By
From the * Institut National de la Santé et de la Recherche Médicale U 404 "Immunité et
Vaccination," Lyon, France; Centre National de la Recherche Scientifique Unité de Recherche
Associée 602, Lyon, France; § Schering-Plough Laboratory for Immunological Research, Dardilly,
France; and
Baylor Institute of Immunology Research, Dallas, Texas 75246
Measles causes a profound immune suppression which is responsible for the high morbidity and mortality induced by secondary infections. Dendritic cells (DC) are professional antigen-presenting cells required for initiation of primary immune responses. To determine whether infection of DC by measles virus (MV) may play a role in virus-induced suppression of cell-mediated immunity, we examined the ability of CD1a+ DC derived from cord blood CD34+ progenitors and Langerhans cells isolated from human epidermis to support MV replication. Here we show that both cultured CD1a+ DC and epidermal Langerhans cells can be infected in vitro by both vaccine and wild type strains of MV. DC infection with MV resulted within 24-48 h in cell-cell fusion, cell surface expression of hemagglutinin, and virus budding associated with production of infectious virus. MV infection of DC completely abrogated the ability of the cells to stimulate the proliferation of naive allogeneic CD4+ T cell as early as day 2 of mixed leukocyte reaction (MLR) (i.e., on day 4 of DC infection). Mannose receptor-mediated endocytosis and viability studies indicated that the loss of DC stimulatory function could not be attributed to the death or apoptosis of DC. This total loss of DC stimulatory function required viral replication in the DC since ultraviolet (UV)-inactivated MV or UV-treated supernatant from MV-infected DC did not alter the allostimulatory capacity of DC. As few as 10 MV- infected DC could block the stimulatory function of 104 uninfected DC. More importantly, MV-infected DC, in which production of infectious virus was blocked by UV treatment or paraformaldehyde fixation, actively suppressed allogeneic MLR upon transfer to uninfected DC-T-cultures. Thus, the mechanisms which contribute to the loss of the allostimulatory function of DC include both virus release and active suppression mediated by MV-infected DC, independent of virus production. These data suggest that carriage of MV by DC may facilitate virus spreading to secondary lymphoid organs and that MV replication in DC may play a central role in the general immune suppression observed during measles.
Measles is a highly contagious disease characterized by
a prodomal illness followed by the appearance of a
generalized macropapular rash, which coincides with the
appearance of the immune response and initiation of virus
clearance. Measles causes a profound immune suppression
(1), which leads to an increased susceptibility to secondary
infections (2, 3), a major cause of children's death in developing countries. Immune suppression during measles starts
at the onset of the rash and may last for several weeks after
recovery. Interestingly, measles vaccination also causes immune suppression (4). In vivo, the loss of immune responsiveness to recall antigens is characterized by loss of tuberculin skin test reactions (5, 6). In vitro, antibody production,
T cell-mediated immune responses (7), as well as NK
cell activity (12) are markedly decreased. Possibly related to
this inhibition is the altered lymphokine production by
blood mononuclear cells from infected patients (13, 14).
Paradoxically, this immune suppression occurs in the context of substantial immune activation (15) which is associated with the induction of a measles virus (MV)1-specific immune response. Effective immune responses both
allow clearance of the virus as well as induction of long-term immunity.
MV is transmitted by aerosol, enters through the respiratory route, and starts replicating within tracheal and bronchial epithelia (18, 19). The virus is subsequently transported from the respiratory tract to the draining lymph
nodes, where virus amplification, during the prodomal
stage, gives rise to giant multinucleated lymphoid or reticuloendothelial cells (i.e., Warthin-Finkeldy cells). These
syncytia, identified in the submucosal areas of tonsils and
pharynx (20), are thought to be a major source of virus
spread to other organs and tissues through the blood stream.
Within the epithelial lining of the conductive airways,
dendritic cells (DC) with functional characteristics of epidermal Langerhans cells (LC) form a continuous network
(21, 22). DC of the respiratory tract, the most potent type
of APC for activation of naive and memory T cells (22),
traffic from the respiratory mucosa to the draining lymph
nodes upon an airway challenge with stimuli such as bacteria (23) or soluble antigen (24). Inasmuch as DC have been
shown to bind and allow replication of viruses such as influenza virus (25) and HIV (26), we postulated that DC
could also be infectable with MV. Here, in vitro infection of cord blood-derived CD1a+ DC with either vaccine or
wild-type strains of MV results within 24-48 h in virus
replication and syncytia formation associated with the release of infectious particles. In addition the MV-infected DC lose their ability to stimulate naive allogeneic CD4+ T
cells. This sensitivity to MV is not restricted to in vitro generated DC as skin-derived DC are a target for MV infection in vitro.
Virus.
The Edmonston (vaccine) and the Hallé (vaccine-like)
strains of MV (American Type Culture Collection, Rockville,
MD) were grown in the African green monkey Vero cell line at
33°C, and cell free supernatant with a virus titer of 107 and 3 × 107 PFU/ml, respectively, was used as virus stock. The wild-type MV strain (LYS-1) was isolated from PBMC taken from a patient with acute measles and passaged once on the permissive B95a
marmoset monkey cell line (gift from Dr. Kobune, National Institute of Health, Tokyo, Japan) (29). Cell free supernatants were
used as virus stock and had a titer of 3 × 105 PFU/ml.
Establishment of DC from CD34 Progenitors.
DC were generated by culturing cord blood CD34+ progenitors in the presence
of 100 ng/ml of rhGM-CSF (specific activity 2 × 106 U/mg;
Schering-Plough Research Institute, Kenilworth, NJ), 2.5 ng/ml
(50 U/ml) of rhTNF- Epidermal Cell Suspension.
Epidermal cells were isolated from
normal skin of patients undergoing plastic surgery, as previously
described (31). Fragments of total skin were incubated for 1-2 h
at 37°C in 0.05% of trypsin (Gibco) in Hanks' balanced medium
supplemented with antibiotics. The epidermis was separated from
the dermis, cut into 1-mm2 fragments from which single cells
were released by repeated pipetting. Epidermal cell suspensions
were enriched for LC by centrifugation for 20 min at 1,400 rpm
over a Lymphoprep gradient. The cells recovered from the interface (referred to as LC-enriched suspensions), containing ~10%
of LC as revealed by morphology and HLA-DR expression, were
used for MV infection experiments.
Infection of DC by MV.
