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Extralymphatic virus sanctuaries as a consequence of
potent T-cell activation
Mike Recher
1,10
, Karl S Lang
1,10
, Alexander Navarini
1,10
, Lukas Hunziker
1,2,10
, Philipp A Lang
1
,
Katja Fink
1
, Stefan Freigang
1
, Panco Georgiev
3
, Lars Hangartner
1
, Raphael Zellweger
1
, Andreas Bergthaler
1
,
Ahmed N Hegazy
1,4
, Bruno Eschli
1
, Alexandre Theocharides
5
, Lukas T Jeker
6
, Doron Merkler
1,7
,
Bernhard Odermatt
8
, Martin Hersberger
9
, Hans Hengartner
1
& Rolf M Zinkernagel
1
T helper cells can support the functions of CD8
+
T cells against persistently infecting viruses such as murine lymphocytic
choriomeningitis virus (LCMV), cytomegalovirus, hepatitis C virus and HIV. These viruses often resist complete elimination and
remain detectable at sanctuary sites, such as the kidneys and other extralymphatic organs. The mechanisms underlying this
persistence are not well understood. Here we show that mice with potent virus-specific T-cell responses have reduced levels and
delayed formation of neutralizing antibodies, and these mice fail to clear LCMV from extralymphatic epithelia. Transfer of virus-
specific B cells but not virus-specific T cells augmented virus clearance from persistent sites. Virus elimination from the kidneys
was associated with the formation of IgG deposits in the interstitial space, presumably from kidney-infiltrating B cells. CD8
+
T cells
in the kidneys of mice that did not clear virus from this site were activated but showed evidence of exhaustion. Thus, we conclude
that in this model of infection, site-specific virus persistence develops as a consequence of potent immune activation coupled with
reductions in virus-specific neutralizing antibodies. Our results suggest that sanctuary-site formation depends both on organ
anatomy and on the induction of different adaptive immune effector mechanisms. Boosting T-cell responses alone may not reduce
virus persistence.
Some viruses are prone to developing persistent infections despite the
presence of virus-specific immune responses
1
. Identified mechanisms
of persistence include mutational escape from T-cell responses, resis-
tance to interferons, expression of cytokine decoy receptors, inhibition
of the cellular antigen-presentation machinery, glycan shielding of
surface glycoproteins and integration into the host’s genome
2–5
. Virus
sanctuaries—persistence of virus in extralymphatic tissues—not only
complicate virus elimination, they also enable reactivation during
immune suppression (for example, after organ transplantation)
6
.
Virus persistence in certain tissues is partly restricted by the expression
of specific receptors on tissue target cells; however, classical lympho-
tropic and/or hepatotropic viruses tend to persist in extralymphatic
organs such as the kidney, brain, lungs, testes and salivary glands
7–12
.
Persistence in these organs cannot always be explained by selective
receptor expression; it also cannot be due simply to continuous virus
spillover from the blood, as peripheral virus isolates differ phylo-
genetically from blood virus isolates and extralymphatic virus has been
detected in individuals without viremia
13
.
Virus persistence and distribution are also shaped by innate resis-
tance mechanisms
14,15
and drug therapy
16
. T helper cells are known to
support CD8
+
T-cell responses that destroy cells infected with persist-
ing viruses such as HIV, hepatitis C virus (HCV), cytomegalovirus
(CMV) and LCMV
17–19
. However, the outcome of chronic HIV,
HCV or CMV infections cannot always be predicted from T-cell
responses
20–22
. Paradoxically, potent immune activation is a risk factor
for end-stage disease in HIV
23
and simian immunodeficiency (SIV)
infections
24
and has been associated with disease severity in cases of
human influenza virus and severe acute respiratory syndrome (SARS)-
coronavirus infection
25,26
. Experimental immunization with T helper
epitopes has even been found to worsen the outcome of SIV infection
in rhesus macaques
27
. We and others have recently described T helper
cell–driven suppression of neutralizing antibody formation after
LCMV infection
28,29
. These findings together indicate that potent
immune activation, which is usually beneficial, can sometimes be
disadvantageous. To help delineate the mechanisms that influence this
equilibrium, we have investigated the roles of specific T cells and
Received 13 August; accepted 25 September; published online 4 November 2007; doi:10.1038/nm1670
1
Institute for Experimental Immunology, University Hospital Zu
¨
rich, Schmelzbergstrasse 12, CH-8091 Zu
¨
rich, Switzerland.
2
Department for Internal Medicine,
University Hospital Basel, 4031 Basel, Switzerland.
3
Department of Visceral and Transplantation Surgery, University Hospital Zu
¨
rich, Ra
¨
mistrasse 100, 8091 Zu
¨
rich,
Switzerland.
4
German Rheumatology Research Center, Charite
´
Platz 1, D-10117 Berlin, Germany.
5
Experimental Hematology, Department of Research, Basel
University Hospital, 4031 Basel, Switzerland.
6
Pediatric Immunology, Center for Biomedicine, University of Basel and University Children’s Hospital of Basel,
Mattenstrasse 28, 4058 Basel, Switzerland and Transplantation Immunology and Nephrology, University Hospital Basel, 4031 Basel, Switzerland.
7
Department of
Neuropathology, Georg August University, Goettingen, Germany.
8
Department of Pathology, University Hospital, Schmelzbergstrasse 12, CH-8091 Zu
¨
rich, Switzerland.
9
Institute of Clinical Chemistry, University Hospital Zu
¨
rich, Ra
¨
mistrasse 100, CH-8091 Zu
¨
rich, Switzerland.
10
These authors contributed equally to this work.
Correspondence should be addressed to M.R. (rechermike@bluewin.ch).
1316 VOLUME 13
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protective antibodies in enhancing or preventing the formation of
extralymphatic persisting virus sanctuaries.
