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TLR-Independent Induction of Dendritic Cell
Maturation and Adaptive Immunity by
Negative-Strand RNA Viruses
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of June 15, 2017.
Carolina B. López, Bruno Moltedo, Lena Alexopoulou,
Laura Bonifaz, Richard A. Flavell and Thomas M. Moran
J Immunol 2004; 173:6882-6889; ;
doi: 10.4049/jimmunol.173.11.6882
http://www.jimmunol.org/content/173/11/6882
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Copyright © 2004 by The American Association of
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References
The Journal of Immunology
TLR-Independent Induction of Dendritic Cell Maturation and
Adaptive Immunity by Negative-Strand RNA Viruses1
Carolina B. López,* Bruno Moltedo,* Lena Alexopoulou,2† Laura Bonifaz,‡ Richard A. Flavell,†
and Thomas M. Moran3*
D
endritic cells (DCs)4 mediate the transition from innate
to adaptive immunity by undergoing a major shift in
their maturation state that involves the up-regulation of
costimulatory molecules, and the secretion of proinflammatory cytokines necessary for the differentiation of effector and memory T
cells and B cells (1). The binding of pathogen-associated molecular patterns (PAMPs) to an assortment of receptors expressed on
the DCs, known as TLRs, constitute the primary mechanism described to trigger DC maturation (2). To date, more than 10 different TLRs have been identified in vertebrates, with specificities
for all major families of microbes (3).
A role for TLRs in monitoring viral infections has been established based on the discovery of TLRs responsive to viral elements. Double-stranded RNA (dsRNA), a viral PAMP generated
during virus replication, binds and activates TLR3 and CpG motifs
found in the DNA of HSV-activator TLR9 (5, 6). Moreover, single-stranded viral RNA (ssRNA), representing the genomic material of many different viruses, triggers the activation of TLR7 and
*Department of Microbiology, Mount Sinai School of Medicine, New York, NY
10029; †Section of Immunobiology, Yale University School of Medicine and Howard
Hughes Medical Institute, New Haven, CT 06529; and ‡Laboratory of Cellular Physiology and Immunology, The Rockefeller University, New York, NY 10021
Received for publication July 13, 2004. Accepted for publication September 22, 2004.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This work was supported by Grants 1R01AI41111 and 1PO1A148204 from the
National Institute of Allergy and Infectious Diseases (to T.M.M.) and by funds
granted by the Charles H. Revson Foundation (to C.B.L.). The statements made and
views expressed, however, are solely the responsibility of the authors.
2
Current address: Centre d’Immunologie de Marseille-Luminy, Centre National de la
Recherche Scientifique-Institut National de la Santé et de la Recherche Médicale,
Campus de Luminy, Case 906, 13288 Marseille Cedex 9, France.
3
Address correspondence and reprint requests to Dr. Thomas M. Moran, Department
of Microbiology, Mount Sinai School of Medicine, One Gustave L. Levy Place, New
York, NY 10029. E-mail address: [email protected]
4
Abbreviations used in this paper: DC, dendritic cell; BMDC, bone marrow-derived
DC; MOI, multiplicity of infection; PAMP, pathogen-associated molecular pattern;
pDC, plasmacytoid DC; RSV, respiratory syncytial virus; SeV, Sendai virus; NP,
nucleoprotein.
Copyright © 2004 by The American Association of Immunologists, Inc.
TLR8 (7–9). Viral envelope proteins (e.g., the respiratory syncytial
virus (RSV) fusion protein) have also been shown to trigger TLR
activation (10 –13). However, the lack of a common structure or
PAMP within these proteins questions the role of these interactions
in the generic recognition of viruses.
Despite the triggering of DC maturation through the binding of
viral components to TLRs, the role of TLR signaling in eliciting
antiviral adaptive immune responses is unknown. In humans, the
virus-responsive TLR7, TLR8, and TLR9 are expressed in a restricted subset of DCs, the plasmacytoid DCs (pDCs), which are
specialized in secreting large amounts of type I IFN after virus
recognition (14, 15). Controversy exists regarding the ability of
these cells to stimulate naive T cells. Although CpG-matured
pDCs are able to present Ag and stimulate naive T cells in vitro as
well as in vivo (16 –18), evidence also exists suggesting that pDCs
poorly present Ag and are unable to prime naive T cells (16, 19).