DC obtained from 9-d cultures of cord
blood progenitors were infected for 1 h at 37°C with various MV
strains at a multiplicity of infection (MOI) of 0.05, in a minimum
volume of complete medium. Cells were then rinsed and cultured at 37°C in 6-well plates (Costar Corp., Cambridge, MA) at
a density of 106 cells/ml in 2 ml of complete medium supplemented with 100 ng/ml of rhGM-CSF with or without 2.5 ng/
ml of rhTNF. LC-enriched suspensions obtained from human
skin were infected with either Hallé, Edmonston, or LYS-1 strains
of MV at an MOI of 0.05, and cultured for 3 d at a density of 106
cells/ml in complete medium supplemented with GM-CSF.
May-Grunwald Giemsa Staining.
The morphological analysis
of DC syncytia was performed on the second day of infection.
DC were collected, cytocentrifuged onto glass slides, and stained
with May-Grunwald Giemsa according to routine protocols.
Electron Microscopy.
DC were fixed in the culture plates by incubation for 1 h at room temperature with 2% glutaraldehyde
(vol/vol). The cells were carefully harvested and pelleted by centrifugation for 5 min at 1,200 rpm. The cell pellets were then incubated successively for 30 min in sodium cacodylate 0.1 M, pH
7.4, and glutaraldehyde 2% (vol/vol), then for 30 min in sodium
cacodylate 0.2 M, pH 7.4, and finally for 30 min in sodium cacodylate 0.15 M, pH 7.4, containing 1% OsO4. Dehydration was
then performed by serial 5-min incubations in 30, 50, 70, and
95% ethanol. Finally, cell pellets were included in Epon by impregnation in a mixture containing Epon A (30%) + B (70%) + DMP30 (1.7%) and allowed to polymerize for 48 h at 60°C. Sections (60-80 nm) were made on a microtome (Ultracut-Reichert, Vienna, Austria). Uranyl acetate and Pb citrate were used for contrast. Observation was performed with a Philips EM 300 microscope.
Titration of MV Produced by Infected DC.
DC were either mock-infected or infected for 1 h at 37°C with either Hallé or LYS-1
MV strains at an MOI of 0.05. The cells were washed three
times, resuspended in complete medium supplemented with 100 ng/ml of GM-CSF with or without TNF- Immunostaining of Cell Smears.
Cells were cytocentrifuged for
5 min at 500 rpm onto glass slides and fixed in cold acetone for
10 min. Slides were washed in PBS supplemented with 5% human serum and incubated for 30 min with a polyclonal rabbit
anti-mouse antibody specific for S100 (dilution 1:400) (Dako,
Trappes, France) or normal rabbit serum as control. Specific
staining was revealed using a biotinylated F(ab FACS® Analysis.
Indirect immunofluorescence analysis was
performed according to standard techniques. Briefly, cells were
first incubated for 20 min on ice with 5% normal human AB+ serum to saturate Fc receptors, then stained with the mouse IgG2b anti-MV-hemagglutnin (HA) mAb 55 (32) or an isotype control Ig. Specific binding was revealed by a FITC-conjugated goat
F(ab Internalization of Dextran-FITC.
The ability of DC to internalize dextran-FITC through mannose receptor was tested as previously described (34). DC were harvested using warm PBS containing 0.5 mM EDTA, washed once, and resuspended at 106
cells/ml in complete medium supplemented with Hepes. 200 µl
of cell suspension was preincubated for 1 min at 37°C (or at 0°C for controls) before addition of 20 µl of a 1 mg/ml FITC-dextran solution (Molecular Probes, Eugene, OR). At either 15 or 30 min of incubation, the reaction was stopped by addition of 4 ml
of cold PBS supplemented with 1% FCS, 0.01% NaN3, and cells
were washed four times at 4°C in the same buffer. PI was added
to cells to exclude dead cells, and the uptake of FITC-dextran by
viable cells was analyzed using a FACSscan®.
Purification of Naive CD4+ T Cells from Human Peripheral
Blood.
Mononuclear cells were isolated from adult peripheral
blood by Ficoll Paque gradient centrifugation. Naive CD4+ T
cells were purified as previously described (35) by immunomagnetic depletion using a mixture of mAbs including IOM2
(CD14), ION16 (CD16), B8 12.2 (HLA-DR) (Immunotech,
Marseille, France), OKT8 (CD8) (Ortho), UCHL1 (CD45RO)
(Dako), NKH1 (CD56) (Coulter Corp., Miami, FL), 4G7
(CD19), and mAb 89 (CD40) (36). After two rounds of bead depletion, purity of CD45RA+CD4+ T cells was routinely >95%.
The T cells were either used immediately or frozen at 5 × 106
cells/vial in PBS, 90% FCS, and 10% DMSO until use.
Allogeneic Mixed Lymphocyte Reaction.
DC harvested on day 2 of MV infection (unless otherwise stated) were used as stimulatory cells in allogeneic MLR with naive CD45RA+ CD4+ T
cells. Various numbers of DC were cultured in complete medium with 2 × 104 naive allogeneic CD4+ T cells in triplicate in
round-bottomed 96-well plates (Falcon, Pont de Claix, France).
In some experiments, limiting dilutions of MV-infected DC were
added to cocultures of uninfected autologous DC (104 cells) and
allogeneic CD4+ CD45RA+ T cells (2 × 104 cells). MV-infected
and control uninfected DC were either untreated, UV-irradiated
(0.25 J/cm2) to inactivate the virus, or fixed by a 30-min incubation at 4°C with freshly prepared 1% paraformaldehyde (PF),
washed twice in PBS, quenched by a 30-min incubation with 0.1 M
L-lysine, pH 8, and washed four times before use. In some experiments, supernatants of MV-infected cells either UV-irradiated
(0.25 J/cm2) or left untreated were added to uninfected DC-T
cell cultures. Except for kinetic experiments, cultures lasted for 6 d
and cells were pulsed with 1 µCi of [3H]thymidine for the last 16 h
of culture, harvested, and counted. The results are expressed as
cpm ± SD of quadruplicate wells.
