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Journal of Virology, August 2000, p. 7127-7136, Vol. 74, No. 15
0022-538X/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Mature Dendritic Cells Infected with Herpes Simplex
Virus Type 1 Exhibit Inhibited T-Cell Stimulatory Capacity
Monika
Kruse,1,*
Olaf
Rosorius,2
Friedrich
Krätzer,2
Gerhard
Stelz,2
Christine
Kuhnt,1
Gerold
Schuler,1
Joachim
Hauber,2 and
Alexander
Steinkasserer1
Department of
Dermatology1 and Institute for Clinical
and Molecular Virology,2 University of
Erlangen-Nürnberg, Erlangen, Germany
Received 29 November 1999/Accepted 26 April 2000
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ABSTRACT |
Mature dendritic cells (DC) are the most potent antigen-presenting
cells within the entire immune system. Interference with the function
of these cells therefore constitutes a very powerful mechanism for
viruses to escape immune responses. Several members of the
Herpesviridae family have provided examples of such escape strategies, including interference with antigen presentation and production of homologous cytokines. In this study we investigated the
infection of mature DC with herpes simplex virus type 1 (HSV-1) and the way in which infection alters the phenotype and function of
mature DC. Interestingly, the T-cell-stimulatory capacity of these DC
was strongly impaired. Furthermore, we demonstrated that HSV-1 leads to
the specific degradation of CD83, a cell surface molecule which is
specifically upregulated during DC maturation. These data indicate that
HSV-1 has developed yet another novel mechanism to escape immune responses.
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INTRODUCTION |
Herpes simplex virus type
1 (HSV-1) belongs to the Herpesviridae, a large and
diverse family of vertebrate pathogens including several human
pathogens such as HSV-1 and HSV-2 (23). Members of the
Herpesviridae characteristically have a large
double-stranded DNA genome, and virions consist of an icosahedral
nucleocapsid surrounded by a lipid layer envelope. After the initial
acute infection of the host, HSV-1 establishes a persistent infection. Such persistent infections are established when host immune responses fail to eliminate the virus which is produced during the acute stage of infection.
Dendritic cells (DC) are the most potent antigen-presenting cells and
are also capable of stimulating immunologically naive T cells (4,
8). DC have to mature in order to become potent T-cell
stimulators. As immature DC, they acquire antigens in peripheral tissues and migrate to the secondary lymphoid organs, where, as mature
DC, they present processed peptides very efficiently to rare
antigen-specific T cells. During this maturation, a variety of
different DC-specific genes are induced and expressed. In particular, costimulatory and accessory molecules, as well as major
histocompatibility complex (MHC) class I and class II molecules are
upregulated during DC maturation (4, 8, 30, 37, 38).
In addition to these gene products, CD83, a cell surface protein
representing one of the best markers for mature DC, is specifically upregulated during maturation (35, 39, 40). Although the precise function of CD83 remains elucidated, its upregulation during
maturation, together with the upregulation of the costimulatory molecules, points to an important function. Moreover, we recently demonstrated that inhibition of CD83 cell surface expression on mature
DC leads to a dramatic reduction of their T-cell stimulatory capacity
(19).
DC are essential stimulators of antiviral immune responses. In the
murine system, for instance, DC are the most effective antigen-presenting cells in stimulating recall cytotoxic T-lymphocyte (CTL) responses to Sendai virus, Moloney murine leukemia virus (17), HSV (14), and influenza virus
(22). The ability of DC to also induce a protective immune
response to viral infection has recently been demonstrated by adoptive
transfer with lymphocytic choriomeningitis virus peptide-pulsed DC and
with DC which constitutively expressed the same lymphocytic
choriomeningitis virus epitope (20). DC were also found to
be pivotal for the initiation of anti-influenza virus responses at the
CTL level (13).
The exposure of human DC to influenza virus leads to an efficient
infection in vitro, as shown by the expression of the viral antigens
hemagglutinin (HA) and nonstructural protein 1 (NS1) (7).
However, this infection does not lead to rapid cell death and only
small amounts of infectious virus particles are produced. Interestingly, infected DC, but not macrophages, can induce CTL responses from purified blood CD8+ cells in the absence of
exogenous cytokines (5).
For HSV-1, it has been shown that viral proteins are expressed which
can interfere with the immune response. For instance, ICP47, an
immediate-early protein, inhibits TAP function in human fibroblasts and
keratinocytes (18) by binding via its N terminus to the
cytosolic peptide binding domain of human transporter associated with
antigen presentation (TAP) (1, 33, 34) and thereby inhibits
MHC class I-mediated peptide presentation (36).
