Journal of General Virology (1993), 74, 275~280.
275
Printed in Great Britain
Cell-mediated antiviral response to equine herpesvirus type 1 demonstrated
in a murine infection model by means of adoptive transfer of immune cells
M . A z m i and H . J. Field*
Department of Clinical Veterinary Medicine, University of Cambridge, Madingley Road, Cambridge CB3 0ES, U.K.
Protection against equine herpesvirus type 1 (EHV-1)
infection in a mouse model has been studied by means of
delayed-type hypersensitivity (DTH) and adoptive transfer of immune spleen cells. Mice were found to develop
a positive DTH response to EHV-1 antigen which was
sustained for several months after primary inoculation.
The response was found to cross-react with EHV-4derived antigen. Immune cells (from mice primed with
live or heat-inactivated EHV-1) conferred an enhanced
DTH response on recipients; however, only the immune
cells that were previously primed with live EHV-1 gave
protection against re-infection. The degree of protection
was also dependent on the number of spleen cells
transferred. Immune cells from mice primed with heatinactivated EHV-1 appeared to enhance virus replication
following subsequent inoculation. The serum antibody
response to EHV- 1 appeared to be slightly suppressed in
recipients of spleen cells from mice primed with either
live or heat-inactivated virus. These results support the
important role for cell-mediated responses in protective
immunity to EHV-1 and provide clues to the nature of
protection and immunopathology in the natural host.
Equine herpesvirus type 1 (EHV-1) causes an important
infection in horses associated with a variety of clinical
problems including respiratory distress, abortion (Bryans
& Allen, 1989) and neurological signs (Charlton et al.,
1976; Chowdury et al., 1986). The serologically related
but distinct virus, EHV-4, also causes a common
infection and both EHV-1 and EHV-4 establish latent
infections in horses from which they may be reactivated.
Commercial vaccines for EHV-1 are available; however,
the protection conferred may be incomplete and short
lived. Disease has been reported in animals that have
been vaccinated (Allen & Bryans, 1986) or previously
infected naturally (Grandell et al., 1980). Thus the
epidemiology of EHV and the role of immunoprophylaxis are complex and poorly understood.
Much attention has been given to the humoral
response to EHV-1 and the role of neutralizing antibody
in protection has been investigated in the horse (Allen &
Bryans, 1986; Thompson et al., 1976) and in a hamster
infection model (Stokes et al., 1989). The importance of
celt-mediated responses in immunity to EHV-1 has also
been shown in hamsters (Sentsui et al., 1991). The
hamster infection model, however, has several features
that differ markedly from those seen in the natural
disease. A mu~ine model of EHV-1 infection was
established using intranasal (i.n.) inoculation and the
pathogenesis of the infection in this model has features
which more closely resemble the natural infection. These
features include respiratory sites for virus replication,
viraemia, latency and abortion (Awan et al., 1990, 1991 ;
Field et al., 1992).
In a separate study (M. Azmi & H.J. Field, unpublished) we will report the humoral responses to EHV1 in the murine model. We demonstrated that EHV-1
infection resulted in protective immunity and reduced
viral replication in the respiratory tissues following
secondary infection. The protection in the upper and
lower respiratory tissues was found to vary with the
route of immunization and also protection did not
correlate well with antibody levels suggesting that cellmediated responses may be more important for protection.
The cellular responses in mice to other infections are
well understood and the routine model therefore presents
a suitable one for the investigation of cell-mediated
responses to EHV-1 infection. In the present study we
present direct evidence for the important role of cellmediated immunity (CMI) in establishing protective
immunity to EHV-I infection in the murine i.n. inoculation model and we speculate on the likely importance of these mechanisms in the natural host.
The virus employed was EHV-1 strain AB4 (Awan et
al., 1990). Virus working stocks were grown in rabbit
kidney (RK-13) cells (or Vero cells) as previously
described (Awan et al., 1990) and titrated by a plaque
assay in RK-13 cells. In some cases virus inocula were
heat-inactivated at 56 °C for 30 min.
