Download Dissecting Immune Responses

Appendix 28
Dissecting Immune Responses
Miriam Windsor, Nicholas Juleff, Mandy Corteyn, Pippa Hamblin, Veronica Carr, Paul V Barnett,
Bryan Charleston*
Pirbright Laboratory, Institute for Animal Health, Ash Rd, Woking, Surrey, GU24 0NF, UK.
Abstract:
Objectives: To understand the role of CD4 T cells in protective immune responses against FMDV
infection of cattle. The role of CD4 T cells was investigated during acute infection of naïve animals
and the magnitude of specific CD4 responses was correlated with protection in vaccinated animals.
Materials and methods: Two separate animal studies have been carried out. In the first experiment
naïve animals were depleted of circulating CD4 T cells with monoclonal antibodies before
intradermolingual inoculation with the O UKG 34/2001 strain of FMDV. Neutralizing antibody titres,
clinical signs and memory CD4 T cell responses were measured pre and post challenge.
In the second series of experiments, animals were vaccinated with O serotype vaccine from the UK
emergency vaccine bank. Total and neutralizing anti-FMDV antibody titres were measured at time
points post-vaccination and these titres correlated with the magnitude of the specific T cell
response. One year post vaccination one group of animals was challenged intradermolingually with
homologous virus.
Results: Elimination of CD4 T cells using monoclonal antibody depletion did not influence the
magnitude or duration of clinical signs or reduce the development of neutralizing antibody titres
post-challenge.
There was a correlation between the duration and magnitude of the humoral response with the
magnitude of the cellular immune response. Also, assessing the cellular immune response
improved the ability to predict protection after intradermolingual challenge with homologous virus.
Discussion:
Our preliminary results suggest that the early antibody production in naïve cattle to FMDV is
predominantly a T independent response. Also, the development and control of clinical signs are
not dependent on the presence of circulating CD4 T cells.
Using neutralizing antibody titre combined with specific CD4 T cell proliferative response could
provide an improved correlation with protection from FMDV clinical signs, compared to antibody
titre alone.
Introduction:
Foot and mouth disease virus (FMDV) has a wide host range including all cloven-hoofed animals
and causes an acute vesicular disease in domestic ruminants and pigs, which results in debilitation,
pain and loss of productivity (Alexandersen et al., 2003). Commercial vaccines, which comprise
inactivated virus particles incorporated in adjuvant, are widely used in those parts of the world
where FMD is prevalent (Barnett and Carabin 2002, Barnett et al. 2001). However, despite its
widespread use, the vaccine has a number of shortcomings (Doel 1999). Immunity is relatively
short-lived and hence animals need to be re-vaccinated at 4-6 month intervals. By contrast
available data, although limited, suggest that immunity in animals that have recovered from
infection with FMDV may last up to 3 years (Cuncliffe 1964).
Novel approaches to the development of new vaccines against FMD are limited by a relatively
superficial understanding of the immunology of the disease in the target species. Evidence from
model virus systems in laboratory animals has shown that, while either antibody or T cell-mediated
effector mechanisms can have a dominant role in immunity to different viruses, both arms of the
immune response are often required to maximise protection – antibody to neutralize and remove
free virus, and T cells to inhibit viral replication or to kill-virus-infected cells (Zinkernagel 2002).
The CD4 and CD8 subsets of T cells both have the potential to act as effectors; the former are also
generally required to provide help for antibody production. In the context of FMDV, there are a
number of differences between the immune responses induced by infection and vaccination that
potentially could influence the quality and/or duration of immunity to FMDV.
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By contrast to antibody, knowledge of T cell responses to FMDV is limited. CD4 T cell responses
have been described in infected and vaccinated animals and in both cases were found to be crossreactive between virus serotypes (Collen et al, 1998). CD4 T cells from infected animals recognised
both structural and non-structural proteins. Intriguingly, the dominant viral protein recognized by
vaccinated animals was 3D (Collen et al, 1998), which is a non-structural protein but recently has
been identified as a minor component of intact virions (Newman et al., 1994). Vaccinated animals
also respond to structural proteins and several CD4 T cell epitopes have been identified in these
proteins (Collen et al, 1991; van Lierop et al, 1994 and 1995). One study of vaccinated animals
reported that the responding CD4 T cells express IL-2 and IFN-γ but not IL-4, suggesting a Th1type response (van Lierop et al, 1995). Experiments in T cell-deficient mice have provided evidence
that, in common with a number of other viruses, FMDV can induce T-cell-independent antibody
responses (Borca et al, 1986). Evidence from other viral systems indicates that this is a
consequence of the repetitive arrangement of B cell epitopes on the surface of the viral particles.
