Salmonella Typhi-specific multifunctional CD8+ T cells play a dominant role in protection from typhoid fever in humans
© Fresnay et al. 2016
Received: 24 September 2015
Accepted: 17 February 2016
Published: 1 March 2016
Typhoid fever, caused by the human-restricted organism Salmonella Typhi (S. Typhi), is a major public health problem worldwide. Development of novel vaccines remains imperative, but is hampered by an incomplete understanding of the immune responses that correlate with protection.
Recently, a controlled human infection model was re-established in which volunteers received ~103 cfu wild-type S. Typhi (Quailes strain) orally. Twenty-one volunteers were evaluated for their cell-mediated immune (CMI) responses. Ex vivo PBMC isolated before and up to 1 year after challenge were exposed to three S. Typhi-infected targets, i.e., autologous B lymphoblastoid cell-lines (B-LCL), autologous blasts and HLA-E restricted AEH B-LCL cells. CMI responses were evaluated using 14-color multiparametric flow cytometry to detect simultaneously five intracellular cytokines/chemokines (i.e., IL-17A, IL-2, IFN-g, TNF-a and MIP-1b) and a marker of degranulation/cytotoxic activity (CD107a).
Herein we provide the first evidence that S. Typhi-specific CD8+ responses correlate with clinical outcome in humans challenged with wild-type S. Typhi. Higher multifunctional S. Typhi-specific CD8+ baseline responses were associated with protection against typhoid and delayed disease onset. Moreover, following challenge, development of typhoid fever was accompanied by decreases in circulating S. Typhi-specific CD8+ T effector/memory (TEM) with gut homing potential, suggesting migration to the site(s) of infection. In contrast, protection against disease was associated with low or no changes in circulating S. Typhi-specific TEM.
These studies provide novel insights into the protective immune responses against typhoid disease that will aid in selection and development of new vaccine candidates.
Typhoid fever constitutes a major global health problem, with an estimated 21.7 million cases and 200,000 deaths annually . The development of improved vaccines is necessary, but advances have been delayed by a lack of knowledge of the immunological correlates of protection against Salmonella enterica serovar Typhi (S. Typhi). Since the causative agent of typhoid fever, S. Typhi, is a human-restricted bacteria , current knowledge is limited due to the difficulties associated with performing challenge studies in humans and the lack of adequate pre-clinical models that closely mimic typhoid fever.
Recently, a human challenge model was established by the Oxford Vaccine Group (OVG, University of Oxford) in which naïve participants ingested wild-type (wt) S. Typhi (Quailes strain) [3, 4]. This controlled infection study was modeled after the human typhoid challenge studies performed in the 1960s at the University of Maryland. The Maryland studies improved understanding of typhoid fever [5–8] and resulted in the initiation of the process to license the oral attenuated Ty21a typhoid vaccine , but did not identify the immunological correlates of protection. Although substantial data are available on the immune responses after infection in the field or following vaccination, there are no studies that provide insights into the immunological status before wild-type infection and its possible effects on clinical outcome.
The re-establishment of the human challenge model with wt S. Typhi, and the use of cutting-edge multichromatic flow cytometry allowed us, for the first time, to investigate the pre-challenge immunological status and its correlation with the subsequent clinical outcome. Furthermore, it allowed the initiation of detailed studies of the kinetics and characteristics of the immunological responses occurring following infection with wt S. Typhi.
Several immunization studies with attenuated typhoid vaccine candidates suggested that cell-mediated immunity (CMI), particularly CD8+ effector T cells, constitute a major component in the control of typhoid fever [10, 11]. CD8+ T cells may be involved in destroying infected-host cells through cytolytic activity and/or production of cytokines (e.g., interferon (IFN)-γ, tumor necrosis factor (TNF)-α, interleukin (IL)-17) [12–22].
Recent research on the immune responses after oral immunization with Ty21a have revealed persistent multiphasic, multifunctional (simultaneous production of multiple cytokines) responses to antigenic presentation by class Ia HLA and by the more conserved and less polymorphic non-classical HLA-E molecules [13, 19, 20, 22].