RESULTS
Induction and function of CD8
+
T cells depends on T helper cells
C57BL/6 mice infected with low-dose (200 plaque-forming units
(PFU)) LCMV (WE strain), rapidly cleared the virus from all organs
tested within 10 d (Fig. 1a). Infection of extralymphatic organs was
absent or low, even at the peak of virus replication. To allow the virus
to spread widely in the mouse before the onset of the CD8
+
T cell–
derived immune response, we transiently depleted CD8
+
T cells by
administering monoclonal antibodies 1 or 2 d before infection (that is,
on day –2 or day –1) with LCMV-WE (standard protocol, used
throughout this manuscript). Ten days after infection, mice that had
been depleted of CD8
+
T cells showed virus replication in all organs
tested (Fig. 1a). After their initial depletion, CD8
+
T cells reemerged
in the blood around day 10 after infection and then expanded, peaking
at day 20 after infection as assessed by fluorescence-activated cell
sorting (FACS) analysis of peripheral blood (Fig. 1b). This expansion
of CD8
+
T cells was almost completely virus specific, as up to 80% of
these CD8
+
T cells were tetramer-specific for known immunodomi-
nant virus epitopes (GP33-41, GP276-284 and NP396-404) (Fig. 1c;
for representative FACS plot, see Supplementary Fig. 1a online). The
CD8
+
T cells were mostly of a CD62L
lo
phenotype, indicating
activation (Fig. 1d). Most CD8
+
T cells expressed granzyme B, and
a small fraction produced interferon-g (IFN-g) directly ex vivo with-
out further restimulation (Fig. 1e; for representative FACS plots see
Supplementary Fig. 1b). To determine the importance of T helper
cells in CD8
+
T-cell priming and proliferation in our model, we
infected Cd4
–/–
mice with 200 PFU LCMV-WE using the standard
protocol described above. Expansion of CD8
+
T cells in peripheral
blood, as detected by FACS, was suppressed in Cd4
–/–
mice (Fig. 1b).
CD8
+
T cells were less activated, as downregulation of the selectin
CD62L was impaired (Fig. 1d). Granzyme B–expressing CD8
+
T cells
were also relatively reduced in the absence of T helper cells (Fig. 1e
and Supplementary Fig. 1b). By contrast, adoptive transfer of naive,
virus-specific T-cell receptor transgenic CD4
+
T helper cells (Smarta
cells
30
) before LCMV infection of C57BL/6 mice (C57BL/6+Smarta)
enhanced CD8
+
T-cell activation. CD62L downregulation and direct
ex vivo granzyme B expression were both greater than in control
C57BL/6 mice (Fig. 1d,e). Transfer of Smarta T helper cells augmented
formation of IFN-g by T helper cells (Supplementary Fig. 1c). At the
same time that CD8
+
T cells peaked in mouse blood samples, the mice
lost weight (Fig. 1f) and developed hepatitis, as indicated by elevated
serum bilirubin concentrations and augmented liver transaminase
activity in serum (Fig. 1g,h). Both weight loss and hepatitis correlated
markedly with the number of available T helper cells (Fig. 1f–h).
Thus, CD8
+
T-cell expansion and expression of effector functions
(granzyme B and IFNg) correlated with available T-cell help.
Delayed virus clearance in mice with potent T-cell responses
As expected, Cd4
–/–
mice infected with LCMV showed high viral titers
in blood at all time points measured (Fig. 2a). However, virus
elimination from the blood was, paradoxically, delayed in mice with
high T–helper cell responses (C57BL/6+Smarta) compared with con-
trol C57BL/6 mice (Fig. 2a). The impaired virus clearance in mice
7
6
5
4
<3
80
60
40
20
0
Virus organ titer
(log
10
ml
–1
)
CD8
+
cells
(% of blood lymphocytes)
CD8
+
Tetramer
+
cells
(% of CD8
+
cells)
CD62L
lo
CD8
+
cells
(% of CD8
+
cells)
3
Spleen Liver Kidney Lung Brain
20
15
Anti-CD8
Time after infection (d)
10
5
0
120
110
100
90
80
70
01020
Time after infection (d)
30 40
0 102030
60
80
60
40
20
0
40
20
6
1,500
1,000
500
0
5
4
3
2
1
<0.1
GP276 GP33 NP 396
C57BL/6+Smarta
C57BL/6
Cd4
–/–
C57BL/6+Smarta
C57BL/6
Cd4
–/–
0
CD8
+
T-cell function
(% of CD8
+
cells)
Weight
(% of starting weight)
C57BL/6+Smarta
C57BL/6
Cd4
–
/–
Serum bilirubin
(mg dl
–1
)
Serum ALT
(units per L)
C57BL/6+Smarta
C57BL/6
Cd4
–/–
abcd
hgfe
Figure 1 T helper cell–dependent CD8
+
T-cell response against distributed virus. (a) C57BL/6 mice were depleted of CD8
+
T cells at days –2 and –1 (’)
or left untreated (&) before infection with 200 PFU LCMV-WE. Organ viral titers were measured 10 d later (n ¼ 3). (b) C57BL/6 mice (’), C57BL/6 mice
supplemented with 10
7
Smarta splenocytes on day –1 before infection (C57BL/6+Smarta, &)orCd4
–/–
mice () were all transiently CD8
+
T-cell depleted
with monoclonal antibodies administered on days –2 and –1 before infection, then infected with 200 PFU LCMV-WE on day 0 or left uninfected (E).