Female BALB/c mice were inoculated i.n. at 4 to 5
0001-1254 © 1993 SGM
19
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276
Short communication
weeks old with l0 s to 107 p.f.u, of EHV-1 (in 40 gl
inoculum). When appropriate, uninfected RK-13 cell
lysate was used for mock infection of control mice. Mice
were killed at various times post-inoculation (p.i.). The
whole lungs and turbinate bones were homogenized and
tested for the presence of virus (log10 p.f.u, per organ) by
titration in R K cell monolayers.
Virus was purified to produce target antigen for the
ELISA, for testing the delayed-type hypersensitivity
(DTH) response and for the preparation of hyperimmune
sera. Infected cell suspensions were sonicated for 3 rain
and centrifuged at 3000 r.p.m, for 10 min. The supernatant and the cell-free virus supernatant were subjected
to ultracentrifugation at 28000 r.p.m, for 2 h at 4 °C.
The resultant pellet was collected in a small amount of
T N E (Tris, NaCI, EDTA) buffer with repeated pipetting
to break the pellet, and then sonicated for 30 s. The virus
suspension was gently layered on top of a 60 ml 10 to
50 % potassium tartrate linear density gradient in T N E
buffer. The gradient was centrifuged at 20000 r.p.m, for
3 h at 4 °C. The white opalescent band was carefully
aspirated from about the middle of the gradient. The
virus suspension was further diluted 1:4 in T N E buffer
before being subjected to centrifugation at 28 000 r.p.m.
for 1 h at 4 °C. The pelleted virus was resuspended in a
small amount of T N E buffer, titrated by means of plaque
assay and stored at - 7 0 °C.
For the preparation of anti-EHV- 1 hyperimmune sera,
BALB/c mice were inoculated subcutaneously (s.c.) with
107 p.f.u, purified EHV-1 (inactivated at 56 °C for 30 min
and emulsified with Freund's complete adjuvant) grown
in RK-13 cells. Another three inoculations were given at
2 week intervals using Freund's incomplete adjuvant.
Sera were collected 14 days after the final booster (using
live virus without adjuvant), pooled and stored at
- 2 0 °C. Antibody titres were determined by means of
ELISA. Preimmune or normal mouse sera were collected
from uninfected mice.
Donor spleen cells were obtained from syngeneic
BALB/c mice which had been inoculated i.n. 3 weeks
previously with 5 x l0 G p.f.u, live EHV-1 or with the
same amount of heat-inactivated virus. Groups of mice
were inoculated i.n. with uninfected RK-13 cell lysate
and these were used as a source of control donor cells.
Twenty-one days after the priming inoculation, the
spleens were removed aseptically and pooled before
being gently homogenized. The cell suspensions were
filtered through fine sterile muslin gauze and centrifuged
at 1500 r.p.m, for 10 min. The cells were treated with
distilled water for 15 s to lyse erythrocytes. The osmotic
balance was then restored in RPMI-1640 medium and
the cells were washed twice. Approximately 5 ml samples
of the cell suspensions (containing 107 cells) were added
and allowed to adhere to Petri dishes by incubation at
37 °C for 1 h. The adherent cells consist ofmacrophages.
Non-adherent cells (mostly lymphocytes) were harvested
and pooled. The cell suspensions were centrifuged at
1500r.p.m. for 10rain and resuspended in a small
volume of RPMI. The number of cells and their viability
were determined by counting in a haemocytometer using
the trypan blue exclusion method. The concentration of
the cells was adjusted in RPMI as required for
inoculation. The viability was > 95 %. Two-hundred gl
of the cell suspension was injected into the tail vein of
recipient mice. Control mice received spleen cells from
the mice inoculated with uninfected R K cell lysate. Virus
challenge inoculations were carried out several hours
later on the same day of spleen cell transfer.
