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Deficiency in the guanine nucleotide exchange factor DOCK8 causes a human immunodeficiency syndrome associated with recurrent sinopulmonary and viral infections. We have recently identified a DOCK8-deficient mouse strain, carrying an ethylnitrosourea-induced splice-site mutation that shows a failure to mature a humoral immune response due to the loss of germinal centre B cells. In this study we turned to T-cell immunity to investigate further the human immunodeficiency syndrome and its association with decreased peripheral CD4+ and CD8+ T cells. Characterisation of the DOCK8-deficient mouse revealed T-cell lymphopenia, with increased T-cell turnover and decreased survival. Egress of mature CD4+ thymocytes was reduced with increased migration of these cells to the chemokine CXCL12. However, despite the two-fold reduction in peripheral naïve T cells, the DOCK8-deficient mice generated a normal primary CD8+ immune response and were able to survive acute influenza virus infection. The limiting effect of DOCK8 was in the normal survival of CD8+ memory T cells after infection. These findings help to explain why DOCK8-deficient patients are susceptible to recurrent infections and provide new insights into how T-cell memory is sustained.
The recent discovery of the human DOCK8 immunodeficiency syndrome has highlighted DOCK8 and the DOCK super-family of guanine nucleotide exchange factors as important mediators of normal immune function [1-4]. As a group, members of the DOCK family interact with Rho GTPases to regulate pathways that are typically involved in the cytoskeletal rearrangements required to maintain cell structure, migration and adhesion; but the details of their individual functions are largely unknown [3, 5]. The DHR1 (CZH1) domain in DOCK proteins has been shown to bind phosphatidylinositol (3,4,5)-triphosphate (PIP3) generated by PI3K activity, thus promoting localized membrane binding and activation of DOCK proteins , while the DHR2 (CZH2) domain binds to Rho-family GTPase proteins (Rac, Cdc42) and provides catalytic GEF activity to stimulate these proteins and promote integrin reorganization and adhesion, lamellipodia formation, cell polarization, phagocytosis or cell fusion [3, 4]. DOCK8 had been associated with Cdc42 and found to localize to lamellipodia in fibroblasts; but its role in immune function is largely unknown .
Recently described patients who are deficient in DOCK8 fail to mount an effective immune response and are susceptible to recurrent sinopulmonary and viral infections, hyper-IgE, atopy and allergic disease and some types of cutaneous cancer and lymphoma [1, 2]. The majority of patients have low absolute counts of both CD4+ and CD8+ T cells in the blood and impaired proliferation of the cells following mitogen stimulation with a modest decrease in the production of the anti-viral cytokines IFN-γ and TNF-α. These facts suggest a defect in T-cell immunity; yet the nature of any T-cell dysfunction in the absence of DOCK8 remains to be defined.
DOCK8 was also recently identified as being critical for a functional B-cell response in a screen for N-ethyl-N-nitrosourea (ENU) induced mutations that cripple the maturation, but not initiation, of antibody production . Two independent strains (CPM and pri) were found to have loss-of-function DOCK8 mutations causing the cell intrinsic loss of germinal center (GC) B cells during affinity maturation . Further analysis showed that naïve DOCK8-deficient cells are unable to recruit the intercellular adhesion molecule ICAM-1 into the B-cell immune synapse during recognition of membrane-bound antigen. This deficit may explain the impact on GC B-cell survival, which depends on binding antigen and integrin ligands on follicular dendritic cells .
In this study we use the DOCK8-deficient mice to gain insights into the clinical and laboratory findings of DOCK8 immunodeficiency in humans, extending our study of B-cell immunity to examine the T-cell compartment. Like humans, DOCK8-deficient mice have decreased peripheral T cells. Although development of T cells is normal, mature single positive CD4+ T cells accumulate in the thymus and show increased chemotaxis to CXCL12; peripheral CD4+ and CD8+ T cells show increased turnover and decreased survival. Despite these defects, DOCK8-deficient animals are able to generate a comparatively normal primary T-cell response to immunisation with Modified Vaccinia Ankara (MVA) or infection with influenza. However, the memory recall response is significantly reduced due to decreased survival of memory CD8+ T cells. These results may explain the relentless pattern of recurrent infection in the human DOCK8 immunodeficiency syndrome, and may provide clues to how T-cell memory is generated and sustained.