In addition, all virus-related TLRs can be stimulated by inactivated
viruses or viral RNA surrogates (4, 7, 8), but live virus infection is
needed for the optimal induction of DC maturation and immunity
against ssRNA viruses (20, 21). These observations argue that the
predominant role for TLR signaling during virus infection might
be to elicit innate immune responses rather than to generate adaptive immunity.
Thus, we set out to investigate whether the induction of DC
maturation and adaptive immunity to viral infections could be triggered independently of TLR signaling. To do this, we took advantage of the paramyxovirus Sendai virus (SeV) strain Cantell, a
particularly strong inducer of mouse DC maturation (22). We have
previously demonstrated that the potent DC maturation-inducing
activity of SeV is not dependent on the secretion of the antiviral
cytokine type I IFN, which has been related to the amplification
and preservation of DC maturation after infection with other viruses, such as Newcastle disease virus (21). SeV allowed us to
visualize the requirements for DC maturation after infection with
live viruses independently of the contribution of type I IFN signaling. Our data demonstrate that TLR3 or the TLR adaptor protein MyD88, necessary for the signaling of most TLRs, are not
0022-1767/04/$02.00
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TLR signaling leads to dendritic cell (DC) maturation and immunity to diverse pathogens. The stimulation of TLRs by conserved
viral structures is the only described mechanism leading to DC maturation after a virus infection. In this report, we demonstrate
that mouse myeloid DCs mature normally after in vivo and in vitro infection with Sendai virus (SeV) in the absence of TLR3, 7,
8, or 9 signaling. DC maturation by SeV requires virus replication not necessary for TLR-mediated triggering. Moreover, DCs
deficient in TLR signaling efficiently prime for Th1 immunity after infection with influenza or SeV, generating IFN-␥-producing
T cells, CTLs and antiviral Abs. We have previously demonstrated that SeV induces DC maturation independently of the presence
of type I IFN, which has been reported to mature DCs in a TLR-independent manner. The data presented here provide evidence
for the existence of a novel intracellular pathway independent of TLR-mediated signaling responsible for live virus triggering of
DC maturation and demonstrate its critical role in the onset of antiviral immunity. The revelation of this pathway should stimulate
invigorating research into the mechanism for virus-induced DC maturation and immunity. The Journal of Immunology, 2004,
173: 6882– 6889.
The Journal of Immunology
essential for the induction of costimulatory molecule expression or
the secretion of cytokines by mature DCs after in vitro or in vivo
infection and that their absence has no significant impact on the
generation of antiviral Th1 immunity in mice.
Materials and Methods
Viruses and mice
SeV Cantell and influenza A virus were grown, purified, and inactivated as
described (20, 22). RSV (A2 strain) was grown in mycoplasma-free Vero
cells. C57BL/6 and MHC class I⫺/⫺ mice were purchased from Taconic
Farms (Germantown, NY). MyD88⫺/⫺ mice were kindly provided by Dr.
S. Akira (Osaka University, Osaka, Japan) and Dr. R. Steinman (Rockefeller University, New York, NY). Age- and sex-matched animals were
used in all experiments. The animals were housed in pathogen-free conditions and the experiments were performed according to institutionally
approved protocols.
Bone marrow-derived DC (BMDC) preparation and treatment
DC isolation from lymph nodes and staining
Lung-draining lymph nodes were harvested 48 h after mice infection. Tissues were incubated 20 min with collagenase (Liberase Blendzymes;
Roche, Indianapolis, IN). Washed single cell suspensions were incubated
for 4 h with brefeldin A (BD Biosciences, San Diego, CA). Cells were
stained with allophycocyanin-conjugated anti-CD11c and FITC-labeled antiCD86 Abs (BD Biosciences). Intracellular staining for IL-12 was performed using the Cytofix/Cytoperm kit from BD Biosciences and PE-conjugated anti-IL-12 Ab (BD Biosciences). Infected, mock-infected, or LPStreated BMDCs were stained 24 h after treatment with FITC-conjugated
anti-CD86 or anti-CD80 Abs (BD Biosciences). Flow cytometry was performed in a XLS-Coulter station (Beckman Coulter, Miami, FL). Data was
analyzed with Flowjo software.