An antigen was prepared as above except that virus
was grown in Vero cells to minimize contamination with
cross-reactive cellular antigens. Twenty lal of the inoculum, containing 106 erstwhile p.f.u. EHV-1 (heatinactivated), was inoculated intradermally into the left
ear pinna of mice under general anaesthesia. A similar
quantity of mock-infected cell lysate suspension was also
inoculated at the same time into the right ear pinna as a
control. Mice primed with uninfected cell lysate were
inoculated with virus antigen as an additional control.
All inoculations were carried out in quadruplicate. The
ear skin thickness was measured at daily intervals with
an engineer's micrometer screw gauge. The details of this
technique have been previously described (Nash et al.,
1980).
An ELISA was developed to detect antibodies reactive
with EHV-1 antigens. Briefly, twofold serial dilutions of
test sera were added to antigen-coated 96-well plates,
then allowed to react with a goat anti-mouse (IgG)
antibody conjugated to horseradish peroxidase, using
2,2'-azino-bis(3-ethylbenzthioline-6-sulphonic acid) (Sigma) as the substrate. The absorbance was read at dual
wavelength mode absorbance 405 to 492 nm using a
Titertek Multiskan multichannel photometer (Flow
Laboratories). Antibody titres were determined graphically. This enabled the values of log10 dilution to be read
from the curve that corresponded to an absorbance value
>~ the mean of eight wells of preimmune sera (_+ 3 s.I).).
As a positive control, hyperimmune mouse serum was
included on each plate and preimmune or normal mouse
sera were used as negative controls. The statistical
significance of differences between groups of data was
determined using the two-tailed Student's unpaired ttest.
Mice were inoculated with EHV-1 i.n. or intravenously
(i.v.) and re-inoculated by the same route at 2 months.
After a further 2 months, a D T H response to EHV-1
antigen was measured. One day after the test injection a
positive response to EHV-1 antigen was shown with a
greater magnitude in mice inoculated i.n. compared to
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45
(a)
40
35
30
25
20
15
8 10
x 5
O,
6
I
(a)
Lungs
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277
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Time p.i. (days)
2
Fig. 1. DTH response in mice previously inoculated i.n. or i.v. with live
EHV-1. The mice were inoculated either i.n. (11) or i.v. ([B) with 2 x 106
p.f.u. EHV-1 or given uninfected RK cell lysate (A). To test for DTH
response, mice were inoculated 2 months later intradermally into ear
pinnae with (a) EHV-1 or (b) EHV-4 antigens. Data points represent
mean with s.D. ear thickness increase from groups of four mice.
those i n o c u l a t e d i.v. (Fig. 1 a). H o w e v e r , the difference
was n o t significant ( P > 0.05) a n d n o t a p p a r e n t o n d a y 2
after antigen i n o c u l a t i o n . T h e mice were also f o u n d to
r e s p o n d to an a n t i g e n p r e p a r a t i o n derived f r o m E H V - 4
which is a distinct b u t serologically related virus (Fig.
lb).
M i c e were p r i m e d b y i.n. i n o c u l a t i o n o f live E H V - 1 ,
h e a t - i n a c t i v a t e d E H V - 1 o r a m o c k - i n f e c t e d R K cell
lysate. Live v i r u s - i n o c u l a t e d mice d e v e l o p e d clinical
signs a n d s o m e died, a c c o r d i n g to the i n o c u l u m dose.
M i c e i n o c u l a t e d with h e a t - i n a c t i v a t e d virus or uninfected
cell lysate s h o w e d no clinical signs a n d n o n e died. T h r e e
weeks later, surviving d o n o r mice were used to p r o v i d e
spleen cells for t r a n s f e r to recipient mice. T h e n u m b e r o f
cells t r a n s f e r r e d r a n g e d f r o m 8 x 106 to 4 x 107 a m o n g the
different experiments. R e c i p i e n t mice were challenged
later on the d a y o f i.n. t r a n s f u s i o n with live EHV-1 a n d
the clinical signs noted. M i c e were killed o n d a y s 3 a n d
5 after transfer a n d the virus titres in lungs a n d t u r b i n a t e
b o n e s were m e a s u r e d . I n s o m e cases D T H was m e a s u r e d
3 o r 4 weeks later b y m e a n s o f a skin test a n d a n t i b o d y
levels in s e r u m were also d e t e r m i n e d b y m e a n s o f an
ELISA.