To establish the effects of DOCK8 deficiency on T cells in humans we counted naïve CD45RA and activated effector/memory CD45RO CD4+ and CD8+ T cells in the blood of patients of different ages. This showed a reduction in both naïve and effector/memory numbers, although the latter were less affected and even normal in some patients (Figure 1A). However, it is difficult to interpret the significance of these results or establish the causes of immunodeficiency when T-cell phenotype is continually altered by persistent infection and antigenic stimulation. Therefore, we turned to a recently described ENU mutant, CPM, carrying homozygous null alleles of DOCK8 . The CPM splice site mutation truncates the native protein, removing the DHR2 GEF domain. To investigate the effects of DOCK8 deficiency on T cells, we enumerated CD4+ and CD8+ T cells in the secondary lymph organs of DOCK8-deficient CPM mice (Figure 1B-E).
Examination of spleen, mesenteric lymph nodes (MLN) and peripheral lymph nodes (PLNs) revealed a 2-3 fold reduction in CD4+ and CD8+ T cells compared with WT C57BL/6 controls (Figure 1B-D). Total CD4+ and CD8+ T cells were reduced in the blood as a fraction of total lymphocytes (Figure 1E, upper panel) and in absolute terms, where CD4+ counts were 5.28 ×105/ml (± SE 0.541) in CPM versus 14.53 ×105/ml (± SE 2.132) in WT (p =0.014), and CD8+ 3.32 ×105/ml (± SE 0.240) in CPM versus 5.62 ×105/ml (± SE 0.404) in WT (p=0.008). In contrast, the absolute number of activated CD44hi/CD62Llo CD4+ and CD8+ T cells was more similar in the spleen and PLNs and 1.5-2 fold higher in the MLN of CPM mice compared with that of WT (Figure 1B-D). The increased fraction of activated CD44hi CD8+ and CD4+ T cells was also seen in the blood of CPM mice (Figure 1E, lower panel and data not shown). We conclude that DOCK8 deficiency in mice leads to peripheral T-cell lymphopenia predominantly affecting the number of naïve rather than activated cells, reminiscent of the phenotype in the human immunodeficiency syndrome.
Studies in DOCK8-deficient patients have demonstrated decreased T cell activation and proliferation, but are inconclusive regarding function due to the difficulty in isolating naïve T cells from the peripheral blood [1, 2]. Therefore we analyzed the response of DOCK8-deficient T cells from CPM to antigenic stimulation to see if loss was associated with a defect in activation or differentiation. Overnight activation of naïve DOCK8-deficient mouse T cells with anti-CD3 showed no difference in the upregulation of CD69 and CD25 on CPM and WT CD4+ and CD8+ T cells (Figure 2A); moreover, when stimulated with anti-CD3 and anti-CD28 in vitro, CPM and WT CD4+ and CD8+ T cells underwent the same number of divisions, after five days, in culture (Figure 2B). These findings show that DOCK8-deficient T cells from uninfected mice are able to respond normally to antigenic stimulation and proliferate freely in vitro.
To explore T cell differentiation we analyzed the cytokines produced by activated/memory CD45RO+ CD4+ T cells from the blood of DOCK8-deficient patients, parents and controls stimulated ex vivo with PMA and ionomycin. A higher proportion of activated cells secreting TH2 cytokines IL4 or IL13 were observed in 3/5 and 4/5 DOCK8-deficient patients respectively, a finding that may be linked to the phenotype of hyper-IgE in DOCK8 immunodeficiency whilst, the number of cells secreting IFN-γ was variable (Suppl Figure 1A). However, the degree to which these phenotypes reflect ongoing infection, over-stimulation/exhaustion or other secondary effects caused by the immunodeficient state was again uncertain. Therefore, we asked if CPM CD4+ T cells could differentiate normally under TH1 or TH2 polarising conditions in vitro. Under these conditions, Tbet and GATA3 were induced appropriately (Suppl Figure 1B). IFN-γ production by TH1 cells was equivalent, and only the proportion of IL4 secreting cells was higher in CPM mice under TH2 conditions (Suppl Figure 1C and 1D). It is possible that DOCK8 and integrin dependent effects on the signalling threshold of the TCR or other receptors, or small differences in the basal and induced levels of T-bet and GATA-3, might contribute to such a bias in humans and mice. Overall, however, the differentiation of effector cell subsets is not intrinsically unimpaired and therefore unlikely to contribute to the state of immunodeficiency.