CD8
+
T cells in peripheral blood were monitored at the indicated time points by FACS analysis (n ¼ 5–10). (c) Twenty days after infection of C57BL/6
mice as described above, reemerging CD8
+
T cells were tested for LCMV specificity using tetramer staining for three immunodominant epitopes
(GP33-41, GP276-284 and NP396-404) and FACS analysis (n ¼ 4–9). (d,e) CD62L expression (d) or direct ex vivo granzyme B (black) and IFN-g (gray)
expression (e) of CD8
+
T cells were measured by FACS 15 d (d)or20d(e) after infection (n ¼ 5). (f–h) C57BL/6 mice (’), C57BL/6+Smarta mice (&)
or Cd4
–/–
mice () were infected with LCMV according to the standard protocol (n ¼ 4–5). Mouse body weight was measured at the indicated time points
(f). (g,h) 20 d after LCMV infection, serum bilirubin concentrations and serum amino liver transaminase (ALT) activity were measured (n ¼ 5).
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with potent T-cell responses was mostly due to a failure to clear the
virus from the kidneys and lungs (Fig. 2b–f). Compared to Rag1
–/–
mice, which are deficient in T cells and B cells, virus clearance was
1,000–10,000 times greater in the spleen and liver of C57BL/6+Smarta
mice, although no significant difference was measured in the kidneys
and lungs (Fig. 2b–f). The difficulty in eliminating the virus from the
kidneys was still observed 50 d after infection of C57BL/6+Smarta
mice, whereas virus control in the lungs and brain was comparable to
that of control C57BL/6 mice at this time point (Supplementary
Fig. 2a,b online). Notably, no virus could be detected in the blood at
this time point (Fig. 2a). The observed site-specific virus elimination
in C57BL/6+Smarta mice was not due to site-specific T helper cell
responses, as T helper cell activation and function were comparable in
the liver, kidneys and lungs (Supplementary Fig. 2c).
In summary, potent T-cell responses were apparently effective in
lowering viral burden in some organs but did not clear virus from
other tissues.
Virus persistence correlates with delayed antibody formation
LCMV-infected C57BL/6 mice developed measurable neutralizing
antibodies by 30 d after infection (Fig. 3a). In keeping with our earlier
results
28
, the formation of neutralizing antibodies was, paradoxically,
delayed and reduced in mice supplemented with Smarta T helper cells,
to a similar extent as in mice lacking CD4
+
T cells (Fig. 3a). To test
whether the delayed antibody formation was responsible for the
observed site-specific virus persistence, we analyzed virus elimination
in mice deficient in antibody formation. Aicda
–/–
mice lack activation-
induced cytidine deaminase (AID) and therefore do not undergo
7
6
5
4
3
7
P = 0.001
P < 0.001
P = 0.07
P = 0.07 P > 0.1
P = 0.062
P = 0.005 P < 0.001
P = 0.012
6
5
4
3
<3
<3
<3
2
0102030
Time after infection (d)
C57BL/6
C57BL/6
Rag1
–
/–
Rag1
–/–
C57BL/6+Smar
ta
C57BL/6+Smar
ta
C57BL/6
Rag1
–
/–
C57BL/6+Smar
ta
C57BL/6
Rag1
–
/
–
C57BL/6+Smar
ta
C57BL/6
Rag1
–/–
C57BL/6+Smar
ta
40 50
<1.7
Blood viral titer
(log
10
PFU ml
–1
)
Liver viral titer
(log
10
PFU)
Brain viral titer
(log
10
PFU)
Spleen viral titer
(log
10
PFU)
7
6
5
4
3
7
6
5
4
3
<3
7
6
5
4
3
<3
7
6
5
4
3
Kidney viral titer
(log
10
PFU)
Lung viral titer
(log
10
PFU)
P < 0.001
P < 0.001
P < 0.001 P < 0.001P > 0.1 P > 0.1
abcd
fe
Figure 2 Organ-specific viral persistence enhanced by potent
cellular immune responses. (a) Viral blood titers of C57BL/6
mice (’), C57BL/6 mice supplemented with Smarta T helper
cells (C57BL/6+Smarta, &)orCd4
–/–
mice infected with LCMV
according to the standard protocol (
) were analyzed at the
indicated time points (n ¼ 3–5). (b–f) Twenty days after LCMV
infection using the standard protocol, viral titers in the spleen
(b), liver (c), brain (d), kidneys (e)andlungs(f) were measured
in individual Rag1
–/–
mice, C57BL/6 mice and C57BL/6+Smarta
mice as indicated (individual values are shown).
5
7
P = 0.015
P = 0.015
P < 0.001
P < 0.001
P = 0.001
P = 0.03
P = 0.02
P = 0.06
6
5
4
3
<3
0 10203040
Time after infection (d)
50 60 Spleen Liver Brain Kidney Lung
4
3
2
1
<1
Neutralizing titer
(–(log
2
) × 10)
Viral organ titer
(log
10
PFU)
Viral organ titer
(log
10
PFU)
LCMV-Arm organ titer
(log
10
PFU)
Spleen
NS
Liver Brain Kidney Lung Spleen Liver Kidney Lung
Spleen Liver Brain Kidney Lung
7
6
5
4
3
<3
7
6
5
4
3
<3
P = 0.030
P = 0.003
P > 0.1
P = 0.014
7
6
5
4
3
<3
Viral organ titer
(log
10
PFU)
ab c d
e
Figure 3 Site-specific viral persistence as a consequence of delayed antibody responses. (a) C57BL/6 mice
(’), C57BL/6 mice supplemented with transgenic virus–specific CD4
+
T cells (C57BL/6+Smarta, &), Cd4
–/–
mice (), Aicda
–/–
mice (E) and VI10Yen mice (B) were depleted of CD8
+
T cells on days –2 and –1, then
infected with 200 PFU LCMV-WE on day 0. (a) At the indicated time points, neutralizing antibody formation
was assessed by neutralization assay (n ¼ 4–9). (b–e) Aicda
–/–
mice were transiently depleted of CD8
+
T
cells on days –2 and –1, then infected with 200 PFU LCMV-WE (b) or LCMV-Armstrong (LCMV-Arm) (d)on
day 0. LCMV-WE–infected Rag1
–/–
mice were analyzed as a control (c). Alternatively, Aicda
–/–
mice that had
not been transiently depleted of CD8
+
T-cells were infected with 2 10
6
PFU LCMV-WE (e). Viral organ
titers were measured by plaque assay 30 d (b)or50d(c,d,e) after infection (individual values are shown).
