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Blood, Vol. 96 No. 1 (July 1), 2000:
pp. 210-217
IMMUNOBIOLOGY
Interferon- and - inhibit the in vitro differentiation of
immunocompetent human dendritic cells from CD14+ precursors
Bradford L. McRae,
Taro Nagai,
Roshanak Tolouei Semnani,
Jean Maguire van Seventer, and
Gijs A. van Seventer
From the Committee on Immunology, Department of Pathology, Division
of Biological Science, University of Chicago, Chicago, IL; BASF
Bioresearch, Worcester, MA; and Tsukiyono Hospital, Gunma, Japan.
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Abstract |
Dendritic cell (DC) precursors and immature DC reside in epithelium
where they encounter pathogens and cytokines, which stimulate their
differentiation. We hypothesized that type-I interferons (IFN- and
- ), cytokines that are produced early in the innate immune response
against viruses and some bacteria, may influence DC differentiation and
function. To examine this possibility, we used an in vitro model of DC
differentiation in which initial culture of human CD14+
monocytes with granulocyte-macrophage colony-stimulating factor (GM-CSF) and interleukin (IL)-4 generates immature DC, and subsequent culture with tumor necrosis factor (TNF)- drives the final
development into mature DC. We found in this model that IFN-/,
added from the initiation of the culture on, significantly reduced the
survival and altered the morphology and differentiation of DC.
TNF--dependent maturation of IFN--treated immature DC led to
cells with reduced expression of CD1a, CD40, CD54, and CD80 when
compared with mature DC controls. IFN-/-treated DC further had a
reduced capacity to induce naive Th-cell proliferation through
allostimulation or anti-CD3 monoclonal antibody stimulation. In
addition, IFN-/-treated DC secreted less IL-12 upon stimulation
with Staphylococcus aureus Cowan strain or with
CD4+ T cells, and this decrease correlated directly with
their inability to support CD4+ T-cell secretion of
IFN- , even though T-cell lymphotoxin production was unaffected.
These findings indicate that type-I IFNs can influence the generation
of acquired immune responses by modifying T-helper cell differentiation
through the regulation of DC differentiation and function.
(Blood. 2000;96:210-217)
© 2000 by The American Society of Hematology.
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Introduction |
Dendritic cells (DC) are highly specialized
antigen-presenting cells (APC) required for primary T-cell activation
and T-cell-dependent immunity.1 Immature DC reside within
nonlymphoid tissues, where they actively capture and process antigen
and where contact with proinflammatory cytokines and bacterial products
induces their maturation. Upon activation, DC migrate via the afferent
lymphatics from peripheral tissues, such as the skin and intestine, to
secondary lymphoid tissues.2 Mature DC relocate to the
T-cell zone of lymph node parenchyma where they reside as potent
stimulators of naive T cells, presumably because of their high
expression of costimulatory molecules and production of cytokines such
as interleukin (IL)-12.2,3 CD4+ T cells that
recognize their specific antigen in the context of major
histocompatibility complex (MHC) class II presented on DC are retained
in the lymph node, where they undergo expansion and differentiation.
Unstimulated CD4+ T cells migrate through the medullary
sinus and back into the blood.
In recent years, several cytokines have been identified that
support in vitro maturation of DC from bone marrow or blood-derived progenitor cells. CD14+ mononuclear cells isolated from
peripheral blood and cultured with granulocyte-macrophage
colony-stimulating factor (GM-CSF) and IL-4 are efficient APC with
morphology and cell surface molecule expression typical of immature
DC.4 Alternatively, CD34+ progenitors cultured
with IL-3 or stimulated through CD40 also differentiate into functional
immature DC.5,6 In both instances, tumor necrosis factor
(TNF)- induces, in vitro and in vivo, further DC maturation
and significantly enhances the capacity of DC to stimulate resting T
cells by increasing the expression of costimulatory molecules and
cytokines, promoting migration of DC to draining lymph nodes, and
down-regulating DC antigen capture and processing.4,7 Other
factors that can similarly drive DC maturation include
lipopolysaccharides and IL-1.8,9 Thus, pathogen-derived
components and cytokines produced at sites of infection by the innate
immune response can contribute to DC maturation and, as a result,
regulate the ability of DC to stimulate naive T cells in draining lymph nodes.