P r i m i n g with live virus p r o d u c e d a p r o t e c t i v e effect in
5
Time p.i. (days)
Fig. 2. Virus infectivity in mice following adoptive transfer of spleen
cells from donor mice primed with live or heat-inactivated EHV-1 and
inoculation with live EHV-1. Donors were primed with live or heatinactivated EHV-I, or uninfected RK cell lysate (to produce 'normal'
spleen cells). (a) Recipients were given 1 x 107 ([]) or 4 x 107 ([]) spleen
cells from mice primed with live EHV-1, or 4 x 107 'normal' ( I ) spleen
cells, then inoculated i.n. with 5 x 106 p.f.u. EHV-1. (b) Recipients were
given 2 x 107 spleen cells from mice primed with heat-inactivated EHV1 then inoculated i.n. with 2 x 106 p.f.u. EHV-1. The virus titres in
turbinates with 'normal' ( . ) or immune ([~) spleen cells, and lungs
with 'normal' ([]) or immune ([]) spleen cells, were measured on days
3 and 5 p.i. Bars represent mean with S.D. virus titre from groups of
four mice.
recipient mice t h a t received 4 x 107 spleen cells. A l t h o u g h
there was no difference in virus titre on d a y 3, b y d a y 5,
there was a clear r e d u c t i o n in virus in b o t h lungs a n d
n a s a l t u r b i n a t e s (Fig. 2 a). T h e r e d u c t i o n s were significant
( P < 0"05) a n d the degree o f virus r e d u c t i o n c o r r e l a t e d
with the n u m b e r o f cells transferred. W h e n the n u m b e r
o f cells was r e d u c e d to < 107 the r e d u c t i o n in virus titre
was n o t significant ( d a t a n o t shown). T h e r e d u c t i o n s in
virus titre were also reflected in r e d u c e d clinical signs in
the i n o c u l a t e d mice. These r e d u c e d signs b e c a m e p a r ticularly a p p a r e n t b e y o n d the 5th d a y p.i, T h u s the
transfer o f spleen cells c o n f e r r e d p r o t e c t i v e i m m u n i t y o n
the recipients. The transfer o f spleen cells was m o r e
effective in r e d u c i n g virus titres in lungs (at least 2 log10 )
c o m p a r e d to t u r b i n a t e s ( a p p r o x i m a t e l y 1 log10 ). T h e
spleen weights in recipients were f o u n d to c o r r e l a t e w i t h
p r o t e c t i o n . O n d a y 5 p.i., the mice t h a t received 4 x 107
i m m u n e spleen cells h a d the heaviest spleens (128_+
11.2 mg) f o l l o w e d b y mice given 1 x 10 v i m m u n e spleen
19-2
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278
Short communication
2
1.8
1.6
1-4
1.2
I
(a)
I
I
I
I
1
0.8
0-6
0.4
.o'~ 0.2
.,~
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i
I
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I
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< 1.6
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1.2
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0.6
0-4
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0
5
10
15
20
Time p.i. (days)
I
25
30
Fig. 3. Antibody response in mice following adoptive transfer of spleen
cells from donor mice primed with live or heat-inactivated EHV-1 and
i.n. inoculation with live EHV-I. (a) Donors were primed with live
EHV- 1 ([]) or uninfected RK cell lysate ( • ) . The recipients were given
4 x 107 spleen cells then inoculated with 5 x 106 p.f.u. EHV-l. (b)
Donors were primed with heat-inactivated EHV-1 ([~) or uninfected
R K cell lysate ( • ) . The recipients were given 2 x 107 spleen cells then
inoculated with 2 x 106 p.f.u. EHV-1. Data points represent mean with
S.D. antibody titre from groups of four mice.
cells (107 + 12.4 mg). The weight of spleens from mice
that received spleen cells from mice primed with cell
lysate was 82_+9"6 mg and the weight of normal spleens
ranged from 80 to 100 rag.