In this study we show that HSV-1-infected mature DC do not propagate
infectious virus particles and, interestingly, that these infected DC
have a dramatically reduced T-cell-stimulatory capacity. Furthermore,
we demonstrate that HSV-1 leads to the specific degradation of CD83, a
cell surface molecule which is specifically upregulated on mature DC,
indicating a new strategy for viral immune evasion.
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MATERIALS AND METHODS |
Cells and culture.
Peripheral blood mononuclear cells
(5 × 107) were isolated from healthy donors by
sedimentation in Ficoll-Hypaque (Pharmacia Biotech, Uppsala, Sweden)
and cultured in RPMI 1640 (BioWhittaker, Walkersville, Md.)
supplemented with 1% human plasma from a single AB donor, glutamine
(300 µg/ml), penicillin-streptomycin (20 µg/ml each), and 10 mM
HEPES (pH 7.5) (Sigma, Deisenhofen, Germany). DC precursors were seeded
onto immunoglobulin G (IgG)-coated (10 µg of Ig-globulin per ml from
the Cohn fraction [Sigma]) culture dishes for 1 h. A first
nonadherent fraction was harvested after 1 h, and a second was
harvested after a further 7 h. Immature DC were generated using
the cytokines granulocyte-macrophage colony-stimulating factor (GM-CSF)
(800 U/ml [Novartis Research Institute, Vienna, Austria]) and
interleukin-4 (IL-4) (1,000 U/ml [Genzyme Corp., Cambridge, Mass.]).
Fresh medium (3 ml) containing 4,000 U of GM-CSF and 5,000 U of IL-4
was added to the cell culture on day 3. On day 5, nonadherent cells
were collected, counted, and transferred into new dishes and the final
DC maturation was induced by adding tumor necrosis factor alpha (2.5 ng/ml [Boehringer Ingelheim, Vienna, Austria]), prostaglandin
E2 (1 ng/ml [Cayman Chemical, Ann Arbor, Mich.]), GM-CSF
(200 U/ml), and IL-4 (250 U/ml) to the medium.
HSV-1 infection of mature DC.
Mature DC were prepared from
healthy donors, washed, and resuspended in RPMI 1640 at a density of
0.5 × 106 to 1 × 106 cells/ml. The
cells were inoculated either with infectious HSV-1 (strain ang)
(21) at a multiplicity of infection (MOI) of 0.01, 0.1, or 1 or with UV-inactivated HSV-1 for 1 h at 37°C. Following three
washes, 0.5 × 106 cells were transferred into
six-well plates and cultivated for a further 24 h. Control
experiments were performed in the presence of cycloheximide (CHI)
(Sigma) at a final concentration of 10 µg/ml.
FACS analyses.
The mature DC phenotype was analyzed by
fluorescence-activated cell sorting (FACS) using the following
monoclonal antibodies: CD83 (Immunotech, Marseilles, France); CD25,
CD80, and CD86 (Becton-Dickinson, Heidelberg, Germany); and CD40, CD95,
MHC class I, and MHC class II (PharMingen, Hamburg, Germany). As
isotype controls, IgG1a, IgG2a and IgG2b were used (Becton-Dickinson)
and run in parallel. Then 10 h postinfection the HSV-1 infection
efficacy was determined using a VP16-specific antibody in combination
with intracellular FACS staining (3).
Reverse transcription-PCR.
Total cellular RNA was isolated
from DC (106) infected with HSV-1 at a MOI of 0.01, 0.1, or
1, using TRIzol (Gibco-Life Technologies, Eggenstein, Germany).
Subsequently, the RNA was reverse transcribed into single-stranded cDNA
by using avian myeloblastosis virus reverse transcriptase as specified
by the manufacturer (Roche Molecular Biochemicals, Mannheim, Germany).
These cDNAs were used as PCR templates to amplify the viral transcripts
ICP27, ICP8, and glycoprotein G (gG), as well as CD83- and S14-specific fragments.
The following PCR primers were used: ICP27 sense
(5'-CGAGACCAGACGGGTCTCCTGG-3') and antisense
(5'-GCAGACACGACTCGAACACTCCTG-3'), ICP8 sense
(5'-CTCGGTAACGACCAGATACAGGAGG-3') and antisense
(5'-CCAAGACGGCAACCACCATCAAGG-3'), gG sense
(5'-CATGCCAAGTATTGGACTGGAGGAG-3') and antisense
(5'-CACAGGTGTGTCGCCATCGCAC-3'), CD83 sense
(5'-GTTATTGGAGGGTGGTGAAGAGAGG-3') and antisense
(5'-GTGAGGAGTCACTAGCCCTAAATGC-3'), and S14 sense
(5'-GGCAGACCGAGATGAATCCTCA-3') and antisense
(5'-CAGGTCCAGGGGTCTTGGTCC-3'). The following PCR cycling
profile (30 cycles) was used: 95°C for 60 s, 60°C for 60 s, and 72°C for 90 s. The reaction products were analyzed on 2%
agarose gels and visualized by ethidium bromide staining.
