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Kennedy Institute of Rheumatology, London, United Kingdom
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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production. The negative
effect of CD14+ cells may also be IL-10 mediated, IL-10
being lost from the culture supernatants of CD14-depleted PBMC and
rIL-10 reversing the effect of this depletion. On the other hand, mRNA
for the p35 and p40 subunits of IL-12 is still induced in CD14-depleted
cultures. The expansion of CD3-CD56+ cells was
also inhibited by CTLA4-Ig, indicating a role for CD80/86. B
lymphocytes were not required for the expansion of
CD3-CD56+ cells, whereas removal of MHC class
II+ cells from CD14-depleted cultures resulted in a
complete abrogation of these responses. Expansion of
CD3-CD56+ cells was reconstituted in MHC class
II-depleted cell cultures by adding back monocyte-derived dendritic
cells. These results indicate that the responses of
CD3-CD56+ NK cells to LPS may be driven by a
MHC class II+ B7+ CD14- peripheral
population, most likely blood dendritic cells. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, IL-12,
and TNF- (3, 4, 5, 6). During the course of our studies on the responses of human T cells to LPS, we observed that, in addition to CD3+ cells, a large population of CD3- cells was expanded in practically all the LPS-responsive individuals analyzed. Captivated by these results, we tried to define which other peripheral mononuclear cell population was expanding upon LPS challenge. As it is well known that human B cells and myeloid cells do not proliferate in response to LPS (2, 7), we thought it logical to test whether LPS could induce the proliferation and expansion of human CD3-CD56+ NK-like cells. The analysis of PBMC stimulated in vitro with LPS provided the experimental evidence that the expanding CD3- cells expressed the CD56 marker, thus indicating their likely NK nature.
NK cells play a significant role in immune responses to exogenous pathogenic agents, as well as in defense against cancer cells (8, 9). Further studies have shown that they can also influence the adaptive immune system and direct the pattern of T cell responses in autoimmune diseases (10, 11). More recently, the mechanisms involved in target cell recognition by NK cells have been studied in great detail, particularly with regard to the role of MHC class I-specific inhibitory receptors (12, 13, 14).
The physiological stimuli normally required for the promotion and
regulation of NK cell proliferation are, however, less well
characterized. Human
CD3-CD56+ cells have
previously been expanded in vitro using combinations of cytokines,
usually IL-12 with either IL-2 or IL-7 (15, 16).
Furthermore, stimulation of CD14+
monocyte-depleted low density lymphocytes with high concentrations of
IL-2 alone has also been reported to enhance the expansion of both
CD3-CD56+ and
CD3+CD56+ populations
(17). Depletion of CD14+ monocytes
from PBMC also results in the expansion of
CD3+CD56+ cells on
stimulation with IL-1, IL-2, IFN-, and anti-CD3, or with a
combination of IL-2 and IL-12 (18, 19). These results
suggested that, although myeloid cell-derived growth factors were
required for the expansion of CD56+ cells,
CD14+ monocytes could have an inhibitory
effect.
In this study we, therefore, investigated the roles of myeloid cells, and their released factors in modulating the expansion of CD3-CD56+ (NK) cells by LPS.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PBMC were prepared from the blood of 10 normal, healthy individuals by standard separation on Ficoll-Hypaque. In all cases, informed consent was sought and ethical permission was granted. Cells were cultured in RPMI 1640 supplemented with 10% freshly prepared autologous serum. Serum was not heat inactivated, as this can impair LPS responses (20). Cells were stimulated as 1- to 2-ml cultures in 14-ml polyethylene tubes (Falcon) at a concentration of 1 x 106/ml for between 5 and 9 days in total. LPS, serotype 026:B6 (Sigma, Poole, U.K.), was used at 1 µg/ml final concentration and was freshly reconstituted for each experiment. In some experiments, human rIL-10 (PharMingen, San Diego, CA) was added to cultures at a final concentration of between 0.25 and 10 ng/ml. For measurement of proliferation, 100-µl aliquots of cells (equivalent to 1 x 105 of starting population) were transferred to 96-well U-bottom plates and pulsed overnight with [3H]thymidine at 0.5 µCi/well. Further aliquots of cells were taken for flow-cytometric analysis.