MV Infects DC and Induces Syncytia Formation. Cultured
DC, harvested on day 9 of culture of CD34+ progenitors
with GM-CSF-TNF-
We subsequently examined whether human epidermal LC were susceptible to
MV infection. To this end, LC-enriched and keratinocyte suspensions were cultured for 24 h with GM-CSF and infected with the vaccine or wild-type strains of MV. 3 d after infection with each MV strain, cell fusion and syncytia
formation was observed in LC-enriched epidermal cells,
but not in keratinocyte-enriched suspensions (data not
shown). FACS® showed that LC-enriched suspensions
(which contained 24% of DR+ LC cells) yielded ~25% of
HA+ cells, while no HA+ cells could be found in keratinocyte-enriched suspension (not shown). Furthermore, all
syncytia were stained specifically with the anti-S100 antibody, demonstrating that they were LC-derived. (Data obtained with Edmonston are shown in Fig. 1, g and h).
These data indicate that DC from epithelial tissues such as
the skin are susceptible to MV infection in vitro.
MV replication in host cells results in expression at the surface of the infected cells of MV glycoproteins, including
the HA and the fusion protein, which are both required for
fusion and syncytia formation (37). To determine the percentage of in vitro generated DC infected with MV, cell
surface expression of HA was determined by flow cytometry analysis of immunofluorescence staining with the HA-specific antibody, mAb 55. Since DC syncytia have a large
size and a poor viability, only MV-infected DC present as
single cells could be analyzed with this method. Cell surface expression of HA on viable DC was maximal on day 2 of infection with all strains of virus tested. HA expression
was reproducibly detected on 70-100% of DC infected
with MV-Hallé (Fig. 2 b) and on 20-40% of DC infected
with MV-LYS-1 (Fig. 2 c) or Edmonston (not shown).
Double immunofluorescence analysis revealed that >90%
of HA+ cells are CD1a-positive after infection with Hallé
(Fig. 2 d) and >75% are after LYS-1 (Fig. 2 e). Thus, all
three strains of MV can infect an important proportion of
DC and induce some of these to form syncytia.
Transmission electron microscopic analysis of DC at
day 2 of infection with either Hallé or LYS-1 confirmed
the presence of syncytia containing several nuclei with a
polylobular shape characteristic of a DC nucleus (Fig. 3 a).
Typical paramyxovirus nucleocapsid structures were observed in the cytoplasm (Fig. 3 b) and occasionally in the
nuclei of DC syncytia. Electron-dense structures reminiscent of viral glycoproteins could be identified at sites of
membrane rufflings of the DC syncytia showing virus budding with release of virions (Fig. 3 c). Images of DC apoptosis (Fig. 3 d) and cell clasmatosis resulting from detachment of syncytia fragments (Fig. 3 e) could be also observed
with each MV strain. These data demonstrate that infection
of DC with wild-type and vaccine strains of MV is associated with a complete replication cycle ultimately leading to
DC lysis.
We next examined whether infection of DC with MV
yields infectious virions. At various days after infection
with either the Hallé or LYS-1 MV strains at an MOI of
0.05, the titer of infectious virus released into the supernatant was determined by a plaque assay, using as indicator
cells the B95a marmoset B cell line permissive for each vaccine and wild-type strain of MV. As shown in Fig. 4, DC
could efficiently replicate MV and yield infectious particles
with a titer of ~104 PFU/ml on day 3 of infection, irrespective of the strain.
Since DC have a
unique capacity to activate naive T cells, we asked whether
MV infection would affect their ability to activate naive allogeneic CD4+ T cells. Various numbers of DC obtained
on day 2 after infection with either MV-Hallé or MV-
LYS-1 as well as day 2 mock-infected DC were added to
naive CD45RA+CD4+ T cells and T cell proliferation was
analyzed 6 d later. As few as 30 mock-infected DC were
able to stimulate allogeneic T cell proliferation; in contrast
up to 104 DC infected with either Hallé, LYS-1 (Fig. 5 A),
or the Edmonston strains (not shown) were unable to induce T cell proliferation. Kinetic studies performed between days 2 and 6 showed that MV-infected DC were
unable to induce the proliferation of allogeneic T cells, at
all times tested (Fig. 5 B). Note that T cell viability was not
affected. A 1-h contact between DC and MV (at MOI
ranging from 0.1 to 0.001) was sufficient to fully abrogate
the allostimulatory capacity of DC. The inhibition of allogeneic MLR by DC was observed only when DC were infected with live MV but not when DC were pulsed with
UV-inactivated MV, irrespective of the MV strain (Fig. 5 C). These data showed that MV replication in DC blocks
their ability to support the proliferation of allogeneic CD4+
T cells.
The percentage of MV-infected
cells which remained viable (as determined by Trypan blue
dye exclusion) after MV infection, was 70% on day 2 and
30% on day 4 after infection, as compared to 90% on day 2 and 60% on day 4 for uninfected DC.
We first examined whether MV infection of DC affected
mannose-receptor-mediated endocytosis, which is involved
in endocytosis of glycosylated antigens and represents one
pathway of antigen uptake by DC (34). Dextran-FITC internalization by MV-infected DC was examined on day 2 of infection. The percentage of single cells able to internalize dextran-FITC and the intensity of fluorescence was
comparable for mock-infected or MV-infected DC (Fig. 6
A), indicating that the uptake of antigen through the mannose receptor was not affected by MV-infection of DC.
Kinetic studies of DC viability were next performed on
days 2, 3, and 4 after infection to determine whether inability to support T cell proliferation was due to DC death
or apoptosis induced by MV infection. Double stainings of
DC cultures for Annexin V and PI were performed on days
2, 3, and 4 after infection in order to evaluate the percentage of intact viable cells (Annexin V We next examined whether inhibition of the allostimulatory capacity of DC induced by MV-infected DC was due to the release of infectious virus
which inactivated the uninfected DC or T cells. As shown
in Fig. 7 A, supernatants of day 2 MV-infected DC (with a
virus titer of 104 PFU/ml) were able to induce inhibition
of CD4+ T cell proliferation in response to allogeneic DC.
In contrast, UV-irradiated supernatant of MV-infected DC
(containing <10 PFU/ml) had no inhibitory effect (Fig. 7
B) indicating that spreading of infectious virions produced
by MV-infected DC to uninfected DC or T cells could
contribute to the inhibition of T cell proliferation.
The efficiency of MV-infected DC to transfer inhibition
of allogeneic MLR was further analyzed by testing the ability of limiting numbers of MV-infected DC to alter the allostimulatory function of uninfected DC. As shown in Fig.
7 C, addition of as few as 10 Hallé-infected DC to uninfected DC-T cell cultures was sufficient to abrogate the
proliferation of allogeneic T cells. Thus, MV-infected DC
could block the allostimulatory function of uninfected
DC, even when present at low numbers (i.e., 0.1% of total
DC) at least partly through the release of infectious virus.