Determination of viral particle numbers using a plaque test.
Mature DC were infected with HSV-1 for 24 h at an MOI of 0.01, 0.1, or 1. The cells were subsequently washed twice with
phosphate-buffered saline (PBS) and lysed by three freeze-thaw steps.
Cell debris was removed by centrifugation, and the viral titer produced
by the infected DC was determined in the supernatants. Cell
supernatants were diluted six times (10
1 to
106), and 100 µl of each dilution was added for 1 h at 37°C to confluent Vero cells (5 × 106
cells/well) in 24-well plates. After virus particle adsorption, 1 ml of
minimal essential medium (Gibco-BRL) containing 10% fetal calf serum
(Gibco-BRL) was added. After 48 h, the medium was removed and
cells were fixed for 2 min using 5% (vol/vol) formalin in water. Then
cells were washed five times with water, incubated with 1% (vol/vol)
crystal violet in water for 2 min, and washed again five times with
water. The viral titers were determined by plaque counting.
Allogeneic T-cell proliferation.
Human PBMC were isolated
from buffy coats, and the T-cell fraction was purified by rosetting
with neuraminidase-treated sheep red blood cells as described by Bender
et al. (6). T cells (2 × 105/well) were
cocultured with DC, which had been inoculated with infectious virus or
with UV-inactivated virus or left untreated, for 4 days in 200 µl of
RPMI 1640 supplemented with 5% human serum from a single AB donor in
96-well plates. In addition, the allogeneic mixed leukocyte reaction
(MLR) assay was performed using a mixture of uninfected and infected DC
as follows. A total of 200, 900, 2,900, or 9,900 uninfected or
HSV-1-infected DC were added to 100 HSV-1-infected DC and cocultured
with T cells as described above. Then the cells were pulsed with
[3H]thymidine (1 µCi/well) (Amersham, Braunschweig,
Germany) for 8 to 16 h. The culture supernatants were harvested
onto glass fiber filters (Printed Filtermat A; Wallac, Turku, Finland)
by using an ICH-110 harvester (Inotech, Dottikon, Switzerland), and [3H]thymidine incorporation was determined using a
microplate counter (Wallac).
Protein gel electrophoresis and immunoblotting.
Mature DC
(0.5 × 106) were harvested, washed twice with PBS,
and then inoculated with infectious virus or with UV-inactivated virus,
treated with CHI, or left untreated. The cells were collected by
centrifugation and solubilized in gel-loading buffer (50 mM Tris-HCl
[pH 6.8], 2% sodium dodecyl sulfate [SDS], 10% glycerol, 5%
-mercaptoethanol, 0.1% bromphenol blue), cell extracts were separated by SDS-polyacrylamide gel electrophoresis, and the proteins were transferred to nitrocellulose membranes (Amersham). The membranes were blocked with 10% dry milk (Nestlé, Glendale, Calif.) in TBST (150 mM NaCl, 10 mM Tris-HCl, 0.05% Tween 20 [pH 8.0]) at 4°C
overnight, incubated for 2 h at 25°C with a monoclonal anti-CD83 antibody (Dianova, Hamburg, Germany), washed five times in TBST, and
incubated with the secondary antibody coupled to peroxidase (Dianova)
at a dilution of 1:10,000 in TBST plus 5% dry milk for 1 h at
25°C. Specific bands were visualized using an enhanced chemiluminescence detection system (Amersham).
Immunofluorescence microscopy.
The expression plasmid
p3CD83, coding for the entire CD83 molecule, was microinjected into the
nuclei of Vero cells, cultured for 20 h. Subsequently, cells were
inoculated with infectious HSV-1 at an MOI of 1 for a further 7 h.
