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Lipopolysaccharide Stimulates the Proliferation of Human CD56⁺CD3^- NK Cells: A Regulatory Role of Monocytes and IL-10¹

Kennedy Institute of Rheumatology, London, United Kingdom

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
NK cells recognize and kill tumor cells and normal cells, and these play an important role in immune defense in cancer, infectious disease, and autoimmunity. NK killing is regulated by positive or negative signals derived from the interaction of surface receptors with ligands on the target cells. However, the mechanisms controlling the proliferation and maintenance of NK cells in normal human individuals are less clearly defined. In this study, using an entirely autologous system, we demonstrate that human peripheral blood CD3^-CD56⁺, killer cell-inhibitory receptor (KIR)-expressing cells proliferate and expand in response to LPS. These responses are enhanced in the presence of anti-IL-10 receptor-blocking Abs or on the removal of CD14⁺ cells from the cultures. This enhancement is also reflected in substantial increases in cytolytic activity and IFN- 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

Top Abstract Introduction Materials and Methods Results Discussion References

 
Lipopolysaccharide has pleiotropic effects on many different cell types, particularly those of the myeloid lineage, including monocytes/macrophages and dendritic cells (1, 2). LPS has, more recently, been shown to stimulate human T cells and cause bystander activation of mouse CD8⁺ T cells (3, 4). LPS also prevents cell death in superantigen-activated T cells and enhances the clonal expansion of Ag-specific T cells in mice (5, 6). In this way, LPS is considered as an important link between innate and acquired (Ag-specific) immunity. LPS is thought to stimulate T cells indirectly in all the above models, acting principally on myeloid cells and involving the production of soluble factors such as IFN-, 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.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Preparation and stimulation of PBMC

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.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
LPS induces CD56⁺ cell proliferation

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⁺CFSE^low (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).

View larger version (46K): [in this window] [in a new window]   FIGURE 1. CD56+CD3- cells proliferate in response to LPS. PBMC from a representative donor were labeled with CFSE, as described in Materials and Methods, and either left in medium alone or stimulated with LPS (1 µg/ml) for the number of days shown. Cells were then counterstained with anti-CD56 and anti-CD3 to detect cell division in these populations. Similar results were obtained in three further individuals.

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.

View larger version (17K): [in this window] [in a new window]   FIGURE 2. Expression of KIR receptors on CFSE low (proliferating) cells and CFSE high (nonproliferating) cells. PBMC from three responsive individuals were stimulated for 8 days with LPS. The proportion of CFSE high and low expressing CD56+ or KIR receptor+ cells is shown within PBMC after gating on CD3-negative cells. Abs used were A3 (KIR2DL2), DX27 (anti-KIR2DL2/3 and anti-KIR2DS2), DX9 (anti-KIR3DL1), and DX31 (KIR3DL2). Isotype controls were a combination of mouse IgG1, -2a, and -2b, each at the relevant concentration.

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).

View larger version (48K): [in this window] [in a new window]   FIGURE 3. Opposing effects of neutralizing anti-IL-12 and anti-IL-10 receptor Abs on proliferative responses and the expansion of CD56+ cells. PBMC from two individuals were cultured with LPS (1 µg/ml) in the presence of anti-IL-12, anti-IL-10 receptor, or isotype-matched control mAbs (mouse IgG1 and rat IgG2a for IL-12 and IL-10R, respectively). A, Proliferation was measured after 8 days in cultures stimulated with LPS (solid bars) or left in medium alone. B, Expansion of CD56+ and CD3+ cells was measured on day 8 by FACS analysis in parallel cultures to the proliferation assay. These experiments were performed with an additional four individuals, and consistent results were obtained.

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).

View larger version (47K): [in this window] [in a new window]   FIGURE 4. A, Depletion of CD14+ monocytes enhances LPS-induced proliferation. PBMC or CD14-depleted PBMC were stimulated with LPS, and proliferation was measured between 6 and 7 days of culture. B, Phenotypic analysis of cultures from A was conducted by flow cytometry after 8 days. Similar results were observed in four additional individuals on at least two occasions.

