Perforin-dependent CD4+ T-cell cytotoxicity contributes to control a murine poxvirus infection.

Abstract

CD4(+) T cells are generally regarded as helpers and regulators of the immune response. Although cytolytic CD4(+) T cells have been described, whether those generated during the course of a viral infection play a role in virus control remains unknown. Here we show that during acute infection with ectromelia virus, the mouse homolog of the human virus of smallpox, large numbers of CD4(+) T cells in the draining lymph node and liver of resistant mice have a cytotoxic phenotype. We also show that these cells kill targets in vivo in a perforin-dependent manner and that mice with specific deficiency of perforin in CD4(+) T cells have impaired virus control. Thus, perforin-dependent CD4(+) T-cell killing of infected cells is an important mechanism of antiviral defense.

Full text

Page 1

Perforin-dependent CD4+T-cell cytotoxicity contributes
to control a murine poxvirus infection
Min Fanga, Nicholas A. Sicilianob, Adam R. Herspergerb, Felicia Roscoea, Angela Hua, Xueying Maa,
Ahamed R. Shamsedeena, Laurence C. Eisenlohrb, and Luis J. Sigala,1
aImmune Cell Development and Host Defense Program, Fox Chase Cancer Center, Philadelphia, PA 19111; andbDepartment of Microbiology and
Immunology, Jefferson Medical College, Thomas Jefferson University, Philadelphia, PA 19107
Edited by Bernard Moss, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, and approved May 16, 2012
(received for review February 6, 2012)
CD4+T cells are generally regarded as helpers and regulators of
the immune response. Although cytolytic CD4+T cells have been
described, whether those generated during the course of a viral
infection play a role in virus control remains unknown. Here we
show that during acute infection with ectromelia virus, the mouse
homolog of the human virus of smallpox, large numbers of CD4+T
cells in the draining lymph node and liver of resistant mice have
a cytotoxic phenotype. We also show that these cells kill targets
in vivo in a perforin-dependent manner and that mice with specific
deficiency of perforin in CD4+T cells have impaired virus control.
Thus, perforin-dependent CD4+T-cell killing of infected cells is an
important mechanism of antiviral defense.
CD4+T lymphocytes|cytotoxic T lymphocytes|viral immunology|
viral pathogenesis
T
peptides associated with MHC class I (MHC I) proteins and
CD4+T cells recognize peptides associated with MHC class II
(MHC II) molecules. Traditionally, CD4+T lymphocytes have
been described as helpers and regulators of the immune re-
sponse, and cytotoxic T lymphocyte (CTL) effector functions
have been attributed mostly to CD8+T cells. However, the ex-
istence of MHC II-restricted CD4+CTL has been recognized
since the late 1970s (1). CD4+CTL clones have been generated
in humans and mice (2, 3). The presence of CD4+CTL during
primary infection were initially documented in mice that lacked
the normal complement of CD8+T cells (4, 5) or during chronic
or persistent viral infections (6, 7), supporting the notion that
CD4+CTL can be generated by chronic exposure to antigen or
repeated in vitro stimulation. However, CD4+CTL capable of
killing target cells in vivo have been described more recently in
immunocompetent mice infected with the Armstrong strain of
lymphocytic choriomeningitis virus (LCMV) (8) and with West
Nile Virus (9), indicating that CD4+CTL can also be generated
during acute viral infections.
CD8+CTL mainly kill targets by releasing perforin (Prf) and
granzymes, such as granzyme B (GzB) through granule exo-
cytosis (GE) (10, 11). Although human and mouse CD4+T cells
have been shown to lyse cells through Fas-Fas ligand interactions
(12–14), GE-mediated CD4+T-cell cytotoxicity has also been
shown in human infections with HIV, human cytomegalovirus,
Epstein-Barr virus, herpes simplex virus, influenza virus, and
vaccinia virus (2, 15–20).
Regarding a possible role for CD4+CTL in protective im-
munity, previous work by others showed that adoptive transfer of
in vitro primed CD4+CTL protected from lethal influenza in-
fection in a Prf-dependent manner (17). However, whether
CD4+CTL generated during the course of a viral infection play
any role in virus control remains to be demonstrated (3), most
likely because it has been difficult to distinguish from the pro-
tective effects of CD8+CTL.
