reactivity against neuronal antigens has thus far been associated with AD (Dhanwani et al., 2020).

In the case of cardiovascular disease, pools of predicted class II binding peptides derived from apolipoprotein B (APOB) were utilized to identify dominant epitopes, and responses to these APOB epitopes correlated with coronary artery disease severity (Roy et al., 2022). In both neurodegenerative and cardiovascular disease applications, an initial restimulation step was necessary to detect antigen-specific responses, potentially reflective of lower frequency of autoimmune T cells compared to other indications.

Bacterial antigens

Several studies have described MPs encompassing different bacterial targets. The first target was Mycobacterium tuberculosis (MTB), for which a genome-wide screen revealed an unprecedented breadth of responses targeting many known and novel antigens (Lindestam Arlehamn et al., 2013). This resulted in the development of the MTB300 MP (Lindestam Arlehamn et al., 2016), which has been utilized in over 14 different studies since 2020 (Chihab et al., 2023; Day et al., 2021; Du Bruyn et al., 2021; Du Bruyn et al., 2022; Foreman et al., 2022; Hoft et al., 2023; Kauffman et al., 2021; Ogongo et al., 2021; Patankar et al., 2020; Pomaznoy et al., 2020; Riou et al., 2020; Riou et al., 2021; Robison et al., 2021; Sakai et al., 2021; White et al., 2021; Wood et al., 2020; Woodworth et al., 2021). In addition to being used primarily for the characterization of human CD4+ T cell reactivity, this pool has also been used, based on the similarity of peptide binding repertoires, to probe reactivity in non-human primates (Foreman et al., 2022; Kauffman et al., 2021; Mothe et al., 2015; Sakai et al., 2021; White et al., 2021; Wood et al., 2020) and mice (Patankar et al., 2020). Additional MPs were designed to explore the relationship with conservation in other Mycobacteria (Lindestam Arlehamn et al., 2022), to perform a quantitative analysis of TCR and epitope repertoire composition (Glanville et al., 2017; Scriba et al., 2017), to study specific gene deficiencies in humans (Kong et al., 2018; Martinez-Barricarte et al., 2018), to study the role of CD8+ T cell responses (Pomaznoy et al., 2020), and to derive immune signatures associated with transcriptional profiles and different disease outcomes (Singhania, Dubelko, et al., 2021).

B. pertussis is another target where MPs have been described and validated (da Silva Antunes et al., 2020; da Silva Antunes et al., 2023; da Silva Antunes, Babor, et al., 2018; da Silva Antunes, Quiambao, et al., 2021). Briefly, a series of studies defined epitopes encoded in the antigens included in the acellular pertussis (aP) vaccine currently in use (Bancroft et al., 2016). These epitopes were used to generate MPs that revealed a long-lasting polarization of responses as a function of the original priming and unveiled transcriptomic profiles associated with human CD4+ T cell responses to vaccine antigens (da Silva Antunes, Babor, et al., 2018), in addition to clonality assessment of TCR repertoires (Singhania, Pham, et al., 2021). Recently, a whole-genome screening of B. pertussis revealed a highly diverse T cell repertoire and identified epitopes derived from antigens not included in the aP vaccine (da Silva Antunes et al., 2023), which resulted in the development of several MPs that aid in the characterization of CD4+ T cell responses to these antigens. Additional studies described and validated CD4+ T cell human epitopes from the tetanus toxoid protein, which is also in the multivalent tetanus diphtheria aP (TDaP) vaccine (da Silva Antunes et al., 2017).

MPs for flaviviruses

In terms of application of the MP approach to viral targets, early efforts focused on flaviviruses, and in particular on dengue virus (DENV). Initial studies using predicted and experimentally confirmed epitopes spanning the entire proteome of the four DENV serotypes defined a wealth of CD4+ and CD8+ epitopes recognized by T cells derived from individuals previously infected with DENV from different endemic areas, such as Sri Lanka and Nicaragua (Grifoni, Angelo, Lopez, et al., 2017; Grifoni, Moore, et al., 2019; Tian et al., 2019; Weiskopf et al., 2014; Weiskopf et al., 2015; Weiskopf et al., 2016); participants in vaccine studies utilizing a live attenuated vaccine (Angelo et al., 2017); a challenge model (Grifoni, Angelo, Sidney, et al., 2017); and HLA transgenic mice (Weiskopf et al., 2011). This led to the generation of MPs of experimentally defined epitopes covering both CD4+ and CD8+ T cell responses, which were validated in endemic different populations such as Sri Lanka, Nicaragua, India, and Brazil (Grifoni, Angelo, Lopez, et al., 2017; Weiskopf et al., 2015).