Then indirect immunofluorescence studies were performed. Cells were
fixed with 2% paraformaldehyde (Merck, Darmstadt, Germany) and
subsequently permeabilized using 0.1% Triton X-100 (Sigma) for 4 min
and blocked with 1% bovine serum albumin (Sigma) for 30 min. The cells
were then stained for 30 min with a monoclonal anti-CD83 antibody
(Immunotech), an anti-cathepsin D antibody (kindly provided by A. Hille, Göttingen, Germany), or an anti-VP16 antibody. Following
extensive washing steps in PBS, cells were incubated for 30 min with
appropriate secondary antibodies conjugated to Cy2- or Cy3-conjugated
fluorophores (Rockland, Gilbertsville, Pa.). Samples were washed in
PBS, mounted in Mowiol (Calbiochem, Bad Soden, Germany), and analyzed
using an Axiovert-135 microscope (Zeiss, Jena, Germany). Images were recorded with a cooled MicroMax charge-coupled device camera (Princeton Instruments, Stanford, Calif.) and processed using IPLap Spectrum and
Adobe Photoshop software packages. All reactions were performed at room temperature.
DC which were inoculated as described above were also centrifuged onto
polylysine-coated microslides (Menzel-Gläser, Mainz,
Germany) for
30 s at 400 rpm using a Cytospin 3 centrifuge (Shandon,
Pittsburgh, Pa.). CD83 expression was determined using indirect
immunofluorescence as described
above.
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RESULTS |
Phenotypic characterization of mature DC infected with HSV-1.
Mature DC were generated from nonproliferating blood progenitors by
using an improved two-step protocol (6, 24). Briefly, DC
precursors were generated from adherent monocytes cultured in medium
containing 1% human plasma as well as GM-CSF and IL-4. A cytokine
cocktail composed of tumor necrosis factor alpha, prostaglandin E2, GM-CSF, and IL-4 (15; M. Kruse et
al., unpublished data) was then used to generate stable mature DC.
These DC showed the characteristic features of mature DC, e.g.,
stellate cell shape; nonadherence to plastic; high expression of MHC
class I and class II molecules, adhesins, and costimulatory molecules;
and a very potent T-cell-stimulatory activity (data not shown).
Using these cells, we wanted in particular to investigate whether HSV-1
interferes with the phenotype and/or function of DC
as observed with
other viruses such as vaccinia virus (
9) and
measles virus
(
28). FACS analyses of the cell surface expression
of
typical DC markers were performed to monitor the DC phenotype.
Strikingly, 24 h after HSV-1 infection, the surface expression
of
CD83 was drastically reduced (Fig.
1A,
left panel). In sharp
contrast, untreated DC or those which were
inoculated with UV-inactivated
HSV-1 showed no CD83 downregulation
(Fig.
1A, left panel). On
the other hand, HSV-1 had only a limited
effect on the cell surface
expression of MHC class II molecules (Fig.
1A, right panel). Furthermore,
other investigated molecules, including
CD25, CD40, CD80, CD86,
CD95 and MHC class I, revealed no significant
changes in their
cell surface expression (data not shown).
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FIG. 1.
(A) HSV-1 infection induces CD83 downregulation on
mature DC. Mature DC were inoculated either with infectious HSV-1 at a
MOI of 1 or with UV-inactivated virus and analyzed by FACS analysis
after 0, 4, and 24 h. A strong downregulation of cell surface
expression of CD83 was observed (left panel). No such effects could be
observed when DC were left untreated or inoculated with UV-inactivated
HSV-1. In contrast, the expression of MHC class II molecules was not
dramatically affected by the HSV-1 infection (right panel). (B) Time
course of HSV-1-induced CD83 downregulation. At 10 h after the
HSV-1 infection (MOI = 1), a dramatic reduction of the CD83 cell
surface expression is already observed. (C) CD83 downregulation is
HSV-1 titer dependent. Mature DC were infected with HSV-1 at different
MOIs (0.01, 0.1, and 1) and analyzed after 24 h by FACS analysis.
The viral infection rate significantly influences the CD83
expression.
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To expand our studies on the CD83 downregulation, more elaborate time
course experiments were conducted. This analysis clearly
demonstrated
that CD83 downregulation occurs between 4 and 10
h after virus
infection (Fig.
1B). Next we tested if this effect
was MOI dependent.
No downregulation of CD83 was visible at an
MOI of 0.01. Only a slight
effect was observed at an MOI of 0.1,
while a dramatic CD83
downregulation was observed at an MOI of
1 (Fig.
1C). Furthermore,
there was no increase in toxicity upon
HSV-1 infection during the first
24 h, as monitored by propidium
iodide staining and FACS analysis
(data not
shown).
HSV-1 infects mature DC very efficiently.
To determine the
viral infection level, mature DC were inoculated with HSV-1 at an MOI
of 1 and analyzed 10 h postinfection. Using a VP16-specific
antibody in combination with indirect immunofluorescence microscopy, we
demonstrated that most of the DC were infected (Fig.
2A); intracellular FACS analyses showed
that over 90% of the DC were infected (Fig. 2C).
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FIG. 2.