In the present study we investigated the relationship between S. Typhi-specific CD8+ T cell responses before exposure to wt S. Typhi and clinical outcome, i.e., whether the participants who were challenged developed disease or not. We also explored S. Typhi-specific CD8+ T cells responses following challenge, as well as their gut homing potential in relationship with typhoid diagnosis. Finally, we identified the dominant multifunctional S. Typhi-specific response patterns associated with clinical outcome by evaluating the simultaneous production of macrophage inflammatory protein (MIP)-1β, IFN-γ, TNF-α, IL-2 and IL-17A, as well as the expression of the cytotoxicity degranulation marker CD107a . These investigations provide evidence that baseline S. Typhi-specific responses are related to clinical outcome after wt S. Typhi infection and provide novel insights into the immunological responses involved in protection following natural infection and vaccination.
Participants and study design
Twenty-one healthy, male or female participants aged 18–60 years were recruited by the Oxford vaccine Group, Department of Paediatrics, Oxford, UK, to participate in this dose-escalation study. Any participant with a history of typhoid fever or immunization against typhoid fever, or who lived in a typhoid-endemic region for longer than 6 months, was excluded from participation. Only participants with low risk of becoming chronic carriers (including those without gall stones, determined by ultrasound examination of the gallbladder) were included. Participants were challenged orally with a dose of 1–5 × 103 CFU of wt S. Typhi (Quailes strain) suspended in sodium bicarbonate. The S. Typhi Quailes strain, which was used extensively for human challenge studies in the 1960s/1970s was developed by the University of Maryland and used to establish a master cell bank in Oxford. Participants were monitored closely throughout the study. A positive typhoid fever diagnosis was defined based on the following criteria: either a positive blood culture for Salmonella Typhi from day 5 post-challenge or, oral temperature ≥38 °C, persisting continuously for at least 12-h in the absence of anti-pyretic medication, occurring from 72-h after challenge. At the point of typhoid fever diagnosis (TD, as determined by S. Typhi bacteremia or development of a fever >38 °C for ≥12 h) participants were treated with a 2-week course of antibiotics (ciprofloxacin, 500 mg BD). Participants who did not developed typhoid fever (NoTD) received a 2-week course of antibiotics at day 14 post-challenge. Peripheral blood mononuclear cells (PBMC) obtained from 16 selected participants based on cell availability were analyzed in these studies (Additional file 1: Table S1). These included all nine who were not diagnosed with typhoid (NoTD) and seven participants were diagnosed with typhoid (TD).
Written informed consent was obtained and all procedures approved by National Research Ethic Service (NRES), Oxfordshire Research Ethics Comittee A (10/H0604/53) and conducted in accordance with the principles of the International Conference of Harmonisation Good Clinical Practice guidelines.
Specimen collection and isolation of PBMC
Blood was collected from each participant and routine blood hematology was assessed on alternate days after challenge and at typhoid diagnosis. PBMC were separated by Lymphoprep gradient centrifugation (Axis-Shield, Oslo, Norway) cryopreserved in liquid nitrogen following standard techniques within 4 h of initial blood draw within 4 h of initial blood draw and stored in liquid N2. Upon thawing, viability and recovery were measured using trypan blue exclusion and a Guava easyCyte™ flow cytometer (Merck KGaA, Darmstadt, Germany), and cells were rested overnight in complete RPMI (cRPMI: RPMI 1640 media (Gibco, Carlsbad, CA) supplemented with 100 U/mL penicillin (Sigma), 100 µg/mL streptomycin (Sigma), 50 μg/mL gentamicin (Gibco), 2 mM l-glutamine (Gibco), 10 mM HEPES buffer (Gibco) and 10 % heat-inactivated fetal bovine serum (Gemini Bioproducts, West Sacramento, CA) to serve as effector cells in CMI assays.