Our laboratory has been interested in how type-I IFNs (IFN- and
-) may modulate DC development and, as a consequence, influence the
generation of acquired immune responses by altering T-helper cell
differentiation. Of particular relevance to our interest is that
IFN- is now a standard treatment for patients with multiple sclerosis,10,11 which leads to the question of what
immunomodulatory effects a systemic increase in IFN- has on the
generation of DC. IFN-/ are produced by many cell types,
including macrophages, T cells, keratinocytes, and Langerhans cells,
and have a broad range of immunomodulatory effects, including
inhibition of viral replication and stimulation of natural killer cell
activation.12 IFN-/ production during early stages of
the innate immune response against viral infection correlates with
increased viral titers and natural killer cell function in both murine
models and human disease13,14 and has been implicated in
promoting transient immunosuppression.15-17 The role of
IFN-/ in shaping the acquired immune function is, however, not
fully understood.12 Recently, we have shown that type-I
IFNs influence T-helper cell differentiation and function both by
acting directly on the T cell as well as by altering the function of
mature DC.18-20 Specifically, we showed that type-I IFNs
regulate homing receptor expression (E-selectin ligand and L-selectin)
on human CD4+ T cells18 and inhibit
T-cell-mediated CD40-induced secretion of functional IL-12 heterodimer
by DC.19,20 In the present work, we have extended our
studies to demonstrate that type-I IFNs influence the generation of
monocyte-derived DC in vitro by reducing the number of cells that will
differentiate into mature DC and altering the function of
immunocompetent human DC.
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Materials and methods |
In vitro differentiation of DC
CD14+ monocytes were isolated from peripheral blood by
counterflow centrifugal elutriation21 and frozen at
4 × 107 cells/mL. Cells were thawed as needed and
cultured in 6-well tissue culture plates (Costar, Cambridge, MA) at
approximately 5 × 106/mL in complete culture medium
(RPMI-1640 [Gibco/BRL, Gaithersburg, MD] supplemented with 10% fetal
bovine serum, 20 mmol/L L-glutamine, 100 IU/mL penicillin,
and 100 µg/mL streptomycin [BioWhittaker, Walkersville, MD]). IL-4
(biologic activity 108 IU/mg; Pharmingen, San Diego, CA)
and GM-CSF (biologic activity 2 × 108 IU/mg;
Pharmingen) were added to the cells at 30 ng/mL on day 1, day 4, and
day 7 of the culture. Some wells also received recombinant human
(rh)IFN--1a (biologic activity 2 × 108 IU/mg;
provided by Biogen, Cambridge, MA) or IFN-A (biologic activity 2 × 108 IU/mg; Biosource, Camarillo, CA)
on days 1 and 4 of culture at concentrations indicated. On day 6 of
culture, TNF- (Pharmingen) was added at 100 U/mL. Cells were
harvested on day 10 of culture with versene (EDTA) (Biowhittaker),
washed twice with Ca/Mg-free phosphate-buffered saline (PBS), and used
immediately in functional assays. Viable cell count was determined by
trypan blue exclusion. Monocytes were cultured for 48 hours in complete
medium without exogenous cytokines, washed, and used immediately in
functional assays.
Isolation of CD4+, CD45RA+,
CD45RO T cells
Human peripheral blood mononuclear cells from buffy coats of
anonymous healthy donors (Life Source, Glenview, IL) were isolated by
Ficoll gradient centrifugation. Resting
CD4+CD45RA+CD45RO T cells
were obtained by negative selection with antibodies and magnetic beads,
as described.22 CD45RA+ cells were greater than
95% pure by flow cytometric analysis using mouse monoclonal antibody
(mAb): CD45RA-phycoerythrin (PE) (clone B-C15; Biosource) and
CD45RO-FITC (fluorescein isothiocyanate) (clone UCHL1;
Caltag, South San Francisco, CA). Staining of cells with antibodies was
performed according to standard procedures, as described
previously,22 and cells were evaluated using a FACScan (Becton Dickinson, San Jose, CA).
CD4+ T-cell activation
Naive CD4+ T cells (5 × 105/well)
were cultured with allogeneic irradiated DC
(5 × 104/well) in a volume of 1 mL of complete
culture medium for 48 hours in 48-well plates (Costar), which had been
coated overnight at 4°C with 0.25 mL of 1 µg/mL humanized CD3
mAb (h)OKT3 (CDR grafted on human IgG1 23) in
PBS. At the end of the 48-hour stimulation period, the T cells were
resuspended and transferred to 6-well plates (Costar), and 1 mL of
fresh medium was added. Seven days after the initial stimulation, T
cells were counted and restimulated under identical conditions as the
primary stimulation, using fresh DC derived from the same donor used in
the primary stimulation. Supernatants were harvested after the 48-hour
secondary stimulation for analysis of cytokine production.
Enzyme-linked immunosorbent assays
Pairs of mAbs (Pharmingen) were used in sandwich enzyme-linked
immunosorbent assays (ELISAs) to measure lymphotoxin (LT) (sensitivity 0.1 ng/mL), TNF- (sensitivity 0.2 ng/mL), IFN- (sensitivity 0.2 ng/mL), p40 chain of IL-12 heterodimer (sensitivity 0.2 ng/mL), and
p35/p40 (p70) heterodimer of IL-12 (sensitivity 0.1 ng/mL). MaxiSorp
96-well plates (Nunc Inc., Naperville, IL) were coated with capture
mAbs (1-4 ng/mL) overnight at 4°C. The following day, plates were
washed and blocked with 3% bovine serum albumin in PBS at room
temperature for 2 hours. The plates were subsequently washed, and
standards and samples were added to the wells and incubated overnight
at 4°C. Biotinylated secondary mAb (1-3 µg/mL), avidin-peroxidase
(Sigma, St. Louis, MO), and
2,2'-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid (ABTS;
Sigma) were used to quantify cytokine levels, as per the Pharmingen protocol.