-cell–mediated immunity is a major mechanism providing
specific protection against viruses. CD8+T cells recognize
The Orthopoxvirus (OPV) Ectromelia virus (ECTV), the
agent of mousepox, has host specificity for the mouse. This virus
is very similar to variola virus, the agent of human smallpox, the
zoonotic monkeypox virus, and the smallpox vaccine species
vaccinia virus (VACV) (21–25). Following infection in the
footpad, its natural route of entry, ECTV infects the popliteal
draining lymph node (D-LN) and spreads through the lympho-
hematogenous (LH) route to seed visceral organs, mainly the
liver and the spleen. Of interest, ECTV LH spread is usually
used as the textbook example for the many human and animal
natural viruses that, with some variations, spread through the LH
route (26, 27). Although ECTV infects all mouse strains, the
outcome of primary infection following footpad inoculation
varies. Susceptible strains, such as BALB/c, develop mousepox,
which is characterized by uncontrolled replication of the virus in
the liver and spleen. Most susceptible mice die with extensive
liver damage within the first 14 d postinfection (dpi), and those
that survive develop the typical skin rash of poxvirus infections.
In resistant strains, such as C57BL/6 (B6), the virus also infects
the liver and spleen but the mice do not show overt symptoms of
disease because the replication of the virus is controlled by the
combined action of innate and adaptive immune mechanisms
(21, 23, 25, 28–33).
Our previous studies and those of others have shown that
CD4+T-cell help-independent CD8+T-cell responses are es-
sential for the early—and that CD4+T-cell help-dependent Ab
responses are essential for the late but not the early—control of
primary ECTV infections (29, 34–36). Previous work suggested
that, in addition to providing help to B cells, CD4+T cells may
also have a direct effector function to control ECTV but the
specific mechanism was not identified (31). Here we report that
the CD4+T-cell responses to ECTV are very large, particularly
in the LN, draining the primary site of infection, and in the liver.
Moreover, we show that most responding CD4+T cells have
characteristics of CTL because they are GzB+. We also show
that the responding CD4+T cells have in vivo Prf-dependent,
MHC II-restricted CTL activity and that CD4+directly con-
tributes to controlling the virus in a Prf-dependent manner.
Thus, our work is unique in showing that CD4+CTL have an
important role in controlling a viral infection.
Results
ECTV Induced Large Numbers of Potentially Cytolytic CD4+T Cells.
CD11a and CD49d coexpression marks activated CD4+T cells
induced by antigen exposure (37, 38). When we determined the
Author contributions: M.F. and L.J.S. designed research; M.F., N.A.S., A.R.H., F.R., A.H.,
X.M., and A.R.S. performed research; L.C.E. contributed new reagents/analytic tools; M.F.,
N.A.S., A.R.H., L.C.E., and L.J.S. analyzed data; and M.F. and L.J.S. wrote the paper.
The authors declare no conflict of interest.
This article is a PNAS Direct Submission.
1To whom correspondence should be addressed. E-mail: luis.sigal@fccc.edu.
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.
1073/pnas.1202143109/-/DCSupplemental.
www.pnas.org/cgi/doi/10.1073/pnas.1202143109PNAS
| June 19, 2012
| vol. 109
| no. 25
| 9983–9988
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CD4+T-cell response to ECTV in C57BL/6 (B6) mice using
these markers, we found that at 5 dpi ∼60% of the CD4+T cells
in the D-LN were CD11a+CD49d+, indicating strong virus-
specific CD4+T-cell responses. Interestingly, ∼60% of the
CD11a+CD49d+CD4+T cells were also GzB+(Fig. 1A). In the
reverse analysis, ∼95% of the GzB+CD4+T cells were CD11a+
CD49d+, indicating that GzB expression occurs only in antigen
experienced cells (Fig. 1B). At 7 dpi, ∼70% of the CD4+T cells
in the D-LN and liver were CD11a+CD49d+and ∼60% of these
were GzB+. In the spleen ∼40% were CD11a+CD49d+and
40% of these were GzB+but the reverse analysis showed that the
majority of the GzB+but not of the GzB−CD4+T cells in all
organs were CD11a+CD49d+(Fig. S1A). When the total CD4+
T cells were considered, ∼40% in the D-LN but low background
frequencies in the liver or spleen were GzB+at 5 dpi, and 40% in
the D-LN and liver and 15% in the spleen were GzB+at 7 dpi;
however, the absolute number of GzB+cells was higher in the
spleen than in the other organs (Fig. 1C).