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immunoglobulin class switch recombination and affinity maturation,
whereas other B-cell functions, including IgM responses, remain
effective
31
. LCMV-infected Aicda
–/–
mice did not produce neutralizing
antibodies, as shown by a neutralization assay (Fig. 3a), and also did
not produce LCMV glycoprotein–specific IgG responses, as tested by
ELISAs (Supplementary Fig. 4b,c online). Notably, there were only
slight differences in numbers of virus-specific
CD8
+
T cells, distribution of epitopes, pro-
duction of IFN-g, and CD8
+
T-cell cytotoxi-
city between Aicda
–/–
mice and C57BL/6 mice
(Supplementary Fig. 3a–e online). Like
C57Bl/6 mice supplemented with Smarta
cells, the LCMV-infected Aicda
–/–
mice
showed site-specific virus persistence, as
indicated by significantly lower viral titers
in the spleen and liver 30 d after infection
when compared to the kidneys, lungs and
brain (Fig. 3b). In contrast, RAG-deficient
mice had evenly distributed high viral titers
in all organs tested (Fig. 3c). Site-specific
persistence was also found in Aicda
–/–
mice
that were infected with another LCMV strain,
LCMV-Armstrong (Fig. 3d), and in Aicda
–/–
mice infected with high doses of LCMV-WE
without initial transient CD8
+
T-cell
depletion (Fig. 3e).
Reduction of virus sanctuaries by
virus-specific B cells
To analyze whether supplemented B cells
were more effective than T cells at reducing
viral titers, we adoptively transferred either
LCMV-specific CD8
+
T cells or neutralizing
LCMV-specific B cells (see Methods) 30 d
after LCMV infection of Aicda
–/–
mice (for
blood virus kinetics, see Supplementary
Fig. 4a). Viral organ titers were measured
20 d later (see experimental schedule,
Fig. 4a). Fifty days after LCMV infection, Aicda
–/–
mice without cell
transfer showed the lowest viral titers in the liver and brain but had
high titers in the spleen, lungs and kidneys (Fig. 4b–f).Theincreasein
spleen viral titers at day 50 compared to day 30 is probably due to
CD8
+
T-cell exhaustion, which has been described to occur more
rapidly in lymphoid tissue than in other organs
32
. Control C57BL/6
Anti-CD8 LCMV Analysis
Spleen virus titer
log
10
PFU
Liver viral titer
(log
10
PFU)
Kidney viral titer
(log
10
PFU)
Brain viral titer
(log
10
PFU)
Time (d)
50
7
P < 0.001
P < 0.01
P < 0.05
P < 0.05
6
5
4
3
Without
+T cells
+B cells
VI10Yen
C57BL/6
<3
300–2,–1
Tra ns fe r
(B cells versus T cells)
7
6
5
4
3
<3
Without
+T cells
+B cells
VI10Yen
C57BL/6
P < 0.001
P < 0.01
P > 0.1
P > 0.1
7
6
5
4
3
<3
Without
+T cells
+B cells
VI10Yen
C57BL/6
P < 0.01
P > 0.1
P > 0.1
P > 0.1
P < 0.001
P < 0.01
P < 0.05
P > 0.05
7
6
5
4
3
<3
Without
+T cells
+B cells
VI10Yen
C57BL/6
Lung viral titer
(log
10
PFU)
7
6
5
4
3
<3
Without
+T cells
+B cells
VI10Yen
C57BL/6
P < 0.001
P < 0.05
P > 0.05
P > 0.1
abc
ef
d
Figure 4 Reduction of extralymphatic virus sanctuaries associated with LCMV
glycoprotein-specific B-cell responses. Aicda
–/–
mice were transiently depleted of
CD8
+
T cells on days –2 and –1, then infected with 200 PFU LCMV-WE on day 0.
Thirty days after infection, 1 10
7
LCMV-specific neutralizing B cells (+B cells)
or LCMV-specific CD8
+
T cells (+T cells) were adoptively transferred as described
in the Methods. As controls, Aicda
–/–
mice (without), VI10Yen mice (VI10Yen) or
C57BL/6 mice (C57BL/6) were infected with LCMV according to the standard
protocol but did not receive further adoptive cell transfer 30 d after infection. Viral
organ titers were measured 50 d after LCMV infection. (a) Experimental setup.
(b–f) Viral organ titers measured 50 d after LCMV infection in spleen (b), liver
(c), brain (d), kidney (e)andlung(f). Individual values are shown.
20
100
175
150
125
100
75
50
25
0
Liver
Kidney
Lung
75
50
25
PD-1
hi
CD69
hi
CD62L
lo
0
15
10
5
0
60
50
40
30
20
10
0
GP33 GP276 NP396
Tetramer
+
CD8
+
T cells
(% of total CD8
+
cells)
Specific lysis in spleen (%)
Specific lysis in liver (%)
Specific lysis in kidney (%)
Activated CD8
+
cells
(% of CD8
+
cells)
60
50
40
30
20
10
0
60
50
40
30
20
10
Threefold dilutions of effector cells
(starting at effector/target ratio of 100:1)
0
60
50
40
30
20
10
0
60
50
40
30
20
10
0
60
50
40
30
20
10
0
a
d
bc
Granzyme B expression
(MFI of CD8
+
cells)
Figure 5 T cells at sanctuary sites are activated but lack direct cytotoxicity. (a) Fifty days after infection
using the standard protocol, virus was isolated from Aicda
–/–
mouse kidneys. We then injected 200 PFU
of each of two isolates (&,
) from two mice into separate naive C57BL/6 mice. As a control, we used
200 PFU of wild-type LCMV-WE (’). Ten days later, GP33-41, GP276-284 and NP396-404 tetramer-
positive CD8
+
T cells were measured by FACS analysis (n ¼ 5). (b,c) Aicda
–/–
mice were infected with
LCMV according to the standard protocol. Thirty days after infection, CD8
+
T cells isolated from the
liver (black), kidneys (dark gray) or lungs (light gray) were assessed for expression of the indicated
activation markers (b) and for granzyme B expression (c)byFACSanalysis(n ¼ 3–4). (d) Aicda
–/–
mice
were infected with LCMV according to the standard protocol. Thirty days after infection, CD8
+
Tcells
isolated from the indicated organs were analyzed for cytotoxicity in a chromium-release assay. Target
cells were labeled with GP33-41 peptide (’) or unlabeled (&). Each line represents the mean and
s.e.m. for one animal (chromium release measured in duplicates; n ¼ 5–7 experiments).