We next asked if abnormal thymic development might explain the reduction in peripheral T cells. We found no difference in the numbers of developing T cells at the DN or DP stage (Figure 3A) or in the number of mature CD8+ T cells (Figure 3A and 3B); and no consistent differences were seen in CD3, CD5, TCRβ, CD44 or CD25, staining (data not shown). The sole abnormality observed was an accumulation of mature single positive CD4+ T cells, particularly the most mature CD4+CD62LhiCD69lo population (Figure 3A, right panel, and and3B).3B). On its own this may contribute to the peripheral lymphopenia, but cannot explain the deficiency of CD8+ T cells. Labelling CPM and WT mice with BrdU showed no difference in the absolute number of BrdU+ single-positive CD4+ T cells generated over a 40hr period (1.00 × 106 ± SE 0.14 in CPM vs. 1.33 × 106 ± SE 0.18 in WT, p=0.1674; unpaired t-test), implying that accumulation of mature DOCK8-deficient CD4+ T cells is due to delayed egress.
To examine if T-cell lymphopenia in DOCK8 deficiency was cell intrinsic we reconstituted lethally irradiated CD45.1 allotype-marked recipients with 50:50 or 20:80 mixtures of WT (CD45.1) and CPM or WT (CD45.2) bone marrow. Whilst the 50:50 WT:CPM bone marrow generated the expected 1:1 ratio of follicular B cells it failed to reconstitute normal numbers of DOCK8-deficient T cells due to a block in T cell development prior to the DN stage, which was not previously seen in the absence of competition (data not shown). The cause of this competitive disadvantage in establishing early thymic development is unknown; but it is not limiting in non-chimeras where the number of cells at the DN stages of thymic development are similar in CPM and WT mice (Figure 4A). Analysis of 20:80 WT:CPM mixed chimeras showed the same competitive failure to reconstitute thymic development relative to the B cell lineage but no effect of DOCK8 in the transition from DN to DP and mature single-positive cells (Figure 4A & data not shown). However, naïve peripheral T cell numbers were reduced 2-3 fold in CPM compared with WT mice (Figure 4A), which suggests a cell intrinsic defect in peripheral CD4+ and CD8+ survival underlying the immunodeficiency. The absence of CD4+ cell accumulation in the thymus of mixed chimeras also supports the idea that this is not the principal cause of peripheral lymphopenia (Figure 4A). Moreover, the absence of greater numbers of peripheral CPM CD44hi T cells in the mixed chimeras (Figure 4B) implies this is due to homeostatic proliferation in unmanipulated DOCK8-deficient animals, where T cell numbers are low.
B and T cells from DOCK2-deficient mice fail to migrate to a variety of chemoattractants, resulting in the loss of T cell thymic egress in response to S1P, and peripheral lymphopenia [9, 10], therefore we considered whether similar effects could contribute to immunodeficiency in the absence of DOCK8. To assess chemotaxis, we mixed equivalent numbers of mature allotype-marked thymic T cells from CPM and WT and assessed their ability to migrate in response to constitutively expressed cytokines CXCL12, CCL19/21 and S1P. In contrast to DOCK2-deficient mice, the migration of thymic and peripheral T cells to these chemokines was normal, with the exception of a consistent increase in the migration of mature DOCK8-deficient thymic CD4+CD62LhiT cells towards CXCL12, (Figure 5A; p=0.0224 by ANOVA, combining data from 3 experiments). As CXCL12 signalling via CXCR4 is important in lymph node entry , we transferred 50:50 mixtures of mature thymic CPM & WT T cells into WT recipients and plt/plt recipients, which lack expression of the other main T cell attractants CCL21 and CCL19 . In these experiments, the proportion of thymic CPM CD4+ CD62Lhi T cells in peripheral lymphoid organs after 90 mins was increased in comparison to WT cells (Figure 5B). These findings are consistent with an increased chemotactic response to CXCL12 but also confirm that entry and localisation of cells to peripheral lymphoid organs is normal in unimmunised mice.