View larger version (39K): [in this window] [in a new window]   FIGURE 5. Depletion of CD14+ cells enhances CD56+ cell proliferation. PBMC or CD14-depleted PBMC were labeled with CFSE, as described in Materials and Methods. Flow-cytometric analysis was conducted on cells before stimulation (day 0, upper panels), and the loss of CFSE in CD56+ cells was assessed after 8 days of stimulation (day 8, lower panels). Similar results were observed in four additional individuals.

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.

View larger version (21K): [in this window] [in a new window]   FIGURE 6. Enhanced cytolytic activity in CD14-depleted PBMC from weak responders after LPS stimulation. PBMC or CD14-depleted PBMC from a high LPS responder (left-hand panels) or a LPS low responder (right-hand panels) were stimulated for 8 days with LPS and used in cytotoxicity assays against 51Cr-labeled K562 target cells (see Materials and Methods). The inset in the left-hand panel shows lytic activity in CD3-CD56+ cells enriched from PBMC of the high responder after LPS stimulation. Enriched CD3-CD56+ cells were titrated starting at an E:T ratio of 8:1.

IFN- production is enhanced in CD14-depleted PBMC

The 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).

View larger version (15K): [in this window] [in a new window]   FIGURE 7. Enhancement of IFN- production in CD14-depleted cultures. PBMC or CD14-depleted PBMC were left unstimulated or stimulated with LPS. Supernatants were collected after 5 days of stimulation and tested for IFN- by ELISA.

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.

View larger version (34K): [in this window] [in a new window]   FIGURE 8. rIL-10 reverses the effect of monocyte depletion. PBMC or CD14-depleted cells from two individuals were stimulated with LPS in the presence or absence of human rIL-10. Proliferation (A) and the proportion of CD3-CD56+ cells (B) were measured after 8 days of culture.

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.

View larger version (26K): [in this window] [in a new window]   FIGURE 9. PCR analysis of IL-12 mRNA. RNA was extracted from PBMC or CD14-depleted PBMC before and after stimulation with LPS. PCR analysis was conducted using IL-12 p35, IL-12 p40, and actin primers, as described in Materials and Methods. Similar results were obtained in two additional individuals.

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).

View larger version (28K): [in this window] [in a new window]   FIGURE 10. Responses of CD14-depleted cells costimulation and MHC class II+ cells, but are B cell independent. PBMC or CD14-depleted PBMC were treated with CTLA-4Ig for the duration of culture with LPS. CD14-depleted PBMC were also further depleted of either B cells or MHC class II+ (HLA-DR) cells before stimulation with LPS. Proliferation and flow-cytometric analysis were performed after 8 days of stimulation. Similar results were obtained in three additional individuals.

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).

View larger version (37K): [in this window] [in a new window]   FIGURE 11. Reconstitution of NK responses in CD14- class II- cells. Depleted cells were cultured alone or reconstituted with 5% monocyte-derived DC before stimulation with LPS. Proliferation and flow-cytometric analysis were assessed after 8 days of stimulation. Proliferation data (cpm) were as follows: Depleted cultures (CD14-, MHC class-II-), 118 ± 14 cpm and 158 ± 31 cpm for unstimulated and LPS stimulated, respectively; DC-reconstituted cultures, 1442 ± 25 cpm and 5880 ± 294 cpm for unstimulated and LPS stimulated, respectively. Similar results were obtained in three additional experiments.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

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.

	Footnotes

 
¹ M.R.G. is funded by the Wellcome Trust. The Kennedy Institute of Rheumatology is funded by the Arthritis Research Campaign.