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mice had almost completely cleared virus from all organs at this time
point (Fig. 4b–f). LCMV-specific adoptive B-cell transfer into Aicda
–/–
mice was successful, as indicated by the rapid formation of LCMV
glycoprotein–specific IgG (Supplementary Fig. 4b), and was asso-
ciated with significantly reduced viral titers in the kidneys and a trend
toward lower titers in the lungs (Fig. 4e,f). By contrast, T-cell transfer
did not significantly reduce viral titers in the kidneys or lungs
(Fig. 4e,f). Transfer of LCMV glycoprotein–specific neutralizing anti-
bodies into Aicda
–/–
mice did not lower viral titers in the kidneys or
lungs and did not induce measurable glycoprotein-specific IgG
responses, implying that the antibodies were rapidly consumed by
circulating virus particles (data not shown). We also analyzed LCMV-
infected VI10Yen mice. In these mice, most naive B cells are specific
for vesicular stomatitis virus (VSV)
33
. These mice did not produce
LCMV-neutralizing antibodies for up to 50 d after infection (Fig. 3a);
however, in contrast to AID-deficient mice, they eventually formed
LCMV glycoprotein–specific IgG antibodies (Supplementary Fig. 4c).
CD8
+
T-cell function in infected VI10Yen mice was comparable
to that in C57BL/6 mice (data not shown). Compared to those
in AID-deficient mice, viral titers in the kidneys were significantly
lower in VI10Yen mice, mimicking the result of virus-specific B-cell
transfer (Fig. 4b–f).
Together, these results show that the reduction of virus replication
in some organs, such as the kidneys and lungs, correlates with the
amount of LCMV glycoprotein–specific IgG antibodies but not with
T-cell responses. Notably, B-cell transfer into Aicda
–/–
mice did not
augment CD8
+
T-cell activation and function in extralymphatic
organs (Supplementary Fig. 4d–f).
ActivatedbutexhaustedCD8
+
T cells in viral sanctuaries
We next investigated the mechanisms that are involved in the
formation of site-specific viral sanctuaries. First, we analyzed whether
the persisting extralymphatic virus population consisted of
CD8
+
T-cell escape variants and thus resisted T-cell attack. Fifty
days after infection using the standard protocol, we isolated the
virus from the kidneys of two Aicda
–/–
mice. We then injected 200
PFU of each kidney isolate or 200 PFU of wild-type LCMV-WE
separately into naive C57BL/6 mice. Ten days later, we measured the
expansion of LCMV GP33-41–specific, GP276-284–specific and
NP396-404–specific CD8
+
T cells by tetramer staining, finding only
minimal differences (Fig. 5a). Lack of mutation was further confirmed
by sequencing of one representative kidney isolate, which revealed
LCMV-WE wild-type codon usage within and near the immuno-
dominant GP276-284 epitope (data not shown).
Next, we compared the livers, kidneys and lungs of AID-deficient
mice and found that CD8
+
T-cell activation and granzyme B expres-
sion were similar in these organs (Fig. 5b,c). Numbers of virus-specific
CD8
+
T cells and levels of IFN-g were also similar in the kidneys, lungs
and liver (Supplementary Fig. 3b,d), suggesting that differences in
CD8
+
T-cell activation and function could not explain the site-specific
virus persistence observed in Aicda
–/–
mice.
In contrast, CD8
+
T-cell cytotoxicity was lower in the kidneys
than in the liver and spleen (Fig. 5d). Direct cytotoxicity correlated
well with the measured viral titers in kidney versus liver and spleen at
the same time point (Fig. 3). Thus, although we could not demon-
strate differences in the intrinsic activation and function of CD8
+
T cells, the kidney CD8
+
T-cell population as a whole showed low
cytotoxicity, which is indicative of exhaustion and associated with
high viral kidney titers.
Virus persistence at epithelial sites
We performed immunohistological analysis of kidneys 50 d after
infection of mice. In Rag1
–/–
mice, which lack B and T cells, the
virus was mainly located in the kidney tubule epithelium (Fig. 6a, i).
Rag1
–/–
VL4
a
bc
i ii iii iv v vi
vii viii ix x xi xii
xiii xiv xv xvi xvii xviii
xix
xx xxi xxii xxiii xxiv
lgG
CD8
CD4
B-cell transfer
lgG
CD138
VL4
Antibody transfer No transfer
i
ii
iii
VL4
VL4
VL4
RAG
–
AID
–
Aicda
–/–
low dose
Aicda
–/–
high dose
C57BL/6
+Smarta
C57BL /6
VI10Yen
without anti-CD8
Figure 6 Viral persistence at epithelial sites.