Efficient infection of mature DC by HSV-1. Mature DC
were infected with HSV-1 at an MOI of 1 and analyzed after 10 h by
intracellular FACS staining or by indirect immunofluorescence using
HSV-1 VP16-specific antibody. (A and B) VP16 staining (A) and
phase-contrast microscopy (B) of infected DC. (C) More than 90% of the
mature DC were infected after this time.
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Analyses of HSV-1 transcripts in infected mature DC.
On the
RNA level, we next investigated whether HSV-1 was able to express
various classes of viral mRNAs in mature DC. Using specific PCR
primers, we amplified fragments of prototypical immediate-early (ICP27), early (ICP8), and late (gG) transcripts. As shown in Fig.
3A, the immediate-early ICP27 and early
ICP8 gene products were transcribed after 24 h at an MOI of 1. Interestingly, in contrast to ICP27 and ICP8, only very small amounts
of the late gG transcript could be amplified at this MOI (Fig. 3A,
right lanes). These data indicate that mainly immediate-early and early
HSV-1 transcripts are synthesized at larger amounts in mature DC. Next, the levels of CD83-specific mRNA were investigated. As shown in Fig.
3B, there was no difference in CD83 transcription after 4 and 10 h
when HSV-1-infected (MOI = 1) and uninfected samples were
compared. These data clearly show that the downregulation of CD83 from
the cell surface, which was already observed 10 h after HSV-1
infection (Fig. 1B), was not due to an inhibition of CD83
transcription.
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FIG. 3.
RT-PCR analyses of HSV-1- and CD83-specific mRNA. (A)
Total cellular RNA was isolated after 24 h and reverse transcribed
from uninfected and HSV-1 (MOI = 1)-infected mature DC.
HSV-1-specific PCR primers were used to amplify transcripts from
immediate-early (ICP27), early (ICP8), and late (gG) gene transcripts.
Amplification of ribosomal protein S14-specific sequences served as an
internal control. (B) RT-PCR of CD83-specific mRNA. No difference in
the CD83 signals between mock- and HSV-1-infected samples after 4 and
10 h postinfection was detected.
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Mature DC do not support the generation of infectious HSV-1
particles.
Next we checked whether infectious virus particles were
generated in infected cells. Mature DC were thus infected for 1 h at different MOIs (0.01, 0.1, and 1), and 24 h later the viral titer present in the supernatants was analyzed using the plaque test.
Supernatants of infected mature DC were added to HSV-1-permissive Vero
cells. The cells were stained with crystal violet after a 48-h
incubation, and the numbers of lysed cells were determined. Supernatants from HSV-1-infected Vero cells, inoculated at an MOI of
0.01, were included as a control and revealed high viral titers
(106 viral particles/ml) (Fig.
4, control).
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FIG. 4.
HSV-1-infected mature DC do not produce infectious
virus. Mature DC were infected with HSV-1 at a MOI of 0.01, 0.1, or 1. Cell supernatants were analyzed using the RITA-plaque test after
24 h to determine the amount of infectious particles generated
during the infection. No or only low levels of viral particles were
detected in supernatant derived from HSV-1-infected DC. In sharp
contrast, Vero cells inoculated at a MOI of 0.01 generated
106 virus particles per ml (control). Note the logarithmic
scale of the y axis.
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In contrast, we were not able to detect any infectious viral particles
in supernatants from DC which were infected at an MOI
of 0.01, and only
very low levels of infectious particles were
detected in the
supernatant from DC infected at an MOI of 0.1
(approximately 300 particles/ml). Even at an MOI of 1, only 10
3 virus
particles per ml were generated. These 10
3 virus particles
could theoretically also be derived from the
stock initially used to
infect the DC. These data indicate that
HSV-1 does not productively
replicate at high levels in mature
DC.
HSV-1 affects the allogeneic T-cell stimulatory capacity of
DC.
The most distinctive functional characteristic of mature DC is
their ability to stimulate T cells very potently (4, 8, 30,
32). Therefore, we assessed whether HSV-1 interferes with this
stimulatory capacity. Mature DC were either left untreated or
inoculated with infectious HSV-1 or with UV-inactivated HSV-1. These
cultures were then compared in an allogeneic MLR assay. DC treated with
UV-inactivated HSV-1 and untreated DC showed a very high stimulatory
capacity (Fig. 5A). In contrast, DC
inoculated with infectious HSV-1 showed a clear reduction in their
allostimulatory capacity. This effect was particularly striking at a
T-cell-to-DC ratio of 
200:1. These data clearly show that HSV-1
infection interferes with DC function, where the downregulation of CD83 might play a role in this process.