Bachmann and Zinkernagel (1996) have argued that viruses with this property are able to generate
very rapid antibody responses because early expansion of B cells by T cell-independent
mechanisms facilitates an accelerated T-dependent antibody response. If this is correct, it has
important implications for vaccine design since induction of this response would require retention of
a complex antigen configuration in the vaccine. Studies have been initiated to determine the T cell
dependency of antibody response to FMDV in cattle and to investigate the extent to which
immunity in vaccinated animals is dependent on a strong CD4 T cell memory response.
Materials and Methods:
Animals: The animals used in the studies to correlate antibody response with T cell proliferative
response were derived from a herd under a controlled breeding scheme where a selection
according to MHC haplotypes is applied. The animals used showed serological specificity for either
BoLA class II allele DRB3*0701or DRB3*2002 (Collen et al., 2002).
Specific T cell subset depletion studies were performed in commercial out-bred calves.
Clinical scoring: A clinical scoring system based on rectal temperature, the occurrence of lesions on
feet and mouth, lameness and nasal discharge was used.
Monoclonal antibody depletion of T cell subsets: Calves weighing approximately 80kg were treated
over a three day period with 50mg of purified IgG2a monoclonal antibodies against either CD4,
WC1 antigen (expressed on circulating gamma delta T cells) or control antibody (specific for Turkey
Rhinotracheitis virus). Monoclonal antibody depletion was initiated one day prior to challenge.
Lymphocyte proliferation assays: Heparinized blood was collected and peripheral blood
mononuclear cells (PBMC) isolated according to a standard procedure. PBMC were resuspended in
RPMI 1640 with L-glutamine (Gibco), supplemented with 10 % fetal calf serum (Biochrom, UK) and
plated (2 x 105 cells/well) in 96-well round-bottomed microtitre plates (Falcon). Control and test
antigens were added to each well at concentrations described in the results. After five days,
cultures were pulsed with 1 µCi of methyl-3H-thymidine for 18 hours. Cells were collected and
incorporation of 3H-thymidine was measured by liquid scintillation counting.
Flow cytometry: All stages were performed at room temperature. Cells were seeded at 2 x 105
cells/well in a u-bottomed 96 well plate. The cells were washed in FACS Wash Buffer (FWB) by
centrifugation at 1300rpm for 3 minutes. The supernatant was flicked off and the cells vortexed to
resuspend them. Primary antibodies were added at 1µg/ml in a total volume of 25µl. The cells were
incubated for 10 minutes followed by two washes. The cells were then incubated with 25µl of
isotype specific fluorochrome for 10 mins, protected from light, followed by another two washes.
The cells were then diluted into 350µl FWB and analyzed immediately on a FACS machine. If
analysis could not be performed immediately, cells were fixed with 100µl 1% paraformaldehyde,
stored at 2-8°C and protected from light. Recovery of cells was performed by adding 100µl FWB to
wells and centrifuging as above. Live cells were gated according to their forward and side scatter
profiles. Results were analyzed on FACSExpress V3.
CD4 proliferation assays: PBMC were isolated from whole cow blood by density gradient
centrifugation. Blood was diluted 1:1 with D-PBS, underlaid with Histopaque-1083 and centrifuged
for 30 minutes at 2500rpm at room temperature. Subsequent centrifugations were carried out at
8°C. PBMCs were removed from the interface and washed in D-PBS by centrifugation, once at
1800rpm for 10 minutes and twice at 1200rpm for 8 minutes. Prior to the third wash, a 5µl sample
of cells were diluted in 95µl of 0.1% trypan blue for counting.