We then examined whether MV infection of DC could turn on an active suppressor mechanism independently of infectious virus production. Day 2 MV-infected or uninfected DC
were either UV-treated or fixed with PF to inactivate the
virus, before addition in graded numbers to uninfected
DC-T cell cultures. Inactivation of MV in UV- and PF-treated DC was checked after culture for 1 h at 37°C by
the plaque assay performed on whole cultures (DC and supernatant). Both UV-treated and PF-fixed MV-infected
DC contained 0 PFU/ml, showing that both treatments
completely blocked virus replication within the DC as well
as release of infectious virions. As shown in Fig. 8, addition
of up to 104 uninfected DC, either UV-irradiated or PF-fixed, did not affect the proliferation of CD4+ T cells in response to 104 untreated uninfected DC. In contrast, addition of 103 or 104 UV-treated MV-infected DC completely
blocked the allostimulatory effect of uninfected DC (Fig. 8
A). Similar effect was observed with both Hallé-infected or
LYS-1-infected DC. Likewise, PF-fixed MV-infected DC
actively suppressed allogeneic DC-T cell MLR, although complete inhibition of T cell proliferation required addition of 104 PF-fixed MV-infected DC (Fig. 8 B). These
data show that MV infected DC can actively suppress the
allostimulatory function of uninfected DC, independent of
the release of infectious virus.
This study has demonstrated that MV can infect DC isolated from the skin or generated in vitro by culturing hematopoietic progenitors in the presence of GM-CSF and
TNF- An important observation from this study is the demonstration that MV-infected DC can no longer act as stimulatory cells in allogeneic MLR. It is unlikely that syncytia
formation was responsible for the loss of DC stimulatory
function because most infected DC were present as isolated
cells at the time of addition to T cells (and up to day 4 after
MV infection). Moreover, the observation that MV-infected
DC can still function for receptor-mediated endocytosis and that at day 4 after infection intact viable cells still represent 65% of those present in uninfected cultures further
support that the complete inhibition of allogeneic MLR is
not merely due to the death or apoptosis of the infected DC.
Two major mechanisms contributing to the loss of DC
function have been identified in these studies: (a) an efficient transmission of infectious virus by infected DC inactivating uninfected DC or T cells and (b) an active suppression mediated by infected DC, independent of infectious
virus transfer from the DC to uninfected cells. That the release of infectious virions contributes to suppression of T
cell proliferation is supported by the observation that T cell
proliferation in response to allogeneic DC is blocked by addition of supernatant from MV-infected DC containing
infectious virus, but is not affected if these supernatants are
UV inactivated. Noteably, the lack of T cell proliferation is
not due to their death, as 10 and 70% of CD3+CD1a This study provides the first demonstration that both
wild-type and vaccine strains of MV can infect and replicate in DC and suggests that DC may play a central role in
suppression of cell-mediated immune responses during
measles. The relevance of our data with respect to the immune suppression induced by MV infection in vivo remains to be established. As shown in this study, MV can
replicate in CD1a+ DC, which are comparable to the mature interdigitating DC from the T cell areas of secondary
lymphoid organs, as well as in immature epithelial CD1a+
LC. Syncytia generated by MV infection of skin LC and of
in vitro generated DC are likely to represent the in vitro
equivalent of the multinucleated giant cells infiltrating the
nasopharyngeal epithelium and the subepithelial layer on
tonsils (where DC can be found), detected during the prodomal stage of measles (20). In keeping with this, the DC
syncytia located beneath the epithelium of the adenoid of
HIV seropositive asymptomatic patients show active virus
replication (46). As epithelial DC of mucosal tissues are capable of the uptake and transport of infectious microorganisms to draining lymph nodes (22), our finding raises
the possibility that natural MV infection, which follows exposure of mucosal membranes of the respiratory tract to the
virus, is initiated by the uptake and replication of MV
within epithelial LC and DC.
Thus, epithelial DC may represent the initial target of
MV, and serve both as a reservoir for MV infection and as a
vehicle to carry the virus to lymphoid cells in draining
lymph nodes. Such a mechanism may explain the dissemination of infected cells and the general immune suppression
observed in measles. We propose that DC infection by MV
may play a pivotal role in the induction of a primary immune response against MV and in the general immune suppression observed during measles.
(specific activity 2 × 107 U/ml, Genzyme
Corp., Boston, MA), and 25 ng/ml of stem cell factor in RPMI
1640 medium (Gibco, Lyon, France) supplemented with 10% (vol/vol) FCS (Eurobio, Les Ulis, France), 10 mM Hepes, 2 mM
L-glutamine, 5 ×10
5 M 2-mercaptoethanol, 100 U/ml penicillin,
and 100 mg/ml streptomycin (referred to as complete medium), as
previously described (30). The cells, routinely collected at day 9, are composed of 60-90% of CD1a+ DC expressing MHC class-I,
HLA-DR, DP, DQ, and CD46 molecules These cultured CD1a+
cells will be referred to as DC throughout the text.
, and seeded at 2 × 105 cells/well in triplicate wells of round-bottomed microplates. At various time points (day 0, i.e., 2 h, days 1, 2, 3, 5, and 6) after
MV infection, virus production in cell free supernatants was assessed by titration on the adherent B95a marmoset cell line, permissive for both Hallé and LYS-1 strains of MV. The numbers of
plaques were counted at various dilutions of supernatants and the
results were expressed as the number of PFU/ml.
) goat anti-rabbit
IgG antibody (Vector, Biosys, Compiègne, France) and the
streptavidin peroxidase ABC kit (Dako). The enzyme activity was
developed by AEC substrate. The slides were washed and counterstained with hematoxylin.
)2 anti-mouse IgG (H + L) antibody (Zymed, Tebu,
France). Propidium iodine was added before FACS® analysis to
gate out dead cells. Double color immunofluorescence was carried out by sequential incubations of cells with human AB+ serum, MV-HA mAb 55 or an IgG2b control antibody, FITC-conjugated goat F(ab
)2 anti-mouse IgG (H + L), 5% normal
mouse serum, and finally PE-conjugated OKT6 (anti-CD1a)
(Ortho Diagnostic Systems, Raritan, NJ) or PE-conjugated Leu-M3 (anti-CD14) (Becton Dickinson, Inc., Rutherford, NJ) mAbs
or PE-conjugated isotype controls. Double staining for Annexin
V-FITC binding and for cellular DNA using propidium iodide
(PI) was performed for analysis of cell viability, as described (33).