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FIG. 5.
HSV-1 infection decreases the ability of mature DC to
induce an allogeneic T-cell response. (A) DC were inoculated with
either infectious virus or UV-inactivated virus or left untreated,
cocultured with allogeneic T cells for 4 days, and pulsed with
[3H]thymidine for 8 h, and the cell supernatants
were analyzed. DC derived from noninfected ( ) or UV-inactivated
HSV-1 ( ) induced a strong allostimulatory reaction in the primary
allogeneic MLR. In contrast, cells derived from HSV-1-infected DC ( )
showed a strongly reduced ability to induce T-cell proliferation,
particularly at a T-cell-to-DC ratio of 200:1 (MOI = 1). (B)
Increasing numbers of uninfected or HSV-1-infected DC were added to 100 HSV-1-infected DC and subsequently analyzed for their allostimulatory
capacity. Only when a large excess of infected DC were added (ratio of
infected to uninfected DC = 66:1) was an interference with the
allostimulation detectable.
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Next we determined if a mixture of HSV-1-infected and uninfected DC
would interfere with the alloresponse. Thus, increasing
numbers of
uninfected or HSV-1-infected DC (MOI of 1) were cocultured
with 100 HSV-1-infected DC and their T-cell-stimulatory capacity
was analyzed.
Only when a large excess of infected DC was added
(ratio of infected to
uninfected DC = 66:1) was an interference
with the allostimulation
detectable (Fig.
5B). This strongly suggests
that DC must be infected
to downregulate their allostimulatory
function.
HSV-1 induces downregulation of CD83.
Next we addressed
whether the CD83 downregulation was due to molecule uptake into the
cell or to protein degradation. To answer this question, cell extracts
from DC which were inoculated with replication-competent virus or with
UV-inactivated virus, treated with CHI, or left untreated were
separated by SDS-gel electrophoresis. Samples taken after 2 and 24 h were compared. Proteins were transferred to nitrocellulose and
analyzed by Western blotting. Strikingly, cell extracts derived from
HSV-1-infected DC showed an almost complete absence of CD83 after
24 h (Fig. 6A, lane 6). In contrast, no effects were observed in any of the other analyzed samples. As
expected, the CHI treatment induced a slight reduction of CD83 expression after 24 h (lane 4). The same blot was reprobed with an
anti-CD86 antibody, and, as shown in Fig. 6B, no major differences were
observed, indicating that the HSV-1-induced CD83 degradation is
specific and is not due to a general protein degradation phenomenon. Furthermore, 10 h after the HSV-1 infection, no CD83 protein was detectable (data not shown). Taken together, these results and the FACS
analyses clearly support the notion that CD83 degradation occurs during
the first 10 h of viral infection.
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FIG. 6.
HSV-1 infection induces CD83 protein degradation in
mature DC. (A) Total cellular protein extracts (2- and 24-h samples)
from DC which were inoculated either with infectious virus (MOI = 1) or with UV-inactivated virus, treated with CHI, or left untreated
were resolved using SDS-polyacrylamide gel electrophoresis, transferred
onto nitrocellulose membranes, and probed with a CD83-specific
monoclonal antibody. (B) The blot shown in panel A was reprobed using a
CD86-specific antibody.
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HSV-1 infection induces a redistribution of CD83 molecules in
mature DC.
To analyze the data described above, we performed
immunofluorescence studies on a single-cell basis. Mature DC were
either inoculated with virus or left untreated and analyzed after 3 and 23 h, respectively. After fixation, the cells were incubated with a monoclonal antibody specific for CD83 or for MHC class I (Fig. 7). When noninfected DC were analyzed,
there was no difference in the expression of CD83 after 3 and 23 h
(Fig. 7A and B). The same was true for the expression of MHC class I
molecules (Fig. 7E and F). In sharp contrast, a strong reduction of the
CD83 signal was observed in HSV-1-infected DC after 23 h (Fig.
7D). No effect of MHC class I expression was observed (Fig. 7G and H).
These data further support the notion that CD83 is specifically
degraded upon HSV-1 infection.
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FIG. 7.
HSV-1 infection strongly affects the expression of CD83
in mature DC. Mature DC were either inoculated for 1 h with HSV-1
(MOI = 1) or left untreated. After an incubation period of 3 h (A and C) or 23 h (B and D), the cells were fixed and analyzed
by indirect CD83 immunofluorescence. After 3 h, the infected DC
showed only a slight change in their CD83 surface expression (C), but a
dramatic reduction of the CD83 signal intensity was observed after
23 h (D). The same cell population was also analyzed using an MHC
class I-specific antibody (E to H). These analyses did not reveal major
differences between the different cell populations. Bar, 20 µm.