To further address the extent of the CD4+T-cell response
to ECTV, we inoculated B6 mice at different stages of infection
with BrdU intraperitoneally. Three hours later, proliferation by
BrdU incorporation was determined by flow cytometry in various
organs. BrdU incorporation by CD4+T cells peaked at 5 dpi and
remained high at 7 dpi in the D-LN. Incorporation of BrdU in
spleen and liver was low at 5 dpi but increased significantly at
7 dpi. Of interest, the proliferative response was higher in the
D-LN and liver than in the spleen. More important, most of the
CD4+T cells that incorporated BrdU+expressed GzB (Fig.
S1B). We also found that CD4+T cells from mice at 7 dpi and
exposed to infected cells ex vivo expressed surface CD107a, a
sign of degranulation (Fig. S1C). Thus, the CD4+T-cell responses
to ECTV are large, particularly in the D-LN and liver and, con-
sistent with a CTL phenotype, most of the responding CD4+
T cells are GzB+.
VACV MHC II-Restricted CD4+T-Cell Determinant I1L7–21Is also an
ECTV Determinant. To identify ECTV CD4+T-cell determinants,
we screened B6 mice for responses to the previously described
VACV determinants H3L272–286, I1L7–21, and B546–60(39, 40),
which are highly conserved in ECTV. Compared with uninfected
mice, a significantly larger proportion of the splenic CD4+T cells
from infected mice stained with I-Ab-I1L7–21(QLVFNSISAR-
ALKAY in VACV and the tetramer; QLIFNSISARALKAY in
ECTV, bold underlined letters indicate a different amino acid
between VACV and ECTV) but not with tetramers loaded with
H3L272–286, B546–60or control CLIP peptide (Fig. S2A). In addi-
tion, a small but clear population of splenic CD4+T cells from
infected mice produced IFN-γ when stimulated in vitro with the
ECTV variant of I1L7–21(Fig. S2B). Moreover, more CD4+T
cells stained with I-Ab-I1L7–21at 5 dpi in the D-LN and at 7 dpi in
the liver in infected compared with uninfected mice (Fig. 2A).
Furthermore, there was a significantly higher number of I1L7–21
but not H3L272–286specific CD4+T cells in the spleens and livers
at 5 and 7 dpi compared with uninfected mice in ELISPOT assays
(Fig. 2B and Fig. S2C). Hence, the VACV CD4+T-cell de-
terminant I1L7–21,(39) is also an ECTV CD4+T-cell determinant.
MHC II+Cells Are Targets of Virus-Specific Cytolytic CD4+T Cells in
ECTV-Infected Mice. Most cells infected with ECTV in the D-LN
are MHC II+, as revealed by infection with a recombinant ECTV
that expresses enhanced green fluorescence protein (ECTV-
OVA197-386-IRES-EGFP) (Fig. 3A). We next tested whether
ECTV induces CD4+CTL capable of killing MHC II+targets
pulsed with I1L7–21 in vivo. Splenocytes from B6.CD45.1
(CD45.1+) mice stained with 0.8 μM 5-(and 6)-carboxyfluorescein
diacetate succinimidyl ester (CFSElow) and pulsed with I1L7–21
peptide, or stained with 4 μM CFSE (CFSEhigh) and mock-pulsed,
were cotransferred 1:1 into ECTV-infected B6 (CD45.2+) mice at
different days postinfection. Sixteen hours after transfer, the spe-
cific killing of MHC II+and MHC II−CFSElowtargets was de-
termined in different organs (Fig. 3B). At 5 dpi, MHC II-restricted
in vivo killing of target cells pulsed with the I1L7–21peptide was
observed in the D-LN but not in the liver or spleen (Fig. 3C).
However, strong MHC II-restricted specific killing was observed in
the liver and to a lesser extent in the spleen and nondraining
lymph node (ND-LN) at 7 dpi. Thus, the in vivo killing of MHC
GzB
CD4
Uninfected
5.5
CD11a
CD49d
2.4
0.4
GzB
CD4
Infected
59
CD11a
CD49d
61
3.7
A
Infected
37
35
95
B
GzB
CD4
Uninfected
1.0
CD11a
CD49d
9.6
Gated on CD4+ T cells
Gated on CD4+ T cells
CD11a+ CD49d+
CD11a- CD49d-
UninfectedInfected
0
20
40
60
80
% GzB+
P=0.0036
UninfectedInfected
0
20
40
60
80
100
GzB-
GzB+
% CD11a+ CD49d+
P=0.044
C
D-LNND-LNSpleen
infected
Uninfected (LN)
Liver
GzB
CD4
5 dpi
7 dpi
42
41
0.7
3.8
1.4
14
0.5
0.5
37
Gated on CD4+ T cells
Uninfected
5 dpi
7 dpi
Total GzB+ (Log10)
D-LNND-LNSpleenLiver
3
4
5
6
7
P<0.01
P<0.01
P<0.01
P<0.01
P<0.001
P<0.001
P<0.001
% GzB+
D-LNND-LNSpleenLiver
0
10
20
30
40
50
P<0.01
P<0.001
P<0.05
P<0.001
P<0.001
Fig. 1.