Twenty-four hours after transfer, however, the proportions of mature thymic CPM CD4+ and CD8+ T cells were reduced relative to WT (Figure 5C), suggesting that DOCK8 deficiency causes these emigrants to survive less well after localisation to secondary lymphoid organs. We found that expression of the survival factor Bcl-2 is reduced in CPM single positive CD4+ and CD8+ thymocytes, suggesting that DOCK8-dependent activation of anti-apoptotic pathways may contribute to survival at this stage in T cell development (Figure 5D).
Together, the above results suggested that defects in cell survival were the principal reason for the loss of mature T cell populations in DOCK8 immunodeficiency. To evaluate this further, CPM and WT mice were fed BrdU for 15 days and the survival of labelled cells was followed for a further 30 days. In the first phase, BrdU labelling of CD4+ and CD8+ T cells was equivalent or more rapid in the spleen and lymph nodes of CPM compared with WT (Figure 6A-D). This is consistent with a higher level of homeostatic proliferation and the majority of newly labelled cells were CD44hi (data not shown). More importantly, during the washout phase, DOCK8-deficient CD4+ and CD8+ T cells disappeared much faster in all tissues, confirming a survival defect in mature T cells (Figure 6A-D).
During a normal immune response to pathogens in vivo, the activation, proliferation and differentiation of T cells leads to short-term effector cells and long-lived memory cells, which survive after contraction of the effector population and act to curtail recurrent infection. The in vitro experiments, we have described, suggest that the primary effect of DOCK8 might not be on the generation of short-term effector cells but rather the survival of long-lived memory cells. To investigate these possibilities we compared the response of CPM and WT mice to vaccination with replication deficient Modified Vaccinia Ankara virus (MVA). After 9 days there was no significant difference between CPM and WT in IFN-γ, IL2 or TNF-α production by antigen-specific CD8+ T cell directed against MVA (Figure 7A and data not shown) or IFN-γ ELISpot (data not shown), either as percentage of CD8+ T cells or in absolute terms. In three separate experiments, this analysis showed no difference between CPM and WT confirming that the initial phase of the T-cell response is not functionally limited by the lymphopenia in pre-immune animals. Anti-MVA specific CD8+ T cell numbers also remained similar after the contraction phase (Figure 7A); therefore, there was no premature collapse of the effector T-cell population in the absence of DOCK8. However, 42 days after the primary immunisation, during the phase of long-lived memory cell survival, the number of antigen-specific cells was significantly reduced (Figure 7A). The loss of surviving T cells was not due to a block in memory cell differentiation as numbers of memory precursor cells (MPECS), defined as KLRG1low CD127hi (IL7Rhi), were unaltered (Figure 7B). Furthermore, expression levels of memory cell markers CD127+, CD62L+, KLRG1+ and Bcl-2 were similar on MVA-specific WT and CPM T cells at day 9, 14, 42 and after rechallenge supporting this conclusion (Figure 7C and data not shown). After 55 days, the response to rechallenge with MVA in vivo was reduced in DOCK8-deficient mice compared with WT (Figure 7A). However, the fold-increase in the secondary response was proportionate to the number of surviving memory cells (Figure 7A), once again suggesting that it is the survival of memory cells rather than the generation of the secondary effector response that is limiting for CD8+ T-cell immunity in the absence of DOCK8.
To investigate further the nature of DOCK8 deficiency in the maintenance CD8+ memory T cells, we repeated our MVA immunization protocol with chimeric mice reconstituted as before with 20:80 mixtures of WT (CD45.1) and CPM or WT (CD45.2) bone marrow. In this competitive setting, DOCK8 deficiency resulted in reduced primary CD8+ responses to MVA relative to WT cells (Figure 7D). The loss of DOCK8 CD8+ response in this competitive situation is consistent with an underlying defect in survival, which is normally compensated for by increased proliferation in non-chimeras. The greatest effect of competition is on the KLRG1hi short-lived effectors, which are turning over most rapidly (Fig 7D middle panel). During the maintenance phase of memory, day 42, DOCK8 responders were further affected when compared with day 9 MPEC. Together these data support an intrinsic role for DOCK8 in the maintenance of CD8+ T cell memory. The WT and DOCK8 CD8+ memory T cells expressed similar levels of markers such as CD127+ (IL7R) and KLRG1 as well as the survival factor Bcl2 (Figure 7E).