Depletion of cell subsets from PBMC
CD14+ cells were depleted from PBMC using magnetic cell separation system (MACS)3 under standard conditions. Briefly, 2–3 x 107 PBMC were incubated with CD14-conjugated MACS beads, and labeled cells were separated on a MACS column. Depleted cultures always contained less than 0.5% contaminating CD14+ cells. In some experiments, CD14-positive fractions were retained for generation of DC. For sequential depletions, CD14- cells were labeled either with an unconjugated CD19 (Serotec, Columbus, OH) or anti-MHC class II Abs (L243, mouse IgG2a; American Type Culture Collection, Manassas, VA). Cells were then incubated with anti-mouse Ig-conjugated MACS beads and removed on a MACS column. Again, depleted cells contained less than 0.5% contaminating CD19 or MHC class II+ cells. MHC class II-depleted cells were reconstituted in some cases with monocyte-derived DC. Monocyte-derived DC were prepared from the CD14+ cell fraction from the MACS column after leaving overnight and incubation for an additional 5 days in the presence of GM-CSF (Behring, Marburg, Germany) and IL-4 (Sandoz, Basel, Switzerland). In vitro generated DC were, as expected, CD14 negative, CD1b positive, and expressing high levels of MHC class II. In some experiments, CD3-CD56+ cells were enriched for cytotoxicity assays after LPS stimulation. This was done using a MACS NK cell selection kit (Miltenyi Biotec, Bergisch Gladbach, Germany). Cell populations comprising greater than 93% CD3-CD56+ cells with less than 3% contaminating CD3+ cells were obtained by this procedure.
mAbs and FACS analysis
Cells were labeled for analysis with the following mAbs: anti-CD56, FITC (Serotec), or anti-CD56 biotin (PharMingen); anti-CD3, PE, or FITC (Sigma); anti-KIR2DL2 (A3, kindly provided by Dr. E. Ciccone, University of Genoa, Genoa, Italy); and anti-KIR2DL2/3 and anti-KIR2DS2, anti-KIR3DL1, and anti-KIR3DL2 (DX27, DX9, and DX31, respectively; kindly provided by Dr. L. Lanier (21)). Unconjugated Abs were detected using goat anti-mouse Ig PE (Southern Biotechnology Associates, Birmingham, AL), and biotinylated Abs were detected with streptavidin PE (Southern Biotechnology Associates) or streptavidin quantum red (Sigma). A minimum of 10,000 events was measured on a FACScan flow cytometer (Becton Dickinson) and analyzed using the WinMDI software package after gating on total viable cells or blastoid cells on the forward and side scatter profile.
Labeling of cells with CFSE
This was conducted essentially as described elsewhere (22). Briefly, PBMC (2 x 107) were resuspended in 1 ml of RPMI 1640, without serum, and incubated with an equal volume of CFSE (5 µM) for 5 min in a water bath at 37°C. Unincorporated dye was quenched immediately by adding an equal volume of FCS. Cells were then washed in a large volume of RPMI, followed by two washes in RPMI, 10% autologous plasma. The level of CFSE incorporated was tested by FACS analysis after counterstaining cells with anti-CD56 and anti-CD3 Abs (see above). Cells were then stimulated with LPS in the usual manner, and the level of CFSE was monitored.
Cytotoxicity assay
Target cells (1 x 106, K562) were labeled with 100 mCi Na2 (51Cr) (Amersham, Bucks, U.K.) for 90 min at 37°C in RPMI medium containing 10% FCS, washed a total of four times in medium, resuspended in medium with 5% autologous serum, and subjected to cytotoxicity assays. A total of 100 µl of effector cells (serially diluted in RPMI, 5% autologous serum) was added to 5 x 103 labeled targets in 100 µl of medium in 96-well round-bottom plates. No specific target cell lysis was detected in control cultures incubated with LPS alone in the absence of effector cells, confirming that the observed activity was not due to residual LPS in the cultures. The plates were centrifuged briefly and incubated for 4 h. The supernatant was then harvested and counted on a gamma counter. Cytotoxicity was calculated as the percentage of maximal releasable counts (5% Triton X-100) after subtraction of spontaneous release. Spontaneous release was less than 15% of maximum release.