Mice infection, immunization, and Ab detection
Mice were exposed for 30 min to ⬃900 influenza virus particles/min/cm3
of air in an infection chamber (Glas-Col, Terre Haute, IN). For immunizations, mice were injected i.p. with 4 ⫻ 105 DCs or with 107 TCID50 of
influenza or SeV. Detection of antiviral IgG1 and IgG2b Abs in mouse
serum was performed by capture ELISA as described (20).
Spleen cell culture and cytotoxicity
Stimulator splenocytes from naive C57BL/6 or MyD88⫺/⫺ mice were infected with influenza or SeV at an MOI of 40 for 45 min at 37°C and
cultured with spleen cells from immunized animals at a 1:10 stimulator to
responder ratio for SeV-infected cells and a 1:1 stimulator to responder
ratio for influenza virus-infected cells (20). Effector cells from secondary in
vitro cultures were mixed with 51Cr-labeled infected or mock-infected EL4
target cells as described (20). Supernatants were harvested and gamma
radiation was measured. Killing of mock-infected EL4 cells was subtracted
from that observed with infected targets.
Cytokine detection
Supernatants from DC and spleen cell cultures were analyzed by Capture
ELISA for IL-6, IL-1␤, IL-12p40, TNF-␣, IL-4 (R&D Systems, Minneapolis, MN), and IFN-␣ (PBL Biomedical Laboratories, Piscataway, NJ)
following the manufacturer’s protocols. IFN-␥ was measured using Ab
pairs from BD Biosciences.
In vivo CTL assay and tetramer staining
Splenocytes from naive mice were pulsed at 4 ⫻ 107 cells/ml with 20 ␮M
SeV nucleoprotein (NP)324 –332 peptide or without peptide in PBS for 15
min at room temperature. The cells were then labeled at 2 ⫻ 107 cells/ml
with different concentrations of CFSE (Molecular Probes, Eugene, OR; 2.5
or 0.125 ␮M) at 37°C for 30 min. Labeling was stopped with an equal
volume of FBS for 1 min and 1 ⫻ 107 cells of each peptide pulsed and
unpulsed were mixed and injected i.v. into infected and uninfected syngenic mice. After 20 h from the injections, spleens were harvested and
single cell suspensions were prepared and analyzed by flow cytometry.
Percentage of specific killing was calculated as described (24): 100 ⫺ [(%
FIGURE 1. TLR3⫺/⫺ DCs mature normally after infection with SeV. C57BL/6 (wt) and TLR3⫺/⫺ DCs (ko) were either infected with SeV or RSV or
treated with poly(I:C) or LPS. After 24 h of culture, the cells were stained for CD86 or CD80 expression (filled histogram) and analyzed by flow cytometry
(a). Thin line represents isotype controls. Cytokines in the culture supernatants were analyzed after 24 h of culture by capture ELISA (b).
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BMDCs were prepared as previously described (22) and infected with either influenza (multiplicity of infection, (MOI) of 10), SeV (MOI of 0.6 –
10) or RSV (MOI of 2). Control BMDCs were mock infected or incubated
overnight with 10 ng/ml LPS (Sigma-Aldrich, St. Louis, MO) or 50 ␮g/ml
poly(I:C) (Amersham Biosciences, Bucks, U.K.). For maturation studies
the cells were incubated overnight at 1 ⫻ 106 cells/ml in media containing
1% normal mouse serum. Removal of remaining extracellular virus particles was performed as described using neutralizing anti-PR8 influenza
(PY102 and PY210) or anti-SeV Abs (obtained from Dr. A. Portner, St.
Jude Children’s Hospital, Memphis, TN) (23).