(a) Fifty days after LCMV infection, Rag1
–/–
mice,
Aicda
–/–
mice with or without transient CD8
+
T-cell depletion, C57BL/6 mice, Smarta-
supplemented C57BL/6 mice (C57BL/6+Smarta)
or VI10Yen mice were infected with LCMV using
the standard protocol, immunohistology was used
to detect LCMV nucleoprotein (VL4, i–vi), IgG
(vii–xii) CD8 (xiii–xviii) and CD4 (xix–xxiv)in
kidney sections (n ¼ 2–5). (b) Aicda
–/–
mice
were LCMV infected with LCMV according to the
standard protocol. After 30 d, neutralizing
antibodies (monoclonal KL25 antibodies or
polyclonal neutralizing serum) or LCMV-specific
B cells were adoptively transferred. Twenty days
later, expression of IgG, LCMV antigen (VL4)
and plasma cells (CD138) was analyzed by
immunohistology as indicated. (c, i)C57BL/
6+Smarta mice were infected with LCMV
according to the standard protocol. Fifty days
later, immunohistological staining of LCMV-
nucleoprotein (VL4 antibody) was performed
on sections of the distal urinary tract (one
representative section is shown). (c, ii,iii) Aicda
–/–
mice were transiently depleted of CD8
+
T cells
on days –2 and –1, then infected with 200 PFU
LCMV-WE on day 0. As a control, Rag1
–/–
mice
were infected with LCMV on day 0. Fifty days
after infection, immunohistological staining
of LCMV-nucleoprotein (VL4 antibody) was
performed on liver sections (ii) or lung sections
(iii). Scale bars, 200 mm.
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Similar patterns of staining around the kidney tubules were observed
in AID-deficient mice, either with or without transient CD8
+
T-cell
depletion, and in mice transfused with Smarta T cells (Fig. 6a, ii,iii).
Hence, despite a T-cell response, viral distribution in the kidneys
remained the same as in Rag1
–/–
mice. By contrast, C57BL/6 mice had
already efficiently eliminated the virus from their kidneys 50 d after
infection (Fig. 6a, v). Extensive infiltrations of CD8
+
T cells (Fig. 6a,
xiii–xviii) and CD4
+
T cells (Fig. 6a, xix–xxiv) were observed in the
kidneys of all mice tested (except T cell–deficient Rag1
–/–
mice)
independently of virus distribution, confirming that insufficient
T-cell recruitment was not the reason for the epithelial persistence
of virus antigen in Aicda
–/–
mice and mice supplemented with Smarta
T helper cells.
To further analyze virus elimination, we carried out immunohisto-
logical analysis of kidneys from VI10Yen mice, which had low but still
detectable viral titers 50 d after infection (Fig. 4). In these mice, the
virus was found mainly in the kidney interstitial space, rather than in
epithelial cells. (Fig. 6a, vi). IgG staining in these mice localized to the
same interstitial sites, supporting the idea that virus-antibody immune
complexes form there (Fig. 6a, xii). Interstitial IgG deposition was also
found in C57BL/6 mice, implying that interstitial IgG deposition
remained after virus elimination (Fig. 6a, xi). As expected, IgG
staining was not detectable in infected Rag1
–/–
mice and Aicda
–/–
mice (Fig. 6a, vii–ix). When LCMV-neutralizing IgG was injected into
infected Aicda
–/–
mice IgG deposits were found mostly in kidney
glomeruli, but usually not in the interstitial space surrounding the
LCMV-infected epithelia (Fig. 6b), and LCMV kidney titers remained
unchanged (data not shown). Thus, virus elimination from kidneys
correlated with the formation of interstitial IgG deposits, presumably
produced by kidney-infiltrating B cells. Plasma cells could easily be
found within kidneys in immunohistological analyses (Fig. 6b).
Kidney plasma cells were also observed in Aicda
–/–
mice and
C57BL/6 mice, whereas IgG deposition between kidney tubules was
absent in Aicda
–/–
mice (Fig. 6b).
Virus was also detected in the lower urinary tract, where the
epithelium is built of multiple cell layers. There, the virus was detected
mainly in the upper cell layer (Fig. 6c), consistent with virus
propagation in the urinary tract tubular system. Histological analysis
of lung sections revealed similar virus sanctuaries in the bronchiolar
epithelium in Rag1
–/–
mice and Aicda
–/–
mice, indicating that lung
epithelia might show resistance to T-cell attack similar to that seen in
kidney epithelia (Fig. 6c). By contrast, in Aicda
–/–
mice, which had low
liver viral titers, the virus was found mainly in the Kupffer cell
compartment of the liver, whereas virus was detected in liver hepa-
tocytes in Rag1
–/–
mice (Fig. 3b and Fig. 6c).
DISCUSSION
Our experimental data mimic the paradoxical clinical finding that
potent immune activation is sometimes a risk factor for viral persis-
tence and severe disease. CD8
+
T cells, supported by helper T cells,
efficiently cleared the virus from some organs but not other organs.
Virus was particularly persistent in the kidneys and lungs, but more
efficiently cleared from the liver and spleen. We speculate that site-
specific viral persistence develops as a consequence of overactivated and
exhausted T-cell responses combined with slow induction of virus-
specific neutralizing antibodies. Such a combination is typical for
several chronic human virus infections
34
. B cell–deficient mice with
primed T cells specific for g-herpesvirus do not eliminate the virus
from the lungs, but do eliminate it from all other organs, upon rein-
fection
35
. In addition, X-linked lymphoproliferative disease is similarly
characterized by an overreactive cellular immune response combined
with slow induction and low levels of virus-specific antibodies; this
combination might lead to fatal outcomes of Epstein-Barr virus (EBV)
infection in humans and of LCMV infection in mice
36
.Inourmodelof
infection by LCMV, the CD8
+
T-cell response against distributed viral
antigen was strengthened by the help of CD4
+
T cells, as shown pre-
viously
18
. Similar findings exist for SIV and other persisting viruses
17,19
.