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An aliquot of PBMC were isolated irradiated and seeded into u-bottomed 96 well plates at 3 x 103
cells/well in 100µl of media. PBMCs were initially incubated with 500µl of cc30 (IgG1 antibody) in
5mls of D-PBS for 10 minutes at room temperature. The suspension was then washed twice in DPBS and the cell pellet incubated with anti-IgG1 magnetic beads for 10 minutes at room
temperature. After two more washes, cells were positively selected on an LS MidiMACS column.
Cells were counted and seeded over the antigen-presenting cells at a concentration of 1x 105 cells /
well in a volume of 50µl. Antigens were added in triplicate at the appropriate concentrations in a
total volume of 100µl. Pokeweed mitogen was used as a positive control and background was
assessed in wells containing only antigen presenting cells and CD4+ cells.
FMD virus neutralization test: Titres of neutralizing FMD antibodies were measured by the microneutralization assay as described in the OIE Manual in which antibody end point titres are
calculated as the reciprocal of the last serum dilution to neutralize 100 TCID50 of the virus per well.
The same strain of virus was used in the test as in the in vivo cattle potency test.
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Results:
In vitro antigen specific proliferation of peripheral blood mononuclear cells to FMDV antigen is
dependent on the presence of CD4 T cells.
Two animals with established specific proliferative responses to FMDV antigen post vaccination
were selected to determine whether the specific proliferative response was dependent on CD4 T
cells. The results in Figure 1 show the animals have a specific response to FMDV (a). Depletion of
CD 4 T cells from the PBMC results in a loss of a specific proliferative response (b). However, the
addition of purified CD4 cells to irradiated whole PBMC as antigen presenting cells results in a
detectable specific proliferative response (c).
FMD 2 PBMC responses
a
FMD 4 PBMC responses
800000
Mean cpm
600000
400000
200000
0
Antigen
FMD 2 PBMC-CD4 responses
FMD 4 PBMC-CD4 responses
800000
b
Mean cpm
600000
400000
200000
0
Antigen
FMD 2 CD4 +APC responses
c
FMD 4 CD4 + APC responses
Mean cpm
800000
600000
400000
200000
only
Cells
Ova
PWM
Media
only
Cells
Ova
PWM
Media
0
Antigen
Figure 1
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The magnitude of the FMDV specific T cell proliferative response is associated with neutralizing
antibody titre and protection.
a
c
1400
VNT
1200
1600
FMD 1
1000
1400
800
Virus Neutralisation titre
FMD 2
FMD 3
600
FMD 4
400
FMD 5
200
0
0
50
100
150
200
250
300
350
400
1200
FMD6
1000
FMD7
800
FMD8
FMD9
600
FMD10
400
200
Days
0
-13
-10
b
0
1
2
3
4
5
7
8
10
13
Weeks post Vaccination
1, 2 and 4 re-immunised
d
600000
200000
180000
500000
160000
140000
400000
120000
100000
300000
80000
200000
60000
40000
100000
20000
0
0
1
2
3
4
5
6
7
8
9
10
Figure 2
Initial observations relating antibody titre to specific proliferative response of total PBMC of cattle
vaccinated with O Manisa commercial vaccine are shown in Figure 2. (a) Neutralizing antibody
response of five vaccinated cattle assessed for 1 year post-vaccination (MHCII haplotype DRB3*
0701). All animals had a neutralizing antibody titre of 1/45 or less. (b) Specific proliferative
response to FMDV (open bars) and Ovalbumin (closed bars) on the day of challenge (1 Year post
first vaccination). Only animals 2 and 4 were protected from homologous challenge.
(c) Specific virus neutralizing antibody response of five animals (MHCII haplotype DRB3* 2002)
immunized with commercial O Manisa vaccine. All animals were re-vaccinated after 10 weeks and
the specific virus neutralizing antibody response measured 3 weeks later. (d) The specific
proliferative response to FMDV (open bars) and Bovine Herpes Virus (closed bars) 3 weeks post
second vaccination is shown. There is a tendency for animals with high antibody titre to also have
a higher specific proliferative response.
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Depletion of CD4 T cells or WC1+ γδ T cells does not influence the development and resolution of
clinical signs or circulating neutralizing antibody.