resentative flow cytometry plots with gating strategy and summary graphs showing the frequency of GzB expression in CD11a+CD49d+and CD11a−CD49d−
CD4+T cells at 5 dpi. (B) As in A but showing the frequency of CD11a and CD49d coexpression in GzB+and GzB−CD4+T cells. (C) Representative flow cytometry
plots and column graphs showing the frequency and absolute number of GzB expression in CD4+T cells at 5 and 7 dpi. Shown are the significant P values
between the corresponding column vs. the same organ in the uninfected controls. All of the data are representative of two to four similar experiments.
ECTV infection induces a large number of potentially cytolytic CD4+T cells. B6 mice were infected with 3 × 103pfu ECTV or left uninfected. (A) Rep-
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Page 3

II-restricted I1L7–21pulsed targets was consistent with the fre-
quencies of GzB+CD4+T cells in the different organs and with
the kinetics of their response.
In addition to GzB, killing by GE requires Prf. Although GzB-
deficient mice are not readily accessible, Prf-deficient (Prf−/−) mice
can be obtained commercially. Thus, we compared MHC II-re-
stricted killing of targets pulsed with I1L7–21in the livers of B6 and
Prf−/−mice infected with ECTV. Because Prf−/−mice are highly
susceptible to mousepox, all groups were treated with the poxvirus
inhibitor Cyodofovir at 4 dpi. We found that killing was absent in
the Prf−/−mice, indicating that during ECTV infection, MHC II-
restricted killing in vivo depends on the GE pathway (Fig. 3D).
CD4+T Cells Have a Direct Role in Virus Control. Optimal primary
CD8+T-cell responses to ECTV infection are CD4+T-cell–in-
dependent, as demonstrated in CD4-depleted and CD4-deficient
mice (29, 35), and as we also show here in MHC II-deficient
(MHC II−/−) mice (Fig. 4A). Furthermore, although production
of Abs to ECTV requires CD4+T-cell help and anti-ECTV Abs
are required for long-term survival to mousepox, the control of
ECTV at 7 dpi is completely independent of Abs (34, 36). To
determine whether the effector function of CD4+T cells is re-
quired for efficient virus control, we measured virus titers at 7
dpi in B6 mice depleted of CD4+T cells, CD8+T cells, CD4+
and CD8+T cells, and in MHC II−/−mice. The virus titers in the
liver and spleen of the CD4+T-cell–depleted or MHC II−/−mice
were 10- to 100-fold higher than in intact B6 mice, but lower than
in B6 mice depleted of CD8+T cells. Depletion of CD4+and
CD8+T cells resulted in significantly higher virus titers in the
spleen compared with any single depletion (Fig. 4B). Thus, al-
though CD8+T cells appear to be more efficient, the effector
functions of CD4+significantly contribute to control ECTV
loads. We also compared virus titers in CD4+T-cell–depleted, as
well as CD4−/−and CD8−/−, mice to those of CD40−/−and B-
cell–deficient mice, which are unable to make Ab responses but
mount normal CD4+and CD8+T-cell responses (34). Con-
firming the preponderance of effector CD8+T cells but also the
significant contribution of effector CD4+T cells to ECTV con-
trol, we found no differences in the virus titers between CD40−/−,
B-cell–deficient, and wild-type B6 mice. However, there were 10-
to 100-fold higher virus titers (measured only in spleens) (Fig.
4C) of CD4+T-cell–depleted or CD4−/−mice compared with
wild-type B6 mice, and even higher virus loads in CD8−/−mice.
When tested for survival, CD8−/−mice succumbed ∼7 dpi be-
cause of lack of early virus control by CD8+T cells; B-cell–de-
ficient mice and CD40−/−mice succumbed more than 40 dpi. On
the other hand, MHC II−/−mice succumbed ∼20 dpi, indicating
that in addition to their ability to provide help to B cells, CD4+T
cells have a direct role in controlling ECTV (Fig. 4D).