Briefly, the cells (2 × 105) were washed with PBS and resuspended in 200 µl of Annexin V-binding buffer (10 mM Hepes/
NaOH, pH 7.4, 150 mM NaCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM KCl) supplemented with 4 µg/ml of Annexin V-FITC (Bender MedSystems, Vienna, Austria). After 10 min of incubation in the dark, the cells were washed once before addition of 1 µg of PI/ml of cell suspension and incubated for 10 min in the
dark. Single staining using Annexin V-FITC or PI alone were
performed as controls. Flow cytometry analysis was performed
using a FACSstar® plus cytofluorimeter (Becton Dickinson)
equipped with Lysis II software.
, were infected with various MV strains, including the vaccine strains Edmonston (not shown)
and Hallé, and the wild-type strain LYS-1, at an MOI of
0.05. With all virus strains, DC infection resulted in cell fusion and formation of syncytia (Fig. 1, a-c) consisting of giant multinucleated cells (Fig. 1, d-f). Syncytia formation
was constantly observed on day 2 of infection but could
also be detected in some experiments within 24 h of infection. Note that only a fraction of DC formed syncytia, as
~70% of viable DC were present as single cells after 2 d of
infection.
Fig. 1.
MV infection induces syncytia in cultured DC and skin LC. DC derived from day 9 cultures of cord blood CD34+ progenitors were either
mock-infected (a and d ) or infected with either MV-Hallé (b and e) or MV-LYS-1 (c and f ) at an MOI of 0.05. DC syncytia were observed on day 2 of
infection by phase-contrast microscopy analysis (final magnification, ×400) (a-c) and by May-Grunwald Giemsa staining (final magnification ×400 (d );
×1,000 (e and f )). LC-enriched epidermal cell suspensions either uninfected (g) or infected with the Edmonston strain at an MOI of 0.05 (h) were cultured for 3 d in the presence of 100 ng/ml of rGM-CSF. On day 3 of infection, the cells were processed for staining with a polyclonal rabbit antiserum
directed against S100 and counterstained with hematoxylin. No staining was detected with control normal rabbit serum (not shown). (g) Specific staining of LC for S100 in uninfected LC-enriched suspension; (h) S100 expression by LC syncytia in MV-infected LC-enriched suspension. ×400. The results
are representative of five experiments.
[View Larger Version of this Image (108K GIF file)]
Fig. 2.
FACS® analysis of HA expression by CD1a+ DC. DC were
either mock-infected (a) or infected with MV-Hallé (b) or MV-LYS-1
(c). On day 2 of infection, cell surface HA expression by single MV-infected DC was analyzed by indirect immunofluorescence using the MV-HA specific antibody, mAb 55, and an FITC-conjugated F(ab)2 fragment of
goat anti-mouse IgG (shaded histogram). A mouse IgG2b was used as isotype control (white histogram). In double immunofluorescence labeling,
HA+ cells were electronically gated and the percentages of HA+ cells expressing CD1a or CD14 in both Hallé-infected (d) and LYS-1-infected
(e) DC were determined using PE-conjugated anti-CD1a or anti-CD14
mAbs. The dotted lines represent the position of the PE-conjugated control IgG. The data are from a representative experiment out of 10.
[View Larger Version of this Image (28K GIF file)]
Fig. 3.
Electron microscopy analysis of MV replication in DC. Transmission electron microscopic analysis performed on day 2 after infection with
LYS-1, shows the complete replication cycle of MV in DC. (a) Syncytia containing 10 DC nuclei and cytoplasmic structures typical of the viral nucleocapsid (arrow) (×7,600); (b) A higher magnification of the viral nucleocapsid close to a nucleus (×28,000); (c) Viral glycoproteins identified at sites of
membrane rufflings (×34,000); (d) Virus budding leading to the release of intact virions (×46,000); (e) Fragments of DC syncytia resulting from cell clasmatosis, and containing viral nucleocapsids in the cytoplasm (×28,000); and (f) Apoptosis of DC syncytia with condensed chromatin in the nuclei
(×8,600).
[View Larger Version of this Image (163K GIF file)]
Fig. 4.
Productivity of MV
infection of DC. Day 9 DC were
infected with either MV-Hallé
() or MV-LYS-1 (
) at an
MOI of 0.05 and seeded at 2 × 105 cells/well. The supernatants
were harvested at the indicated time of culture and titrated for
the presence of infectious virions in a standard plaque assay using
the permissive cell line B95a. The
results are expressed as PFU/ml.
[View Larger Version of this Image (14K GIF file)]
Fig. 5.
MV infection of
DC abrogates their ability to
stimulate naive CD4+ T cell
proliferation allogeneic MLR.
Naive CD45 RA+ CD4+ T cells
(2 × 104 cells) were cultured for
6 d in triplicate wells with either (A) various numbers or (B) a fixed
number (104) of either mock-
infected () or day 2-Hallé-
infected (
) or LYS-1-infected
(
) allogeneic DC. T cell proliferation was measured either on
day 6 (A) or at various times (B)
of culture by [3H]thymidine uptake during the last 16 h of culture. (C) 2 × 104 naive T cells
were cultured for 6 d with 104
DC which had been either infected with MV or pulsed with
UV-irradiated MV. Thymidine
incorporation was determined
using a
counter. The results,
expressed as mean cpm ± SD,
are from a representative experiment out of three to five.
[View Larger Version of this Image (18K GIF file)]
Fig. 6.
Flow cytometry analysis of dextran-FITC internalization and
Annexin V FITC and PI double staining of DC after MV infection. DC
were either mock-infected (DC/0) or infected with MV (DC/Hallé; DC-LYS-1) at an MOI of 0.05. (A) Dextran-FITC internalization by mock-infected DC (a), DC infected with MV Hallé (b) and DC infected with
MV LYS-1 (c) on day 2 after infection. The cells (2 × 105) were incubated for 15 min in the presence of dextran-FITC either at 37°C (white
histogram) or at 0°C for control (shaded histogram). Dextran-FITC uptake
by viable cells was analyzed by flow cytometry, after exclusion of dead
cells using PI. The numbers in parenthesis represent the percentage of
cells which have internalized dextran-FITC. (B and C) On days 2, 3, and
4 after infection, cell viability was determined by flow cytometry analysis
of double staining with Annexin V-FITC and PI. (B) Dot plot representation of Annexin V-FITC and PI double stainings. The lower left quadrants of each panel show the viable DC (Annexin V PI
). The upper
right quadrants contain the nonviable necrotic DC (Annexin V+ PI+).