After challenge of the monoclonal antibody treated animals with viral inoculum the onset and
duration of clinical signs was similar in all animals. One of the CD4 T cell depleted animals showed
less severe lesions, although the onset and duration of those lesions were consistent with the other
calves. All calves were clinically recovered by 7 days post-challenge except for the presence of
healing lesions (Figure 3). The effectiveness of the monoclonal antibody depletion protocol was
assessed by determining the percentage of the target cell population in circulation at various
timepoints pre and post-treatment. The target populations were effectively depleted for at least 7
days (Figure 4 d-f). All animals had neutralizing antibody titres greater than 1/45 by day 7 postchallenge (Figure 4 a-c).
18
16
14
12
10
8
6
4
2
0
-1
0
1
2
3
4
5
6
7
8
9
10
11
12
Figure 3 Clinical scores post-FMDV challenge of calves depleted of either CD4 or WC1+ γδ T cells.
Control animals — — —; CD4 T cell depleted animals ------;
γδ T cell depleted animals ———
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a
d
RZ 57 ( Depletion control)
T R T 3 C o n t r o l s VN T
800
30
700
25
% lymphocytes
600
500
400
300
200
WC1
20
CD4
15
CD8
10
CD21
5
100
0
0
0
1
2
3
4
5
6
7
9
13
16
21
23
27
-10
30
-5
0
5
CD4 depleted VNT
b
10
15
20
Days post infection
D a y s pos t i nf e c t i on
CD4 Depletion
e
25
800
700
20
% lym p hocytes
600
V N T titre
500
400
300
200
15
10
5
100
0
0
0
1
2
3
4
5
6
7
9
13
16
21
23
27
-5
30
-1
0
4
9
13
W C1 Depletion
Gamma delta depleted VNT
c
7
16
Days post infe ction
Days post infe ction
f
30
800
700
25
% ly m p h o c y tes
600
VN T t itre
500
400
300
200
20
RZ 51
15
RZ52
10
5
100
0
0
0
1
2
3
4
5
6
7
9
Days post infection
13
16
21
23
27
30
-5
-1
0
4
7
9
13
16
Days post infection
Figure 4 Panels (a-c) neutralizing antibody titres post-challenge for control (a), CD4 depleted (b)
and WC-1 depleted (c). Panel (d) shows the percentage of individual mononuclear cell populations
in the peripheral blood animals treated with control antibody. Panel (e) and (f) shows the
percentage of CD4 T cells and WC-1 T cells respectively, after monoclonal antibody treatment.
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Discussion:
Understanding the key elements of the immune response responsible for protection provides a
focus for further vaccine development. However, the correlates of protection involved in resolution
of infection do not necessarily reflect those that would be protective in the presence of pre-existing
vaccine induced immunity.
In the studies reported here, our preliminary results suggest the magnitude and duration of the
neutralizing antibody response are related to the magnitude of the CD4 T cell response. Also, it
would appear that measuring the specific T cell response to FMDV after vaccination may improve
the ability to predict whether animals will be protected from subsequent challenge. Further
development of the assays to measure specific CD4 T cell responses may improve the predictive
capacity of these assays. The usefulness of measuring cell proliferation or interferon gamma
production of whole PBMC populations is compromised because a significant component of the
responses measured are due to bystander activation of other cell populations, for example gammadelta T cells. These difficulties can largely be resolved by analyzing purified cell populations;
unfortunately cell sorting is still a relatively expensive and time-consuming process. Advances in
the development of multiparameter flow cytometry may allow the simultaneous assessment of
multiple cytokines produced from antigen specific CD4 T cells.
Pre-existing antibody provides a mechanism for immediate protection to virus challenge. In the
animals challenged one year after first vaccination, where protection was related to the magnitude
of the T cell response rather than antibody titre, it is possible the presence of previously primed
CD4 T cells may promote a more rapid and effective antibody response.
In contrast, resolution of clinical signs after acute infection with FMDV appears to be independent
of CD4 T cells, WC+ γδ T cells and detectable circulating antibody. These findings corroborate the
ideas of Grubman (2005), who have demonstrated the effectiveness of type-1 interferon in
controlling FMDV infection.
Conclusions:
Further refinement of assays to measure specific T cell responses to FMDV should be performed
alongside vaccine development programmes, to enhance our understanding of protective immune
mechanisms and consequently influence vaccine design.
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