The analysis of T cell responses associated with infection and vaccination with different flaviviruses was further expanded to other members of this viral family, such as yellow fever (YF), Japanese encephalomyelitis (JEV), West Nile virus (WNV), and Zika virus (ZIKV) (Grifoni, Pham, et al., 2017; Grifoni, Voic, et al., 2020; Mateus, Grifoni, Voic, et al., 2020). These pools were utilized to broadly describe patterns of reactivity and cross-reactivity (Grifoni, Pham, et al., 2017; Schouest et al., 2021). Overall, the results illustrated how the MP approach is broadly applicable to viral infection and vaccination targets.

SARS-CoV-2 and coronaviruses

MPs have played a key role in the study and elucidation of T cell responses to SARS-CoV-2 in the context of infection and vaccination. The first description of CD4+ and CD8+ T cell responses in the context of convalescent individuals defined the key characteristics of a successful immune response utilizing not only MPs of overlapping peptides spanning the entire proteome but also MPs based on the use of bioinformatically predicted epitopes (Grifoni, Sidney, et al., 2020; Grifoni, Weiskopf, et al., 2020). These studies highlighted, as previously described for other systems, that MPs based on predicted HLA binders are remarkably effective and recapitulate ≥50% of the total response, while requiring substantially fewer peptides and cells for the analysis (Grifoni, Weiskopf, et al., 2020).

These SARS-CoV-2 pools were made broadly available to the scientific community by distribution to over 110 laboratories in four different continents and were utilized in a large number of studies, resulting thus far in over 80 publications (Ansari et al., 2021; Ansari et al., 2022; Apostolidis et al., 2021; Arunachalam et al., 2022; Banki et al., 2022; Bhuiyan et al., 2022; Blixt et al., 2022; Boland et al., 2022; Bosteels et al., 2022; Bowen, Addetia, et al., 2022; Bowen, Park, et al., 2022; Brasu et al., 2022; Bueno et al., 2022; Cheon et al., 2021; Chiuppesi et al., 2022; Costa et al., 2022; da Silva Antunes, Pallikkuth, et al., 2021; Dan et al., 2021; Dentone et al., 2022; Galvez et al., 2022; Gao, Cai, Grifoni, et al., 2022; Gao, Cai, Wullimann, et al., 2022; Garcia-Valtanen et al., 2022; Gazzinelli-Guimaraes et al., 2022; Geers et al., 2023; GeurtsvanKessel et al., 2022; Goel et al., 2021; Grifoni et al., 2021; Grifoni, Sidney, et al., 2020; Grifoni, Weiskopf, et al., 2020; Hassan et al., 2020; He et al., 2022; Hsieh et al., 2022; Jin et al., 2021; Keeton et al., 2021; Keeton et al., 2022; Keeton et al., 2023; Lederer et al., 2022; Madelon, Heikkila, et al., 2022; Madelon, Lauper, et al., 2022; Mateus et al., 2021; Mateus, Grifoni, Tarke, et al., 2020; Meckiff et al., 2020; Mele et al., 2021; Melo-Gonzalez et al., 2021; Murugesan et al., 2021; Nelson et al., 2022; Ogbe et al., 2021; Painter et al., 2021; Paul et al., 2022; Peluso et al., 2022; Perez-Gomez et al., 2022; Petrone et al., 2021; Petrone, Picchianti-Diamanti, et al., 2022; Petrone, Tortorella, et al., 2022; Pino et al., 2021; Poon, Byington, et al., 2021; Poon, Rybkina, et al., 2021; Premkumar et al., 2020; Ramirez et al., 2022; Rydyznski Moderbacher et al., 2020; Rydyznski Moderbacher et al., 2022; Schultz et al., 2022; Shaan Lakshmanappa et al., 2021; Singh et al., 2022; Soto et al., 2022; Sun et al., 2022; Tarke, Coelho, et al., 2022; Tarke, Potesta, et al., 2022; Tarke, Sidney, Kidd, et al., 2021; Tarke, Sidney, Methot, et al., 2021; Ukey et al., 2022; Valencia et al., 2022; Van Damme et al., 2020; Vikkurthi et al., 2022; Weiskopf et al., 2020; Williams et al., 2023; Yu, Narowski, et al., 2022; Yu, Wang, Garrigan, Goodwin, et al., 2022; Zhang et al., 2022). Although an in-depth review is beyond the scope of this article, the resultant MPs helped clarify a diverse set of topics and concerns related to SARS-CoV-2, including elucidation of responses in the acute phase of infection, responses to vaccination, breakthrough infections, kinetics and features of responses in the memory phase, responses in vulnerable and immunocompromised individuals, responses in healthcare workers, and responses in children.