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.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 

1. Ulevitch, R. J., P. S. Tobias. 1999. Recognition of Gram-negative bacteria and endotoxin by the innate immune system. Curr. Opin. Immunol. 11:19.[Medline]
2. Cella, M., F. Sallusto, A. Lanzavecchia. 1997. Origin, maturation and antigen presenting function of dendritic cells. Curr. Opin. Immunol. 9:10.[Medline]
3. Mattern, T., A. Thanhauser, N. Reiling, K. M. Toellner, M. Duchrow, S. Kusumoto, E. T. Rietschel, M. Ernst, H. Brade, H. D. Flad, et al 1994. Endotoxin and lipid A stimulate proliferation of human T cells in the presence of autologous monocytes. J. Immunol. 153:2996.[Abstract]
4. Tough, D. F., S. Sun, J. Sprent. 1997. T cell stimulation in vivo by lipopolysaccharide (LPS). J. Exp. Med. 185:2089.[Abstract/Free Full Text]
5. Vella, A. T., J. E. McCormack, P. S. Linsley, J. W. Kappler, P. Marrack. 1995. Lipopolysaccharide interferes with the induction of peripheral T cell death. Immunity 2:261.[Medline]
6. Khoruts, A., A. Mondino, K. A. Pape, S. L. Reiner, M. K. Jenkins. 1998. A natural immunological adjuvant enhances T cell clonal expansion through a CD28-dependent, interleukin (IL)-2-independent mechanism. J. Exp. Med. 187:225.[Abstract/Free Full Text]
7. Zubler, R. H., C. Werner-Favre, L. Wen, K. Sekita, C. Straub. 1987. Theoretical and practical aspects of B-cell activation: murine and human systems. Immunol. Rev. 99:281.[Medline]
8. Bancroft, G. J.. 1993. The role of natural killer cells in innate resistance to infection. Curr. Opin. Immunol. 5:503.[Medline]
9. Trinchieri, G.. 1989. Biology of natural killer cells. Adv. Immunol. 47:187.[Medline]
10. Horwitz, D. A., J. D. Gray, K. Ohtsuka, M. Hirokawa, T. Takahashi. 1997. The immunoregulatory effects of NK cells: the role of TGF-ß and implications for autoimmunity. Immunol. Today 18:538.[Medline]
11. Kos, F. J.. 1998. Regulation of adaptive immunity by natural killer cells. Immunol. Res. 17:303.[Medline]
12. Lanier, L. L.. 1998. NK cell receptors. Annu. Rev. Immunol. 16:359.[Medline]
13. Moretta, A., L. Moretta. 1997. HLA class I specific inhibitory receptors. Curr. Opin. Immunol. 9:694.[Medline]
14. Long, E. O.. 1999. Regulation of immune responses through inhibitory receptors. Annu. Rev. Immunol. 17:875.[Medline]
15. Gately, M. K., B. B. Desai, A. G. Wolitzky, P. M. Quinn, C. M. Dwyer, F. J. Podlaski, P. C. Familletti, F. Sinigaglia, R. Chizonnite, U. Gubler, et al 1991. Regulation of human lymphocyte proliferation by a heterodimeric cytokine, IL-12 (cytotoxic lymphocyte maturation factor). J. Immunol. 147:874.[Abstract]
16. Naume, B., M. Gately, T. Espevik. 1992. A comparative study of IL-12 (cytotoxic lymphocyte maturation factor)-, IL-2-, and IL-7-induced effects on immunomagnetically purified CD56⁺ NK cells. J. Immunol. 148:2429.[Abstract]
17. Testa, U., A. Care, E. Montesoro, C. Fossati, G. Giannella, R. Masciulli, M. Fagioli, D. Bulgarini, D. Habetswallner, G. Isacchi, et al 1990. Interleukin-2-dependent long-term cultures of low-density lymphocytes allow the proliferation of lymphokine-activated killer cells with natural killer, Ti / or TNK phenotype. Cancer Immunol. Immunother. 31:11.[Medline]
18. Schmidt-Wolf, I. G., R. S. Negrin, H. P. Kiem, K. G. Blume, I. L. Weissman. 1991. Use of a SCID mouse/human lymphoma model to evaluate cytokine-induced killer cells with potent antitumor cell activity. J. Exp. Med. 174:139.[Abstract/Free Full Text]