In contrast to the results above spleen cells transferred
from donors primed with heat-killed virus produced no
reduction in virus titre in the lungs and turbinate of
recipients following challenge (Fig. 2b). Furthermore,
there appeared to be a significant increase in growth of
virus in lungs and turbinates on day 5 p.i. These
differences were even more apparent when a larger
challenge dose was employed (4 x 106 p.f.u.) causing all
recipient mice to die compared with 25 % mortality in
unimmunized controls and 0 % mortality in mice that
received spleen cells from live virus-primed donors (data
not shown). A dense mononuclear cellular infiltration
with oedema was seen in histological sections of lungs
obtained from these mice on day 5 p.i. Similar observations were made in mice that received spleen cells
from donors primed with live EHV-1 but the cellular
infiltration appeared to be less intense.
Serum antibody responses were measured in the
groups of mice used above. The results (Fig. 3 a) indicate
that a low but measurable antibody response was
recorded consistent with previous observations. When
compared at particular times the antibody levels were
not significantly different between the experimental
groups. It was notable, however, that at five times (days
3, 7, 14, 21 and 28 p.i.) the antibody levels were lower in
the mice receiving immune cells although different mice
were sampled on each occasion.
Similar small reductions in antibody responses were
observed in mice that received spleen cells from the
donor mice primed with heat-inactivated EHV-1 (Fig.
3b). A lower challenge dose was employed (2 x 106 p.f.u.
compared with 5 x 106 p.f.u, used previously) and the
antibody titre obtained was generally lower. As before a
reduced antibody response was observed (days 5, 14, 21
and 28 p.i.) although the differences between individual
time points were not significant.
Surviving mice were also tested for DTH by inoculating a dose of heat-inactivated EHV-1 into the ear
pinna and measuring the increase in skin thickness. The
results, summarized in Table 1, show that recipient mice
acquired an enhanced DTH response with maximal
response obtained from live virus-primed donors and
this DTH response correlated with the number of cells
transferred. The data shown in Table 1 were obtained
from the same group of mice used to measure virus titres
in the organs (Fig. 2 and 3). A positive response was
observed also in mice receiving spleen cells from donors
primed with a single inoculation of heat-inactivated
virus.
One of the important findings to emerge from this
study was that a DTH response developed in mice
inoculated with EHV-1, and the magnitude of response
following i.n. inoculation was higher compared to that
following i.v. inoculation. By contrast, i.v. inoculation
gave a higher antibody response (M. Azmi & H. J. Field,
unpublished). The enhanced CMI response following i.n.
inoculation, despite the lower antibody response, may
explain the observed protective immunity in upper and
lower respiratory tracts against re-infection, suggesting
that the elimination of EHV-1 from the site of replication
is a function of CMI. It was intriguing that a DTH
response was also shown to the EHV-4 antigen which is
known to cross-react with EHV-1 serologically. We have
recently observed both exacerbated responses to and
protection against EHV-1 challenge following primary
inoculation with EHV-4 in mice. This provides evidence
for interaction between the two virus types with a fine
balance between protective and deleterious cell-mediated
responses (M. Azmi & H.J. Field, unpublished); this
may be an important epidemiological factor in the
natural host and is currently being investigated further.