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HSV-1 leads to CD83 degradation in lysosomes.
To further
investigate the CD83 downregulation phenomenon, we used
HSV-1-permissive Vero cells. In contrast to the nonadherent DC, these
adherent Vero cells have the advantage that their intracellular morphology can be studied more easily using immunofluorescence microscopy. Therefore, Vero cell nuclei were microinjected with an
expression plasmid encoding CD83 and cultured for 20 h before being inoculated with HSV-1 at an MOI of 1. The cells were then analyzed by immunofluorescence using monoclonal antibodies specific for
CD83 (Fig. 8A and C) or the viral protein
VP16 (Fig. 8B and D). Noninfected, microinjected cells showed a high
expression and even distribution of CD83 (Fig. 8A). As expected, no
VP16 staining was observed in these noninfected cells (Fig. 8B). In sharp contrast, HSV-1-infected cells showed a dramatic reduction of
CD83 expression and a change in the cellular distribution of CD83 (Fig.
8C). Some fluorescent dot structures were visible in the cytoplasm,
possibly reflecting the loci of CD83 degradation. The VP16 staining,
demonstrating that the cells were indeed infected, is shown in Fig. 8D.
These experiments revealed that the HSV-1-induced downregulation of
CD83 in Vero cells was comparable to the downregulation observed in DC,
indicating that it is feasible to use Vero cells to study HSV-1-induced
effects.
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FIG. 8.
HSV-1 leads to CD83 degradation. The p3CD83 expression
plasmid DNA was microinjected into the nuclei of Vero cells, and the
cells were cultured for further 20 h. They were then either left
untreated or infected with HSV-1 (MOI = 1) prior to a further
incubation period of 7 h. The cells were subsequently analyzed by
double immunofluorescence microscopy using antibodies directed against
CD83 (A and C) and VP16 (B and D). (A) Noninfected Vero cells showed a
characteristic CD83 staining. (C) In contrast, HSV-1 infection affected
the normal CD83 distribution, leading to accumulated intracellular
structures. (D) Indirect VP16 immune fluorescence served as control for
the successful virus infection. More infected cells show less CD83
expression. Compare cells labeled # and * in panels C and D. Bar, 20 µm.
|
|
Next we wanted to determine the cellular compartment in which CD83
might be localized and subsequently degraded. Vero cells
were therefore
microinjected with the CD83-expressing plasmid,
infected with HSV-1,
and stained with anti-CD83 antibody (Fig.
9A) or with an antibody specific for the
lysosomal protein cathepsin
D (Fig.
9B). The colocalization of CD83 and
cathepsin D became
particularly obvious when the individual images were
merged (Fig.
9C). These data strongly suggest that after HSV-1
infection, CD83
is downregulated from the cell surface and localized to
intracellular
lysosomal organelles, where degradation takes place.
View larger version (14K):
[in this window]
[in a new window]
|
FIG. 9.
Subcellular localization of CD83. (A and B) To define
the cellular CD83 localization after HSV-1 infection, cells were
stained with antibodies against CD83 (A) and cathepsin D (B), which
showed a similar intracellular distribution. (C) Merging of the images
in panels A and B indicates that CD83 accumulates in lysosomal
compartments. Bar, 20 µm.
|
|
| |
DISCUSSION |
Viruses have developed a broad range of mechanisms to escape the
host immune response. These include reduced expression of critical
antigen epitopes, genetic variations of MHC epitopes, clonal
exhaustion, production of immunosuppressive cytokines and homologous
cytokine receptors, and downregulation of critical cytokines. From this
list, it has recently become evident that many of the viral escape
mechanisms specifically target DC function. Measles virus infection,
for example, causes apoptosis and syncytium formation in DC, inhibits
IL-12 production, and blocks the T-cell stimulatory capacity of DC
(10, 12, 28). These data may explain why measles virus
infections are often accompanied by a dramatic suppression of the
immune system.
Interestingly, vaccinia virus also inhibits DC maturation and leads to
impaired allostimulatory properties of the infected DC (9a).
Although vaccinia virus has developed so many immune escape mechanisms
(reviewed in reference 29), it induces a strong cellular and antibody-mediated immune response. However, the precise mechanism for the induction of this immune response still requires elucidation. A possible explanation may be provided by so-called cross-presentation. Here, apoptotic vaccinia virus-infected DC may be
taken up by uninfected bystander DC, and the bystander cells could then
present viral antigens to virus-specific cytotoxic T cells. This has
already been described for influenza virus-infected apoptotic cells
(2). These findings clearly demonstrate the dual role of DC
during viral infections. On the one hand, they are able to induce a
specific immune response, while on the other hand, they contribute to
pathogenic effects and immune suppression.