Several additional insights into SARS-CoV-2 infection were also addressed with specific MPs. These include the demonstration of cross-reactive pre-existing memory T cell responses (Mateus, Grifoni, Tarke, et al., 2020) and the demonstration that both CD4+ and CD8+ T cell responses are remarkably preserved in the context of the different variants that originated throughout the pandemic (Tarke, Coelho, et al., 2022; Tarke, Sidney, Methot, et al., 2021). Additional MPs were subsequently generated, based on experimentally defined epitopes, and used to derive immunodiagnostic strategies to address vaccination and infection history (Grifoni et al., 2021; Tarke, Sidney, Kidd, et al., 2021; Yu, Wang, Garrigan, Goodwin, et al., 2022). In parallel, specific MPs were also derived to follow responses to other coronavirus species, such as the main common cold coronaviruses (da Silva Antunes, Pallikkuth, et al., 2021; Tarke et al., 2023; Yu, Narowski, et al., 2022).

Broad application to study responses to human viral pathogens

Although perhaps flaviviruses and SARS-CoV-2 represent “poster child” applications of MPs in the study of viral disease, several additional viral indications have been addressed by the MP approach. These include influenza, where pools addressing different subtypes and antigens have been validated (Meckiff et al., 2020; Poon, Byington, et al., 2021; Yu, Grifoni, et al., 2021), and other common respiratory viruses such as rhinovirus (Grifoni, Mahajan, et al., 2019), human metapneumovirus (HMPV) (Meckiff et al., 2020), human parainfluenza virus (HPIV) (Meckiff et al., 2020), and respiratory syncytial virus (RSV) (Yu, Narowski, et al., 2022), as well as herpesviruses such as Epstein-Barr virus (EBV) (Carrasco Pro et al., 2015; Dan et al., 2016), cytomegalovirus (CMV) (Carrasco Pro et al., 2015; Dhanwani et al., 2021), and varicella zoster virus (VZV) (Voic et al., 2020). Recently, MPs were developed and validated for detection of HIV-specific and CD4+ and CD8+ T cells (Al-Kolla et al., 2022). Murine poxvirus epitopes were first described in the mid-2000s, providing a rationale for the characterization of smallpox vaccines (Pasquetto et al., 2005; Tscharke et al., 2005), followed more recently by the design and validation of specific MPs broadly encompassing orthopox- and monkey pox (MPOX)-derived epitopes for monitoring of infection and vaccination (Grifoni et al., 2022).

NOTE : Appropriate informed consent is necessary for obtaining and use of human study material.

This protocol describes how to produce MPs. The initial steps encompass synthesis of peptide sets that are produced and lyophilized, as common practice in the field. They can be purchased from any supplier. However, all MPs described in this protocol were generated from peptides synthesized by TC Peptide Lab or Mimotopes. Each peptide is then solubilized and a pool generated by combining equivalent amounts of each solubilized peptide. The resulting pool is then re-lyophilized. If necessary, the procedure is repeated until a white, reasonably fluffy powder (lyocake) is obtained. Any synthesized peptides can be pooled for use into an MP. Generally, given the high cost of large sets of peptides, purified preparations are not necessary, and the of use crude peptides is acceptable. In the vast majority of cases, these will achieve >70% purity. Further, large-scale synthesis of crude peptide can usually be obtained in microtube racks, which allow efficient dilution of peptide stocks using, for example, multichannel pipettors. Purified peptides are excellent if resources are available or use necessitates. The final MP amount required should be calculated based on the number of samples or assays that need to be tested, but we recommend starting this protocol with 1 to 2 mg per individual peptide.