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TLR-INDEPENDENT INDUCTION OF DC MATURATION BY VIRUSES
peptide pulsed in infected mice/% unpulsed in infected mice)/(% peptide
pulsed in uninfected/% unpulsed in uninfected)] ⫻ 100. To determine the
percentage of T cells bearing the specific anti-SeV-NP324 –332 peptide
H-2Kb receptor, splenocytes or lung single cell suspensions were obtained
10 –12 days after infection with SeV and costained with anti-CD8 FITC Ab
and PE-labeled H-2Kb tetramers carrying the SeV NP324 –332 peptide (National Institutes of Health tetramer core facility).
Statistical analysis
Statistical analysis was performed by using t test, paired two sample
analysis.
Results
TLR3 signaling is not required for DC maturation after
infection with SeV
The first evidence for viral-induced TLR signaling was the demonstration that the synthetic dsRNA analog poly(I:C), as well as
isolated viral dsRNA, could induce the expression of costimulatory
molecules on B cells and the secretion of cytokines from APCs
through TLR3 signaling (4). Since then, a number of seemingly
contradictory studies have attempted to define the role of TLR3 in
dsRNA-induced cell activation (8, 21, 25–27). Concurring with
reports by others (8, 21, 25), we observe that the up-regulation of
costimulatory molecules and the secretion of IFN-␣ are independent of TLR3 signaling when poly(I:C) is used to treat DCs. In
contrast, the secretion of IL-12 and TNF is impaired in the absence
of this receptor (Fig. 1).
To test the role of TLR3 in the triggering of DC maturation after
live virus infection, DCs were infected with SeV. LPS and RSV
were used as controls for TLR3-independent induction of DC maturation as they have been shown to trigger the secretion of cytokines through binding to TLR4 (10, 28). As shown in Fig. 1, normal costimulatory molecule up-regulation and cytokine secretion
is observed after SeV infection of TLR3⫺/⫺ DCs when compared
with wild-type DCs, mimicking the effect seen when TLR3-independent stimuli (LPS or RSV) are used. These results demonstrate
that TLR3 stimulation is not necessary for the triggering of DC
maturation after infection with SeV.
DC maturation after SeV infection is independent of MyD88mediated signaling
In mice, TLR7, TLR8, and TLR9 are expressed in myeloid DCs,
pivotal in the initiation of adaptive immunity (29). To test the
requirement for TLR7, 8, or 9 stimulation in the induction of myeloid DC maturation, we took advantage of mice deficient in
MyD88, an adaptor protein essential for their signaling (7, 8).
MyD88 binds to members of the IL-1R-associated protein kinases
and signals further via the TNFR-associated factor-6 (7, 9).
MyD88⫺/⫺ DCs show normal up-regulation of CD80 and CD86
following SeV infection (Fig. 2a). IL-12, IL-6, TNF, and IFN-␣
secretion by infected DCs is also completely independent of
MyD88 after SeV infection (Fig. 2b). Additionally, and in agreement with previous reports (22), the induction of proinflammatory
cytokine secretion by SeV-infected DCs depends on virus replication as shown by the absence of cytokines in response to UVinactivated virus (Fig. 2b). In contrast, the induction of cytokine
secretion by the TLR-activators RSV and LPS is completely abolished in MyD88⫺/⫺ cells and RSV replication is not a requirement
for the stimulation of cytokine secretion by DCs (Fig. 2b). The
independence of MyD88 signaling for the secretion of type I IFN
and other proinflammatory cytokines and for the up-regulation of
costimulatory molecules, is observed as early as 6 h postinfection
(Fig. 2c and data not shown) and is independent of the dose of SeV
used (Fig. 2d and data not shown). These data demonstrate that
signaling by the adaptor protein MyD88 is not necessary for the
induction of DC maturation after infection with SeV.
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FIGURE 2. MyD88⫺/⫺ DCs mature normally after infection with SeV. C57BL/6 and MyD88⫺/⫺ DCs were infected with either live or UV-inactivated
SeV or RSV, or treatment with LPS. After 24 h of culture, the cells were stained for CD86 or CD80 expression (filled histogram) and analyzed by flow
cytometry (a). Thin line represents isotype controls. Cytokines in the culture supernatants were analyzed after 6 (c) or 24 (a, c, and d) h of culture by capture
ELISA (b). Data are representative of more than three experiments.