This dual role has also been reported for human immunodeficiency virus
type 1 infections. In this case, DC not only elicit virus-specific CTL
responses but also transport the virus from the periphery to the T
cells in the lymph nodes, where human immunodeficiency virus type 1 replicates (11). Furthermore, hepatitis C virus seems to
impair the allostimulatory capacity of DC, as assessed by examination
of peripheral blood DC isolated from hepatitis C virus-infected
patients (16). These reports all clearly demonstrate that DC
are very important targets for viruses in their struggle for survival
against the immune response.
Here we report that HSV-1-infected mature DC are inhibited in their
T-cell-stimulatory capacity. Furthermore, we show that mature DC do not
generate infectious virus particles. On the transcriptional level,
mainly immediate-early and early viral transcripts were detected.
Therefore, we speculate that proteins encoded by these gene products
might be responsible for the interference with DC function. However, we
have not yet identified the molecular mode of action of this
HSV-1-mediated inhibitory effect. In contrast to mature DC, immature DC
can also be infected efficiently at lower MOI, i.e., 0.01; strikingly,
these cells never reach the final state of maturation and are therefore
very poor T-cell stimulators (M. Kruse, unpublished results). In
support of these findings, a study published during the final
preparation of this paper reported that HSV-1 interferes with the
maturation of immature DC (27).
Interestingly, when we investigated the phenotype of mature DC infected
with HSV-1, we observed a very striking feature, namely, the loss of
CD83 cell surface expression. However, the expression of other typical
DC molecules, including CD25, CD40, CD80, CD86, CD95, and MHC class I
and class II, was not influenced. CD83 expression is induced during DC
maturation and represents one of the best-known markers for mature DC
(35, 39, 40). Although the function of CD83 is still
unknown, the fact that its surface expression is strongly upregulated
during DC maturation, along with costimulatory molecules, indicates
that it has an important function during the immune response.
Noteworthy is the finding that inhibition of CD83 cell surface
expression leads to a dramatic reduction of DC-mediated T-cell
stimulation (19). Using a specific inhibitor of the
eukaryotic initiation factor 5A, a protein which is involved in the
nuclear export pathway of specific RNAs (25, 26), we showed
that the CD83 cell surface expression on DC could be prevented and that
their allostimulatory capacity was strongly reduced (19).
A typical feature of HSV-1 infection is the loss of protein synthesis
due to disruption of polysomes and degradation of mRNA (31).
However, the loss of CD83 cell surface expression was shown not to be
due to CD83 mRNA degradation. The dramatic downregulation of the CD83
protein was already observed 10 h after infection. Nevertheless,
at this time point there was no difference between the uninfected and
infected samples when CD83 mRNA was analyzed. In contrast, HSV-1 led to
a complete and specific degradation of CD83 protein in mature DC. Since
we observed a colocalization of CD83 and cathepsin D, this degradation
is probably lysosome mediated. Since mainly immediate-early and early
viral gene transcripts are generated in mature DC, we speculate that
these viral gene products might be involved in the degradation of CD83.
A detailed analysis of this hypothesis will be performed in a future
study. Although the inhibitory effect of HSV-1 on mature DC function cannot be exclusively attributed to the degradation of CD83, this inhibition represents an additional new viral immune system escape mechanism for HSV-1.
Peripheral DC and Langerhans cells reside close to keratinocytes,
fibroblasts, and epithelial cells, the major cells in which HSV-1
replication occurs. Therefore, it is conceivable that DC and Langerhans
cells encounter HSV-1, either directly through infection or indirectly
via uptake of infected cells, which could lead to an antiviral immune
response in both cases. Our data suggest that HSV-1 interferes with the
function of mature DC by preventing the T-cell activation by these most
powerful antigen-presenting cells. Consequently, caution should be
applied to the use of HSV-1-derived vectors currently being developed
as tools for vaccination strategies to be implemented in human clinical trials.
| |
ACKNOWLEDGMENTS |
This work was supported by the Deutsche Forschungsgemeinschaft
(SFB 466) and the Wilhelm Sander-Stiftung (96.042.2).
We thank Heidi C. Joao for critical reading of the manuscript.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Department of
Dermatology, University of Erlangen-Nürnberg,
Hartmannstrasse 14, D-91052 Erlangen, Germany. Phone:
49-9131-853-3734. Fax: 49-9131-853-5799. E-mail:
kruse{at}derma.med.uni-erlangen.de.
| |
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Abstract
Introduction