CD4+ T-cell proliferation
Proliferation assays were performed using standard techniques, as
described.24 Briefly, purified naive CD4+ T
cells (5 × 104 cells/well) were cultured in
complete culture medium in 96-well flat-bottom tissue culture plates
(Costar) with irradiated DC or irradiated monocytes. Seventy-two hours
later, cultures were pulsed with 25 µL/well of
[3H]thymidine solution (5 mCi/mL, 2 mCi/mmol specific
activity; New England Nuclear, Boston, MA) and harvested 24 hours later on glass fiber filters. For mixed lymphocyte cultures, purified naive
CD4+ T cells (1 × 105 cells/well) were
cultured in complete culture medium in 96-well round-bottom tissue
culture plates (Costar) with allogeneic DC and were pulsed during the
last 24 hours of a 6-day culture with 25 µL/well of
[3H]thymidine solution. Incorporation of radioactive
label was measured by a liquid scintillation counter. Results are
expressed as the arithmetic mean counts per minute of triplicate
cultures. Accessory cell-independent CD4+ T-cell
proliferation was obtained by stimulation with the combination of
phorbol myristate acetate (PMA) (1 ng/mL) and phytohemagglutinin (PHA)-M (Life Technologies, Gaithersburg, MD). Anti-CD80
(IF1) and anti-CD86 (3D1), both generously provided by Genetics
Institute (Cambridge, MA), were used at a final concentration of 10 µg/mL.
Flow cytometry
The murine antihuman mAbs used to stain DC included: anti-CD1a clone
B-B5 (Biosource), anti-CD14 clone B-A8 (FITC conjugated; Biosource),
anti-CD40 clone EA-5 (a gift from Dr T. Lebien, University of
Minnesota, Minneapolis, MN), anti-CD54 (intercellular adhesion molecule
[ICAM]-1) clone 84H10 (a gift from Dr P. Mannoni, INSERM U 119, Marseille, France), anti-CD80 clone BB1 (FITC conjugated; Pharmingen),
anti-CD86 clone IT2.2 (FITC conjugated; Pharmingen), anti-MHC class I
clone 5H7 (a gift from Dr S. Woodle, University of Chicago, Chicago,
IL), antiMHC class II clone IVA12 (a gift from Dr J. D. Capra,
University of Texas Southwestern Medical Center, Dallas, TX), and
anti-CD83-FITC (Immunotech, Marseille, France). Isotype controls
were used either unconjugated (ICN, Costa Mesa, CA) or directly
conjugated to FITC (Ancell, Bayport, MN). Staining of cells with
antibodies was carried out according to standard procedure,
as described previously,22 and the cells were evaluated
using a FACScan. Propidium iodide (PI) was used to exclude dead cells
from analysis.
Light microscopy photographs
Monocyte-derived DC were generated as described earlier. Cells were
harvested on day 10 of culture and transferred with culture supernatant
to fibronectin-coated (20 µg/mL; Life Technologies) glass chambers
(Bioptech, Butler, PA) for overnight culture. The next day,
differential interference contrast images were obtained with a Zeiss
Axiovert TV 100 microscope (Thornwood, NY) using a × 63 NA 1.4 Plan-Apochromat oil objective and a 12-bit Micromax 1300 Y CCD camera
(Princeton, Raleigh, NC). Digital images were subsequently converted
into TIFF format and imported into Adobe Photoshop 4.0 (San Jose, CA)
for editing and printing.
Induction of IL-12 secretion from DC
Naive CD4+ T cells (5 × 105) were
cultured in a volume of 2 mL of complete culture medium with
5 × 104 irradiated allogeneic DC,DC-IFN-, or
DC-IFN- for 48 hours in 24-well plates (Costar), which had been
coated overnight at 4°C with 0.5 mL of 1 µg/mL of CD3 mAb
OKT323 in PBS. In some experiments, DC or DC-IFN-
(5 × 105) were stimulated for 48 hours with 0.1%
Staphylococcus aureus Cowan strain (SAC) (Sigma).
Statistical analysis
Variations among culture conditions were examined by paired
t test.
