Mucosal-associated invariant T (MAIT) cell responses in Salmonella enterica serovar Typhi strain Ty21a oral vaccine recipients.

Abstract

Mucosal–associated invariant T (MAIT) cells are unconventional innate-like T cells abundant in human mucosal tissues and are associated with protective responses to microbial infections. MAIT cells have the capacity for rapid effector functions, including the secretion of cytokines and cytotoxic molecules. In this study, we examined the longitudinal circulating MAIT cell response to the live attenuated oral vaccine Ty21a (Ty21a) against Salmonella enterica serovar Typhi (S. Typhi). We enrolled healthy adults who received a course of oral live-attenuated S. Typhi strain Ty21a vaccine and assessed peripheral blood MAIT cell longitudinal responses pre-vaccination, and at seven days and one-month post-vaccination, using flow cytometry, cell migration, and tetramer decay assays. We showed that following vaccination, circulating MAIT cells were lower in frequency, but were more activated, and had higher levels of gut-homing marker integrin α4β7 and chemokine receptors CCR9 and CCR6, suggesting the potential of MAIT cells to migrate to mucosal sites. We found no significant differences in MAIT cell functionality, cytotoxicity and T-cell receptor avidity, except in TNF expression, which was higher post-vaccination. We show that MAIT cell immune responses are modulated post-vaccination against S. Typhi. This study contributes to our understanding of MAIT cells’ potential role in oral vaccination against bacterial mucosal pathogens.

Introduction

Mucosal-associated invariant T (MAIT) cells are innate-like αβ T cells defined by the expression of an invariant α chain, generally Vα7.2 linked to Jα33, Jα12, or Jα20, and a limited repertoire of T cell receptor β (TCRβ) chains in humans [1, 2]. MAIT cells are restricted by the non-classical MHC-related molecule 1 (MR1) and respond to vitamin B metabolites derived from bacterial and fungal species [3]. MAIT cells have been associated with protection and antibacterial immune defense in various bacterial infections, including Legionella longbeachae, Mycobacterium tuberculosis, Mycobacterium bovis, Francisella tularensis, Escherichia coli, Vibrio cholerae, and Klebsiella pneumoniae [4–8]. Exposure to enteric bacteria in clinical studies, such as in challenge studies with S. Typhi and S. Paratyphi A [9, 10], and in vaccine trials with Shigella flexneri [11], MAIT cell activation, proliferation, and homing to mucosal sites were observed. MAIT cell functions include the capacity to secrete TNF, IFN-γ, IL-17, as well as Granzyme B upon activation [12, 13]. MAIT cells are potential targets for improving vaccine responses [14] as they bridge the adaptive and innate immune responses against bacterial infections and are donor-unrestricted (not restricted by MHC polymorphism) [15].

Emerging evidence supports a role of MAIT cells in the response to vaccination [9, 10, 16, 17]. In participants vaccinated with an attenuated strain of Shigella dysenteriae, MAIT cell activation was observed in those who had mounted LPS-specific IgA antibody-secreting cell responses [11]. In bacterial challenge studies, MAIT cells are activated and decline in blood upon human challenge with S. Typhi [10]. A recent study revealed that MAIT cells play a crucial role in the initial priming of adaptive T cell immune responses to antigens encoded by the ChAdOx1 viral vaccine vector in both mice and humans [17]. However, it remains unclear whether this finding extends to other vaccine modalities such as the orally administered attenuated S. Typhi strain Ty21a, a commercially available vaccine for typhoid fever [18]. Therefore, the objective of our study was to examine the longitudinal circulating MAIT cell responses in a cohort of Ty21a recipients. We showed that Ty21a vaccination resulted in changes in MAIT cell frequency, activation, cytokine production and homing markers expression.

Materials and methods

Subjects

Eleven healthy participants took part in this study, including six males (aged 42 ± 15 years) and five females (aged 40 ± 10 years). All participants provided written informed consent. Four doses of a single oral capsule of Ty21a (Vivotif, PaxVax) were taken on day 1, 3, 5 and 7. Blood samples were collected from each subject at study enrollment (day 0), and approximately 7 days and 1 month after the last dose of vaccine (day 7- and 1-month post-vaccination, p.v.). Venous blood was collected and centrifuged over a Lymphoprep (STEMCELL Technologies Inc) density gradient using a standard protocol to isolate peripheral blood mononuclear cells (PBMCs). PBMCs in cryogenic vials were placed immediately into an isopropanol freezing container (Nalgene Mr Frosty) and were cryopreserved at −80°C until used for immunologic assays. Study procedures were reviewed and approved by the Institutional Review Board of the University of Utah (IRB #84287).