Autologous Epstein-Barr virus (EBV)-transformed lymphoblastoid cell line (B-EBV cells) and autologous blasts were generated from the PBMC of each participant isolated before challenge. B-EBV cells were obtained by incubation of the PBMC with EBV-containing supernatant from the B95-8 cell line (ATCC CRL1612) and cyclosporine (0.5 ug/mL; Sigma-Aldrich, Saint-Louis, MO) at 37 °C with 5 % CO2 for 2–3 weeks. PHA-activated blasts were prepared by incubating PBMC with 1 μg/ml PHA (Sigma-Aldrich, St. Louis, MO) in cRPMI for 24 h, followed by washing and culture in cRPMI containing 20 IU/ml recombinant human IL-2 (rhIL-2; Roche, Indianapolis, IN) for 7 days. The HLA classical class I-defective B cell line 721.222.AEH which expresses non-classical class-I HLA-E molecules was provided by Dr. D. Geraghty  and cultured in cRPMI supplemented with 200 mU/ml hygromycin B (Sigma-Aldrich).
S. Typhi infection of stimulator cells
Stimulating cells were infected by incubation for 3 h at 37 °C with wt-S. Typhi strain ISP1820 at a multiplicity of infection of 7:1 in RPMI without antibiotics, washed three times with cRMPI and incubated overnight with cRPMI containing 150 μg/ml gentamicin. Cells were washed and infection with S. Typhi was confirmed by staining with anti-Salmonella common structural Ag (CSA-1) (Kierkegaard and Perry, Gaithersburg, MD) and analysis by flow cytometry as previously described .
Ex-vivo stimulation of effector cells
PBMC were thawed and rested in cRPMI overnight at 37 °C in 5 % CO2 before stimulation with S. Typhi-infected stimulating cells. Uninfected target cells and Staphylococcal enterotoxin B (SEB; 10 μg/ml) were used as negative and positive controls, respectively. Target cells were gamma-irradiated (6000 rad) and co-cultured with PBMC (effector:stimulator ratio 5:1) in the presence of the FITC-conjugated anti-CD107a (BD Biosciences) monoclonal antibody (mAb). After 2 h, the protein transport blockers monensin (1 µg/ml, Sigma) and brefeldin A (2 µg/ml; Sigma) were added to the PBMC and cells were incubated overnight at 37 °C in 5 % CO2. For the kinetics analysis, because of the variable responsiveness at baseline in the different participants, the net values (day x post-challenge—day 0) were used to normalize the data.
Immunostaining with 14-color panel mAbs
Following stimulation, PBMC were harvested, washed in 1X PBS and stained for dead-cell discrimination using Yellow Live/Dead viability kit (Invitrogen, Carlsbad, CA). Cells were then washed with wash buffer (PBS 1 % FCS) and non-specific Fc receptor binding was blocked with human immunoglobulin (3 µg/mL; Sigma) for 10 min at room temperature (RT). Cells were surface stained with mAbs against CD14-BV570 (M5E2, Biolegend), CD19-BV570 (HIB19, Biolegend), CD3-BV650 (OKT3, Biolegend), CD4-PE-Cy5 (RPA-T4, BD), CD8-PerCP-Cy5.5 (SK1, BD), CD45RA-biotin (HI100, BD), CD62L-APC-A780 (DREG-56, Ebioscience) and integrin α4β7-A647 (ACT1; conjugated in house) at 4 °C for 30 min. Cells were washed with wash buffer and stained with streptavidin (SAV)-Qdot800 (Invitrogen) at 4 °C for 30 min. Cells were then fixed and permeabilized with Fix and Perm buffers (Invitrogen). Intracellular staining was then performed with mAbs against CD69-ECD (TP1.55.3, Beckman Coulter), IFN-γ-PE-Cy7 (B27, BD), TNF-α-A700 (MAb11, BD), IL-2-BV605 (MQ1-17H12, Biolegend), IL-17A-BV421 (BL168, Biolegend) and MIP-1β-PE (IC271P, R&D) at 4 °C for 30 min. Cells were washed with PBS 1 % FCS, fixed in 1 % paraformaldehyde (PFA) and stored at 4 °C until analyzed. Samples were acquired by flow cytometry using a customized LSRII flow cytometer (BD Biosciences) and analyzed using Winlist v7.0 (Verity Software House, Topsham, ME). CD8+ T cells were selected after a gating strategy involving the exclusion of dead cells and CD3−, CD14+ and CD19+ cells. Absolute numbers of CD3 +, CD8+, CD4+ cells and CD8+ memory subsets cells were calculated by using percentages obtained from flow cytometry analysis related to the absolute number of lymphocytes determined by routine blood count. S. Typhi-specific responses were expressed as net percentage of positive cells (background after stimulation with uninfected cells were subtracted from values obtained with S. Typhi-infected stimulators).