Further evidence for the protective role of immune
cells was obtained directly by means of adoptive transfer
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279
Table 1. The effect of transferring spleen cells from primed donors on the
increase of the ear thickness (DTH response) in recipients after i.n. inoculation
with live EHV-1
Source of spleen cells*
Control (uninfected)§
RK cell lysate
Inoculated mice
Heat-inactivated virus
Live virus
No. cells transferred DTH responset Virusprotection:~
0
8.3 x 106
4 x 10r
25.93
72-85
73.07
•aIJ
-
2 x 107
8-3x 106
4 x 107
107'41
112'33
131"30
-+
+
* Donor micewere primed with the material shownby i.n. inoculation, 3 weeksbeforetransfusion
of spleen cells into recipients.
t Percentage ear thickness (mean from a group of four mice) increase at 24 h followingantigen
inoculation; test carried out 3 weeks after i.n. inoculation with EHV-1.
:~ +, Marked reduction (at least 1 logaop.f.u.) of virus titre in lungs with reduced clinical signs;
- , no reduction of virus titre or clinical signs.
§ Naive mice tested for DTH response with viral antigen.
II NA, Not applicable.
studies. Transfer of immune spleen cells has been
reported to protect against EHV-1 infection, e.g. a
passive transfer of immune spleen cells has been shown
to be protective in hamsters to EHV-1 infection (Sentsui
et al., 1991). However, the route of EHV-1 infection in
that model (intraperitoneal inoculation) and virus target
organs (liver and spleen) are not those normally involved
in the natural equine disease, Rapid clearance of herpes
simplex virus from the ear pinna of mice was shown
following adoptive transfer of imnmne cells (Nash et al.,
1980, 1981). Results of the present adoptive cell-transfer
experiments showed that the recipients were protected
against challenge with EHV-1, if the mice were given
spleen cells from mice primed with live EHV-1, although
the elimination of infectious virus was found to be less
effective in the upper compared to the lower respiratory
tract. However, it appeared that the number of cells
transferred is a critical factor in effecting protection since
protection was not obtained in mice that received < 107
immune spleen cells.
The results of adoptive cell-transfer experiments in
which donors were primed with heat-inactivated virus
showed that the recipients were not protected against
challenge with EHV-1; indeed virus replication was
enhanced. An intense cellular infiltration in lungs may
have resulted from antigen expression in the infected
tissue focusing activated cells with consequent release of
inflammatory mediators, This was consistent with the
profound inflammatory exudate observed in the sections
of lungs obtained from recipients. We also found that the
cell number may be also a crucial factor in the
enhancement of virus replication.
Transfer of cells from donors primed with live or heatinactivated virus was found to enhance the D T H
response compared to that seen following transfer from
control donors. The magnitude o f response was attri-
buted to the number of spleen cells being transferred as
well as to the type of antigen used for the stimulation of
the donors. However, this did not correlate with
protection from infection.
Serum antibody responses were obtained in recipients
that received spleen cells from donors primed with live
or heat-inactivated EHV-1, Compared to the control
mice (which received spleen cells from mice primed with
uninfected cell lysate) a slight suppression of serum
antibody response was noted at several different time
points although the difference between individual timepoints was not significant. Lower antibody response in
these cases may correspond to the induction of active
cytotoxic CD8 + lymphocytes in the population of
transferred spleen cells, which could have a suppressor
role reflected in the reduced antibody production.
Results described in this study demonstrate the
potential of the murine model for the study of the
immune response to EHV. Further experiments are in
hand to define specific populations of immune cells
involved in inducing the responses to EHV-1 and EHV4 and the effector mechanism involved in virus clearance.
These results suggest mechanisms by which these
responses may be involved in EHV pathogenesis in the
horse and how these responses may be used in the design
of immunoprophylactic treatments.
We wish to thank Drs A. A. Nash, J. S. Gibson and A. R. Awan for
their helpful discussion of data and Ms Alana Thackray for her expert
technical assistance. M. Azmiwas supported by a scholarship from the
Univerisiti Pertanian Malaysia and a grant from the Jowett Trust,
Cambridge University. We gratefully acknowledgesubstantial support
for this work by the Equine Virology Research Foundation.
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(Received 27 July 1992; Accepted 7 October 1992)
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