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Results |
IFN- and - reduce cell recovery of monocyte-derived DC
The cytokine environment at the time of primary T-cell activation
has been shown to regulate Th-cell differentiation.25 We
postulated that the cytokines generated during the innate immune response may similarly contribute to the environment that influences DC
maturation, migration, and function. To test this hypothesis, we
generated functional DC in vitro in the presence or absence of IFN-
or -. Immature DC were generated by culturing CD14+
-elutriated monocytes for 10 days in the presence
of GM-CSF and IL-4. Addition of TNF- on day 6 induced further DC
maturation and resulted in large adherent cells with multiple dendritic
processes making up the "veils" that are characteristic of mature
DC (Figure 1A, top). The large majority of
cells cultured with GM-CSF, IL-4, TNF-, and 1 ng/mL IFN-
(referred to as DC-IFN-) (data not shown) or IFN- (referred to
as DC-IFN-) (Figure 1A, bottom) did not spread out as much as the
control DC and had a much reduced or even absent veiled appearance.
Furthermore, fluorescence-activated cell sorter analysis
of both forward scatter and side scatter showed a pattern for
DC-IFN- and - that was clearly distinct from control DC (Figure
1B and data not shown). Most strikingly, there was a dose-dependent
reduction in the number of viable cells recovered (as determined by
trypan blue exclusion) at day 10 from DC generated with either IFN-
or IFN- compared with control DC (Figure
2A). The reduced cell number in IFN- and
- cultures was thought most likely to reflect a decrease in DC
survival and not inhibition of DC expansion because DC generated from
monocytes do not undergo much cell division in our culture conditions.
To test this possibility, we further analyzed the IFN-/-induced decrease in viable DC number by performing flow cytometric studies following PI staining of cells taken directly out of culture. This
"no wash" analysis did not use any centrifugation step, and therefore it avoided the possible preferential loss of dead cells that
may occur during a wash step. Using this method, we found that although
the viability of IFN--treated DC (cultured at 1 ng/mL) was never
less than 80% at day 10, it was always slightly and significantly
decreased compared with control DC (Figure 2B). The mean percentages of
PI-positive-staining cells at day 10 were 12.5% and 8.0%,
respectively (n = 5, P < .05). Thus, it is possible that
the large reduction in day-10 recovery in DC-IFN- might have
resulted from the cumulative effects of a decreased viability of
IFN-- and --treated DC.
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| Fig 1.
Type-I IFNs alter the morphology of DC derived from
CD14+ precursors.
Mature DC were produced in vitro by culture with GM-CSF, IL-4, and
TNF-, as described in Materials and methods. IFN- was added to
the wells at a final concentration of 1 ng/mL on days 1 and 4 of
culture. (A) Photographs of DC were taken on day 10 of culture, as
described in Materials and methods: control DC (A, top),
IFN--cultured DC (A, bottom). Arrows indicate
dendritic processes within veils, and the 20-µm reference bar is
identical for both panels. (B) On day 10 of culture, cells were
harvested and subjected to flow cytometric analysis to evaluate
light-scattering properties by forward (x-axis) and side (y-axis)
scatter. Nonviable cells were eliminated from analysis using propidium
iodide. Results are representative of 5 independent experiments.
IFN- was added at days 1 and 4 of culture at the indicated
concentrations.
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| Fig 2.
Dose-dependent reduction in viable cells with type-I
IFNs.
(A) Coculture with type-I IFNs reduced cell recovery of
monocyte-derived DC. Relative viable cell recovery of DC at day 10 of
culture is presented. Type-I IFNs were added at days 1 and 4 of culture
at the indicated concentrations. n = number of independent
experiments; *P < .05,
**P << .01 as determined by paired
t test. (B) IFN- induced a small but significant decrease in
cell viability after 10 days of culture. On day 10 of culture,
cells were harvested and subjected to flow cytometric analysis to
evaluate the percentage of dead cells with propidium iodide (PI)
staining. Numbers indicate the percentage of PI-positive cells. Results
are 2 representatives of 5 independent experiments.
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IFN--treated monocyte-derived DC express less CD1a, CD40, CD54,
and CD80
Although the finding that type-I IFNs can influence cell survival is
of considerable interest, this study focused on defining the functional
properties of the viable DC remaining in culture. The cell surface
expression of viable DC recovered after 10 days in culture was analyzed
to identify any phenotypic changes due to the presence of IFN-.
Elutriated monocytes (CD14+,
CD1a) expressed high levels of MHC class I and II,
showed low levels of ICAM-1, and were negative for CD40, CD80, CD86,
and CD83 (data not shown). As reported previously, monocyte-derived
mature DC (CD14, CD1a+) expressed high
levels of ICAM-1, CD40, and CD86, and intermediate levels of
CD80.4 Interestingly, although DC-IFN- similarly lost
CD14 expression, they failed to uniformly up-regulate CD1a (Figure
3). Levels of ICAM-1 and CD40 were
consistently lower on DC-IFN- than on DC derived in the absence of
IFN-. Furthermore, IFN- prevented optimal up-regulation of CD80
while inducing a slightly higher expression of CD86, a costimulatory
molecule associated with mature DC (Figure 3).26 Expression
of MHC class II and the mature DC marker CD8327 was not
altered by culture with IFN- (Figure 3). The above described changes
in surface phenotype by coculture with IFN- were observed only at
the higher concentration of 1 ng/mL (Figure 3; compare upper and lower
panels). Interestingly, although at the lower (0.1 ng/mL) concentration
of IFN-, no distinct cell surface phenotype was observed, some of
the functional effects can still be seen (see Results).