The lower right quadrants represent the apoptotic DC (Annexin V+ PI
).
(C) Histogram representation of the percentages of intact viable, apoptotic, and necrotic DC among total DC present at various time of culture
of either uninfected (
), Hallé-infected (
), or LYS-1-infected (
) DC.
[View Larger Version of this Image (36K GIF file)]
PI
), early apoptotic
cells (Annexin V+ and PI
), and necrotic cells (Annexin
V+ PI+ cells) as previously described (33). As shown in Fig.
6, B and C, the percentage of intact viable DC decreased
from ~70% on day 2 to 50% on day 4 after MV infection,
as compared to 80% on day 2 and 75% on day 4 in parallel
uninfected DC cultures. The percentage of apoptotic cells
in MV-infected DC cultures increased from 10% on day 2 to 35% on day 4 as compared to 10% in uninfected DC
cultures. The percentage of necrotic cells was <20% in
both uninfected and MV-infected DC cultures until day 4. Thus, MV caused 25% of DC to undergo apoptosis after
4 d of infection but the percentage of viable cells was still
65% compared to that of uninfected cultures. These data
show that the loss of the allostimulatory function of DC,
observed as early as day 2 of MLR (i.e., corresponding to
day 4 of DC infection) could not be attributed to the death or apoptosis of the DC.
Fig. 7.
Inhibitory effect of
MV-infected DC and of DC supernatants in allogeneic MLR.
(A and B) Supernatants from either
mock-infected (), day 2 Hallé-
infected (
), or LYS-1-infected
(
) DC were either untreated
(A) or UV-irradiated (B) and added
to cocultures of various numbers
of uninfected DC and 2 × 104
naive allogeneic CD4+ T cells.
(C) Various numbers of either
uninfected (
), Hallé-infected (
), or LYS-1-infected (
) DC
were cocultivated for 6 d with
104 uninfected DC and 2 × 104
allogeneic CD4+ T cells. T cell
proliferation was analyzed on day
6 of culture by thymidine uptake
over the last 16 h of culture. The
results are expressed as mean
cpm ± SD of triplicate wells.
[View Larger Version of this Image (18K GIF file)]
Fig. 8.
UV-treated or PF-fixed MV-infected DC can inhibit T cell
proliferation in allogeneic DC-T cell MLR. Uninfected () or day 2 LYS-1-infected (
) DC were either UV-treated (A) or PF-fixed (B) before addition in graded numbers to cultures containing 104 DC and 2 × 104 CD45RA+CD4+T cells. T cell proliferation was analyzed on day 6 of
culture by [3H]thymidine uptake for the last 16 h of culture. The results, expressed as mean cpm ± SD of triplicate wells, are representative of one
experiment out of three.
[View Larger Version of this Image (14K GIF file)]
. Both vaccine and wild-type strains of MV undergo a complete replication cycle in DC, as demonstrated
by the presence of the viral nucleocapsid in the cytoplasm,
HA expression at the cell surface, cell fusion leading to syncytia, and virus budding releasing infectious virions. DC
differ from lymphocytes and monocytes for replication of
MV (38) inasmuch as they do not require activation.
While in vitro generated DC represent mature DC, as determined by high levels of CD80, CD83, and CD86, skin
LC clearly represent immature resting DC. Yet, it is possible that the DC growth and survival factor, GM-CSF (42),
may be provided by keratinocytes during the culture, thus
possibly providing the necessary activation allowing MV
replication in resting DC. Further T cell signals, provided
during allogeneic DC-T cell coculture, did not result in
higher virus replication (not shown).
T cells
express HA on day 4 of cocultivation with LYS-1-infected or Hallé-infected DC, respectively (not shown). The observation that T cell unresponsiveness was achieved even
when MV-infected DC were present in limiting numbers
(i.e., as low as 0.1% of the total DC) in the allogeneic
DC-T cell cultures suggests that an efficient replication of
MV in DC is sufficient for virus transmission to T cells
and/or uninfected DC. Alternatively, MV-infected DC exert an active inhibitory effect on allogeneic MLR induced by uninfected DC, inasmuch as they can transfer inhibition even when production and release of infectious
virions has been blocked by either UV irradiation or PF
fixation before addition to uninfected DC-T cell culture.
Thus, active suppression by MV-infected DC can occur independently of the release of infectious virus, possibly through cell-cell interaction between infected DC and uninfected DC or T cells. In this context, MV glycoproteins
expressed on a limited number of infected PBMC have
been shown to transduce a negative signal blocking the
proliferation of uninfected cells in response to various stimuli (43). More recent studies reported that MV-infected
PBMC which were UV-inactivated could block allogeneic MLR and demonstrated that both HA and the fusion protein, F, expressed on the surface of MV were required for
suppression (44). Thus, the mechanism of immune suppression induced by MV infection of DC appears to be
complex and multifactorial, and most likely includes virus
spreading to T cells as well active suppression mediated by
infected DC. In addition, alterations of the DC function
may also contribute to the loss of DC function. Although
we found no decrease in MHC class I or class II molecules
or in the costimulatory molecules B7.1, B7.2, or CD40
(data not shown) which could explain the loss of immunostimulatory function of DC, it remains possible that MV infection of DC can alter the antigen-presenting function of
these cells through modification of their cytokine profile. In this respect, downregulation of IL-12 production has
been reported after MV infection of monocytes in vitro
(45). Experiments are in progress to determine the mechanism of the active suppression mediated by MV-infected DC.
Address correspondence to Dominique Kaiserlian, INSERM U404 Immunité et Vaccination, Ex-Batiment Institut Pasteur, Avenue Tony Garnier, 69365 LYON CX 07, France. Phone: 33-4-72-72-25-56; FAX: 33-4-72-72-25-67; E-mail: kaiserlian{at}lyon151.inserm.fr
Received for publication 10 February 1997 and in revised form 7 July 1997.