19. Satoh, M., S. Seki, W. Hashimoto, K. Ogasawara, T. Kobayashi, K. Kumagai, S. Matsuno, K. Takeda. 1996. Cytotoxic or ß T cells with a natural killer cell marker, CD56, induced from human peripheral blood lymphocytes by a combination of IL-12 and IL-2. J. Immunol. 157:3886.[Abstract]
20. Meszaros, K., S. Aberle, M. White, J. B. Parent. 1995. Immunoreactivity and bioactivity of lipopolysaccharide-binding protein in normal and heat-inactivated sera. Infect. Immun. 63:363.[Abstract]
21. Soderstrom, K., B. Corliss, L. L. Lanier, J. H. Phillips. 1997. CD94/NKG2 is the predominant inhibitory receptor involved in recognition of HLA-G by decidual and peripheral blood NK cells. J. Immunol. 159:1072.[Abstract]
22. Wells, A. D., H. Gudmundsdottir, L. A. Turka. 1997. Following the fate of individual T cells throughout activation and clonal expansion: signals from T cell receptor and CD28 differentially regulate the induction and duration of a proliferative response. J. Clin. Invest. 100:3173.[Medline]
23. De Waal Malefyt, R., C. G. Figdor, R. Huijbens, S. Mohan-Peterson, B. Bennett, J. Culpepper, W. Dang, G. Zurawski, J. E. de Vries. 1993. Effects of IL-13 on phenotype, cytokine production, and cytotoxic function of human monocytes: comparison with IL-4 and modulation by IFN- or IL-10. J. Immunol. 151:6370.[Abstract]
24. Kang, K., M. Kubin, K. D. Cooper, S. R. Lessin, G. Trinchieri, A. H. Rook. 1996. IL-12 synthesis by human Langerhans cells. J. Immunol. 156:1402.[Abstract]
25. Kotake, S., Jr H. R. Schumacher, R. L. Wilder. 1996. A simple nested RT-PCR method for quantitation of the relative amounts of multiple cytokine mRNAs in small tissue samples. J. Immunol. Methods 199:193.[Medline]
26. Mattern, T., H. D. Flad, L. Brade, E. T. Rietschel, A. J. Ulmer. 1998. Stimulation of human T lymphocytes by LPS is MHC unrestricted, but strongly dependent on B7 interactions. J. Immunol. 160:3412.[Abstract/Free Full Text]
27. De Waal Malefyt, R., J. Abrams, B. Bennett, C. G. Figdor, J. E. de Vries. 1991. Interleukin 10 (IL-10) inhibits cytokine synthesis by human monocytes: an autoregulatory role of IL-10 produced by monocytes. J. Exp. Med. 174:1209.[Abstract/Free Full Text]
28. Peritt, D., S. Robertson, G. Gri, L. Showe, M. Aste-Amezaga, G. Trinchieri. 1998. Differentiation of human NK cells into NK1 and NK2 subsets. J. Immunol. 161:5821.[Abstract/Free Full Text]
29. Hayes, M. P., F. J. Murphy, P. R. Burd. 1998. Interferon--dependent inducible expression of the human interleukin-12 p35 gene in monocytes initiates from a TATA-containing promoter distinct from the CpG-rich promoter active in Epstein-Barr virus-transformed lymphoblastoid cells. Blood 91:4645.[Abstract/Free Full Text]
30. Verhasselt, V., C. Buelens, F. Willems, D. De Groote, N. Haeffner-Cavaillon, M. Goldman. 1997. Bacterial lipopolysaccharide stimulates the production of cytokines and the expression of costimulatory molecules by human peripheral blood dendritic cells: evidence for a soluble CD14-dependent pathway. J. Immunol. 158:2919.[Abstract]
31. Ito, T., M. Inaba, K. Inaba, J. Toki, S. Sogo, T. Iguchi, Y. Adachi, K. Yamaguchi, R. Amakawa, J. Valladeau, et al 1999. A CD1a⁺/CD11c⁺ subset of human blood dendritic cells is a direct precursor of Langerhans cells. J. Immunol. 163:1409.[Abstract/Free Full Text]