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| Fig 3.
Cell surface phenotype comparison of DC and
DC-IFN- generated with 0.1 ng/mL or 1 ng/mL IFN-.
DC and DC-IFN- were harvested on day 10 of culture, washed twice,
and prepared for flow cytometry as described in Materials and methods.
Light lines represent staining of DC generated in the absence of
exogenous IFN-, whereas bold lines indicate staining of DC-IFN-
generated with 0.1 ng/mL or 1 ng/mL IFN-, as indicated. Unlabeled
isotype controls are depicted in upper left corner panels, and direct
FITC-conjugated isotype control for CD83 staining is depicted in the
panels directly left of CD83. Specificity of the staining with specific
mAbs is as indicated below the panels. Results are representative of 5 to 7 independent experiments.
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In summary, DC that develop in the presence of IFN- express to a
varying degree lower levels of CD1a, CD40, CD54, and CD80, but
otherwise exhibit a cell surface phenotype with low CD83 and high CD86
and MHC class II that is characteristic of mature DC.
DC-IFN- and DC-IFN- have diminished capacity to support
naive CD4+ T-cell proliferation
Competent APC provide the costimulatory signals and soluble
factors necessary for T-cell differentiation and expansion. We used a
naive Th-cell stimulation model to determine whether IFN- or -
treatment of DC influenced the ability of these cells to provide T-cell
costimulation. In this model, immobilized anti-CD3 mAb is used
specifically to trigger the TCR/CD3 complex, while DC cells added to
the culture provide costimulatory signals and soluble factors. This
stimulation model allows analysis of the costimulatory capacity of the
DC in isolation, as compared with a direct allogeneic stimulation
model, which would measure both the antigen presentation and
costimulatory capacity of the DC. (It should be noted that although
allogeneic DC are used in this model, any possible allospecific T-cell
response induced by the DC will provide only an extremely minor
contribution to antigen receptor stimulation as compared with that
induced by cross-linking with immobilized anti-CD3 mAb.) Naive
CD4+ T cells proliferated well in response to immobilized
hOKT3 and DC and required fewer DC to support optimal proliferation
than did CD4+ T cells stimulated with DC-IFN- or
-IFN- (Figure 4A,B). DC-IFN- and
- often could stimulate only 50% to 80% of the proliferative response observed with normal DC. When blocking mAbs were used to
disrupt specific costimulatory pathways, DC-IFN--induced T-cell proliferation was found to be largely dependent upon CD86
costimulation, whereas both CD80 and CD86 costimulation contributed to
normal DC-induced T-cell proliferation (Figure 4C). These data suggest that DC-IFN- depend largely on CD86 to support proliferation of
naive T cells.
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| Fig 4.
DC-IFN- and DC-IFN- have a decreased capacity to
support naive CD4+ T-cell proliferation.
Naive CD4+ T cells were stimulated with immobilized
anti-CD3 and either DC, DC-IFN- (A), or DC-IFN- (B) at the
indicated ratios. IFN- and IFN- were added at indicated
concentrations at days 1 and 4 of the culture. APC were washed to
remove cytokines used for in vitro differentiation. No exogenous
cytokines were added to the cultures for measurement of
[3H]-thymidine incorporation. After 3 days
in culture, wells were pulsed with 1 µCi of
[3H]-thymidine per well. Twenty-four hours later, plates
were harvested and incorporated [3H]-thymidine was
measured. DC-induced T-cell proliferation was significantly different
from DC-IFN--induced and DC-IFN--induced T-cell proliferation
at a 1:0.1 ratio (n = 4; P < .05 as determined by a
paired t test). Results depicted are representative of 4 independent experiments. (C) Naive CD4+ T-cell
proliferation induced by DC-IFN- (1 ng/mL) was not dependent on
CD80 costimulation. Anti-CD80 and anti-CD86 were added at the beginning
of the proliferative assays at a final concentration of 10 µg/mL.
Data are expressed as the percentage of inhibition of proliferation in
the presence of isotype-matched control Ig at a T-cell to DC ratio of
1:0.1. Results shown are the mean and SEM of 3 independent
experiments.
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A characteristic function of mature DC is their capacity to
generate a mixed lymphocyte reaction. The data provided in Figure 5 show that DC-IFN- also have a reduced
capacity to induce a mixed lymphocyte reaction. The day-6 proliferative
response of naive CD4+ Th cells to allogeneic DC-IFN-
(1.0 ng/mL) was significantly reduced when compared with control DC
(P << .01, n = 4).
