Institute of Medical Microbiology and Hygiene, Department of Virology, Building 47, University of the Saarland, Kirrberger Strasse, 66421 Homburg/Saar, Germany
Correspondence
Andreas Meyerhans
Andreas.Meyerhans{at}uniklinik-saarland.de
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ABSTRACT |
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INTRODUCTION |
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The expression of tissue-specific homing receptors on the surface of effector/memory lymphocytes and their temporal recirculation in blood after antigen encounter allows a simple quality control of immunization routes. By intracellular cytokine staining of antigen-triggered leukocytes directly from blood, it is possible to quantify antigen-specific CD4+ and CD8+ T cells and to characterize their differentiation phenotype and their homing commitments. A 6 h duration of antigenic ex vivo activation ensures that only specific effector and memory CD4+ and CD8+ T cells are detected (Heintel et al., 2002). Naïve T cells, in contrast, need longer times of activation to differentiate and produce cytokines. The additional analysis of CD45RA/RO and CD27 expression allows us to typify the differentiation status, while via CCR7, CD62L and
4
7, their homing to lymph nodes and the gut can be identified, respectively (Mackay, 1999
; Sallusto et al., 1999
).
Poliovirus is a prototype for gastrointestinal viral infections and is close to worldwide eradication due to highly effective vaccines. Poliovirus is a single-stranded RNA virus of the family Picornaviridae and may cause an acute paralytic disease in non-immune humans (Bodian & Horstmann, 1965). It is transmitted via the faecaloral route. It initially penetrates through the M cells of the intestinal epithelium (Sicinski et al., 1990
) and replicates in the Peyer's patches. The virus then circulates through the blood and may enter the central nervous system where it causes paralysis (Bodian & Horstmann, 1965
). Mucosal immunity is of particular importance in protection against poliovirus infection (Faden et al., 1990
; Ogra, 1968
). Protection is mediated through neutralizing antibodies and at least four neutralizing antibody epitopes have been identified (Minor, 1990
; Minor et al., 1990
). However, poliovirus-specific T cells are an important component in poliovirus defence. This was demonstrated by adoptive transfer experiments in poliovirus-receptor transgenic mice, which showed that only primed B cells together with polyclonal poliovirus-specific T cells protected from a lethal intravenous wild-type poliovirus challenge (Mahon et al., 1995
).
For several prevalent and threatening infections, i.e. human immunodeficiency virus (HIV), a mucosal antiviral cellular immune response is considered to be an integral part of any preventative vaccine candidate. From the vast number of vaccine studies it seems apparent that mucosal homing could only be achieved with a mucosal antigenic trigger. However, it seems possible that subsequent non-mucosal booster immunizations may break mucosal commitment as has been suggested (Kantele et al., 1999). To examine whether virus-specific T cells after mucosal vaccination may be boosted via a non-mucosal immunization and whether the mucosal commitment would be broken or not, a group of nine volunteers was followed after poliovirus vaccination. All volunteers have been vaccinated more than 10 years ago with an oral live-attenuated vaccine (OPV). Their cellular immune response to a booster immunization with an inactivated poliovirus vaccine (IPV) was followed with respect to poliovirus-specific T cell frequencies, proliferation, differentiation phenotype and homing commitment. Consistently, an expansion of gut-homing Th1 effector memory responses was observed suggesting that the location of the initial immune trigger determined the subsequent homing properties of the effector T cells.
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METHODS |
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Poliovirus neutralization assay.
Serial twofold diluted serum samples (dilution from 1 : 4 to 1 : 4096) were examined in 96-well flat-bottomed culture plates (Nunc). Diluted sera were mixed with 100 TCID50 of poliovirus types 13 each (in-house serotyped patient isolates) and incubated for 1 h at 37 °C with 5 % CO2 in a standard cell incubator. Green monkey kidney (GMK) cells (2·5x104) were added per well and cultured for 4 days. The cells were fixed with 1 % glutaraldehyde and stained with 1 % crystal violet in PBS. The poliovirus-induced cytopathic effect was analysed by light microscopy. The neutralizing antibody titre was considered to be the highest dilution of serum that protected the duplicate cultures from the cytopathic effect. The variance between repeated titre determinations was within one dilution step.
Poliovirus-specific proliferation of peripheral blood mononuclear cells.