Prf Deficiency Exclusively in CD4+T Cells Results in Deficient ECTV
Control. To investigate whether CD4+CTL are required to ef-
ficiently control ECTV, we depleted B6 mice of CD4+T cells,
0.57
2.08
InfectedUninfected
0.581.99
CD4
I-Ab-I1L7-21
D-LN
Liver
A
BDMSO
H3L272-286
I1L7-21
DMSO
H3L272-286
I1L7-21
Spleen 5 dpi
0 2 4 6 8 10
# of spots
Liver 7 dpi
P=0.0142
DMSO
H3L272-286
I1L7-21
Spleen 7 dpi
0 100
200
P=0.0006
# of spots
0200400
P=0.0001
# of spots
Gated on CD4+ T cells
Fig. 2.
ECTV determinant. B6 mice were infected with 3 × 103pfu ECTV or left
uninfected. (A) The D-LN and liver cells stained with I-Ab-I1L7–21tetramer at
7 dpi. (B) ELISPOT assay for CD4+T cells from indicated organs at 5 or 7 dpi
cultured with DMSO (negative control), ECTV H3L272–286, ECTV I1L7–21and
ConA (positive control). P values that were significant when comparing with
DMSO control are shown. Data are representative of two or three similar
experiments. A picture of the ELISPOT plate is shown in Fig. S2.
The VACV MHC II-restricted CD4+T-cell determinant I1L7–21is also an
55
22
19
3.8
CFSE
Cells
MHC II +
MHC II -
43%
CD45.1
CD45.1
CFSE
IC
CFSE
I-Ab
Uninfected Infected
B
A
EGFP
IC
3 dpi
EGFP
I- Ab
UninfectedECTV-WTECTV-EGFP
0.4
99
0
0
0.6
99
0
0.1
0.6
77
0.2
22
73
27
0.1
0
40
60
0
0
CD
Specific killing (%)
D-LN ND-LNSpleenLiver
0
10
20
30
40
50
Specific killing (%)
5 dpi
D-LN ND-LNSpleen Liver
0
10
20
30
40
7 dpi
P=0.0088
B6 Prf-/-
0
10
20
30
40
Fig. 3.
infected mice. (A) B6 mice were infected with 3 × 103pfu ECTV expressing
GFP or left uninfected. At 3 dpi, cells from the D-LN were stained with isotype
control or anti-MHC II mAb. Data are representative of three similar experi-
ments. (B) Strategy for the determination of in vivo MHC II-restricted killing.
At the indicated days postinfection, the mice were cotransferred with a 1:1
mixture of 107naive splenocytes from B6-CD45.1 mice labeled with 4 μM CFSE
(CFSEhigh)or labeled with 0.8 μM CFSE (CFSElow) and pulsed with I1L7–21. After
16 h the mice were killed and the cells double-stained for CD45.1 to facilitate
identification of the transferred cells and for MHC II or isotype control (IC).
The ratio of CFSEhighand CFSElowcells in the CD45.1+MHC II+and CD45.1+
MHC II−cells was determined by flow cytometry and specific killing calculated
as detailed in Methods. (C) B6 mice. Specific killing at 5 and 7 dpi in the D-LN,
ND-LN, spleen, and liver as indicated. (D) Specific killing in the liver of the
indicated mice at 7 dpi. The mice had been treated with the anti-OPV drug
cidofovir to prevent death of the Prf−/−mice.
MHC II+cells are targets of virus-specific cytolytic CD4+T cells in ECTV-
Fang et al.PNAS
| June 19, 2012
| vol. 109
| no. 25
| 9985
IMMUNOLOGY

Page 4

harvested their splenocytes, mixed them with magnetically pu-
rified CD4+T cells from B6 or Prf−/−mice in an 84:16 ratio, and
transferred them into SCID B6 mice. Two weeks after re-
constitution, the mice had normal proportions of CD4+and
CD8+T cells in the blood (Fig. 5A). Three weeks after re-
constitution, the mice were challenged with ECTV. At 7 dpi,
a similarly large proportion of CD4+T cells in the livers of the
reconstituted SCID and control B6 mice expressed GzB (it
should be noted that there are no good commercial Abs to
identify Prf+cells by flow cytometry) (Fig. 5B). However, the
virus titers in the liver and spleen of SCID mice reconstituted
with Prf−/−CD4+T cells were significantly higher than in mice
reconstituted with wild-type CD4+T cells (Fig. 5C). Thus, in
mice that had a specific deficiency of Prf in CD4+T cells, the
control of ECTV was defective. These data are unique in dem-
onstrating that CD4+CTL generated during the course of a viral
infection play a significant role in controlling the virus.