Antigenic stimulation and incubation

Viable S. Typhi Ty21a were obtained by dissolving a vaccine capsule (Vivotif) in 10 ml brain heart infusion (BHI) media and incubating overnight at 37°C. The bacteria were then sub cultured (1 : 10) in BHI media for 4 h (O.D = 0.4), harvested and stored in 50% glycerol at −80°C. For phenotypic analysis of MAIT cells, PBMCs were thawed from −80°C and were seeded (2 × 10⁶ cells/well) in complete medium in 96-well U bottom plates. Cells in each well were stimulated at a multiplicity of infection (MOI) of 100 with heat-killed S. Typhi Ty21a (killed by incubation at 95°C for 30 min) to mimic the immune response of MAIT cells in vivo, consistent with a prior study that utilized similar methods and elicited MAIT responses in Ty21a vaccinees [19]. Unstimulated control wells were treated with complete medium (RPMI 1640 with 10% FBS, 1% penicillin/streptomycin, and 15 mM HEPES). Cells were then incubated at 37° in 5% CO₂. After an overnight incubation, brefeldin A (3 μg/ml) (BD GolgiPlug; BD Biosciences) was added to each well, and the plate was incubated for a further 4 h at 37° in 5% CO₂ prior to cell staining for flow cytometry_. In pilot experiments, we used fixed E. coli stimulation of PBMCs as a positive control [20, 21]. However, due to limitations in the number of PBMCs available from each subject, we did not use positive controls for subsequent experiments.

Flow cytometry

Prior to surface staining, PBMCs were labeled with Zombie UV Fixable Viability Dye (BioLegend). The cells were then washed and incubated with fluorochrome-conjugated antibodies and the 5-OP-RU tetramer (NIH Tetramer Core) for 40 min at room temperature (RT). Before surface staining, the cells were incubated in a human Fc block (BD) for 15 min at RT [22]. Details of viability dye and antibodies used are listed in Supplementary Table 1. For intracellular cytokine analysis, surface staining was followed by fixation, permeabilization, and staining using the Foxp3/transcription factor staining buffer set (eBioscience). Compensation beads (BD Biosciences) were used to create compensation matrices, and Fluorescence minus One (FMO) control were used to identify populations of interest. We defined MAIT cells as live CD45⁺ CD19⁻ TCRγδ⁻ CD4⁻ CD3⁺ CD8⁺ CD161⁺ Vα7.2⁺ MR1-5-OP-RU tetramer⁺ (Figs 2–4). No significant differences were observed in the frequencies of CD4+ Vα7.2⁺ MR1-5-OP-RU tetramer⁺ MAIT and CD4⁺ CD8⁺ Vα7.2⁺ MR1-5-OP-RU tetramer⁺ MAIT cells in post-vaccination samples compared to pre-vaccination samples (supplementary figure 2G and H). A total of 10⁶ events per sample were acquired using Cytek Aurora (Cytek Biosciences) and analyzed using FlowJo software v10 (Tree Star Inc, Ashland, OR).

Figure 2.

MAIT cell frequency and activation post-vaccination. (A) MAIT cell gating strategy and representative flow cytometry plots from unstimulated samples. (B) MAIT cells frequency as percentage of CD³⁺ T cells and (C) as percentage of CD⁸⁺ T cells. (D) absolute MAIT cell counts (CD³⁺ MAIT) pre-vaccination and post-vaccination. (E and F) Frequency of activated MAIT cells expressing CD38 and CD69 pre- and post-vaccination. Black circles are unstimulated donors. The data are representative of two independent experiments out of a total of two. *P <0.05, **P <0.01, ***P <0.001 in Wilcoxon signed-rank test (paired samples).

Figure 3.

MAIT cells effector function post-vaccination. PBMCs obtained from vaccine recipients (n = 11 per group) were stimulated with heat-killed S. Typhi at MOI of 100 and intracellular expression of (A) TNF (B) IFN-γ, (C) IL-17, (D) granzyme B and (E) Perforin in MAIT cells (CD³⁺ 5-OP-RU tetramer+ CD8+) were analyzed using flow cytometry (blue circle shows unstimulated and red circle shows stimulated data). Data were expressed as mean ± SEM. The data are representative of two independent experiments out of a total of two. Paired comparisons were made using Wilcoxon matched-pairs signed-rank test using Prism v9 (GraphPad). *P < .05, **P < .01, ***P < .001.