Materials

• Peptides (TC Peptide Lab or Mimotopes)
• DMSO (Sigma, cat. no. D2650)
• Distilled water
• • Suitable vessel (e.g., 15- or 50-ml conical tube)
• Lyophilization unit (e.g., FreeZone^® Benchtop Freeze Dryer, 4.5L Capacity, Labconco^®, cat. no. 720401000)
• Soft, lint-free cloth or paper towel
• 50-ml tube holder (Fast-Freeze^™ Tube Holder 50 ml, Labconco^®, cat. no. 7379500), –80°C
• Lyophilizer flasks (Fast-Freeze^™ Flask 900 ml, Labconco^®, cat. no. 7540901), –80°C
• Filter paper (Fast-Freeze^™ Filter Paper 50 ml, Labconco^®, cat. no. A-75448)
• Flask adaptors (Fast-Freeze^™ Flask Adaptors 3/4" to 3/4", Labconco^®, cat. no. 7457200)
• Tubes for aliquoting (e.g., 0.2-ml PCR tubes)

Peptide preparation and pooling

1.Resuspend peptides intended for use in MPs at a high concentration (≥20 mg/ml) in 100% DMSO.

2.Pool a uniform and desired amount of each peptide in a suitable vessel, such as a 15- or 50-ml conical tube.

3.Dilute pooled sample 2:1 with distilled water (i.e., maximum 15 ml total volume in a 50-ml conical tube).

4.Place diluted sample in a –80°C freezer for 24 hr.

Lyophilization

5.Operate the lyophilization unit as per the manufacturer's specifications. Verify that all parameters have been set as prescribed by the manufacturer.

6.Remove the accessory drying chamber (manifold) from the collector chamber lid and, using a soft, lint-free cloth or paper towel, wipe the port gasket(s) and sealing surfaces of the drying chamber/manifold and collector chamber lid to remove any dirt or contaminants that could cause a vacuum leak or contaminate the sample.

7.Check that each sample valve is closed or in the “vent” position and then start the instrument collector and allow the refrigeration system to reach its specified operating temperature (e.g., –105°C). Start the vacuum when collector temperature is at –40°C or colder.

8.Moving quickly to avoid thawing, retrieve sample from the freezer (see step 4) and, using a pre-chilled 50-ml tube holder, place the samples into a pre-chilled lyophilizer flask. Cover the samples with a piece of filter paper and then attach the flask to the lyophilizer with a flask adaptor.

9.Slowly open the port valves to commence lyophilization.

10.Check sample and system periodically to ensure proper temperature and pressure are maintained and that the sample has not thawed.

11.When the sample has completely dried, close the pressure valve and remove the flask from the lyophilizer.

12.Resuspend the sample in water using the same initial volume of water as was used to dilute the original sample (see step 3; i.e., 2 parts water to 1 part sample in DMSO) and then refreeze the sample.

13.After the sample is frozen as in step 4, return it to the lyophilizer, repeating steps 5 to 8 as necessary. When the sample is completely dry, close the valve and carefully remove the flask from the machine.

MP resuspension and storage

14.Resuspend the lyophilized MP in 100% DMSO down to 1 mg/ml peptide.

15.Perform additional dilutions with water if concentrations <1 mg/ml are necessary.

16.Aliquot diluted pools into small aliquots (e.g., in 0.2-ml PCR tubes; volume of 25 or 50 µl is optimal) to prevent freeze-thaw cycles.

17.Store aliquoted MPs at –20°C and inspect over time for appearance. If precipitation is noted, re-lyophilize the MPs.