Discussion
Previous studies in mousepox-resistant B6 mice suggested that in
addition to their helper functions, CD4+T cells may have an
effector role in controlling ECTV infection (31). At the time, the
authors did not investigate the mechanism but proposed it did
not involve cytotoxicity. Here we found that the CD4+T-cell
responses to ECTV are very strong and dominated by cells that
express GzB, in particular in the D-LN and liver, where at the
peak of the response they reached ∼40% of the total CD4+T
cells. To our knowledge, such extensive expression and tissue
compartmentalization of a cytolytic molecule in CD4+T cells
during a viral infection has not been documented.
The large proportion of GzB+CD4+T cells that we detected
in the D-LN and liver also suggested that CD4+T-cell responses
might be more extensive than generally thought. Our analysis of
the kinetics of the CD4+T-cell response by BrdU incorporation
confirmed that the CD4+T-cell responses are massive in the D-
LN and liver but smaller in the spleen. Our results also indicated
that the majority of the responding CD4+T cells (CD11a+
CD49d+) are GzB+in the D-LN and liver and, to a smaller ex-
tent, also in the spleen. Although it is commonly thought that the
antiviral CD4+T-cell responses are quantitatively much smaller
than those of CD8+T cells, our data are consistent with a recent
report suggesting that CD4+T-cell responses are much larger
than previously appreciated and showing that the anti-LCMV
CD4+T-cell responses detected in peripheral blood lymphocytes
comprised 50% of the CD4+T cells (38). Nevertheless, the
CD4+T-cell responses to I1L7–21, the most dominant determi-
nant in our experiments and which is also the most dominant
determinant yet indentified for VACV (39), consisted of no more
than 1.5% of the total CD4+T cells. This finding contrasts with
the CD8+T-cell response where 5–8% of the total CD8+T cells
A
D
C
B
Spleen
Liver
B6
B6, CD4 depl
MHC II-/-
B6, CD8 depl
B6, CD8 & CD4 dep
B6
B6, CD4 depl
MHC II-/-
B6, CD8 depl
B6, CD8 & CD4 depl
0246810
n.s.
p=0.0001
p=0.02
p=0.002
n.s.
p<0.0001
p=0.0008
n.s.
B6
CD40-/-
B cell deficient
B6, CD4 depl
CD4-/-
CD8-/-
068
Log10 virus titers
Log10 virus titers
2
4
p=0.007
p=0.002
n.s.
n.s.
n.s.
020406080
0
20
40
60
80
100
B6
CD8-/-
MHC II-/-
B cell deficient
dpi
Survival (%)
<PE-A>: IFN
GzB
IFN-γ
Uninfected B6 MHC II-/-
ECTV 7 dpi
Gated on CD8+ T cells
0
99
0
0.6
0
38
26
36
0
49
21
30
Fig. 4.
mice were infected with 3 × 103pfu ECTV. CD8+T-cell responses were de-
termined in the spleen at 7 dpi. Data correspond to a pool from three mice
and is representative of three similar experiments. An uninfected B6 mouse
is shown as control. (B) Virus titers in the indicated organs of the indicated
mice at 7 dpi. mAb 2.43 and GK1.5 were used to deplete CD8+and CD4+T
cells, respectively. (C) As in B but with the indicated mice. (D) Survival of the
indicated mice following ECTV infection. Data correspond to three mice per
group and is representative of three similar experiments.