Mann–Whitney tests and linear regression analysis were performed using Prism v7.02 (Graphpad software, La Jolla, CA). Areas under the curve were measured using the trapezoidal method (GraphPad Prism v7.02). P values <0.05 were considered significant. Some comparisons were based on multiple outcomes from the same individual as indicated in the text. In these cases the same individual provided information on cytokine production and/or CD107a expression levels after stimulation with three types of stimulations (EBV-B, AEH, and blasts), and these responses we evaluated with regard to clinical outcome (i.e., TD vs. NoTD patients). In these analyses we accounted for the correlation between multiple outcomes from the same person using a mixed effects model fit by maximum likelihood, including a random effect for person, using SAS 9.3 (Cary, NC). We found by performing simulations that this correlation model provided more accurate statistical inference than models with more complex correlation structure given the small sample sizes.
Baseline S. Typhi-specific CD8+ T cell responses correlate with clinical outcome after challenge
CD8+ T cells have been reported to be a major component of the CMI response against S. Typhi [11–13, 16, 22]. Thus, we first explored whether CD8+ T cell responses in healthy participants at baseline could predict clinical outcome after challenge with 1–5 × 103 CFU of wt S. Typhi. All nine volunteers who did not develop typhoid disease (NoTD group) and seven participants who did develop typhoid disease (TD group) were evaluated. Peripheral blood mononuclear cells (PBMC) isolated from volunteers before challenge (day 0) were stimulated in vitro with S. Typhi-infected autologous B-EBV, S. Typhi-infected autologous blasts or S. Typhi-infected AEH cells (the latter to measure HLA-E-restricted CMI). PBMC were evaluated using 14-color multiparametric flow cytometry and CD8+ T cells were divided based on their expression of CD62L and CD45RA into naïve T (TN; CD62L+CD45RA+), T central memory (TCM; CD62L+CD45RA−), T effector memory (TEM; CD62L−CD45RA−), and T effector memory CD45RA+ (TEMRA; CD62L−CD45RA+) subsets. No significant differences were observed at baseline between the TD and NoTD groups in the absolute number of white cells, lymphocytes, CD3, CD8 and CD4 memory subsets (Additional file 1: Figure S1). S. Typhi-specific responses were further characterized by co-expression of the T cell activation marker CD69 and the cytotoxicity degranulation marker CD107a, as well as the production of IFN-γ, TNF-α, MIP-1β, IL-17A and IL-2.
Distinct clinical outcomes are accompanied by discrete S. Typhi-specific responses after challenge
Enhanced gut homing potential of S. Typhi-specific CD8+ T cells in participants diagnosed with typhoid disease
S. Typhi-specific CD8+ TM cells are primarily multifunctional (MF)
Studies on typhoid fever immunity in humans have been largely restricted to patients residing in endemic areas and subjects immunized with licensed and experimental typhoid vaccines. In contrast, virtually no information is available on the immune status, particularly regarding CMI, prior to infection and on the immunological correlates of protection following oral exposure to wt S. Typhi. The re-establishment of the human challenge model with wt S. Typhi provided a unique opportunity to directly assess the impact of S. Typhi-specific CMI responses before pathogen exposure to the subsequent development of, or protection from, typhoid disease. Herein we show that higher S. Typhi-specific baseline responses are associated with protection against typhoid fever and delayed time to diagnosis and characterized these responses in great detail. If confirmed by future studies involving additional volunteers, these novel observations suggest an important role of anti-S. Typhi specific multifunctional CMI responses in protection from disease. This information has the potential to greatly accelerate the development of novel new generation typhoid vaccines.