Figure 4.

Increased homing markers and chemokine receptor expression in MAIT cells post vaccination. PBMCs obtained from vaccine recipients (n = 11 per group) were stimulated with heat-killed S. Typhi at MOI of 100 and surface expression of (A) Integrin α4β7, (B) CCR9, (C) CD103, (D) CCR4, (E) CCR5, (F) CCR6, (G) CXCR5 expression in MAIT cells was measured using flow cytometry. Data were expressed as mean ± SEM. The data are representative of two independent experiments out of a total of two. Paired comparisons were made using Wilcoxon matched-pairs signed-rank test.

Cell migration assay

To assess migratory properties of MAIT cells, PBMCs were stimulated with heat-killed S. Typhi Ty21a overnight, washed, and resuspended in RPMI 1640 medium supplemented with 0.1% bovine serum albumin (Thermo Fisher Scientific, MA, USA). Cells were seeded in the upper chamber of a 6-well transwell plate inserted with a pore size of 3 μm (Thermo Fisher Scientific, MA, USA) at a density of 1 × 10⁶ cells/well. Cells were allowed to migrate against a gradient of 150 ng/ml recombinant chemokines CCL20/MIP-3α and CCL25/TECK-3 (Peprotech, NJ, USA) in RPMI 1640 medium supplemented with 0.1% bovine serum albumin for 4 h at 37°C [23]. Cells migrated into the bottom chamber were collected, washed in FACS buffer (phosphate-buffered saline with 2% fetal bovine serum), stained with fluorochrome-conjugated anti-CD8a, anti-TCRVα7.2, anti-CD161 antibodies and anti-human MR1 5-OP-RU Tetramer (NIH Tetramer Core Facility), and MAIT cells were counted using Countbright^TM absolute counting beads (Thermo Fisher Scientific, MA, USA) and flow cytometry.

Tetramer staining and decay assay

Tetramer staining and decay assay were performed as previously described [9]. Briefly, PBMCs were stained with live/dead fixable viability dye efluor 780 (eBiosciences, Thermo Fisher Scientific, MA, USA) for 15 min at room temperature (RT), followed by an incubation with 5-OP-RU MR1 tetramer for 40 min at RT. Excess tetramer was washed off and cells were re-suspended in FACS buffer with or without 20 μg/ml anti-MR1 antibody (clone 26.5, Biolegend). Samples were left at 37°C and periodic samples were taken, washed and fixed immediately. CD3 staining was performed after collecting all samples and cells were analyzed using flow-cytometry.

ELISAs

For enzyme-linked immunosorbent assays (ELISA), we used S. Typhi LPS antigen and assessed plasma antibody responses (immunoglobulin [Ig] M, IgA, and IgG). ELISAs were performed as previously described [24]. Briefly, microplates (nunc-maxisorp flat-bottom 96-well plates, Invitrogen) were coated with 1 μg/ml LPS, and plasma was added at a dilution of 1 : 200 for IgG and 1 : 100 for IgA and IgM. Bound antibodies were detected with anti-human IgG, IgA, and IgM conjugated with horseradish peroxidase (Jackson ImmunoResearch), and plates were developed by adding 100 μl/well TRB substrate for 10 min in the dark. Development was stopped by the addition of 100 μl/well of 0.2 N H₂SO₄ and OD read at 450 nm in an ELISA microplate reader (Multiskan Ascent; Thermo Labsystems).

Statistical analysis

Paired comparisons were performed using Wilcoxon matched-pairs signed-rank test using Prism v9 (GraphPad). P values are two-tailed and considered significant at P < .05.

Results

Ty21a vaccination resulted in a decrease in frequency of circulating MAIT cells but an increase in activation

Analysis of plasma antibody responses to S. Typhi LPS in Ty21a vaccine recipients revealed that vaccine elicited significantly higher IgA and IgG antibody responses at 7 days and 1-month post-vaccination compared to day 0 (pre-vaccination) (Fig. 1A and B). No significant differences were found in IgM antibody responses (Fig. 1C). These results are consistent with prior studies of Ty21a vaccination [25–27].

Figure 1.