CD4+T cells have a direct role in virus control. (A) B6 and MHC II−/−
A
CD8
CD4
WT B6 WT CD4+
130.2
Prf-/- CD4+
SCID reconstitution
CD4
B
C
GzB
WT B6 WT CD4+
Prf-/- CD4+
SCID reconstitution
Naive B6
ECTV infection 7 dpi
0.2 44 53 52
75
12
14
74
0.2
12
15
70
0.2
15
WT B6 WT CD4+ Prf-/- CD4+
SCID reconstitution
Liver
2.5
3.0
3.5
4.0
4.5
5.0
P=0.0002
WT B6 WT CD4+ Prf-/- CD4+
SCID reconstitution
Log10 pfu
Spleen
2.5
3.0
3.5
4.0
4.5
5.0
P=0.012
Virus titers
Reconstitution
CD4+ T cell response (Gated on CD4+ T cells)
Fig. 5.
control. SCID mice were reconstituted with 108of a 84/16 mixture of sple-
nocytes from CD4-depleted B6 mice and magnetically purified CD4+T cells
from B6 or Pfr−/−mice. (A) CD4+and CD8+T cells in peripheral blood 2 wk
after reconstitution. Data correspond to one representative mouse per
group. A B6 control mouse is also shown. (B) Three weeks after reconsti-
tution, SCID mice and B6 controls were infected with ECTV. Plots show ex-
pression of GzB and CD4 in gated CD4+T-cell from the livers at 7 dpi. Data
correspond to one representative mouse/group. (C) Virus titers at 7 dpi in
spleen and liver of reconstituted SCID and control B6 mice. Data are repre-
sentative of two experiments with three mice per group each.
Prf deficiency exclusively in CD4+T cells results in deficient ECTV
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are directed toward the dominant determinant B8R20–27(35, 41).
One possibility is that for CD4+T cells, the total response
spreads among a much broader set of determinants. Alterna-
tively, the dominant determinant may remain to be discovered.
The existence of CD4+CTL in humans and mice is well
documented (2, 3). Furthermore, it has been shown that ex vivo
generated CD4+CTL can protect mice from influenza virus in-
fection (17). However, whether the cytolytic CD4+T cells that
develop during the course of a viral infection can play a signifi-
cant protective role remained elusive (3), most likely because of
their overlapping function with NK cells and CD8+T cells. Our
finding that most of the antiviral CD4+T cells in the D-LN and
liver expressed GzB, a molecule that is required for GE cyto-
toxicity, and that many ECTV-infected cells were MHC II+,
offered the possibility of addressing this important issue. Our
analysis demonstrated that the antiviral CD4+CTL kill targets
through GE in an MHC II-restricted manner, confirmed that
CD4+T cells have a direct effector function in controlling ECTV
infection, and demonstrated that the enhanced control of ECTV
by CD4+T cells requires expression of Prf. To our knowledge,
this demonstration that the CD4+CTL that develop during the
course of a viral infection have a significant role in controlling
a virus is unique. Many viruses that naturally infect humans and
animals spread through the LH route and infect (MHC II+)
dendritic cells and macrophages in the D-LN (26). Thus, it is very
possible that the cytolytic activity of the CD4+T cells also plays
a role in their control. Thus, our results have important impli-
cations for our understanding of the functions of the immune
system. Moreover, because CD4+CTL are so prevalent, the
ECTV model offers a tractable system to identify the molecular
mechanisms that result in their induction in vivo and this may be
important for the development of novel vaccines.
Methods
Mice. The Fox Chase Cancer Center Institutional Animal Care and Use Com-
mittee approved the experimental protocols involving animals. Homozygous
B6.129S-H2dlAb1-Ea
(MHC II−/−),B6.129S2-Igh-6tm1Cgn/J
B6.129S2-Cd4tm1Mak/J
(CD4−/−), B6.129S2-Cd8atm1Mak/J
Prf1tm1Sdz/J(Prf−/−), and B6.129P2-Cd40tm1Kik/J(CD40−/−) were originally
purchased from the Jackson Laboratories and bred at the Fox Chase Cancer
Center Laboratory Animal Facility in specific pathogen-free rooms. Geno-
typing was according to the vendor’s protocols. B6 mice were purchased
from Taconic Farms or the Jackson Laboratories. CB6.CB17-Prkdcscid/SzJ (SCID
mice lacking T and B cells in C57BL/6J background) mice were purchased
from the Jackson Laboratory. B6-LY5.2/Cr (B6-CD45.1) were purchased from
the Frederick National Laboratory for Cancer Research. For reconstitution,
splenocytes from B6 mice depleted of CD4+T cells 3 d before were mixed in
an 84/16 ratio with magnetically purified CD4+cells (Miltenyi Biotech;
Automacs and CD4+beads following the manufacturer’s instructions) from
wild-type B6 or Prf−/−mice and 108transferred intravenously into SCID mice.
(B-cell–deficient),
(CD8−/−), C57BL/6-
Infections. For infections, sex-matched 8- to 12-wk-old animals were used.