The Journal of Immunology
DCs mature normally after SeV infection in MyD88⫺/⫺ mice
To corroborate our in vitro observations, we analyzed the induction of DC maturation, in vivo, after intranasal infection of
MyD88⫺/⫺ mice with SeV. Two days after infection with influenza or SeV, mature DCs with up-regulated costimulatory molecules and producing proinflammatory cytokines, can be obtained
from the lymph nodes of infected animals (Ref. 30 and data not
shown). As shown in Fig. 3a, mature DCs with up-regulated CD86
can be seen in the lung-draining lymph nodes of wild-type and
MyD88⫺/⫺ mice 2 days after infection with SeV. Moreover, a
similar increase in the expression of IL-12 p40/p70 is observed in
DCs from wild-type and MyD88⫺/⫺ mice upon virus infection
despite a difference in the basal level of expression of this cytokine
among these animals (Fig. 3, b and c). These results demonstrate
that following SeV infection, DCs equivalently matured can be
distinguished in the lung-draining lymph nodes of wild-type and
MyD88⫺/⫺ mice.
MyD88⫺/⫺ mice expand IFN-␥ secreting cells and CTLs after
virus infection
To analyze the role of MyD88 signaling in the development of
antiviral immunity, we infected mice with SeV and analyzed the
primary generation of CTLs, as well as the development of antiSeV Abs. Both wild-type and MyD88⫺/⫺ mice expand CTLs that
eliminate target cells carrying the immunodominant SeV peptide
(NP324 –332), in vivo, 7 days after the infection (Fig. 4a). However,
less efficient killing is observed in MyD88⫺/⫺ mice consistent with
a reduction in the number of CD8⫹ cells bearing SeV-NP-specific
TCR, as detected by staining with NP-peptide bearing tetramers (Fig.
4b). Wild-type and MyD88⫺/⫺ mice develop equivalent amounts of
total virus-specific IgG Abs (data not shown) with a slight bias toward
the Th2-isotype IgG1 in MyD88⫺/⫺ mice (Fig. 4c).
Our laboratory has a well-characterized in vivo model for influenza infection (31). We took advantage of this model to study
the role of MyD88 signaling in the development of virus-specific
humoral and memory cellular responses. After infection with aerosolized live influenza virus, known to induce potent Th1 immunity
(31, 32), control and MyD88⫺/⫺ mice develop equivalent amounts
of total virus-specific IgG Abs (data not shown) with a slight bias
toward the Th2-isotype IgG1 in MyD88⫺/⫺ mice (Fig. 4d), similar
to that observed in SeV-infected animals (Fig. 4c). Moreover,
splenocytes from MyD88⫺/⫺ and wild-type mice restimulated in
vitro with influenza virus 14 days after the infection, secrete comparable levels of the Th1 cytokine IFN-␥ together with a remarkable production of the Th2 cytokine IL-4 only by MyD88⫺/⫺ cells
(Fig. 4e). A small reduction in the clearance of both viruses is
observed in MyD88⫺/⫺ animals when compared with wild-type
mice 7 days postinfection (data not shown), but both groups of animals show complete recovery from infection as measured by recovery
of lost weight and virus clearance from the lungs 10 days postinfection (data not shown). Interestingly, despite the secretion of IL-4 and
the Th2-biased Ab production, normal development of memory CTL
precursors is observed in MyD88⫺/⫺ mice (Fig. 4f). Thus, although
MyD88⫺/⫺ mice show minor differences in the in vivo response to
both, SeV and influenza virus, a normal immune adaptive response is
generated that leads to complete recovery from infection.