We also examined T-cell responsiveness in DOCK8-deficient cells during pathogenic infection by challenging CPM and WT mice with mouse adapted influenza strain x31(A/HK-x31 (H3N2)). Using daily weight loss as a clinical parameter we found no significant difference in disease severity or the ability to survive the acute infection through the nadir of infection at day 6 to recovery by day 10 (Figure 8A). 11 of 12 WT mice survived the challenge, while 9 of 12 CPM mice survived. Seven days post immunization, there was no significant difference between WT and CPM in IFN-γ production and numbers of LAMP1hi CD8+ T cells to T-cell restricted epitopes with similar expression profiles of CD127+, KLRG1 and CD62L+ (Figure 8B and data not shown). Numbers of MPECS were also produced to normal levels (data not shown). These findings again indicate that the immunodeficiency in the absence of DOCK8 is not due to an inability to mount an effective response to acute infection. To assess the memory response we bled mice 56 days after primary infection and counted LAMP-1hi CD8+ T cells responsive to conserved internal ‘flu T-cell restricted epitopes (Figure 8B). Whilst WT mice produced a robust response to x31, memory T cells were not sustained above background in CPM mice; these data were substantiated by blood ELISpot (data not shown). Together these findings show that the functions of DOCK8 are not limiting during primary T-cell immune response against viral infection but are critical for maintenance of memory to vaccination and viral infection.
The results presented here contribute to our understanding of how deficiency in DOCK8 can increase susceptibility to disease, particularly viral infection [1, 2]. Our findings show that DOCK8 operates at multiple steps in cellular immunity, but its main effects are on T-cell survival, and it is most limiting in the survival of memory T cells. The inability to generate high affinity antibodies to pathogens due to a failure to sustain GC B cells, shown previously, and the absence of T-cell memory shown here may explain the increased susceptibility of human DOCK8 immunodeficiency syndrome patients to recurrent infections [1, 2, 7].
The relative paucity of peripheral T cells in DOCK8-deficient mice is not due to defective thymic development. Although DOCK8-deficient T cells have a selective disadvantage in populating early thymic development relative to WT cells in mixed chimeras this is not limiting in natural mutants or single bone marrow chimeras. B-cell development is normal in the mixed chimeras and so competitive effects on the early thymus might be due to the defective survival, proliferation and migration of T cell precursors. However, at terminal thymic development, DOCK8-deficient single positive CD4+ T cells accumulate in the thymus and these CD4+ cells show increased migration to SDF-1/CXCL12. Emigration from the thymus is mediated primarily by trafficking to S1P, and whilst CXCL12-dependent signals have been suggested to play a role in thymic egress it remains to be formally demonstrated in physiological circumstances [12, 13].
The discovery that mature naive thymic CD4+ T cells show increased migration to CXCL12 does raise the interesting possibility of a connection between DOCK8 and the human WHIM syndrome, which is a combined immunodeficiency defined by warts, hypogammaglobulinemia, immunodeficiency, and myelokathexis causing neutropenia . WHIM and DOCK8-deficient patients share a similar spectrum of recurrent respiratory and cutaneous viral infections  although not all clinical features are comparable. T cells from patients with the WHIM syndrome also show augmented T-cell responses to CXCL12, and a substantial number of these patients have mutations in the CXCR4 chemokine receptor gene [14, 15]. DOCK8 may, therefore, be a regulator of CXCR4 signalling and speculatively, WHIM patients with no obvious abnormality in CXCR4 dependent signalling may have mutations in DOCK8-dependent signalling pathways [15, 16].
In theory, the decreased number of naïve T cells in the peripheral T-cell compartment could be due to defects in thymic egress, differentiation and proliferation or the increased death of cells. However, our data suggest that decreased survival is the principal reason for the loss of T cells in DOCK8-deficient animals. The increased turnover of DOCK8-deficient T cells in vivo and the increase in activated or memory-like (CD44hi/CD62Llo) T cells is likely to be caused by lymphopenia-induced proliferation, which occurs when T cells in lymphopenic hosts divide and irreversibly acquire a ‘memory-like’ phenotype . The increase in CD44hi/CD62Llo T cells is not present in mixed chimeras where cell numbers are normal, and so, in this case, the loss of naive peripheral DOCK8 T cells must be due to an intrinsic defect in survival rather than forced differentiation and division. More importantly, lymphopenia affecting the primary repertoire does not seem to be limiting in the primary response to vaccination or infectious challenge, when proliferation can presumably compensate for the reduction in T cells, whereas the most striking effect of DOCK8 deficiency is on the survival of memory CD8+ T cells.