Inhibition experiments using mAbs or CTLA-4Ig
Anti-IL-12 mAbs (C8.6 and C8.1, both mouse IgG1; kindly provided by Dr. Giorgio Trinchieri, Wistar Institute, Philadelphia, PA) were used in combination at 10 and 2 µg/ml final concentration, respectively. Anti-IL-10R Ab (3F9, rat IgG2a; kindly provided by Kevin Moore, DNAX, Palo Alto, CA) was used at 5 µg/ml. Isotype-matched control Abs, MOPC-21 (mouse IgG1; Sigma) and AFRC (rat IgG2a, anti-dog MAC 1), were used at the same equivalent concentration as above. CTLA-4Ig (Genetics Institute, Boston, MA) was used at a final concentration of 10 µg/ml. All neutralizing reagents were added 2 h before stimulation with LPS and were present for the duration of culture.
Enzyme-linked immunosorbent assay
Supernatants were taken from unstimulated PBMC or CD14-depleted
PBMC, or after 24 h of stimulation with LPS. IL-10 was measured by
sandwich ELISA using unconjugated mouse anti-human IL-10 (945A5D11;
Cambridge Bioscience, Cambridge, U.K.) as a capture reagent and
biotinylated mouse anti-human IL-10 (945A5A10; Cambridge
Bioscience) to detect. For IFN- measurement, supernatants were taken
from PBMC or CD14-depleted cultures after 5 days and tested in an ELISA
as above using the Ab pair NIB42 for capture and 4SB3 biotin to detect
(PharMingen). The bound capture reagents were detected using
HRP-conjugated streptavidin (Southern Biotechnology Associates),
followed by tetramethylbenzidine substrate.
PCR analysis
RNA was extracted from PBMC or CD14-depleted PBMC using the RNAeasy kit (Qiagen, Chatsworth, CA), according to manufacturer’s instructions, and reverse transcribed under standard conditions. Amplification of cDNA was conducted using the following primers: IL-12 p35 sense primer, 5'-CTTCACCACTCCCAAAACCTG-3' (nt 281–302) (23); IL-12 p35 antisense primer, 5'-AGCTCGTCACTCTGTCAATAG-3' (nt 813–792) (23); IL-12 p40 sense primer, 5'-CCACATTCCTACTTCTC-3' (nt 822–839) (24); IL-12 p40 antisense primer, 5'-GTCTATTCCGTTGTGTC-3' (nt 1077–1060) (24); ß-actin sense primer, 5'-GGGTCAGAAGGATTCCTATG-3' (nt 1365–1384) (25); and ß-actin antisense primer, 5'-CTCCTTAATGTCACGCACGATTTC-3' (nt 2307–2284) (25). Samples were amplified as follows: IL-12 p35, 35 cycles with denaturation at 94°C for 30 s, annealing at 56°C for 30 s, and extension at 72°C for 1 min; IL-12 p40, 35 cycles, denaturation at 94°C for 30 s, annealing at 52°C for 30 s, and extension at 72°C for 1 min; ß-actin, 25 cycles, denaturation at 94°C for 30 s, annealing at 55°C for 30 s, and extension at 72°C for 1 min. Products were run on a 2% agarose gel and visualized using software.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Initial experiments showed that CD56+ cells
increase in number, become blastoid in appearance, and acquire the
activation marker CD25 on stimulation with LPS. We therefore wanted to
show that such an increase actually involved cell division and was not
simply, for example, an LPS-induced effect on cell survival or a
modulation of CD56 expression. To do this, we labeled PBMC with CFSE, a
fluorescent dye, which is distributed equally between daughter cells on
each cell division, resulting in a progressive loss of membrane
fluorescence with time during cell proliferation (22).