Virus-infected DCs deficient in MyD88 signaling induce
polarized Th1 immune response in mice
MyD88 is a required element in the signaling through IL-1 and
IL-18 receptors, both members of the Toll/IL-1R containing receptor family (33, 34). Therefore, the less efficient development of
CD8⫹ cells bearing SeV-NP-specific TCR (Fig. 4b) and a nonpolarized memory immune response in MyD88⫺/⫺ mice may result
from a failure of these cytokines to promote the differentiation of
CD4⫹ T cells into a Th1 phenotype (35). To unambiguously determine the role of MyD88 in the initiation of antiviral immunity,
its function in APCs must be distinguished from its effect in T
cells. We immunized mice by adoptively transferring ex vivo infected DCs to distinguish the effect of MyD88 on these cells from
that on T cells. DCs, which are abortively infected by influenza
and SeV, have been shown to efficiently induce protective antiviral
immunity (23). BMDCs were infected with influenza or SeV and
injected i.p. into wild-type animals. Both IgG1 and IgG2b Abs are
produced when control or MyD88⫺/⫺-infected DCs are used for
immunization (Fig. 5a). No evidence for a Th2 (IgG1)-biased response is seen when immunizing with infected MyD88⫺/⫺ DCs as
observed in directly immunized MyD88⫺/⫺ mice (Fig. 4a). In addition, high levels of IFN-␥ were secreted by splenocytes from
animals immunized with infected wild-type or MyD88⫺/⫺ DCs
(Fig. 5b), corresponding with the development of CTLs (Fig. 5c).
We were unable to detect IL-4 production in any of the cultures.
Similarly, adoptively transferred TLR3⫺/⫺ DCs efficiently induced
antiviral CTLs when infected with influenza or SeV (data not
shown).
To validate this model, we analyzed the contribution of crosspresentation to the immune response generated by this method.
This phenomenon occurs when Ag is transferred from dead cells to
endogenous cells which present antigenic peptides on their own
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FIGURE 3. Normal maturation of MyD88⫺/⫺ DCs after SeV infection
in vivo. C57BL/6 (wt) and MyD88⫺/⫺ mice were infected intranasally with
10 ID50 of SeV. Two days later, the lung-draining lymph nodes were collected and stained for FACS analysis. CD11c⫹ cells were gated. Cells from
mock-infected (gray line) or infected (black line) animals were stained for
CD86 (a). Isotype control is represented in the filled histogram. Cells were
also intracellularly stained for IL-12p40/p70 (b). The percentage of IL12p40/p70 expression over the basal level presented on noninfected cells is
shown in c.
6885
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TLR-INDEPENDENT INDUCTION OF DC MATURATION BY VIRUSES
MHC class I molecules and could bias the interpretation of results
obtained when DCs are used for immunization (36). Control or
MHC class I⫺/⫺ DCs were infected with SeV and injected into
C57BL/6 mice. As shown in Fig. 6, a and b, MHC class I⫺/⫺ DCs
are very inefficient in generating CTL only partially eliminating
target cells carrying the immunodominant SeV peptide NP324 –332.
This correlates with more than a 70% reduction of CD8⫹ cells
bearing a TCR specific for the SeV NP324 –332 peptide as detected
by staining with NP324 –332-conjugated tetramers (Fig. 6c). These
data demonstrate that at least 50% of the CTLs generated after DC
immunization are the result of direct presentation of viral Ags by
the injected DCs, and allowed us to demonstrate that DCs deficient
in TLR3 or MyD88 can induce normal antiviral adaptive immunity
in mice.
Discussion
DC maturation, necessary for the transition from innate to adaptive
immunity, can be triggered by TLR signaling after recognition of
microbial structures. Because a number of TLRs can signal in response to viruses, we sought to determine whether the virus-triggered TLR signaling is essential for the maturation of myeloid DC,
and the subsequent induction of antiviral immunity.
TLR7 and TLR8 are receptors for ssRNA, the genetic material
of many viruses including those used in this study (7–9). In hu-
mans, the expression of these TLRs is restricted among the DC
populations to pDCs, and most studies analyzing their role in cell
activation by viruses have only focused on this DC subpopulation
(7, 8). In mouse, TLR7 and TLR8 are also expressed in myeloid
DCs, the primary cells responsible for the induction of adaptive
immunity (29). As signaling through these receptors has been
shown to be dependent on MyD88, our results using MyD88⫺/⫺
BMDCs demonstrate that in mice TLR7 and TLR8 are not needed
for maturation of myeloid DCs in response to SeV. The failure of
inactivated SeV to trigger DC maturation strengthens the argument
that TLR7 and TLR8, which have been shown to be stimulated by
inactivated virus, are not necessary for myeloid DC maturation.