Changes in S. Typhi specific antibody response post-vaccination. Plasma ELISA’s against S. Typhi LPS (A–C) IgA, IgG and IgM antibody responses pre-vaccination (day 0) and post-vaccination (day 7 and 1 month). The data are representative of two independent experiments out of a total of two. *P < .05, **P < .01, in Wilcoxon signed-rank test (paired samples).

Because of the potential importance of MAIT cells against S. Typhi infections and as potential targets for vaccine development, we assessed whether immunization with the Ty21a typhoid vaccine elicits MAIT cell immune responses. PBMCs from Ty21a vaccinees were isolated and stimulated with heat-killed S. Typhi Ty21a, and MAIT cell frequency and phenotype were analyzed using flow cytometry. We defined MAIT cells as live CD45⁺ CD19⁻ TCRγδ⁻ CD4⁻ CD3⁺ CD8⁺ CD161⁺ Vα7.2⁺ MR1-5-OP-RU tetramer⁺, and we used fluorescence minus one (FMO) controls of the MR1-5-OP-RU tetramer and the 6-FP-tetramer for gating MAIT cells (Fig. 2A). We found that the frequencies of circulating MAIT cells as a proportion of total CD3⁺ T cells and as a proportion of total CD8⁺ cells significantly decreased at day 7 and at one-month post-vaccination compared to pre-vaccination (Fig. 2B and C). Absolute MAIT cell counts were also lower both at day 7 and at one-month post-vaccination compared to pre-vaccination (Fig. 2D and Supplementary Table 2).

We next measured the expression of activation markers CD38 and CD69 in MAIT cells. We found that the frequencies of CD38⁺ MAIT cells (Fig. 2E) increased at one-month post-vaccination (CD38 positive population of Vα7.2⁺ CD161+ cells were also higher at one-month post-vaccination compared to pre-vaccination (Supplementary Fig. 1)). No significant differences between post-vaccination and pre-vaccination were found in CD69⁺ MAIT cells (Fig. 2F). Conventional CD4⁺ and CD8⁺ T cells frequencies increased post-vaccination, consistent with a previous publication, [28] and the frequencies of CD14⁺ monocytes decreased in circulation post-vaccination (Supplementary Fig. 2). No significant differences were found in B cells, CD4⁺ CD8+ MAIT cells, CD4⁺ MAIT cells and TCRγδ cells at the time points assessed post-vaccination compared to pre vaccination (Supplementary Fig. 2). Taken together, we found that vaccination with Ty21a impacted circulating MAIT cell frequency and activation.

Ty21a vaccination resulted in higher frequencies of TNF expressing circulating MAIT cells

We next examined whether vaccination with Ty21a induces MAIT cells to express effector molecules such as IFN-γ, TNF, IL-17a and cytotoxic molecules. Using flow cytometry, we found a significant increase in the frequencies of MAIT cells intrinsically expressing TNF at day 7- and 1-month post-vaccination compared to pre-vaccination, only in the unstimulated condition (Fig. 3A). This indicates that the intrinsic TNF responses in MAIT cells increased following vaccination. IFN-γ expressing MAIT cells also were non-significantly higher (P = .08) post-vaccination in unstimulated conditions (Fig. 3B and Supplementary Fig. 3). No significant differences were found in IL-17a, granzyme B, or perforin expression among the groups (Fig. 3C–E). IFN-γ, TNF and perforin expressing MAIT cells significantly increased in response to ex-vivo S.Typhi stimulation compared to unstimulated condition at the pre-vaccination timepoint. However, except for TNF at 1-month post-vaccination, we did not observe any significant differences between unstimulated and stimulated conditions at any of the post-vaccination timepoints (Fig. 3A).

To determine whether vaccination changes the TCR avidity of the circulating MAIT cell population for 5-OP-RU–MR1 complexes, we stained PBMCs with 5-OP-RU tetramers, and assayed the rate of tetramer dissociation by incubating cells over a time course with an anti–MR1 blocking antibody [9]. We hypothesized that vaccination could selectively enhance or activate a subset of high-avidity MAIT cells. Despite the observed decrease in MAIT cell numbers post-vaccination, an increase in avidity would suggest preferential survival or expansion of high-avidity clones. In the presence of anti-MR1 antibody, tetramer dissociated rapidly over an hour interval in both pre- and post-vaccination groups. Using the MFI of MAIT cells bound to tetramer and a two-phase (fast and slow) decay model for our analysis, we saw minimal differences in rate of tetramer dissociation in pre-vaccinated and post-vaccinated samples [29] (Supplementary Fig. 4), suggesting that vaccination did not lead to a change in circulating MAIT cell TCR avidity.