MPs are a complex mixture of multiple peptides, and therefore, it is unfeasible to perform mass spectrometry analysis on MPs for quality purposes. The quality of MPs is generally assessed by testing their bioactivity in a T cell assay. This protocol describes how to assess the quality of MPs and their use in an immunological assay to quantify antigen-specific responses. Specifically, the immunological assay described herein is an AIM assay using peripheral blood mononuclear cells (PBMCs) as biological source material. This protocol also describes the steps of thawing and washing cryopreserved PBMCs prior to cell culture and stimulation and how to perform flow cytometry staining for membrane markers. This will allow identification of co-stimulatory receptors that are expressed on activated T cells after antigen-specific stimulation with MPs. Although this protocol is focused on an AIM assay, MPs can be used in other assay modalities to assess antigen-specific responses and/or validate MP quality, such as ICS or ELISPOT assays and assays using whole blood instead of PBMCs.

Materials

• Frozen PBMCs

• HR5 medium (see recipe), 4°C

• Benzonase (Sigma, cat. no. E8263)

• MPs (prepared as in Basic Protocol 1; e.g., Spike MP, CD4RE MP, CD8RE MP, EBV MP; see Table 2)

• Anti-CD40 antibody (Miltenyi Biotech, cat. no. 130-094-133)

• Phosphate-buffered saline (PBS; GIBCO BRL, cat. no. 10010-023), 4°C

• MACS buffer (see recipe), 4°C

• 37°C water bath

• Conical polypropylene tubes (e.g., 50 ml; Falcon), 4°C

• Standard tabletop centrifuge, 4°C

• 96-well U-bottom cell culture−treated plate (Grenier Bio-One, cat. no. 655180)

• Flow cytometer (e.g., ZE5, Bio-Rad)

• Additional reagents and equipment for counting cells (see Current Protocols article: Phelan & May, 2015) and for preparing a cocktail of conjugated antibodies (see Table 3)

NOTE : All solutions and equipment coming into contact with cells must be sterile, and proper sterile technique should be used accordingly, including use of a laminar flow hood (Labconco Purifier BSC Class II or equivalent).

NOTE : All culture incubations are performed in a 37°C, 5% CO₂ incubator unless otherwise specified.

Thawing of PBMCs

1.Submerge cryovials with frozen PBMCs in a 37°C water bath. Hold the vials in the bath until all but a tiny bit of ice remains in the vials.

2.Transfer the cells into cold conical propylene tubes (e.g., 50 ml) with cold culture medium (10 ml HR5 medium + 20 µl benzonase per vial used) and centrifuge the cells for 7 min at 180 × g , 4°C.

3.Count the cells and determine cell viability.

4.Add HR5 medium to adjust the cell concentration to 10 × 10⁶ cells/ml.

Cell stimulation

5.Prepare MP stimulation solution in HR5 medium at the desired 2× concentration. Adjust the volume depending on the number of samples used per experiment.

6.Plate in a 96-well U-bottom cell culture−treated plate, with 100 µl stimulus per well.

7.Plate 100 µl cells at a concentration of 10 × 10⁶ cells/ml (see step 4) to plate 1 × 10⁶ cells per well.

8.Optional: If the membrane staining (see step 10) will include staining for the activation marker CD40L, then add anti-CD40 antibody (0.5 µg/ml) to the solution for each donor before incubation.

9.Incubate plate for a total of 24 hr at 37°C, 5% CO₂.

Antibody surface staining and flow cytometry

10.To perform membrane staining after 24 hr of stimulation, prepare a cocktail of conjugated antibodies as shown in Table 3 using cold diluent and concentrations recommended by manufacturers.

11.After stimulation, spin 96-well U-bottom plate for 2 min at 300 × g , 4°C.

12.Wash plate by aspirating supernatant, adding 200 µl cold PBS per well, and centrifuging 2 min at 300 × g , 4°C.

13.Aspirate PBS wash and add 100 µl cold antibody mix from step 10 to each well.

14.Incubate for 30 min at 4°C. Protect from light.

15.After incubation, add 100 µl cold MACS buffer per well and spin plate for 2 min at 300 × g , 4°C.

16.Wash plate twice by aspirating supernatant, adding 100 µl cold MACS buffer per well, and spinning plate for 2 min at 300 × g , 4°C.

17.Aspirate supernatant and resuspend cells in 100 µl cold PBS per 1 × 10⁶ cells plated.

18.Acquire samples in a flow cytometer.