Peripheral blood mononuclear cells (PBMC) were isolated from 10 ml heparinized venous blood by Ficoll-Hypaque density centrifugation (PAA). Cells were adjusted to 1x106 cells ml1 in RPMI 1640 supplemented with 10 % human AB serum (Sigma). Alternatively, serum-free medium (AIM-V; Gibco) was used for cultivation. Triplicate cultures of 1x105 PBMC in 100 µl culture medium were seeded in 96-well round-bottomed microtitre plates (Greiner). PBMC were stimulated from day 5 to day 6 in the presence of 10 µl poliovirus antigen ml1 (equal volumes of complement fixation reagents for poliovirus types 13; BioWhittaker). Stimulation with 50 ng 2FE tetanus toxoid (Chiron Behring) ml1 was used as a positive control. Twelve hours before harvesting, 1 µCi [3H]thymidine (Amersham) per well was added to the PBMC. Cells were harvested on glass fibre filters (Wallac), fused with scintillator wax (Wallac) and [3H]thymidine uptake was measured in a scintillation counter (Wallac).
Stimulation of poliovirus-specific CD4+ and CD8+ T cells within whole blood.
Human venous heparinized blood was collected at days 0, 7, 14 and 21 after immunization. Whole blood was stimulated with titrated amounts of poliovirus antigen (equal volumes of complement fixation reagents for poliovirus types 13, BioWhittaker) in the presence of 1 µg CD28 and
CD49d ml1 (clones CD28.2 and 9F10, respectively; Becton Dickinson) as previously described (Heintel et al., 2002
). As a negative control, blood cells were incubated with the complement fixation reagent control, which did not contain any poliovirus protein (BioWhittaker). Initially we also used IPV (Chiron Behring) and produced poliovirus ourselves from the supernatants of poliovirus-infected GMK cells for stimulation, which gave similar results. As a positive control, cells were stimulated with SEB at 2·5 µg ml1 (Toxin Technologies). Cells were incubated in polypropylene tubes at 37 °C at 6 % CO2 for a total of 6 h. During the last 4 h, 10 µg Brefeldin A (Sigma) ml1 was added to block extracellular transport of cytokines. Thereafter, the blood was treated with 2 mM EDTA for 15 min, the erythrocytes subsequently lysed and leukocytes fixed for 10 min using a Becton Dickinson lysing solution following the manufacturer's instruction. Cells were washed once with FACS buffer (PBS plus 5 % filtered FCS, 0·5 % BSA, 0·07 % sodium azide) and either immediately processed for FACS analysis or left overnight at 4 °C.
Determination and characterization of poliovirus-specific T cells.
Fixed leukocytes were permeabilized with 2 ml FACS buffer containing 0·1 % saponin (Sigma) for 10 min at room temperature. Thereafter, they were immunostained for 45 min at room temperature in the dark using saturating conditions for the following phycoerythrin (PE)- or FITC-labelled monoclonal antibodies: CD4 (clone RPA-T4),
CD8 (clone RPA-T8),
IFN-
,
IL-4,
IL-2,
CD45RA,
CD45RO,
CD27 (all purchased from Becton Dickinson). The PC5-labelled
CD4 (clone 13B8.2),
CD8 (clone B9.11) and the PE-labelled
CD69 (clone TP1.55.3) antibodies were purchased from Coulter-Immunotech. Cells were washed once with 3 ml FACS buffer and fixed in 1 % paraformaldehyde. At least 20 000 CD4+ or CD8+ lymphocytes were analysed on a FACScan (Becton Dickinson) using the CellQuest Software version 3.1. The percentage of poliovirus-specific T cells was calculated by subtracting the frequency of IFN-
-positive cells in the control stimulation from the frequency in the poliovirus antigen stimulation.
The quantification of homing receptor-positive poliovirus-specific T cells was performed by indirect staining since directly fluorescent-labelled antibodies were not available. For the detection of 4
7 and CD62L, mouse monoclonal antibodies were used as the primary antibodies and a fluorescent-labelled rabbit anti-mouse as secondary antibody. To ensure homing-receptor specificity, poliovirus-specific T cell stimulation in these cases was performed in the absence of the mouse-derived co-stimulatory antibodies
CD28 and
CD49d, and cell staining for homing-receptor detection was performed before staining for CD4 and IFN-
.
Stimulated cells were incubated for 45 min at room temperature with the mouse monoclonal antibodies specific for the gut-homing integrin 4
7 (clone ACT-1, kindly provided by Millennium, Cambridge, MA, USA, and Alf Hamann, Berlin) or the lymph node-homing selectin CD62L (clone Dreg-56; Becton Dickinson). After washing the cells with FACS buffer, a PEcyanin-5 (PC5)-labelled rabbit F'ab anti-mouse Ig (DAKO-Cytomation) was used as secondary antibody. To quantify the cytokine receptor CCR7 surface expression, cells were incubated with the rat
CCR-7 antibody (clone 3D12, kindly provided by Martin Lipp, Berlin) and subsequently with a donkey anti-rat FITC-labelled secondary antibody (Coulter-Immunotech). Incubation conditions to detect the other antigens were as described above.