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| Fig 5.
DC-IFN- have reduced capacity to induce a mixed
lymphocyte reaction.
Naive CD4+ T cells were stimulated with either allogeneic
DC or DC-IFN- (0.1 ng/mL or 1 ng/mL) at the indicated ratios. DC
were washed to remove cytokines used for in vitro differentiation. No
exogenous cytokines were added to cultures for measurement of
[3H]-thymidine incorporation. After 5 days in culture,
wells were pulsed with 1 µCi of [3H]-thymidine per
well. Twenty-four hours later, plates were harvested and incorporated
[3H]-thymidine was measured. Means of triplicate
measurements are shown. A representative of 4 independent experiments
is shown.
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DC-IFN- and DC-IFN- produce decreased levels of IL-12
IL-12 is a critical cytokine secreted by DC and influencing naive
Th-cell differentiation. Previous studies using our naive Th-cell
differentiation model demonstrated that anti-CD3 mAb-activated naive
CD4+ T cells stimulated DC secretion of the p40 subunit of
the IL-12 heterodimer.19 In these studies, the induction of
DC secretion of biologically active p35/p40 (p70) IL-12 heterodimer was
indicated by the finding that neutralizing anti-p70 IL-12 antibodies
prevented the T-cell IFN- secretion normally observed in this model,
even though p70 itself remained below the level of detection of our ELISA.19
We determined the influence of IFN- and - on the capacity of DC
to secrete IL-12 during Th-cell activation. Mature DC produced significant levels of p40 IL-12 when stimulated with CD4+ T
cells (Figure 6A), and we observed this
secretion to be significantly inhibited by the addition of IFN- and
- during the DC differentiation process. As expected from our
previous work,17 both DC and DC-IFN- and - failed
to secrete detectable levels of p70 IL-12 in this Th-cell stimulation
model (data not shown). Monocytes also did not produce detectable
levels of IL-12 in this assay (Figure 6A). To be able to determine the
levels of secreted p35/p40 (p70) IL-12 by DC, we stimulated the DC with
0.1% SAC for 48 hours. The results in Figure 6B show that, consistent
with their lower secretion of p40 IL-12, DC-IFN- are significantly
inhibited in their capacity to secrete p70 IL-12 in comparison
with control DC.
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| Fig 6.
DC-IFN- and DC-IFN- have reduced capacity to
secrete IL-12.
(A) DC (5 × 104), DC-IFN- (0.01 ng/mL or 1 ng/mL), or DC-IFN- (0.01 ng/mL or 1 ng/mL) were cultured with
5 × 105 naive CD4+ T cells and
immobilized anti-CD3. DC were washed to remove cytokines used for in
vitro differentiation. No exogenous cytokines were added to stimulation
cultures. Supernatants were harvested after 48 hours of the primary
stimulation. The results indicated with * show that p40
IL-12 production was significantly different from control DC p40 IL-12
production (n = 3-4, *P < .05 as determined
by paired t test). Results shown are the mean and SEM. (B)
DC-IFN- had reduced capacity to produce p70 IL-12 in response to
activation with 0.1% SAC. DC (5 × 104) or
DC-IFN- (1 ng/mL) were cultured with 0.1% SAC. DC were washed to
remove cytokines used for in vitro differentiation. No exogenous
cytokines were added to stimulation cultures. Supernatants were
harvested after 48 hours of stimulation. Results indicated with
* show that p70 IL-12 production was significantly
different from control DC p70 IL-12 production (n = 3;
*P < .05 as determined by paired t test). Results
shown are the mean and SEM.
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Thus, these and our previous results indicate that culture with IFN-
and - during DC differentiation prevents the development of DC with
full capacity to secrete functional IL-12.
DC-IFN- have a greatly reduced capacity to provide naive Th-cell
costimulation that leads to the generation of IFN--secreting Th
cells
The ability of monocytes, DC, and DC-IFN- (1 ng/mL) to support
activation and differentiation of anti-CD3 mAb-stimulated naive
CD4+ T cells was analyzed by measuring the T-cell cytokine
secretion profile at 2 time points during primary and secondary
stimulation. We previously reported that mature DC in this model
generate Th cells that secrete only Th1 cytokines, such as IFN- and
LT.19 In addition, the IFN- secretion observed in this
model was shown to be IL-12 dependent, whereas LT secretion was found
to be IL-12 independent. The results in Figure 7
demonstrate that monocyte costimulation failed to generate any
IFN-- or LT-secreting Th cells in either the primary or secondary
stimulation. In contrast, mature DC costimulation supported IFN-
production, which could be detected at low levels by 48 hours after the
primary stimulation and at much higher levels after the second
stimulation (Figure 7, top). DC-IFN-
could not induce IFN- production during primary T-cell activation,
and induction of IFN- secretion was at significantly lower levels
when compared with DC during the secondary stimulation. Of particular
interest was the finding that both DC populations induced similar
levels of T-cell LT secretion (Figure 7, bottom). Thus, although
DC-IFN- can induce normal levels of IL-12-independent T-cell LT
secretion, they are deficient in stimulating IL-12-dependent IFN-
secretion when compared with DC derived without IFN-. This finding
is consistent with the decreased secretion of IL-12 that was observed
in this DC population (Figure 6A,B). The failure of DC-IFN- to
induce significant IFN- secretion did not lead to the induction of
secretion of the Th2 cytokines IL-4, IL-5, and IL-13 (data
not shown).