Results for a representative donor are shown in Fig. 1
. LPS stimulation resulted in a
progressive loss of CFSE predominantly in CD56+
cells, this being detected as early as day 5 (Fig. 1, second
row). The level of CFSE expression decreased further by day 7, and
a progressive expansion of the resulting
CD56+CFSElow
(proliferating) cell population was observed (day 5, 3.2%; day 7,
17.4%; and day 9, 48.7%). A small proportion of
CD3+ cells was also proliferating, reaching 7.6%
by day 9 (Fig. 1, right-hand panels). These were also
contained CD56+ cells (data not shown). These
results show that CD56+ cells are the predominant
cell population proliferating in response to LPS. The data presented
also provide strong evidence that LPS does not stimulate all
CD3-CD56+ cells. Only a
small fraction of these cells is undergoing cell division on day 5
(3.2% in the experiment shown), and the majority retain the level of
CFSE observed in unstimulated cells or cells tested on day 0 (Fig. 1, upper and left panels). The data reported in this
figure also indicate that LPS stimulation up-regulates the expression
of CD56 in proliferating cells as well as in nondividing
CD56+ cells (Fig. 1, second,
third, and fourth left panel).
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LPS drives the proliferation of CD3-KIR+ cells
As the
CD3-CD56+ phenotype
normally defines NK-type cells in humans, it was important to confirm
that the cells stimulated by LPS expressed KIR receptors. To do this,
we again loaded PBMC with CFSE, stimulated them with LPS, and analyzed
the proportion of CD56+ or
KIR+ cells within both dividing (CFSE low) or
nondividing (CFSE high) populations after gating on
CD3- cells. Results from three LPS-responsive
individuals are shown in Fig. 2. As
expected and similarly to Fig. 1
, a high proportion of
CD56+CD3- cells was
present in the dividing cell (CFSE high) population in all three
individuals (Fig. 2). Separate experiments revealed that the
proliferating CD56+CFSE low cells also expressed
CD16 and CD94 (data not shown). Again, similarly to Fig. 1, some of the
CD3-CD56+ cells remained
in the nondividing fraction, indicating that not all of these cells
could proliferate in response to LPS. We used four different reagents
recognizing KIR2DL2, KIR2DL2/3, KIR3DL1, and KIR3DL2. All of these were
expressed in the CFSE high, nondividing population in all three
individuals shown, with the exception of KIR3DL2, which was not present
in donor 3, using isotype-matched control Ab as a reference (Fig. 2).
The expressed KIR were all present in the dividing (CFSE low) cell
population after stimulation with LPS (Fig. 2). These data confirm that
LPS stimulates cells of NK phenotype. Furthermore, LPS stimulates a
cross-section of CD56+CD3-
cells and does not act preferentially on cells with a particular KIR
phenotype in the individuals tested.
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It has been reported that the proliferative responses of human
PBMC to LPS are IL-12 dependent (26), and IL-12 has been
shown to enhance the proliferation of CD56+ cells
from human PBMC responding to exogenously introduced growth factors
such as IL-2 and IL-7 (15). We therefore wanted to know
whether the expansion of CD56+ cells
(CD3-CD56+ and
CD3+CD56+) was dependent on
IL-12. To do this, we cultured cells with neutralizing anti-IL-12
or with isotype-matched controls for the duration of LPS stimulation.
Results from two of eight LPS-responsive individuals tested are shown
in Fig. 3A. LPS-induced
proliferation of PBMC was inhibited in the presence of anti-IL-12
mAb compared with the isotype-matched control (>75% inhibition in
both individuals shown), confirming the results of others
(26).
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A). The same effects were
observed using a neutralizing anti-IL-10 mAb, but anti-IL-10R
Abs were selected for use, as these were effective at lower
concentration (data not shown).
Since we described that LPS principally expanded
CD56+ cells (mainly
CD3-CD56+), we tested
whether the substantial changes in proliferation were mirrored by
changes in the proportion of these cells in the cultures (Fig. 3B). Anti-IL-12 mAbs caused a large reduction in the
proportion of CD56+ cells
(CD3-CD56+ and
CD3+CD56+) in the cultures
(Fig. 3B).