TLR3 that responds to the virus replication intermediary dsRNA, is
also expressed on myeloid DCs. This receptor signals by MyD88dependent and -independent pathways forcing us to investigate its role
in the induction of DC maturation by SeV separately from the MyD88
dependent TLRs. Our results demonstrate that TLR3 signaling is not
needed for the induction of myeloid DC maturation by SeV and
agrees with accumulating data arguing that TLR3 signaling is not
necessary for the induction of DC maturation and immunity by other
viruses (9, 21, 37). Furthermore, DCs from MyD88⫺/⫺ and wild-type
mice showed identical up-regulation of costimulatory molecules and
an increase in IL-12 production following in vivo infection with SeV,
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FIGURE 4. MyD88⫺/⫺ mice generate a nonpolarized immune response against influenza and SeVs. C57BL/6 (wt) and MyD88⫺/⫺ (ko) mice were
infected intranasally (i.n.) with SeV (a– c) or with aerosolized influenza virus (d–f). Six days after SeV infection, peptide pulsed and unpulsed CFSE-labeled
cells were injected into the animals. Histograms show the number of CFSElow unpulsed or CSFEhigh peptide-pulsed cells 20 h after injection (a). Results
are representative of two experiments. Average percentage of specific killing from two experiments is also shown. Ten days after infection lymphocytes
from the lung were stained for CD8⫹ cells bearing the SeV NP peptide-specific TCR using PE-labeled tetramers (b). Two weeks after virus infection,
anti-SeV (c) and influenza virus (d) Abs were measured from the mouse sera (1/5000 and 1/1000 serum dilution shown for SeV and influenza virus,
respectively), ⴱ, p ⬍ 0.006. Splenocytes from infected mice were restimulated in vitro (e and f). Cytokine secretion was determined by capture ELISA after
3 days of culture (e). CTL generation was determined after 5 days of culture by a 51Cr release assay (f).
The Journal of Immunology
corroborating the dispensable role of MyD88 signaling in the induction of DC maturation after infection with SeV observed in vitro.
To complement these studies, we show that the development of
an efficient antiviral adaptive immune response is independent of
TLR3 or MyD88-dependent activation. Although a slight Th2 bias
in the antiviral immune response is observed in MyD88⫺/⫺ mice,
probably resulting from a deficiency in IL-1 and IL-18 signaling,
viral clearance, the production of IFN-␥ by T cells, and cytotoxic
T cell expansion are not significantly different from control animals. The normal functioning of MyD88⫺/⫺ DCs is supported by
experiments adoptively transferring virus-infected MyD88⫺/⫺
DCs into wild-type mice. No Th2 bias was observed under these
conditions and MyD88⫺/⫺ DCs induced cytokines, Ab and CTL
generation are indistinguishable from the animals infused with virus-infected wild-type DCs. Studies using MHC I⫺/⫺ DCs validated the model system by showing that a significant part of the
response observed following adoptive transfer is elicited by the
injected cells and not by cross-priming or in vivo uptake of virus
or viral proteins. The independence of MyD88 signaling for the
normal development of antiviral immunity concurs with the reported normal antiviral immunity in children carrying a mutation
in IL-1R-associated protein kinase-4, a crucial downstream element in the signaling pathway that uses the adaptor MyD88 (38).