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| Fig 7.
Costimulatory signals provided by DC-IFN- are
insufficient for optimal IFN- production by CD4+ T
cells.
A total of 5 × 104 monocytes, DC, or DC-IFN-
were cultured with 5 × 105 naive CD4+ T
cells and immobilized anti-CD3. APC were washed to remove cytokines
used for in vitro differentiation. No exogenous cytokines were added to
stimulation cultures. Supernatants were harvested from the primary and
secondary stimulations at the indicated time points. IFN- secretion
(top) and lymphotoxin (LT) secretion (bottom) were measured by ELISA.
Results indicate that DC-IFN- induced IFN-
production was significantly different from IFN- production induced
by DC (n = 4; *P < .05 as determined by paired t
test). Results shown are the mean and SEM.
|
|
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Discussion |
We have found that type-I IFNs disrupt the developmental program of
monocytes that is initiated by GM-CSF and IL-4, and which leads in a
TNF--dependent manner to mature DC. This disruption appears to
occur at least at 2 levels: (1) by decreasing viable cell recovery at
the end of the culture, and (2) by altering the phenotypic and
functional characteristics of the remaining viable cells. In regard to
the first issue, a small but significantly higher (5% to 10%) level
of cell death was observed at the end of culture in DC-IFN-
compared with control DC. We hypothesize that either the cumulative
effects of such increased cell death over the 10-day culture period may
account for the large (> 50%) decrease in viable cell recovery that
occurred or that early during DC differentiation, a large reduction in
viable cells occurred that is not adequately reflected in the viability
at the end of the culture. The results from the phenotypic and
functional studies were determined on viable cells and, therefore, the
alterations induced by IFN- and - that were observed in these
studies were likely to be caused by mechanisms independent from those
resulting in decreased cell recovery. An important phenotypic
alteration induced by IFN- coculture was a change in the expression
of costimulatory molecules, most notably a decrease in the
up-regulation of CD1a, CD40, ICAM-1, and CD80 expression. Functionally,
cells cultured under conditions normally leading to mature DC were
altered by coculture with IFN- and - such that: (1) they provided
less optimal costimulation for naive Th-cell proliferation, which for DC-IFN- was more dependent on CD86 costimulation than for control DC; and (2) they secreted less IL-12, which led to a decrease in
Th-cell IFN- secretion. Interestingly, both DC populations, generated with or without IFN-, were comparable in their
costimulatory capacity to induce Th-cell secretion of LT. The decrease
in costimulatory capacity of DC-IFN- to induce Th-cell
proliferation was also observed in direct allogeneic responses.
The precise mechanism(s) involved in the observed IFN-- and
IFN--induced alterations have not been identified, but several possible pathways have been eliminated. In this model, type-I IFNs do
not modulate expression of CD95 or CD95L, suggesting that Fas-mediated
apoptosis is probably not responsible for the enhanced death of
IFN--treated DC progenitors (data not shown). TNF- potentiates
DC development, at least in part through up-regulation of GM-CSF
receptor.28 Consequently, we investigated the possibility that IFN- might negatively regulate expression of this receptor, and
found that IFN- did not inhibit GM-CSF receptor levels (as determined by the CD116 staining) on the surface of DC progenitors (data not shown). We cannot, however, rule out the possibility that
IFN- may disrupt some other component of the GM-CSF signaling pathway. Similarly, we cannot exclude that the effect of coculture with
type-I IFNs on DC differentiation is in part mediated through effects
on other cell types present in the initial starting population of
elutriated monocytes.
It has been difficult to classify definitively the cells derived in the
presence of IFN-. We have referred to these cells as dendritic
because of their loss of the monocyte marker CD14 and their expression
of CD83, despite the decreased expression of CD1a (Figure 3).
Furthermore, DC-IFN- expressed high levels of adhesion molecules
associated with DC such as ICAM-3, CD11b, and CD11d (data not shown).