1 Abbreviations used in this paper: DC, dendritic cells; HA, hemagglutinin; LC, Langerhans cells; LYS-1, wild-type MV strain; MOI, multiplicity of infection; MV, measles virus; PF, paraformaldehyde; PI, propidium iodide.We are grateful to doctors and colleagues from clinics and hospitals in Lyon who provided us with umbilical cord blood samples and human skin from plastic surgery. We thank Dr. Simone Peyrol for her precious help in electron microscopy studies. We also thank Dr. Bernard Verrier (UMR103 CNRS -Biomerieux) for his gift of blood sample from a measles patient.
1. |
Oldstone, M.B.A..
1996.
Virus-lymphoid cell interactions.
Proc. Natl. Acad. Sci. USA.
93:
12756-12758
|
2. | Beckford, A.P., R.O.C. Kaschula, and C. Stephen. 1985. Factors associated with fatal cases of measles. A retrospective autopsy study. S. Afr. Med. J. 68: 858-863 [Medline]. |
3. | Morley, D.. 1969. Severe measles in the tropics. Br. Med. J. 1: 297-300 [Medline]. |
4. | Starr, S., and S. Berkovitch. 1964. Effect of measles, gamma-globulin-modified measles and vaccine measles on the tuberculin test. N. Engl. J. Med. 270: 386-391 . |
5. | von Pirquet, C.P.. 1908. Das Verhalten der kutanen tuberkulin Reaktion Wahrend der Masern. Dtsch. Med. Wochenschr. 34: 1297-1300 . |
6. | Tamashiro, V.G., H.H. Perez, and D.E. Griffin. 1987. Prospective study of the magnitude and duration of changes in tuberculin reactivity during complicated and uncomplicated measles. Pediatr. Infect. Dis. J. 6: 451-454 [Medline]. |
7. | Whittle, H.C., A. Bradley-Moore, A. Fleming, and B.M. Greenwood. 1973. Effects of measles on the immune response of Nigerian children. Arch. Dis. Child. 48: 753-755 [Medline]. |
8. | Coovadia, H.M., M.A. Parent, W.E. Loening, A. Wesley, B. Burgess, F. Hallett, P. Brain, J. Grace, J. Nardoo, P.M. Smythe, and G.H. Vos. 1974. An evaluation of factors associated with the depression of immunity in malnutrition and in measles. Am. J. Clin. Nutr. 27: 665-669 [Medline]. |
9. | Whittle, H.C., J. Dossetor, A. Oduloju, A.D.M. Bryceson, and B.M. Greenwood. 1978. Cell-mediated immunity during natural measles infection. J. Clin. Invest. 62: 678-684 [Medline]. |
10. | Arneborn, P., and G. Biberfeld. 1983. T lymphocyte subpopulations in relation to immunosuppression in measles and varicella. Infect. Immun. 39: 29-37 [Medline]. |
11. | Hirsch, R.L., D.E. Griffin, R.T. Johnson, S.J. Cooper, I. Lindo de Soriano, S. Roedenbeck, and A. Vaisberg. 1984. Cellular immune responses during complicated and uncomplicated measles virus infections of man. Clin. Immmunol. Immunopathol. 31: 1-12 [Medline]. |
12. | Griffin, D.E., B.J. Ward, E. Jauregui, R.T. Johnson, and A. Vaisberg. 1990. Natural killer cell activity during measles. Clin. Exp. Immunol. 81: 218-224 [Medline]. |
13. | Crespi, M., J.K. Struthers, A.N. Smith, and S.F. Lyons. 1988. Interferon status after measles virus infection. S. Afr. Med. J. 73: 711-712 [Medline]. |
14. | Ward, B.J., R.T. Johnson, A. Vaisberg, E. Jauregui, and D.E. Griffin. 1991. Cytokine production in vitro and the lymphoproliferative defect of natural measles virus infection. Clin. Immunol. Immunopathol. 61: 236-248 [Medline]. |
15. | Griffin, D.E., B.J. Ward, E. Jauregui, R.J. Johnson, and A. Vaisberg. 1989. Immune activation during measles. N. Engl. J. Med. 320: 1667-1672 [Abstract]. |
16. | Griffin, D.E., and B.J. Ward. 1993. Differential CD4 T cell activation in measles. J. Infect. Dis. 168: 275-281 [Medline]. |
17. | Griffin, D.E., B.J. Ward, E. Jauregui, R.J. Johnson, and A. Vaisberg. 1990. Immune activation during measles: interferon-gamma and neopterin in plasma and cerebrospinal fluid in complicated and uncomplicated disease. J. Infect. Dis. 161: 449-453 [Medline]. |
18. | Sakaguchi, M., Y. Yoshikawa, K. Yamanouchi, T. Sata, K. Nagashima, and K. Tadeka. 1986. Growth of measles virus in epithelial cells and lymphoid tissues of cynomolgus monkeys. Microbiol. Immunol. 30: 1067-1073 [Medline]. |
19. | Black, F.L., and S.R. Sheridan. 1960. Studies on attenuated measles-virus vaccine. N. Engl. J. Med. 263: 165-169 [Medline]. |
20. | Warthin, A.S.. 1931. The occurrence of numerous large giant cells in the tonsils and pharyngeal mucosa in the prodomal stage of measles. Arch. Pathol. 11: 864-874 . |
21. | Holt, P.G., M.A. Schon-Hegrad, and J. Oliver. 1988. MHC class II antigen bearing dendritic cells in pulmonary tissues of the rat. Regulation of antigen presentation activity by endogenous macrophage populations. J. Exp. Med. 167: 262-265 [Abstract]. |
22. | Holt, P.G., M.A. Schon-Hegrad, M.J. Phillips, and P.G. McMenamin. 1989. Ia-positive dendritic cells form a tightly meshed network within the human airway epithelium. Clin. Exp. Allergy. 19: 567-601 . |
23. | McWilliams, A.S., D. Nelson, J.A. Thomas, and P.G. Holt. 1994. Rapid dendritic cell recruitment as a hallmark of the acute inflammatory response at mucosal surfaces. J. Exp. Med. 179: 1331-1336 [Abstract]. |
24. | Xia, W., C. E. Pinto, and R. L. Kradin. 1995. The antigen presenting activities of Ia+ dendritic cells shift dynamically from lung to lymph nodes after an airway challenge with soluble antigen. J. Exp. Med. 181: 1275-1283 [Abstract]. |
25. | Bhardwaj, N., A. Bender, N. Gonzalez, L. K. Bui, M.C. Garrett, and R. Steinman. 1994. Influenza virus-infected dendritic cells stimulate strong proliferative and cytolytic responses from human CD8+ T cells. J. Clin. Invest. 94: 797-807 [Medline]. |
26. | Langhoff, E., F. Terwilliger, H.J. Bos, K.H. Kalland, M.C. Poznansky, O.M.L. Bacon, and W.A. Haseltine. 1991. Replication of human immunodeficiency virus type 1 in primary dendritic cell cultures. Proc. Natl. Acad. Sci. USA. 88: 7998-8002 [Abstract]. |
27. | Macatonia, S.E., R. Lau, S. Patterson, A.J. Pinching, and S.C. Knight. 1990. Dendritic cell infection, depletion and dysfunction in HIV-infected individuals. Immunology. 71: 38-45 [Medline]. |
28. | Cameron, P.U., M.G. Lowe, F. Sotzik, A.F. Coughlan, S.M. Crowe, and K. Shortman. 1996. The interaction of macrophage and nonmacrophage tropic isolates of HIV-1 with thymic and tonsillar dendritic cells in vitro. J. Exp. Med. 183: 1851-1856 [Abstract]. |
29. | Kobune, F., H. Sakata, and A. Sugiura. 1990. Marmoset lymphoblastoid cells as a sensitive host for isolation of measles virus. J. Virol. 64: 700-705 [Medline]. |