HR5 medium

• 1000 ml RPMI (Omega, cat. no. RP-21)
• 50 ml Heat Inactivated Human Serum AB (GeminiBio, cat. no. 100-512)
• 10 ml penicillin-streptomycin (GeminiBio, cat. no. 400-109)
• 10 ml L-glutamine (GlutaMAX; Gibco, cat. no. 35050061)
• Store ≤1 month at 4°C

MACS buffer

• 500 ml PBS, pH 7.4 (Invitrogen, cat. no. 10010-0230)
• 2.5 g bovine serum albumin (BSA; Sigma, cat. no. A-3294; store at 4°C)
• 2 ml 0.5 M EDTA
• Filter through 0.2-µm filter
• Degas for 30 min
• Store ≤1 month at 4°C

Background Information

To evaluate the targets of T cell responses to diverse pathogens or allergens, testing peptide pools with hundreds of peptides may be necessary. However, challenges associated with pooling and testing a large number of peptides can hinder the identification and quantification of T cell responses. Here, we developed a sequential lyophilization method to increase peptide concentrations and reduce solvent toxicity for further use in biological assays and assessment of antigen-specific responses such as, but not restricted to, AIM assays.

AIM assays are based on TCR-dependent upregulation of co-stimulatory immune molecules, independent of cytokine response, and have been successfully used in various studies to detect T cells specific to viruses, bacteria, or allergens. In our work, we used, as an example, SARS-CoV-2 MPs (see Understanding Results), which have been applied in more than 80 studies thus far, and OX40, 4-1BB, and CD69 as activation markers. In this assay, CD4+ T cell responses are measured utilizing the OX40+CD137+ dual-marker combination, and CD8+ T cell responses are measured utilizing the 4-1BB+CD137+ dual-marker combination. Other activation markers and/or marker combinations can be used as discussed previously (da Silva Antunes et al., 2017; Dan et al., 2016; Reiss et al., 2017).

Critical Parameters

To avoid contamination or vacuum leaks prior to starting the MP lyophilization (Basic Protocol 1), remove any baffling (if present) and wipe with a soft cloth or paper towel to remove any remaining moisture within the drying chamber, and also ensure that the collector chamber and drain line are free of any residual moisture. It is recommended that all the samples be covered using a filter paper, parafilm, or Kimwipe as an extra precaution to avoid contamination of samples during the freeze-drying process.

Because of the intrinsic solubility and numbers of epitopes, there can be great variability in solubility and viscosity between different MPs, and even different batches of the same MP. Additionally, storage conditions and multiple freeze-thaw cycles can affect the stability of the peptides in the MP. To maximize cellular responses and lengthen the shelf-life of the pool, once an MP is removed from the freezer, it should be stored ≤2 weeks at 4°C or refrozen. Avoid more than five freeze-thaw cycles.

When assessing MP reactivity (Basic Protocol 2), the most important and mandatory control in the AIM assay is a control for specificity or negative control. For that purpose, all assays should be performed with a DMSO stimulation control in parallel, at a concentration matching the same exact concentration of MPs used. It is highly recommended to perform this control in triplicate if enough PBMCs are available. The DMSO signal should then be subtracted from the MP-specific signal.

The second most important control in this assay is a positive control. This control is used to ensure that detection of signal is observed by validating the expression and/or upregulation of the different activation markers. Typical positive controls used in this assay are polyclonal stimulation agents such as phytohemagglutinin (PHA), phorbol 12-myristate 13-acetate (PMA)/ionomycin, α-CD3/CD28, or staphylococcal enterotoxin B (SEB).

Another control, although optional, is the use of an MP that is not associated with the biological target that the user is interested in, typically called an irrelevant MP control. This allows for an internal assay control for the donor response or sample being interrogated. Typical MPs used as irrelevant controls are MPs associated with generalized responses across a human population, like those elicited after vaccination (e.g., pertussis, tetanus, or SARS-CoV-2) or after infection with ubiquitous pathogens (e.g., CMV or EBV).