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RESULTS |
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Poliovirus antigen specifically stimulated CD4+ T cells from IPV-immunized individuals. The control antigen failed to induce any relevant cytokine release. No IFN- production before or after IPV immunization was detected in the CD8+ T cells. Induction of IFN-
in CD4+ T cells indicates a Th1 phenotype. Th2 cytokines, like IL-4, were below the detection limit (data not shown). A dot plot of a flow cytometric analysis for a representative volunteer is shown in Fig. 3
(A). Fourteen days post-immunization, 0·37 % of CD4+ T cells produced IFN-
. Only 0·09 % of CD4+ T cells produced IFN-
before immunization at day 0.
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IPV booster immunization induces poliovirus-specific gut-homing effector memory lymphocytes
To characterize the differentiation phenotype of the observed poliovirus-specific Th1 lymphocytes, the reciprocal expression of the memory markers CD45RA and CD45RO was analysed. The majority of poliovirus-specific T cells were of a memory phenotype, as shown by expression of CD45RO and lack of CD45RA (e.g. 86·41±7·89 % RO+ versus 22·82±8·90 % RA+, Table 1). CD27 was analysed as a maturation marker of T cells (Hamann et al., 1997; Hintzen et al., 1993
). Most of the poliovirus-specific Th1 cells were CD27+ (87·40±11·85 %, Table 1
), demonstrating that these were not fully mature effector T cells. Finally, the chemokine receptor CCR7 was used to divide the observed poliovirus-specific lymphocytes into central memory T cells or CCR7 circulating effector memory lymphocytes (Sallusto et al., 1999
). Of the poliovirus-specific Th1 cells, 91·77±7·17 % were CCR7 (Table 1
), showing that they were homing to peripheral tissues and not to lymph nodes.
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DISCUSSION |
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To our knowledge, this is the first description of a gut-homing (4
7+) poliovirus-specific T cell effector memory response in humans. It is well documented that the neutralization of poliovirus infection is mainly due to antibodies. In particular, mucosal IgA responses from gut-homing B cells seem to be required for efficient protection, their induction being dependent on mucosal priming (Herremans et al., 1999
). However, transfer experiments in poliovirus-receptor transgenic mice have demonstrated an important role for virus-specific T cells. Mice were only protected from infection when primed B cells were transferred together with polyclonal poliovirus-specific T cells. The transfer of each leukocyte population alone was not protective. Subsequent experiments with T cell clones showed further that Th1 cells can mediate the protection in vivo through their helper activity for humoral immunity (Mahon et al., 1995
).
The detection, quantification and characterization of virus-specific T cells directly from blood via intracellular cytokine staining is a rapid and reliable means to follow not only virus infections but also vaccinations. While the induction of a cellular immune response may easily be seen and a quantitative comparison of different vaccination schedules performed, a direct correlation with protection is at present not possible. This would require the establishment of correlates between the levels of response and protection from infection or disease development, as has been done for antibody titres. To date, such a correlate has been defined only for CMV-induced disease in immunosuppressed patients after renal transplantation. When the frequency of CMV-specific CD4+ T cells dropped below 0·25 % of the CD4+ T cell population, the patients lost immune control over CMV and had to be treated with antiviral agents (Sester et al., 2001).
It has become increasingly apparent that an efficient antiviral immune response requires the combination of various effector and helper functions. Similar elegant experiments, as mentioned above for poliovirus infections in transgenic mice, have been performed in other mouse virus systems. For example, cooperativity between neutralizing antibodies and T cells is necessary to prevent persistent lymphocytic choriomeningitis virus infections (Baldridge et al., 1997). Likewise, CD4+ T cells, CD8+ T cells and B cells are involved in protection against Friend retrovirus infection (Dittmer et al., 1999
). In consequence, this means that the improvement of new vaccination strategies always should include the testing of the comprehensive adaptive immune response.
In conclusion, peripheral booster immunization with an inactivated vaccine can expand gut-homing memory CD4+ T cell responses in mucosally vaccine-primed humans. Thus the mucosal commitment induced by an attenuated oral poliovirus vaccine is not broken, even after an extended time period of 10 years. The ease by which such T cell responses can now be measured directly from the blood of vaccinees should encourage investigators to incorporate the respective assays into their vaccine trials.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Received 18 December 2003;
accepted 29 January 2004.