We found that the presence of virus-specific IgG responses was
associated with virus clearance and with changes in virus distribution
from epithelial cells to the interstitial space. Other models of persistent
virus infection have similarly proposed that B-cell responses are
associated with virus elimination
37,38
. Histological analysis showed
that T cells efficiently infiltrated virus-infected epithelia, in a pattern
reminiscent of ‘autoreactivity’ (for example, interstitial nephritis,
bronchiolitis or Sjo
¨
gren-like disease, which might convert into overt
autoimmune disease). Although we did not detect CTL-escape var-
iants at extralymphatic sites up to 50 d after infection, the observed
colocalization of CD8
+
T cells with the virus might eventually create
sites conducive to the formation of CTL-escape mutants
39
. Lympho-
mas associated with chronic viral infection, which often arise in
extralymphatic tissues, could be a consequence of ongoing extralym-
phatic immune responses. Together, our findings might explain why
chronic virus infections, lymphomas and autoimmunity have been
described as forming a triple association
40
.
In conclusion, the successful and complete elimination of a dis-
tributed persistence-prone viral infection requires efficient interplay
between cellular and neutralizing humoral responses. Whereas T-cell
responses can effectively clear the virus from both lymphatic and more
‘solid’ extralymphatic organs such as the liver, IgG responses are
especially needed in organs with tubular epithelial microanatomy,
such as the kidneys and lungs. The relative contributions of these two
adaptive arms of the immune system also regulate immunopatho-
logical disease mediated by chronic inflammation. Thus, boosting of
T-cell responses alone might not reduce virus persistence
41
and might
eventually induce immunopathology or autoimmunity
36
.Ourresults
indicate that virus replication, tropism and sanctuary formation
depend on the specific virus and on organ anatomy and physiology,
but also on the induction, kinetics and efficacy of different adaptive
immune effector mechanisms.
METHODS
Mice. All mice were bred and maintained under specific pathogen–free
conditions, and experiments were performed in accordance with institutional
and Swiss national guidelines. Animal care protocols were approved by the
animal experiment committe of Zu
¨
rich, Switzerland.
Cd4
–/–
, VI10Yen–B-cell transgenic, TgH(KL25)–B-cell transgenic, 318–T-cell
transgenic, Smarta1–T-cell transgenic, Rag1
–/–
and Aicda
–/–
mice all have a
C57BL/6 genetic background. All mice were purchased from the Institute for
Labortierkunde, Faculty of Veterinary Medicine, University Zu
¨
rich-Irchel.
Virus. LCMV strains WE and Armstrong (Arm) were originally gifts
(see Acknowledgments) and were propagated on L929 or BHK21 cells.
Mice were infected with 2 10
2
or 2 10
6
PFU of LCMV injected
intravenously as indicated.
Detection of viral and neutralizing antibody titers. LCMV viral titers were
detected by a plaque-forming assay on MC57 fibroblasts as described
42
.
Detection of neutralizing activity against LCMV in mouse sera has also been
described
42
. Briefly, neutralizing activity against LCMV-WE was measured in a
plaque-reduction assay. The neutralizing titer was defined as the log
2
dilution
that caused a half-maximal reduction in plaques, with the reduction being
measured in comparison to the number of plaques formed by similar amounts
of the virus incubated with control sera from uninfected mice or medium
alone. A titer of o1 indicates no detectable neutralization at an initial serum
predilution of 1:10.
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In vivo depletion of CD8
+
T cells. Cell depletions were performed using rat
monoclonal antibodies specific for CD8 (YTS 169.4) as described
28
.The
hybridomas were initially obtained from H. Waldmann. We injected 200 ml
of purified CD8
+
T cell–depleting antibodies intraperitoneally on days –2
and –1 before LCMV infection. The efficiency of depletion was always
confirmed by FACS analysis of peripheral blood cells using fluorescein
isothiocyanate (FITC)-, phycoerythrin (PE)-, or allophycocyanin (APC)-
labeled CD8-specific antibodies (Pharmingen).
Markers of liver injury. We measured serum activity of alanine aminotrans-
ferase (ALT) as a marker of hepatocellular injury and total serum bilirubin
concentrations using the serum multiple analyser (Ektachem DTSCII, Johnson
and Johnson) or the Roche Modular Analytics Analyser with pyridoxal
phosphate (Roche).
Adoptive cell transfer. Aicda
–/–
mice were transiently depleted of CD8
+
T cells
and infected with LCMV by the standard protocol, as described in Results.
After 30 d, either 1 10
7
MACS-sorted naive CD8
+
T cells from LCMV-
specific T-cell receptor–transgenic mice (318 mice) or in vivo CD8
+
Tcell–
depleted naive TgHKL25 splenocytes (from LCMV-specific B cell–knock-in
mice
33
) were adoptively transferred. Immunohistology and measurement of
organ viral titers and CD8
+
T-cell function were performed 20 d later.
LCMV-neutralizing monoclonal antibodies (KL25, 1.5 mg) or polyclonal
LCMV-neutralizing hyperimmune serum (200 ml) were transfused intrave-
nously as a control 30 d after LCMV infection of Aicda
–/–
mice.
LCMV-glycoprotein GP1-specific IgG measurements. The detection of
LCMV-glycoprotein GP-1–specific IgG by ELISA has been described
28
.
Fluorescence-activated cell sorting analysis of T-cell activation. Single-cell
suspensions of the different organs were prepared. Cells were then incubated
with fluorescent antibodies specific for PD-1 (PE-labeled, eBioscience), CD69
and CD62L (FITC-labeled, both BD Pharmingen), counterstained with fluores-
cent CD8-specific antibodies (APC-labeled, BD Pharmingen) and analyzed
using a FACSCalibur system (BD Pharmingen). For CD4
+
T-cell activation,
antibodies to CD69, CD62L and CXCR3 (BD Pharmingen) were used.