The CD8+ T cell immune response can be divided into three phases : the first phase is of growth and differentiation of naïve CD8+ T cells into effector cells with lytic activity and cytokine production; the second is a phase of 10-20 fold contraction leading to a smaller population of long lived memory cells, which arise by the differentiation of effectors [19, 20]; and the third, a phase in which the memory cells are maintained in the host [20, 21]. Long-lived CD44+CD62LhiCD127+ central memory T cells (TCM) arise after 5 days following a typical infection and survive within secondary lymphoid organs where they can respond swiftly in a recall response [20-23] The generation of the memory cells and the expansion of the immune response depend on TCR signalling as well as costimulatory molecules including CD28, CD40, 4-1BB, CD27, ICOS, and/or OX40, IL2 and inflammatory cytokines including IL-12 and IFN-α [24-29]. The normal response following MVA vaccination or influenza infection and the initial generation of memory cells suggests that DOCK8 is not limiting downstream of these pathways. The long-term survival of memory CD8+ T cells does not require the presence of antigen but is dependent on cytokines, particularly IL-15 and IL-7 [30-33]. However, the absence of defects in lymphopenia-induced homeostatic expansion and NK cell development (Greg Crawford, unpublished observations) which depend on IL-7 and IL-15  argues against a deficiency in cytokine signalling as the cause of CD8+ T-cell memory failure in the absence of DOCK8.
Since DOCK8 mutant B cells are unable to cluster ICAM-1 following antigen-receptor signalling it is tempting to speculate that similar defects in integrin-dependent signalling might affect the memory T-cell response. Although ICAM-1-dependent interactions between T cells and DCs play a role in the formation of CD8+ memory T cells [34, 35], the absence of ICAM-1/LFA-1 interactions has already been described as having its greatest effect on the survival of memory cells [36-38]. It is interesting to speculate that a dependence on external signals from within the cellular niche might also account for the low Bcl-2 expression in DOCK8-deficient single positive thymocytes and their decreased survival on transfer into the periphery.
How DOCK8-dependent reorganization of the cytoskeleton could affect lymphocyte survival is also not yet clear. However, a decreased memory response to viral infections is also reminiscent of Wiskott-Aldrich Syndrome (WAS), which is an X-linked, immunodeficiency syndrome principally caused by mutations in the WAS protein (WASP) gene . WASP regulates the actin cytoskeleton in a variety of immunological functions including cell migration and the formation and the preservation of T-cell memory . WASP and WASP-family verprolin-homologous (WAVE) proteins are fundamental actin-cytoskeleton reorganizers found in eukaryotes. The conserved function across species is to receive upstream signals from Rho-family GTPases and then activate the actin-related protein (ARP) 2/3 complex, leading to rapid actin polymerization [41, 42]. There are a number of striking similarities between WAS and DOCK8-deficiency including peripheral T-cell lymphopenia, a lack of marginal zone B cells and poor immunological synapse formation in WASP-deficient B cells as a consequence of impaired BCR and integrin signalling [43, 44]. Cdc42, the probable target of DOCK8, is an important regulator of WASP function . Therefore it is an intriguing possibility that DOCK8 augments the formation of active Cdc42, at the membrane interface thereby signalling through WASP. Further investigation of such pathways and their relationship to the human immunodeficiency syndromes will provide important insights into how T-cell memory is sustained.
Human studies were approved by the institutional review board of the National Institute of Allergy and Infectious Diseases in accordance to Declaration of Helsinki principles. Mouse experiments were performed under UK Home Office License or in accordance UCSF Institutional Animal Care and User Committee approved protocols. CPM mice were generated by ENU-mutagenesis ; C57BL/6 plt/plt mice were used for transient transfer experiments . Bone marrow chimeras, C57BL/6 (CD45.1+) mice were irradiated with 2 fractions of 4.5 Gy spaced by 3 hours and reconstituted with 5-10×106 bone marrow cells, comprising 50:50 or 20:80 mixtures of C57BL/6 (CD45.1+) bone marrow and either CPM or C57BL/6 (CD45.2+) cells and allowed to reconstitute for 8–10 weeks.