Increased proliferation in the presence of anti-IL-10R Ab was also
mirrored by an overall increase in CD56+,
involving both CD3- and
CD3+ populations compared with the
isotype-matched control (Fig. 3B). The proportion of
CD56+CD25+ cells also
increased in both donors shown and in two further individuals in the
presence of anti-IL-10R Ab compared with isotype-matched control
(data not shown).
Expansion of CD3-CD56+ cells is enhanced in monocyte-depleted cultures
As both IL-10 and IL-12 are secreted by monocytes
(CD14+) upon LPS stimulation, we wanted to
investigate the role of these cells in supporting the proliferation and
expansion of CD3-CD56+
cells. Depletion experiments were conducted comparing the responses of
intact PBMC or monocyte-depleted PBMC with LPS. Cells were left
undepleted or were depleted using magnetic beads conjugated to an
anti-human CD14 mAb. Cell preparations were then cultured for 8
days in the presence or absence of LPS. Proliferative responses of PBMC
to LPS vary between different individuals on the basis of stimulation
index, as described previously (3, 26). We therefore
tested the effect of monocyte depletion on different individuals with
strong or weak responses to LPS. Results from three representative
individual donors (one high and two low responders) are shown in Fig. 4A. Proliferative responses
were enhanced in individuals originally making high responses to LPS
(donor 1). More surprisingly, proliferation was also enhanced in the
three individuals making only weak responses within PBMC (donors 2 and
3). An increase in stimulation index was observed after CD14 depletion
for all of the individuals shown. This was due to both a decrease in
background proliferation and an enhancement of LPS-induced
proliferation. The observed increase in LPS-induced proliferation was
reflected in an increase in the proportions of
CD3-CD56+ cells (Fig. 4B). We also observed an increase in the total number of
CD3-CD56+ cells recovered
after LPS stimulation of CD14-depleted cultures in four of seven
individuals tested (mean cell number before
stimulation/106 cells, 1.25 x
105, range 1.06–1.46 x
105; mean cell recovery after LPS stimulation,
1.79 x 105, range 1.27–2.21 x
105). Furthermore, the enhancement of
CD56+ cell proliferation in the absence of
CD14+ cells was confirmed by testing the CFSE
staining profile after stimulation with LPS. The level of CFSE in
CD56+ cells was then determined by FACS analysis
(Fig. 5). As expected, unstimulated
CD56+ cells (day 0) had uniformly high levels of
CFSE, both for PBMC and CD14-depleted populations (Fig. 5, upper
panels). Similarly to Fig. 1, a clear CD56+
CFSE low/negative population was detected in PBMC after 8 days of
stimulation with LPS (Fig. 5, lower left panel). The
proportion of cells in this population was clearly enhanced after
depletion of CD14+ cells, indicating enhanced
proliferation (Fig. 5, lower right panel).
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The studies reported to date do not address the question of
whether the cells expanded by LPS have cytolytic activity, a key
feature of NK cells. Moreover, as depletion of monocytes resulted in an
enhanced proliferation of CD56+ cells, we were
interested to test whether these cells had any alteration in their
cytolytic activity. To do this, we compared the ability of PBMC or
CD14-depleted fractions to lyse K562 target cells in a chromium release
assay. Fig. 6 shows lytic activity after
LPS stimulation of PBMC or CD14-depleted PBMC from representative high
and low LPS responders. Both PBMC and CD14-depleted populations from
the high responder efficiently lysed K562 cells after stimulation with
LPS, specific lysis being observed at E:T ratios as low as 2.5:1, and
an enhancement occurring in CD14-depleted cultures (Fig. 6, left). That CD3+ cells were not
responsible for the observed cytolytic activity was also confirmed
using CD3-CD56+ cells
enriched from the PBMC of this donor by magnetic bead separation after
stimulation with LPS (see Materials and Methods)
(inset, Fig. 6, left). For the low responder,
only weak lytic activity was observed in unfractionated PBMC (Fig. 6, right). This was, however, dramatically enhanced in
CD14-depleted PBMC, over 40% lysis being observed at E:T ratio of 80:1
and clearly detectable at lower ratios. Furthermore, only marginal
lysis of K562 cells was observed in unstimulated cultures of PBMC or
CD14-negative cells, indicating that the killing activity was LPS
driven. This indicates that CD14+ cells are not
required, and may often be inhibitory, for the induction of NK
cytolytic activity on stimulation with LPS.