30. |
Caux, C.,
B. Vanbervliet,
C. Massacrier,
C. Dezutter-Dambuyant,
B. de Saint-Vis,
C. Jacquet,
K. Yoneda,
S. Imamura,
D. Schmitt, and
J. Banchereau.
1996.
CD34+ progenitors
from human cord blood differentiate along two independent
dendritic cell pathways in response to GM-CSF + TNF-![]() |
31. | Peguet-Navarro, J., C. Dalbiez-Gauthier, C. Dezutter-Dambuyant, and D. Schmitt. 1993. Dissection of human Langerhans cell allostimulatory function: the need for an activation step for full development of accessory function. Eur. J. Immunol. 23: 376-380 [Medline]. |
32. | Giraudon, P., and T. F. Wild. 1985. Correlation between epitopes on hemagglutinin of measles virus and biological activities: passive protection by monoclonal antibodies is related to their hemagglutination inhibiting activity. Virology. 144: 46-58 [Medline]. |
33. | Vermes, I., C. Haanen, H. Steffens-Nakken, and C. Reutelingsperger. 1995. A novel assay for apoptosis. Flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein-labeled Annexin V. J. Immunol. Methods. 184: 39-51 [Medline]. |
34. | Sallusto, F., M. Cella, C. Danieli, and A. Lanzavecchia. 1995. Dendritic cells use macropinocytosis and the mannose receptor to concentrate macromolecules in the major histocompatibility complex class II compartment: downregulation by cytokines and bacterial products. J. Exp. Med. 182: 389-400 [Abstract]. |
35. | Caux, C., C. Massacrier, C. Dezutter-Dambuyant, B. Vanbervliet, C. Jacquet, D. Schmitt, and J. Banchereau. 1995. Human dendritic Langerhans cells generated in vitro from CD34+ progenitors can prime naive CD4+ T cells and process soluble antigen. J. Immunol. 155: 5427-5435 [Abstract]. |
36. | Vallé, A., C.E. Zuber, T. Defrance, O. Djossou, M. De Rie, and J. Banchereau. 1989. Activation of human B lymphocytes through CD40 and interleukin 4. Eur. J. Immunol. 19: 1463-1468 [Medline]. |
37. | Wild, T.F., E. Malvoisin, and R. Buckland. 1991. Measles virus: both the haemagglutinin and fusion glycoproteins are required for fusion. J. Gen. Virol. 72: 439-442 [Abstract]. |
38. | Hyypia, T., P. Korkiamäki, and R. Vainionpää. 1985. Replication of measles virus in human lymphocytes. J. Exp. Med. 161: 1261-1271 [Abstract]. |
39. | Osunkoya, B.O., A.R. Cooke, O. Ayeni, and T.A. Adejumo. 1974. Studies on leucocyte cultures in measles. I. Lymphocyte transformation and giant cell formation in leucocyte cultures from clinical cases of measles. Arch. Gesamte Virusforsch. 44: 313-322 [Medline]. |
40. | Lucas, C.J., J.C. Ubels-Postma, A. Rezee, and J.M.D. Galama. 1978. Activation of measles virus from silently infected human lymphocytes. J. Exp. Med. 148: 940-952 [Abstract]. |
41. | Sullivan, J.L, D.W. Barry, S.J. Lucas, and P. Albrecht. 1975. Measles infection of human mononuclear cells. I. Acute infection of peripheral blood lymphocytes and monocytes. J. Exp. Med. 142: 773-784 [Abstract]. |
42. | Witmer-Pack, M. D., W. Olivier, J. Valinsky, G. Schuler, and R.M. Steinman. 1987. Granulocyte/macrophage colony-stimulating factor is essential for the viability and function of cultured murine epidermal Langerhans cells. J. Exp. Med. 166: 1484-1498 [Abstract]. |
43. | Yanagi, Y., B.A. Cubitt, and M.B.A. Odstone. 1992. Measles virus inhibits mitogen-induced T cell proliferation but does not directly perturb the T cell activation process inside the cell. Virology. 187: 280-289 [Medline]. |
44. |
Schlender, J.,
J.J. Schnorr,
P. Spielhoffer,
T. Cathomen,
R. Cattaneo,
M.A. Billeter,
V. ter Meulen, and
S. Schneider-Schaulies.
1996.
Interaction of measles virus glycoproteins
with the surface of uninfected peripheral blood lymphocytes
induces immunosuppression in vitro.
Proc. Natl. Acad. Sci.
USA.
93:
13194-13199
|
45. | Karp, C.L., M. Wysocka, L.M. Wahl, J.M. Ahearn, P.J. Cuomo, B. Sherry, G. Trinchieri, and D.E. Griffin. 1996. Mechanism of suppression of cell-mediated immunity by measles virus. Science (Wash. DC). 273: 228-231 [Abstract]. |
46. | Frankel, S., B.M. Wenig, A.P. Burke, P. Mannan, L.D.R. Thompson, S.L. Abbondanzo, A.M. Nelson, M. Pope, and R.M. Steinman. 1996. Replication of HIV-1 in dendritic cell-derived syncytia at the mucosal surface of adenoid. Science (Wash. DC). 272: 115-118 [Abstract]. |