Ty21a vaccination resulted in an increased expression of tissue-homing chemokine receptors and integrin on MAIT cells

Given that we found lower frequencies of circulating MAIT cells post-vaccination, it is possible that MAIT cells home to other compartments, such as the gut or other mucosal tissues. To address this possibility, we measured the expression of integrin α4β7 and CCR9 molecules, gut-homing markers known to be found on MAIT cells [30–33]. We found that the proportion of MAIT cells expressing integrin α4β7 increases at day 7 post-vaccination upon stimulation (Fig. 4A and Supplementary Figs 3 and 5), and the proportion of MAIT cells expressing CCR9, were higher one-month post-vaccination in unstimulated and stimulated conditions (Fig. 4B). We found similar results when we analyzed integrin α4β7 and CCR9 molecules in Vα7.2⁺ CD161+ population and when we analyzed median fluorescent intensity (MFI) of integrin α4 and integrin β7 (Supplementary Fig. 5). Next, we used a trans-well chemotaxis assay to assess the potential migration of MAIT cells in the presence of CCR9 ligand CCL25/TECK-3. We found an observed increase in MAIT cell migration in response to CCL25 at one-month post-vaccination compared to pre-vaccination, although this analysis was limited by the insufficient PBMC sample size at this later time point (P = 0.25, Supplementary Fig. 6). It is important to note that this increase was not statistically significant. We also found higher percentages of MAIT cells expressing CCR4, CCR5 and CCR6 chemokine receptors, which are involved in tissue homing [12, 34–36], one-month post-vaccination (Fig. 4C–E). No significant differences were observed in MAIT cells expressing CD103/αE β7 integrin (skin/liver homing [30]) expression (Fig. 4F) and MAIT cell chemotaxis for CCR6 ligand MIP-3α (Supplementary Fig. 6). Furthermore, given recent identification of CXCR5⁺ follicular helper-like MAIT cells [22], we found a higher proportion of stimulated MAIT cells at post-vaccination time points to be positive for CXCR5⁺ (Fig. 4G). Taken together, we found evidence that the decreased proportion of MAIT cells in blood may be associated with increased homing and migration into tissues following Ty21a vaccination.

Discussion

MAIT cells have been implicated in protective responses to microbial infections and, due to their abundance in humans, capacity for rapid effector functions, and lack of restriction by donor genotype, could be attractive vaccine targets [14, 15]. While previous studies have described MAIT cell response to systemic vaccinations in humans [11, 37], mice [8] and macaques [38], there are very limited data on how MAIT cells respond to mucosal vaccines. In this longitudinal study, we evaluated MAIT cell response seven days and one-month after oral S. Typhi strain Ty21a vaccination, and found changes in circulating MAIT cell frequency, activation and homing markers.

We found that circulating MAIT cells are reduced in frequency and activated following Ty21a vaccination. In humans, similar reductions in MAIT cell frequencies have been observed in the blood of children following Vibrio cholerae infection [7], patients with active Mycobacterium tuberculosis [39], and people living with human immunodeficiency virus (HIV) infection [40]. Our results are in line with a prior study demonstrating that oral challenge of participants with wild-type S. Typhi results in a sharp decline of circulating MAIT cells 48 and 96 h after typhoid diagnosis, with MAIT cells highly activated and co-expressing CCR6 and CCR9 homing markers [10]. Similarly, our results are also consistent with a study of subjects receiving an attenuated Shigella vaccine, which also demonstrated alterations in MAIT cell frequency and activation [11]. One possible explanation for the lower frequencies of MAIT cells in circulation post-vaccination is their migration to mucosal tissues, where they may be recruited to contribute to innate effector responses [41]. In support of this, we found that MAIT cells post-vaccination had increased expression of several mucosal-homing chemokine receptors, including CCR6, CCR9 and integrin α4β7. However, we did not measure the expression of chemokines like CCL25 (for CCR9) or adhesion molecules like MAdCAM-1 (for α4β7 binding) in the gut, and in the case of insufficient homing signals, MAIT cells might fail to enter the tissue, leading to the retention of α4β7⁺ or CCR9⁺ MAIT cells in circulation. Further work is needed to determine the contribution and activity of MAIT cells homing to the gut post-Ty21a vaccination.