Mice were infected in the left footpad with 25 μL PBS containing 3 × 103pfu
unless otherwise indicated. When indicated, 400 μg cidofovir (Vistide) was
injected 4 dpi intraperitoneally.
Cell Culture, ECTV Moscow, Virus Production, and Determination of Virus Titers.
Cellculture, ECTV Moscow, virus production, and determination ofvirus titers
were done as described previously (42). ECTV-OVA197-386-IRES-EGFP was
produced by homologous recombination as described previously (43) by in-
troducing a C-terminal fragment of chicken ovalbumin coding for amino
acids 197–386, followed by the internal ribosomal entry site from Enceph-
alomyocarditis virus in front of EGFP.
Flow Cytometry. BrdU incorporation, isolation of liver lymphocytes, and in-
tracellular staining was done as described previously (34, 42, 43). For MHC
class II tetramer staining, APC labeled I-Ab-CLIP (PVSKMRMATPLLMQA),
-H3L272–286,-I1L7–21, and -B546–60tetramers were prepared by the National
Institute of Allergy and Infectious Diseases tetramer facility and used
according to their standard protocol. Peptide restimulation was performed
for 6 h with bone marrow-derived primary dendritic cells obtained as de-
scribed previously (44), and pulsed with 2 μg/mL peptide.
In Vivo Killing Assays. For in vivo killing assays, single-cell suspensions of
lymphocytes from naive B6.Ly5.2 (CD45.1) mice (National Cancer Institute)
were split into two. One set was labeled with 4 μM CFSE (CFSEhigh) and the
second set was labeled with 0.8 μM CFSE (CFSElow) and pulsed with 1 μg/mL
I1L7–21peptide). The two populations were mixed in a 1:1 ratio and 2 × 107
total cells were injected intravenously into recipient mice. Lymphocytes were
isolated 16 h later from lymph nodes, livers, and spleens. The presence of
CFSElow, CD45.1+, CFSEhigh, MHC II+and MHC II−cells was determined by flow
cytometry. Specific lysis was calculated as (percentage CFSEhigh/percentage
CFSElow). Percentage specific lysis = [1 − (ratio unprimed/ratio primed) × 100].
At least 100,000 cells were analyzed by flow cytometry using a LSR II system
(BD Biosciences).
IFN-γ ELISPOT Assay. Groups of three B6 mice were infected in the footpad
with 3 × 103pfu ECTV. Cells isolated from the livers and spleens from each
group were pooled (n = 3) and 5 × 105/well incubated at 37 °C and 5% CO2
for 18 h in 96 well ELISpot plates (Millipore) together with 2 × 105spleno-
cytes from naive mice that had been pulsed with 10 μg/mL of the indicated
peptide. IFN-γ+T-cell responses were assayed using IFN-γ ELISPOT (BD) as
directed by the manufacturer. Spots were counted using ImmunoSpot
software (Cellular Technology).
In Vivo Depletions. Depletion of CD4+and CD8+T cells was performed by
intraperitoneal inoculation of 200 μg anti-CD4 mAb GK1.5 or 200 μg anti-
CD8 mAb 2.43 1 d before ECTV infection. The efficiency of depletion was
>95%, as determined by flow cytometric analysis of white blood cells one
day after Ab inoculation.
Data Displayed and Statistical Analysis. Unless indicated, all displayed data
correspond to one representative experiment of at least three similar
experiments with groups of three to five mice. When indicated, the LNs,
livers, and spleens from the mice in a group were pooled. Statistical analysis
was performed using Prism (GraphPad Software) software. For survival
studies, P values were obtained using the Log-rank (Mantel-Cox) test. All
other statistical analyses were performed using an unpaired two-tailed
Student t test or the Mann–Whitney test as applicable. When applicable,
data are displayed with mean ± SEM.
ACKNOWLEDGMENTS. We thank Fox Chase Cancer Center Laboratory
Animal, Flow Cytometry, and Tissue Culture Facilities for their services;
Ms. Lucy Aburto for technical assistance; Ms. Holly Gillin for assistance in
the preparation of the manuscript; and the National Institute of Allergy
and Infectious Diseases (NIAID) Tetramer Facility for MHC II tetramers.
This work was supported by NIAID Grants R01AI048849, R01AI065544, and
5U19AI083008 (to L.J.S.) and NCI Grant P30CA006927 (to the Fox Chase
Cancer Center).
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