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production is enhanced in CD14-depleted PBMCThe data presented to date indicate that LPS induces, although at different levels, the expansion of CD3-CD56+ cells with killing activity. More importantly, removal of CD14+ cells drastically enhanced the expansion, proliferation, and killing activity of these cells. Thus, depletion of CD14 cells amplified the action of LPS on CD3-CD56+ cells.
It is known that IFN- is produced by LPS-stimulated PBMC and by
IL-12-driven CD3-CD56+ NK
cell clones (3, 28). We therefore tested the effect of
CD14 depletion on IFN- production in response to LPS. Fig. 7 shows the amount of IFN- produced in
cultures of PBMC or monocyte PBMC after LPS simulation in two poor
responders to LPS. Only small amounts of IFN- were detected in
LPS-stimulated PBMC from both individuals. This was, however,
dramatically enhanced after LPS stimulation of monocyte-depleted
cultures (Fig. 7).
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As a consistent increase in CD56+ cell
expansion was observed both in anti-IL-10R-treated PBMC and
CD14-depleted PBMC, we wanted to confirm that
CD14+ cells were indeed the major producer of
IL-10 in LPS-treated PBMC. To do this, we depleted the PBMC from five
different individuals of CD14+ cells and measured
the amount of IL-10 in the supernatants after 24 h of stimulation.
Table I shows that depletion of
CD14+ cells results in a dramatic reduction in
the amount of IL-10 produced in LPS-stimulated cultures, approaching
background, unstimulated levels in all the individuals tested.
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shows the effect of rIL-10 on the
responses of two individuals. LPS stimulated proliferation in
CD14-depleted cultures in both individuals, and this was inhibited in a
concentration-dependent manner by rIL-10 (Fig. 8A). As
expected, the stimulation of proliferation by LPS was reflected in an
increase in the proportion of
CD3-CD56+ cells,
particularly those expressing high levels of CD56 (Fig. 8B,
left-hand panels). rIL-10 inhibited both the expansion of
CD3-CD56+ cells and the
increased CD56 expression, these approaching the levels observed in
unstimulated cells for both the individuals shown (Fig. 8B,
central and right-hand panels). IL-10 therefore
has a negative effect on the expansion of
CD3-CD56+ cells in
response to LPS even in CD14-depleted cultures.
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We have shown that IL-12 is required for the LPS-mediated
stimulation of CD3-CD56+
cells. As monocytes are capable of producing IL-12 in response to LPS,
we tested whether this could still be produced in CD14-depleted
cultures. We were unable to measure IL-12 p70 protein by ELISA in
either PBMC or CD14-depleted cells after LPS stimulation. This lack of
soluble factor may have been to IL-12 being produced only in low
amounts and/or rapidly utilized in these cultures. We therefore decided
to test for IL-12 p35 and p40 mRNA in both PBMC and CD14-depleted
cultures before and after LPS stimulation. Initially, we performed a
time course of IL-12p35 and p40 mRNA in PBMC between 0 and 20 h of
stimulation and found increased expression of both mRNAs after 8 h
of stimulation. We then compared the expression of these mRNAs in PBMC
or CD14-depleted cells in unstimulated cells or after 8 h of
stimulation with LPS. Fig. 9 shows PCR
analysis of IL-12 p35 and p40 mRNAs from a representative individual.
IL-12 p35 PCR resulted in two major products of 531 and 420 bp, the
smaller product lacking exon 3 of the published genomic sequence (U.