Another optional control is the inclusion of PBMC samples from donors that are not expected to respond to a specific MP (negative donors), such as using cells from HIV-negative donors to test reactivity to an HIV MP. This control will identify if any undesired reactivity of an MP or issues with the MP synthesis occurred (e.g., lyophilization contamination). Alternatively, PBMC samples from donors that are expected to respond to a specific MP can be included (positive or control donors), such as donors with a particular indication or donors who have experienced a previous clinically diagnosed disease or undergone a particular vaccination schedule (e.g., pertussis, tetanus, or SARS-CoV-2). This control can be particularly valuable in the monitoring and assessment of longitudinal response stability (da Silva Antunes, Sutherland, et al., 2021).

Troubleshooting

Fluctuations in the vacuum readings as well as a loud suctioning grunt during the machine startup is often an indicator of a vacuum leak in Basic Protocol 1. This can be resolved by making sure all the knobs on the valves are in the “Vented” position, which closes the valves, allowing the drying chamber pressure to drop to working conditions. It is important to also inspect the lid as well as the gaskets on the chamber/manifold for proper placement and/or any damage or degradation that may cause unwanted vacuum leaks.

If samples begin to thaw before being placed into the freeze-drying unit, it is necessary to have them refrozen to facilitate freeze-drying efficiency and avoid sample loss. Liquid nitrogen can be employed to help with this after a short period in the –80°C freezer. Flasks can be further insulated using a neoprene cozy and/or aluminum foil to minimize sample thawing. The cozy can also function to protect the vessel from breakage in case it falls.

Although the Labconco^® FreeZone^® Benchtop Freeze Dryer is our lyophilizer of choice, solvent removal from peptide pools could be performed using a standard SpeedVac concentrator, which will remove a large amount of solvent relatively quickly. However, this type of device does not do as complete a job as a freeze dryer and thus may result in issues with DMSO toxicity in the final sample.

Regarding the AIM assay (Basic Protocol 2), if using frozen PBMCs that are not thawed properly, viability and cell recovery can be compromised, and the cells may not perform optimally in the AIM assay. In general, cells should be thawed quickly and kept cold while still in the presence of DMSO. Cells with DMSO intercalated into their membranes are very fragile and must be pelleted and handled gently.

When performing the surface staining steps, it is important to minimize the exposure to light, as the antibodies used for this assay are conjugated with fluorochromes that are sensitive to light. The dilution for each antibody depends on the flow cytometer used to acquire the sample and on the manufacturers’ recommendations. As such, it is advisable to perform an antibody titration to assess the optimal dilution based on the instrument. Similarly, fluorochrome combinations are machine dependent and should be chosen according to the flow cytometer used for the specific experiment.

When interpreting results, some MPs can be used to simultaneously detect CD4+ and CD8+ T cell responses, whereas others are optimized to detect exclusively CD4+ or CD8+ T cell responses. If unsure, please refer to the details about the specificity of the MP you are using.

The number of peptides that can be pooled into a single MP depends on several factors that affect solubility, such as the number of peptides with a high proportion of hydrophobic amino acids, the size of the peptides, and the presence of salts in the peptide preparations, to name a few. As a general rule, MPs should not exceed 300 to 400 peptides. However, a strategy to test a higher number of peptides could be to use multiple pools for simultaneous stimulation directly in the culture assay (Yu et al., 2022).

Understanding Results

In this section, we describe and visually outline the detection and characterization of antigen-specific responses to SARS-CoV-2 after employing the AIM assay, as mentioned above. To detect SARS-CoV-2 T cell reactivity, an MP of overlapping peptides spanning the entire Spike sequence and two MPs optimized for detection of non-Spike (remainder of SARS-CoV-2 proteome) reactivity in CD4+ and CD8+ T cells were employed (CD4RE and CD8RE, respectively). The combination of these pools allowed us to discriminate four groups of subjects with different SARS-CoV-2 infection and COVID-19 vaccine statuses, as described below. An EBV MP was used as a control (Fig. 1). CD4+ and CD8+ T cell responses were calculated as percent of total CD4+ (OX40+CD137+) or CD8+ (CD69+CD137+) T cells, respectively. The background was removed from each stimulation by subtracting the signal from wells stimulated with DMSO. The gating strategy utilized is shown in Figure 2, as well as an example of reactive CD4+ and CD8+ T cell responses to SARS-CoV-2 Spike, CD4RE, and CD8RE MPs and EBV MP from a representative donor.