It has been recently described that the multifunctional quality of the responses is critical for protection against pathogens [23, 25–28]. In the current study we provide evidence that S. Typhi-specific MF CD8+ TEM and TEMRA cells are the major effector subsets associated with protection against typhoid fever, as well as delayed time to disease onset. Of note, while MF S. Typhi-specific CD8+ cells in NoTD participants were dominant both before and after challenge, this dominance was seen in TD participants only after challenge. MIP-1β was shown to play a role in CTL activity and in controlling infection in HIV non-progressors [29–31]. We previously described the co-production of MIP-1β with IFN-γ, TNF-α, IL-17A and IL-2 following vaccination with Ty21a  and MIP-1β was reported to be produced by PBMC obtained from S. Typhi-infected convalescent patients . Herein, we show that production of MIP-1β is a major feature of S. Typhi-specific MF populations suggesting that co-production of MIP-1β with other cytokines is a key component in protection against S. Typhi. These results highlight that a strong MF component of CD8+ TEM responses both at baseline and after challenge, is associated with disease protection and delayed time to diagnosis.
Following challenge we observed decreases in the percentages of circulating S. Typhi-specific TEM cells in TD participants, suggesting that these cells might be leaving the systemic circulation during the incubation phase of the disease. Similar to the decreases in lymphocyte counts reported post-challenge in TD participants , we also observed a decline in the absolute numbers of CD3+, CD8+ and CD4+ T cells. Integrin α4β7 expression plays an essential role in the selective homing of T cells to the gut, the site of entry for S. Typhi [16, 33–36]. After challenge, TD participants show a decrease in total integrin α4β7+ CD8+ TEM and in both integrin α4β7− and α4β7+ S. Typhi-specific TEM cells. However, 48 h after diagnosis, the proportion of S. Typhi-specific MF integrin α4β7+ TEM in TD subjects was higher than in integrin α4β7− cells. This shift in the balance of single-positive vs. MF S. Typhi-specific CD8+ TEM in circulation might be the result of activation and expansion of MF S. Typhi-specific integrin α4β7+ CD8+ TEM cells due to the ongoing infection in the gut and other lymphoid tissues. Taken together, these results highlight that strong MF CD8+ TEM responses with the potential to home to the gut as well as to other lymphoid tissues are associated with disease protection and delayed time to diagnosis. These results complement and expand our previous findings showing that vaccination against S. Typhi elicited integrin α4β7+ and integrin α4β7− CD8+ effector T cells [14, 16, 17, 24]. These observations also suggest that S. typhi-specific TEM cells migrate not only to mucosal sites, but also presumably to secondary lymphoid tissues, where they may reduce S. typhi replication during the incubation phase, delaying disease onset. Moreover, post-challenge decreases in circulating S. Typhi-specific T cells are proportional to the levels present at baseline, suggesting that the higher the pool of S. Typhi-specific T cells available, the higher the number of these cells that are recruited to the sites of inflammation.
Several studies in the murine S. typhimurium model have shown that the balance of suppressive regulatory T cell (Treg) and pro-inflammatory T cell responses influence bacterial clearance or persistence . We have recently evaluated in these challenged volunteers the hypothesis that the development of Treg responses  following exposure to wt S. Typhi could be responsible, at least in part, for the decrease of S. Typhi-specific TEM responses. We observed that TD participants exhibited up-regulation of the gut homing molecule integrin α4β7 pre-challenge, followed by a significant down-regulation post-challenge consistent with Treg homing to the gut, as well as up-regulation of activation molecules post-challenge. We also showed that depletion of Treg results in increased S. Typhi-specific cytokine production by CD8+ TEM in vitro. These observations suggest that the tissue distribution of activated Treg, their characteristics and activation status may play a key role in typhoid fever, possibly through suppression of S. Typhi-specific effector T cell responses .