MAIT cells produce effector molecules such as IFN-γ, TNF and cytotoxic molecules required for killing and eliminating bacteria-infected cells. In our study, we found increased frequencies of TNF expressing circulating MAIT cells post-vaccination but no significant differences in cytotoxic molecules. This highlights the context-dependent functional differences of MAIT cells, as tissue-resident MAIT cells can have different phenotypes and cytokine production than those in the circulation [42, 43]. We also found that while pre-vaccination samples showed significant increases in TNF-α, IFN-γ, and perforin upon ex-vivo stimulation with heat-killed S. Typhi, this was not observed in post-vaccination samples. We used heat-killed S. Typhi to mimic the immune responses of MAIT cells in vivo. Notably, post-vaccination MAIT cells, in the absence of stimulation, had a higher level of activation marker CD38, and correspondingly, higher levels of expression of cytokines (TNF in particular). The reason for this phenomenon is unclear and could partly be due to a non-antigen-driven activation of MAIT cells that persists following mucosal vaccination. According to Boulouis et al. [44], pre-vaccination and post-vaccination levels of MAIT cells correlated positively with the magnitude of the immune response to the BNT162b2 mRNA SARS-CoV-2 vaccine indicating non-antigen driven activation of MAIT cells. The Ty21a vaccination might have primed the immune system in a way that enhances the cytokine response, leading to a baseline activation state of MAIT cells and resulting in higher expression levels of the activation markers.

Our study has a number of limitations. First, due to the preliminary nature of this study, we had a small sample size, although this did not preclude us in observing significant changes in MAIT cell response post-Ty21a vaccination. Studies with larger cohorts are required to confirm and expand our findings. Second, our study was limited to the description of MAIT cells in blood. Given our findings of increased homing markers and migration, future studies examining tissue-resident MAIT cells are needed. Third, the overall MAIT cytokine response was low upon ex-vivo stimulation with heat-killed S. Typhi, likely due to the heat sensitivity of MAIT cell-activating antigens, such as riboflavin metabolites [20, 45]. We lacked sufficient cells to evaluate MAIT effector functions with alternative known and potent stimulations (e.g. MR1 ligand 5-OP-RU or cytokines). Fourth, a limitation of our tetramer dissociation assay is the time points used for measuring MAIT cell avidity. Our current experimental design included points beyond 1 h but given the relatively fast kinetics of MAIT TCR-MR1 interactions, shorter time intervals (e.g. within 1 h) may provide a more precise assessment of avidity.

Despite these limitations, our study expands the limited body of evidence on MAIT cell responses to oral vaccination. Specifically, we provide new insights into MAIT cell dynamics in the context of S. Typhi vaccination: (i) Unlike previous studies focused on acute infection [9, 10], this study investigated longitudinal vaccine-induced changes in MAIT cells which may inform the development of MAIT-targeted vaccines strategies. (ii) We show that vaccination modulates MAIT cell TNF production and migration markers, suggesting that vaccines can have longer-term impact on MAIT cell responses. Given the global burden of enteric infections, there is an urgent need for more effective mucosal vaccines, particularly oral formulations [46]. Our findings provide an important step towards the understanding and harnessing of MAIT cell function, which may offer an important therapeutic strategy to improve mucosal immunity.

Author contributions

Shubhanshi Trivedi (Formal analysis [lead], Investigation [lead], Methodology [lead], Writing—original draft [lead], Writing—review & editing [lead]), Olivia J. Cheng (Methodology [supporting], Writing—review & editing [supporting]), Ben J. Brintz (Formal analysis [supporting], Writing—review & editing [supporting]), Richelle C. Charles (Methodology [supporting], Resources [supporting], Supervision [supporting], Writing—review & editing [supporting]), and Daniel T. Leung (Conceptualization [supporting], Funding acquisition [lead], Resources [supporting], Supervision [lead], Validation [supporting], Visualization [supporting], Writing—review & editing [supporting])

Supplementary data

Supplementary data is available at OXFIMM Journal online.

Conflict of interest: None declared.

Funding

This work was supported in part by the National Institutes of Health (grant number K24 AI166087 to D. T. L.); and by the Dr Thomas D. Rees and Natalie B. Rees Presidential Endowed Chair in Global Medicine to D. T. L.

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files.

Ethics and consent

All participants provided written informed consent. Study procedures were reviewed and approved by the Institutional Review Board of the University of Utah (IRB #84287).