As expected, Spike SARS-CoV-2-specific T cells can be detected in individuals who have been vaccinated with COVID-19 vaccines (non-infected and vaccinated; I−V+) or infected with SARS-CoV-2 (infected and non-vaccinated; I+V−) or both (infected and then vaccinated; I+V+). Conversely, no responses are observed in individuals who have been neither infected nor vaccinated (non-infected, non-vaccinated; I−V−). Importantly, CD4RE and CD8RE responses are only detected in convalescent subjects (I+V− or I+V+) and not in unexposed (I−V−) or vaccinated (I−V+) subjects, as expected, given that all vaccinated donors included in this study received exclusively Spike-based mRNA vaccines. Additionally, EBV reactivity was detected and observed at equal levels across all the groups. Overall, these data demonstrate the attributes of the MP approach and versatility of MP design to detect antigen-specific responses to SARS-CoV-2 and EBV, complex pathogens composed of multiple antigens. Importantly, due to the intrinsic nature and diversity of the epitopes selected, the MPs employed in our work allowed us to discriminate responses specific to COVID-19 vaccination or SARS-CoV-2 infection, highlighting the potential use of MPs as immunodiagnostic tools. For complete details on the use and execution of this approach for this particular experimental setting and/or interpretation of the data, please refer to prior work (Yu, Wang, Garrigan, Goodwin, et al., 2022).

Time Considerations

On average, it takes about 16 to 18 hr per run to completely lyophilize a sample and two runs in total, with an additional dilution step, to extract all the DMSO from a peptide sample (Basic Protocol 1). For the lyophilization of any MP, if time is of the essence, the drying process can be accelerated first with a SpeedVac concentrator, and then the lyophilizer can be utilized to further remove residual DMSO, though this does incur the potential risks involved with additional sample handling. Use of a SpeedVac must also proceed with caution, as concentrating too quickly risks loss of sample along with the solvent.

The AIM assay (Basic Protocol 2) requires 2 days to be completed. The thawing of PBMCs should take <1 hr, although more time may be needed if more than 10 samples are used at the same time. If using fresh PBMCs, plan ahead to allocate time for blood separation before PBMCs are processed. The T cell culturing period with MPs should be exactly 24 hr. The surface staining should be performed the following day, immediately after stimulation. If not possible, the plate should be removed from the incubator and placed at 4°C until the surface staining is performed. After staining, samples should be immediately acquired in a flow cytometer machine. However, fixation of the PBMC samples can be performed after the last step of the protocol to extend the period before sample acquisition.

Acknowledgments

Research reported in this publication was supported by the National Institute of Allergy and Infectious Diseases (NIAID) of the National Institutes of Health (NIH) under award number U19 AI142742 and contract numbers 75N93019C00001, 75N93019C00065, and 75N93019C00066. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH.

Author Contributions

Ricardo da Silva Antunes : Data curation; formal analysis; investigation; methodology; visualization; writing—review and editing. Daniela Weiskopf : Data curation; formal analysis; funding acquisition; investigation; writing—original draft; writing—review and editing. John Sidney : Conceptualization; data curation; formal analysis; investigation; methodology; writing—original draft; writing—review and editing. Paul Rubiro : Investigation; methodology; writing—original draft. Bjoern Peters : Funding acquisition; investigation; resources; writing—review and editing. Cecilia S. Lindestam Arlehamn : Data curation; formal analysis; funding acquisition; investigation; methodology; writing—original draft; writing—review and editing. Alba Grifoni : Conceptualization; data curation; formal analysis; funding acquisition; investigation; methodology; writing—original draft; writing—review and editing. Alessandro Sette : Conceptualization; funding acquisition; investigation; methodology; resources; supervision; writing—original draft; writing—review and editing.

Conflict of Interest

A.S. is a consultant for Gritstone Bio, Flow Pharma, Arcturus Therapeutics, ImmunoScape, CellCarta, Avalia, Moderna, Fortress, and Repertoire. A.G. is a consultant for Pfizer. La Jolla Institute for Immunology (LJI) has filed for patent protection for various aspects of MP design. All other authors declare no conflict of interest.

Data Availability Statement

The MPs described in this article will be made available to the scientific community upon request, depending on material availability and following execution of a material transfer agreement (MTA), by contacting A.S. (alex@lji.org).

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