Johansson, personal communication), (29). As expected,
IL-12 p35 mRNA was expressed in unstimulated cells and induced further
after 8 h of stimulation with LPS in PBMC. IL-12 p35 mRNA was also
present in PBMC depleted of CD14+ cells, and this
increased on LPS stimulation (Fig. 9, upper panel). IL-12
p40 mRNA was absent in unstimulated cells, but was again present in
both PBMC and CD14-depleted PBMC after LPS stimulation (Fig. 9, middle panel). Similar amounts of ß-actin PCR product were
detected in all samples tested. These results indicate that the levels
of IL-12 mRNA increase in CD14-depleted PBMC upon LPS stimulation.
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Proliferative responses of PBMC to LPS have been shown to require
costimulation via B7-CD28, these being inhibited by CTLA-4Ig
(26). CD80 and CD86 are up-regulated on monocyte-derived
DC by LPS, and peripheral blood DC-type cells express or up-regulate
these after overnight culture (30, 31). We therefore
tested the effect of B7 blockade via CTLA-4Ig on proliferation and
CD56+ cell expansion in CD14-depleted cultures.
Results from a representative individual are shown in Fig. 10. As expected, CTLA-4Ig inhibited the
proliferative response of PBMC to LPS (Fig. 10). This inhibition was
reflected in a reduction in the number of
CD3-CD56+ cells in these
cultures. Moreover, the enhanced proliferation and expansion of
CD3-CD56+ cells to LPS in
CD14-depleted cultures were also highly sensitive to treatment with
CTLA-4Ig (Fig. 10).
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). The
additional depletion of CD19+ B cells had no
effect, whereas the depletion of all MHC class II-bearing cells from
CD14-depleted cultures completely abolished proliferation and
CD3-CD56+ cell expansion
in response to LPS (Fig. 10). Similar results were observed in three
further individuals. B7+, MHC class
II+, CD14-,
CD19- cell populations are therefore required to
drive responses to LPS. Monocyte-derived dendritic cells can reconstitute the NK response
We tested the ability of monocyte-derived DC to reconstitute the
CD3-CD56+ response to LPS.
CD14-depleted cultures were further depleted of MHC class II-positive
cells, and were stimulated in the presence or absence of the autologous
monocyte-derived DC. These DC were able to reconstitute the LPS-driven
proliferation and expansion of
CD3-CD56+ cells in
cultures depleted of both CD14 and MHC class II-positive cells (Fig. 11).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The involvement of IL-12 in the induction of the LPS response is perhaps not surprising, because this has previously been shown to synergize with IL-2 to preferentially expand CD56+CD3- cells from PBMC (15). IL-12 alone also augments the cell surface expression of the CD56 molecule, without inducing proliferation (32). More intriguing is the finding that IL-10 plays a powerful negative role in these responses. In view of this, it is likely that, rather than reflecting IL-12 production alone, the magnitude of CD3-CD56+ cell proliferation in response to LPS may be, at least in part, controlled by the relative balance of IL-10 vs IL-12. Such regulation may partially explain why the proliferative response of PBMC to LPS varies in magnitude between different individuals (3, 26).
The control of LPS-induced expansion of
CD3-CD56+ does not rely
solely on IL-12 and IL-10. We describe in this work, in agreement with
previous studies, that the masking of the B7 family molecules with
CTLA-4Ig completely blocks LPS-driven proliferation and, additionally,
the expansion of the
CD3-CD56+ cells. It is
possible that CD3-CD56+
cells can, themselves, receive signals via B7-CD28 ligation. Several
recent lines of evidence support this. First, murine NK cells recognize
and kill B7+ targets, and killing of tumor cell
lines by human NK cells is enhanced on transfection of CD80/CD86
(33, 34). Second, anti-CD28 enhances IL-12-driven
IFN- production in murine NK cells (35). Third, a
subpopulation of human NK cells expresses a particular variant of CD28
(36). Finally, we observe expansion of
CD3-CD56+ cells with LPS
even in cultures that had been depleted of T cells, the other
population that may have been effected by CTLA-4Ig (M. Goodier,
unpublished observations).