In contrast to the results seen in TD participants, we observed that protection against typhoid fever is mostly associated with very low or no changes in circulating S. Typhi-specific TM after challenge. Of note, several of the few NoTD participants showing decreases in S. Typhi-specific TEM were PCR positive or stool positive participants despite the fact that they did not meet typhoid disease diagnosis. These findings suggest that the control and clearance of S. Typhi in NoTD participants might be driven by innate and/or adaptive immune responses in the gut microenvironment, precluding S. Typhi from becoming invasive and causing disease. However, the underlying mechanism(s) of control of S. Typhi infection in the gut microenvironment remain unclear. “Innate-like” T cell types such as TCRγ/δ T cells, NK-T cells and mucosal associated invariant T (MAIT) cells might be involved in protection from typhoid disease. MAIT cells are comprised of CD8+CD4− and CD8−CD4− subsets restricted by the MR1 MHC-related molecule which are widely believed to play an important in mucosal immunity . We have recently shown that MAIT cells from healthy individuals are able to produce IL-17A, IFN-γ and TNF-α when exposed to S. Typhi-infected cells . Ongoing experiments directed to characterize MAIT cell responses in the challenged participants are expected to shed light into the role of these cells following exposure to wt S. Typhi.
Because of the very limited data available regarding immune responses prior to typhoid infection, the reasons for the disparity in baseline responses observed among the participants are unclear. Participants were recruited in the UK, a non-endemic region, have not been vaccinated against typhoid, and therefore have likely never encountered S. Typhi. However, differences in baseline responses could be due to cross-reactive memory responses elicited by previous exposure to other Salmonella serovars [42–44], or other Enterobacteriaceae, including those present in the normal gut microbiota [22, 45–47]. Several studies highlight the importance of the gut microbiota in modulating host immune responses to pathogens or to vaccination [45–47]. We have recently shown that vaccination against S. Typhi caused no alterations of the microbiota, however, individuals displaying early multiphasic CMI responses harbored more diverse communities than late responders . We have initiated studies to identify the interplay between the host immune response, the microbiota and clinical outcome in volunteers challenged with wt S. Typhi. In addition to these acquired immune response differences, genetic determinants like HLA molecules can also be critical in defining the variation in immune responses. For example, the presence of the HLA-DRB1*04:05 allele was recently associated with protection against S. Typhi .
In summary, we have provided the first direct evidence of an association between higher baseline levels of multifunctional S. Typhi-specific CD8+ T cells and protection, as well as delayed time to disease onset, in an oral challenge model with wt S. Typhi in humans. These studies also revealed some of the immunological responses associated with delayed time to disease onset and defined the homing characteristics of the S. Typhi-specific effector and memory CD8+ T cell populations. This information supports the performance of in depth CMI measurements to aid in the early selection of novel vaccine candidates for further development and evaluation in clinical trials.
TCD, CSW, MML, BA and AJP set up the challenge model and generated the clinical data. TCD, CSW, CJ, and CJB collected and processed the PBMC specimens. SF, MAM and MBS conceived and designed the experiments. SF performed the experiments. SF, MAM, LM and MBS analyzed and interpreted the data. All authors contributed to the writing of the manuscript and approved the final version. All authors read and approved the final manuscript.
This work was funded, in part, by a Strategic Translation award from the Wellcome Trust [grant number 092661], the NIHR Oxford Biomedical Research Centre [Clinical Research Fellowships to CSW and TCD], the Jenner Institute, the Oxford Martin School, the European Union [FP7, Marie Curie Research Fellowship to CJB], by NIAID, NIH, DHHS grants R01-AI036525, U19 AI082655 (CCHI) and U19-AI109776 (Center of Excellence for Translational Research [CETR], and a Passano Foundation Clinical Investigator Award (to MAM). SF was funded in part by NIH Fellowship Training Program in Vaccinology T32-AI07524. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Allergy and Infectious Diseases, the National Institutes of Health, the National Health Service, the National Institute for Health Research (NIHR) or the UK Department of Health.
The authors declare that they have no competing interests.
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