Altogether, the data presented argue for a TLR-independent
pathway for myeloid DC maturation that is dependent on virus
replication. The cellular machinery responsible for the induction of
myeloid DC maturation after virus infection remains elusive. Although the dsRNA-dependent protein kinase R has been shown to
mediate these events upon transfection of poly(I:C), we and others
have demonstrated that live virus-induced maturation is independent of protein kinase R (21, 22), arguing that the pathway activated by replicating virus may be separate or at least more complex than what is triggered by dsRNA alone. Further studies are
required to determine the role of other cellular proteins, such as the
TANK (TNFR-associated factor-associated NF-␬B kinase) binding kinase I, the inhibitor of NF-␬B kinase ⑀ (39, 40), NOD (nucleotide-binding oligomerization domain) (41), and the recently
described RNA helicase retinoic acid inducible gene I (42), which
are known to be triggered by dsRNA, in the induction of DC maturation after virus infection.
Additionally, type I IFN produced by all virus-infected cells as
a means to contain viral replication and spreading has been shown
to be required for optimal DC maturation after infection with Newcastle disease virus (21). We have previously shown that SeVinduced maturation of myeloid DC is independent of exogenous
type I IFN signaling (22). In addition, SeV efficiently blocks the
IFN signaling pathway in infected cells (43). We have noted that
viruses that strongly activate the type I IFN synthesis machinery,
such as SeV strain Cantell, are also the best inducers of DC maturation (22) suggesting the existence of shared components in the
induction pathways of type I IFN production and DC maturation.
The difference in the need for exogenous type I IFN signaling for
the complete maturation of DCs after different virus infections may
depend on the strength of the viral activation signal. In light of
these observations, DCs may require, in some instances, exogenous sources of type I IFN for optimal maturation. Type I IFN
released after TLR signaling not reliant on virus replication and
therefore not inhibitable by type I IFN antagonists, may fulfill this
requirement. This would suggest a novel physiologically relevant
role for pDCs as endogenous adjuvants.
Despite our observations, a growing number of examples implicate TLRs in the recognition of diverse viruses, including the
binding of the RSV F protein to TLR4 (10) and the measles virus
H protein to TLR2 (11). Interestingly, even though these interactions result in activation of APCs and the secretion of cytokines,
they do not always result in the development of efficient antiviral
immunity. This is well exemplified by the ineffective immune response generated against RSV (44, 45). Moreover, no defined
common structure seems to be involved in the binding of these
viruses to TLRs leaving open the possibility that these obligatory
intracellular pathogens escape or conveniently use the scrutiny of
TLRs. Interestingly, RSV induces a MyD88-dependent secretion
of cytokines by DCs but fails to up-regulate costimulatory molecules (Fig. 3). This unusual TLR-mediated maturation of DCs by
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FIGURE 5. Influenza and SeV-infected MyD88⫺/⫺ DCs induce a polarized Th1 immune response in mice. C57BL/6 mice were immunized i.p.
with ex vivo-infected C57BL/6 (f) or MyD88⫺/⫺ (p) DCs. Control
C57BL/6 (wt) mice were directly immunized with influenza (Flu) or SeV.
Two weeks after virus infection anti-influenza or SeV Abs were measured
from the mouse sera (1/1000 serum dilution shown) (a) and splenocytes
were restimulated in vitro. IFN-␥ secretion was determined by capture
ELISA after 3 days of culture (b). CTL generation was determined after 5
days of culture by a 51Cr release assay (c).
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TLR-INDEPENDENT INDUCTION OF DC MATURATION BY VIRUSES
RSV could be fundamental for the generation of the impaired
adaptive immune response against this virus that allows multiple
re-infections during childhood.
In conclusion, we have demonstrated that SeV can induce efficient DC maturation and immunity in a TLR3- and MyD88-independent manner. This pathway, which requires live virus, implies
a novel mechanism for virus detection and initiation of an antiviral
immune response. Its further characterization should allow a better
understanding of the mechanisms of viral antagonism that interfere
with the generation of immunity, as well as the identification of
molecular targets amenable to manipulation for the control of antiviral immune responses and the improvement of vaccination
strategies.
Acknowledgments
We acknowledge Dr. Adolfo Garcia-Sastre for critically reviewing this
manuscript, Mount Sinai Hybridoma Shared Research Facility (New York,
NY) for preparation of Abs to Sendai virus, and Luis Muñoz and
Sharon Czelusniak for invaluable technical support.
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