Thus, the IFN- phenotype does resemble that of direct blood-derived
DC.29 Our results further emphasize the heterogeneity of
monocyte-derived DC from the perspective of both the phenotypic markers
and the functional properties.29
Our results demonstrating an inhibitory effect of IFN- on DC
differentiation from monocyte-derived DC progenitors are consistent with other reports on the inhibition of hematopoiesis by IFN-/. McNeill and Killen30 found that poly I-poly C, which
stimulates type-I IFN production, inhibited colony formation in bone
marrow cells. These results were confirmed and extended by other
laboratories to show that type-I IFNs are cytotoxic for progenitor
cells.31 More recently, transient suppression of
hematopoiesis associated with viral infection in both humans and murine
models has been attributed to high levels of
IFN-/.16,17
Recent publications by Wang et al32 and Paquette et
al33 confirm that IFN- greatly reduces the number of DC
that can be generated from monocytes, but no other detrimental effects of IFN- on DC function were reported. However, the effects of IFN- on DC function in those reports cannot be compared directly with our data because of significant differences in both the culture conditions used to generate DC and the purity and phenotype of the
generated DC used in their function assays, as well as the T-cell
populations used to analyze DC function. Specifically, although we used
CD4+ naive T cells, Wang et al32 and Paquette
et al33 both used unseparated T cells for their studies.
Another recent report by Luft et al34 found that IFN-
and TNF- synergize to enhance the transition from immature to mature
DC under serum-free conditions. The differences between our results and
this report may be due to the type of progenitor cell used
(CD14+ versus CD34+, respectively) and the time
point during the differentiation process at which the type-I IFN
treatment was initiated (day 1 at DC progenitor stage versus day 4 at
immature DC stage, respectively). Consistent with the data of Luft et
al,34 we found that IFN- had little effect
on DC morphology when added on day 4 rather than day 1 of culture (data
not shown). Thus, these contrasting results suggest that IFN-/
can have both positive and negative regulatory effects on DC
differentiation depending upon the progenitor cell treated and the
stage during DC maturation at which IFN- is introduced.
Another level of complexity in regard to the role of type-I IFNs in
regulating T-cell immune responses is provided by data from Sprent et
al, who have published a series of studies indicating that type-I IFNs
promote CD8+ memory cell survival35 through the
induction of IL-15 secretion.36,37 Interestingly, these
growth- and survival-promoting effects did not extend to
CD4+ Th cells. In fact, we found that type-I IFNs may lead
to immune deviation by altering naive CD4+ Th-cell
differentiation through the promotion of an IL-10-secreting Th-cell
subset.19
In summary, several proinflammatory cytokines elaborated during the
innate immune response (eg, IL-1 and TNF-) are known to promote
maturation and migration of DC, a step necessary for the eventual
activation of naive CD4+ T cells in regional lymph
nodes.8,38,39 In this way, the early cytokine response to
various pathogens will determine the nature and efficacy of subsequent
T-cell-dependent immunity.40,41 We now show that type-I
IFNs, cytokines either elaborated during the early phase of the innate
host defense or given as therapeutic treatment, can negatively regulate
immune effector functions by influencing DC differentiation and
function. Our data indicate that the presence of type-I IFNs from the
initiation of DC differentiation can alter DC progenitor
differentiation in such a way as to diminish IL-12-mediated
inflammatory responses at sites of infection. These findings may
explain why prolonged IFN-/ production in peripheral tissues and
draining lymph nodes, as occurs during some chronic viral infections,
can lead to immunosuppression.15-17 They may also explain
some of the beneficial effects observed in treatment of patients with
multiple sclerosis with IFN-.10,11
| |
Acknowledgments |
We thank Dr Mark P. Hayes and Valerie Calvert (Food and Drug
Administration, Bethesda, MD) for providing elutriated monocytes, Drs
Susan E. Goelz and Paula Hochman (Biogen Corp., Cambridge, MA) for
providing human recombinant IFN-, and Dr Mary Collins (Genetics
Institute, Cambridge, MA) for the anti-CD80/86 antibodies. Beth
Beilfuss provided expert technical help when needed. Dr Michael Model's help with the light microscopy is greatly appreciated.
| |
Footnotes |
Submitted May 24, 1999; accepted February 22, 2000.
Supported by grant AI34541 from the National Institutes of Health and a
grant from Biogen, Inc. Also supported in part by a postdoctoral
fellowship from the National Multiple Sclerosis Society (B.L.M.).
B.L.M. and T.N. contributed equally to the data presented in this manuscript.
Reprints: Gijs A. van Seventer, Department of Pathology,
University of Chicago, 5841 S Maryland Ave, Room J541A, MC1089, Chicago, IL 60637; e-mail: gvsevent{at}flowcity.bsd.uchicago.edu.
The publication costs of this
article were defrayed in part by
page charge payment. Therefore,
and solely to indicate this fact,
this article is hereby marked
"advertisement"
in accordance with 18 U.S.C.
section 1734.
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References |
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Abstract
Introduction