Human PBL and mouse lymphocytes were isolated and stained as previously described [1, 7]. Antibodies were from B.D. Pharmingen or eBioscience® unless otherwise stated; FITC-conjugated antibodies were against: CD24, BrdU, CD8α, CD62L and CD44 (Caltag). Phycoerythrin-(PE) conjugated antibodies were against: CD5, CD3, CD69, FoxP3, CD62L, CD45.1, CD25 and CD4 (Caltag). PE-Cy7 conjugated antibodies were against B220, CD69, CD8α, CD4 and CD45.1. PE-Cy5.5 conjugated antibodies were against CD8α. Allophycocyanin-(APC) conjugated antibodies were against: CD44 and CD4 (Caltag). APC-Cy7 conjugated antibodies were against CD4. APC-Alexa-Fluor® 750 conjugated antibodies were against: CD45.1. Tricolour-(TC) conjugated CD4 and CD8α (Caltag). BD TruCOUNT tubes were used to enumerate the leukocyte count as per manufacturer’s instructions. Data was collected on FACSCantoTM or LSRII machines (B.D.) and analysed with FlowJo (Tree Star, Inc.) or FACSDiva (B.D.).
Chemotaxis assays were performed as described . To assess T cell activation, splenocytes were plated in 96 well plates coated with anti-CD3 antibody at 10μg/ml (145-2C11) and 10μg/ml anti-CD28 ascites (37.51) in PBS. Cells were incubated at 37°C overnight in RPMI containing 10% FCS. To assess proliferation, CD45.2 WT or CPM lymphocytes were mixed with CD45.1 WT cells to give 2×106 cells/ml and labeled for 8 min at RT with an equivalent volume of CFSE (final 2μM). The reaction was quenched by the addition of FCS and washed 3 times with RPMI-1640 containing 10% FCS. 2×105 CFSE labeled cells were cultured on anti-CD3 and CD28 coated plates for 3 days.
In transient transfer experiments, ~1-2×107 cells per ml were labelled with either 3.3 or 10μM carboxyfluorescein-diacetate –succinimidyl-ester ( C F S E ; Invitrogen / Molecular Probes ) o r 1 0μ M 5-(and-6)-(((4-chloromethyl)benzoyl)amino)-tetramethylrhodamine (CMTMR, Invitrogen/Molecular Probes) in RPMI-1640 containing 2% FCS for 10 to 20 minutes at 37°C then washed by centrifugation through FCS. For cell transfers, recipient mice received ~1-2×107cells by i.v., normalised to account for T-cell number. Recipients were sacrificed at designated times.
BrdU (Sigma/ B.D.) was given at 0.25mg/ml in drinking water and supplemented with 1% glucose (Sigma), for upto 15 days. Cells were fixed in 0.5% paraformaldehyde, permeabilised in 3M HCl/0.5% Tween20 (for 20-min at RT), neutralised with 0.1M Borax (Sigma), washed, and stained with FITC-conjugated anti-BrdU 3D4 (B.D. Pharmingen).
For MVA immunisation or influenza challenge, mice were anaesthetised and immunised intradermally with MVA expressing green fluorescent protein at a dose of 1×106 PFU/mouse (2×107 PFU/ml)  or infected intranasally with 25μl of Influenza X-31 (A/Aichi/68; H3N2) at a dose of 512 HAU/ml. Mice were weighed daily as an indicator of health. Animals reaching 80% of original body weight were humanely culled. Antigen-specific cells in the draining PLNs (MVA) were identified by intracellular staining for IFN-γ (early response) or LAMP-1 (memory) after stimulation with anti-CD3 or epitopes from MVA B8R epitope (sequence TSYFKESV) or a 1μg/well of pooled 20mer peptides spanning conserved internal nucleoprotein and matrix 1 protein epitopes.
Statistical comparisons were performed with Prism 4.0 Software (GraphPad Inc.), using ANOVA or two-tailed paired t-tests for comparisons.
We thank the staff of the Oxford Biomedical Services Unit for animal husbandry and Dr Helen Chapel, Prof. Sarah Gilbert, Prof. Adrian Hill and Dr Ronald Schwartz, for advice and reagents. This work was supported by the Medical Research Council, NIHR Biomedical Research Centre Programme, and the Intramural Research Program of the NIH, NIAID.
Conflicts of Interest The authors declare no financial or commercial conflict of interest.