32. Naume, B., A. C. Johnsen, T. Espevik, A. Sundan. 1993. Gene expression and secretion of cytokines and cytokine receptors from highly purified CD56⁺ natural killer cells stimulated with interleukin-2, interleukin-7 and interleukin-12. Eur. J. Immunol. 23:1831.[Medline]
33. Geldhof, A. B., M. Moser, L. Lespagnard, K. Thielemans, P. De Baetselier. 1998. Interleukin-12-activated natural killer cells recognize B7 costimulatory molecules on tumor cells and autologous dendritic cells. Blood 91:196.[Abstract/Free Full Text]
34. Wilson, J. L., J. Charo, A. Martin-Fontecha, P. Dellabona, G. Casorati, B. J. Chambers, R. Kiessling, M. T. Bejarano, H. G. Ljunggren. 1999. NK cell triggering by the human costimulatory molecules CD80 and CD86. J. Immunol. 163:4207.[Abstract/Free Full Text]
35. Walker, W., M. Aste-Amezaga, R. A. Kastelein, G. Trinchieri, C. A. Hunter. 1999. IL-18 and CD28 use distinct molecular mechanisms to enhance NK cell production of IL-12-induced IFN-. J. Immunol. 162:5894.[Abstract/Free Full Text]
36. Galea-Lauri, J., D. Darling, S. U. Gan, L. Krivochtchapov, M. Kuiper, J. Gaken, B. Souberbielle, F. Farzaneh. 1999. Expression of a variant of CD28 on a subpopulation of human NK cells: implications for B7-mediated stimulation of NK cells. J. Immunol. 163:62.[Abstract/Free Full Text]
37. Heufler, C., F. Koch, U. Stanzl, G. Topar, M. Wysocka, G. Trinchieri, A. Enk, R. M. Steinman, N. Romani, G. Schuler. 1996. Interleukin-12 is produced by dendritic cells and mediates T helper 1 development as well as interferon- production by T helper 1 cells. Eur. J. Immunol. 26:659.[Medline]
38. Siegal, F. P., N. Kadowaki, M. Shodell, P. A. Fitzgerald-Bocarsly, K. Shah, S. Ho, S. Antonenko, Y. J. Liu. 1999. The nature of the principal type 1 interferon-producing cells in human blood. Science 284:1835.[Abstract/Free Full Text]
39. Cella, M., D. Jarrossay, F. Facchetti, O. Alebardi, H. Nakajima, A. Lanzavecchia, M. Colonna. 1999. Plasmacytoid monocytes migrate to inflamed lymph nodes and produce large amounts of type I interferon. Nat. Med. 5:919.[Medline]
40. Robinson, S. P., S. Patterson, N. English, D. Davies, S. C. Knight, C. D. Reid. 1999. Human peripheral blood contains two distinct lineages of dendritic cells. Eur. J. Immunol. 29:2769.[Medline]
41. Biron, C. A.. 1997. Activation and function of natural killer cell responses during viral infections. Curr. Opin. Immunol. 9:24.[Medline]
42. Carbone, E., G. Terrazzano, G. Ruggiero, D. Zanzi, A. Ottaiano, C. Manzo, K. Karre, S. Zappacosta. 1999. Recognition of autologous dendritic cells by human NK cells. Eur. J. Immunol. 29:4022.[Medline]
43. Wilson, J. L., L. C. Heffler, C. Charo, A. Scheynius, M.-T. Bejarano, H.-G. Ljunggren. 1999. Targeting of human dendritic cells by autologous NK cells. J. Immunol. 163:6365.[Abstract/Free Full Text]
44. Fernandez, N. C., A. Lozier, C. Flament, P. Ricciardi-Castagnoli, D. Bellet, M. Suter, M. Perricaudet, T. Tursz, E. Maraskovsky, L. Zitvogel. 1999. Dendritic cells directly trigger NK cell functions: cross-talk relevant in innate anti-tumor immune responses in vivo. Nat. Med. 5:405.[Medline]
45. Smeltz, R. B., N. A. Wolf, R. H. Swanborg. 1999. Inhibition of autoimmune T cell responses in the DA rat by bone marrow-derived NK cells in vitro: implications for autoimmunity. J. Immunol. 163:1390.[Abstract/Free Full Text]

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