For intracellular cytokine staining and T-cell restimulation, splenocytes were
incubated for 5 h with or without LCMV peptide (1 mM) or PMA (50 ng ml
–1
)
and ionomycin (500 ng ml
–1
) for 6 h in RPMI medium supplemented with
10% FBS, b-mercaptoethanol and 5 mgml
–1
brefeldin A (Sigma-Aldrich). The
cells were then harvested, washed once in PBS with 4% FBS and 12.5 mM
EDTA, and stained with PE- or APC-conjugated CD8-specific antibodies or
PE-labeled antibodies specific for CD4 (BD Pharmingen). After washing, cells
were fixed with 4% paraformaldehyde in PBS for 10 min, then permeabilized
using PBS with 4% FBS, 12.5 mM EDTA and 0.1% saponin (Sigma-Aldrich).
For intracellular cytokine staining, cells were incubated with PE-conjugated
granzyme B–specific antibody (Caltag) or APC-labeled IFN-g–specific antibody
(BD Pharmingen) for 30 min at 4 1C. After washing twice with permeabiliza-
tion buffer, we resuspended the cells in PBS with 4% FBS and 12.5 mM EDTA
and analyzed them using a FACSCalibur system (BD Pharmingen).
In vivo cytotoxicity. In vivo cytotoxicity was assayed using C57BL/6 splenocytes
incubated for 1 h with or without LCMV-derived MHC class I GP276-284
peptide and labeled for 10 min with 5 mgml
–1
carboxyfluorescein diacetate
succinimidylester from Molecular Probes (CFSE
high
, peptide-labeled spleno-
cytes) or 0.5 mgml
–1
CFSE (CFSE
low
, unlabeled splenocytes). We injected
10
7
cells of each fraction intravenously into AID-deficient or control C57BL/6
mice 50 d after LCMV infection using the standard protocol described in
Results. The number of CFSE-positive cells remaining in blood 12 h later was
determined by FACS analysis. Specific cytotoxicity was calculated from the cell
counts of LCMV-infected (memory) and noninfected (naive) mice under each
treatment, as 1 – (CFSE
high
memory
/CFSE
low
memory
)/(CFSE
high
naive
/CFSE
low
naive
),
and expressed as a percentage.
51
Cr-release cytotoxicity assay. We performed
51
Cr release assays as
described
43
. Single-cell preparations of organs were prepared and lymphocytes
separated by gradient centrifugation (Lympholyte, Cedarlane). Threefold dilu-
tions of effector cells with a starting effector/target ratio of 100:1 were analyzed.
Immunohistology. Histological analysis was performed on snap-frozen tissue
sections stained with rat monoclonal antibodies against LCMV-nucleoprotein
(VL4), CD8 (53-6.7), IgG (mixture of rat anti-mouse IgG1, IgG2a/b and IgG3)
or CD138. Monoclonal antibodies were detected with goat anti-rat (Caltag)
and alkaline phosphatase-labeled donkey anti-goat (Jackson ImmunoResearch)
secondary antibodies, which we visualized by using naphthol 6-bromo-2-
hydroxy-3-naphtholic acid 2-methoxy anilide (AS-BI) phosphate and new
fuchsin as a substrate. The enzyme reaction yielded a red reaction product.
Kidney sections were stained in some experiments with goat anti-rat secondary
antibody (Caltag), followed by donkey anti-goat labeled with horseradish
peroxidase. Incubation with 3-amino-9-ethylcarbazole (AEC) in the presence
of H
2
O
2
yielded a brown reaction product. Sections were counterstained
with hemalum.
Statistical analysis. Data are presented as mean ± s.e.m. or as single values
(median value visualized as a horizontal line). Values in treatment groups were
analyzed for significant differences using a one-way analysis of variance
followed by a Tukey least-significant difference test. For statistical analysis,
organ viral titers below detection limits were set to 10
1.5
PFU.
Note: Supplementary information is available on the Nature Medicine website.
ACKNOWLEDGMENTS
We thank A.J. MacPherson and D. Pinschewer for discussions and K. Tschannen
for technical assistance. Aicda
–/–
mice were provided by T. Honjo (Kyoto
University). LCMV strains WE and Armstrong (Arm) were originally obtained
from F. Lehmann-Grube (Heinrich Pette Institute) and M. Buchmeier (Scripps
Institute), respectively. This work was supported by Swiss National Foundation
Grants to H.H. (3100A0-100779) and R.M.Z. (3100A0-100068). K.S.L. was
partially supported by the Deutsche Forschungsgemeinschaft LA 1419/1-1. A.B.
holds a PhD fellowship of the Boehringer Ingelheim Fonds. A.N.H. is a fellow of
Graduate School 1121 of the German Science Foundation.
AUTHOR CONTRIBUTIONS
M.R., K.S.L., A.N. and L.H. planned and performed most experiments and wrote
the manuscript; P.A.L. performed experiments during the revision process; S.F.
performed some experiments with AID-deficient mice; A.B. sequenced persisting
kidney virus; A.N.H. helped isolate lymphocytes for cytotoxicity experiments,
K.F., L.H. and R.Z. helped perform experiments with B cell–transgenic mice and
provided materials; B.E. helped perform LCMV glycoprotein–specific ELISAs and
provided materials; D.M. did statistical testing; B.O. performed and supervised
immunohistology; P.G. and M.H. performed and supervised measurement of
serum hepatitis markers; A.T. and L.T.J. contributed to discussions and writing of
the manuscript; and H.H. and R.M.Z. supervised and financed the project and
helped write the manuscript.
Published online at http://www .nature.com/naturemedicine
Reprints and permissions information is available online at http://npg.nature.com/
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ARTICLES
NATURE MEDIC INE VOLUME 13
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NUMBER 11
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NOVEMBER 2007 1323
© 2007 Nature Publishing Group http://www.nature.com/naturemedicine