The opposing influences of IL-10 and IL-12 on CD3-CD56+ cell expansion in our system are intriguing, because different cell types in our cultures may be producing these. We have shown in this study that the stimulation index for proliferation and the expansion of CD3-CD56+ cells are both dramatically enhanced on removal of CD14+ cells, even in individuals normally making weak LPS responses. Different functional characteristics of the CD14+ vs CD14- myeloid cell populations could, therefore, further explain why some donors are poor responders while others are high responders on LPS stimulation (3, 26).
We have also shown, in contrast to the negative role of
CD14+ cells, that a MHC class
II+, non-monocyte, non-B cell population appears
to be driving the response to LPS. We consider a role for pre-existing
activated T cells, which may be both MHC class
II+ and B7-1+, as unlikely,
as the expansion of
CD3-CD56+ cells occurs
even in cultures depleted of CD3+ cells before
LPS stimulation (M. Goodier, unpublished observations). Furthermore, as
shown in this study, this expansion can occur after reconstitution of
CD14- MHC class
II- cells with monocyte-derived DC alone. Prime
candidates in human peripheral blood would therefore be dendritic cell
populations, which are also efficient producers of IL-12 (30, 37). Recent studies have shown the presence of several distinct
DC populations in human peripheral blood (31, 38, 39, 40). One
of these, termed pDC2 (CD11c-, MHC class
II+, CD4+ phenotype), is
responsible for the early production of IFN-ß in response to virus
(38). As IFN-/ß in turn drive IL-12 production
(37), it is possible that this DC population is playing a
role in CD3-CD56+ cell
expansions. Further detailed cell fractionation and reconstitution
experiments will resolve this question.
CD14 is a major cell surface receptor for LPS, which conveys its
biological effects. This raises the question of how LPS is mediating
its effects in cultures that have been depleted of cells bearing
surface CD14. Human monocyte-derived DC respond to LPS by producing
TNF-, IL-6, IL-8, and IL-12 and up-regulating surface expression of
HLA-DR, B7-1, B7-2, and CD40, although these are negative for surface
CD14 (30). The efficient activation of these cells
requires soluble CD14 in the serum (30). A strong
possibility is that soluble CD14 from serum is binding the LPS in our
cultures, thereby acting as a carrier molecule.
NK cells play an important role in immune responses to pathogenic agents such as bacteria and viruses in animal models (8, 41). The mechanisms shown in this study to be involved in the stimulation of human NK cells by LPS, a component of Gram-negative bacteria, may therefore reflect those that normally occur in vivo. Furthermore, in addition to initiating the responses that control infections, it is likely that LPS also activates homeostatic mechanisms important in controlling immune responses.
Human NK cells have recently been shown to kill autologous dendritic cells derived from human monocytes or CD34- progenitor cells in vitro (42, 43). Adoptive transfer studies in mice have, on the other hand, shown that DC can drive the killing of tumor cells by NK in vivo (44). LPS may therefore not only activate DC to drive proliferation and cytokine production in NK cells, but could also effect DC maturation so that they become targets for autologous NK. Such an interaction between NK and DC could, in turn, have profound effects on T cell responses. Several studies have shown that NK can influence the adaptive immune system and direct the pattern of T cell responses in autoimmune diseases (10, 11). Such effects could be mediated indirectly via effects on DC, or NK cells could influence T cells directly, as has been demonstrated in the DA rat model of experimental autoimmune encephalomyelitis, in which they can kill autologous activated T cells (45). Furthermore, subsets of human NK cells have been shown to produce different cytokines that may, in turn, influence the pattern of an immune response. (28). Activated NK cells could, therefore, using diverse mechanisms, modulate the potential for autoreactivity. Further studies on target cell lysis, the stimulatory vs inhibitory NK-receptor repertoire, and the cytokine profile of LPS-activated CD3-CD56+ cells will address these questions.
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Martin R. Goodier, Kennedy Institute of Rheumatology, 1 Aspenlea Road, Hammersmith, London W6 8LH, U.K.
3 Abbreviations used in this paper: MACS, magnetic cell separation system; CFSE, carboxyfluorescein succinimidyl ester; KIR, killer cell-inhibitory receptor.
Received for publication December 20, 1999. Accepted for publication April 19, 2000.
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