Innate immune responses provide the host with its first line of defense against infections. Signals generated by subsets of lymphocytes, including NK cells, NKT cells, and APC during this early host response determine the nature of downstream adaptive immune responses. In the present study, we have examined the role of innate NK cells in an autoimmune model through the use of primary immunization with the myelin oligodendrocyte glycoprotein peptide to induce experimental autoimmune encephalomyelitis (EAE). Our studies have shown that in vivo depletion of NK cells can affect the adaptive immune responses, because NK cells were found to regulate the degree of clinical paralysis and to alter immune adaptive responses to the myelin oligodendrocyte glycoprotein peptide. The requirement for NK cells was reflected by changes in the T cell responses and diminished clinical disease seen in mice treated with anti-NK1.1, anti-asialo GM1, and selected Ly49 subtype-depleted mice. In addition to alteration in T cell responses, the maturational status of dendritic cells in lymph nodes was altered both quantitatively and qualitatively. Finally, examination of TCR Vβ usage of the brain lymphocytes from EAE mice indicated a spectra-type change in receptor expression in NK- depleted mice as compared with non-NK-depleted EAE mice. These findings further establish a recently postulated link between NK cells and the generation of autoreactive T cells.

Increasing evidence has emerged regarding the potential role for NK cells in regulating adaptive immune responses. Early studies in the CMV infection models clearly demonstrated the important regulatory role of NK cells in the CD8-mediated T cell responses (1, 2). In addition to a role in controlling viral infections, NK cells producing IFN-γ have recently been shown to also regulate generation of T cell immunity (3) in a parasitic infection (Toxoplasma gondii). This innate-adaptive interface has been shown to involve a role for NK cells in the regulation of dendritic cell (DC)4 maturation in both humans (4, 5) and mice (5, 6).

In many autoimmune diseases, lymphocytes have been shown to be the effector cells responsible for damage to the tissue target. T cells and B cells, as part of the acquired arm of immunity, have long been known as key inducers of a variety of autoimmune diseases. T cells and B cells have been implicated in uveitis, hemolytic anemia, colitis, myasthenia gravis, lupus, and rheumatoid arthritis among other diseases (7, 8, 9, 10, 11, 12). NK cells, being part of the innate immune system, have been implicated in such autoimmune diseases as diabetes and insulitis, as well as rheumatoid arthritis (13, 14, 15). Experimental autoimmune encephalomyelitis (EAE) is a prototypic autoimmune disease induced in laboratory animals, bearing significant similarities to multiple sclerosis in clinical and histopathological aspects (16, 17). EAE is known to be mediated by CD4+ T cells that recognize peptides derived from encephalitogenic proteins of the CNS. Cytokines, particularly TNF-α, are considered to be the mediators of the pathology that is observed in the CNS with conflicting analyses of their effects reported (18). In the mouse, the disease is characterized by a paralysis proceeding from the hind limbs to the forelimbs. Paralysis initiates within 2 wk of injection of a myelin oligodendrocyte glycoprotein (MOG) peptide (19, 20).

NK cells are known to be a first line of defense in viral infections. In addition, previous studies have suggested that both NK or NK1.1+ T (NKT) cells serve as regulatory cells in some T cell-mediated experimental autoimmune diseases, including murine models of encephalomyelitis (EAE) (21, 22), colitis (23), and diabetes (24). Studies by Ljunggren and colleagues (25, 26) have detailed the contribution of NK cells in primary B cell-mediated autoimmunity. Autoantibodies produced by B cells are the primary cause of disease in a variety of autoimmune conditions, including hemolytic anemia, thyroiditis, stiff man syndrome, pemphigus vulgaris, and systemic lupus erythematosis. Shi et al. (27) demonstrated that MOG failed to induce EAE in IL-18−/− mice. EAE could be observed upon IL-18 administration but disease required the presence of IFN-γ-producing NK cells. These findings established an important, unrecognized link between NK cells and autoimmunity in a primary in vivo model system. More recently, mice lacking CX3CR1 that developed MOG-induced EAE demonstrated a reduced recruitment of NK cells to the CNS (28).

In a another report focused on the possible role of NK cells in the murine EAE model, Zhang et al. (21) noted that in vivo NK cell depletion resulted in exacerbation of clinical symptoms in wild-type C57BL/6 (B6) mice. In addition, these investigators also found that depletion of NK cells resulted in an increased severity of symptoms when disease was induced by passive transfer of a MOG-specific T cell line. However, in this transfer system, it was difficult to directly examine the contribution of the endogenous innate immune system. Analyses of results from such adoptive transfer experiments could be further complicated by the fact that NK cells are known to reject autologous as well as allogeneic cell transfers, potentially resulting in fewer effector cells that are responsible for initiating the histopathological effects on the CNS. These transfer model systems may, in fact, be measuring the effectiveness of cell transfer rather than a role for NK cells in the actual disease pathology.

The results of many of these studies led to the hypothesis that NK cells appear to be involved in changing the balance of immunity by initiating or regulating the intensity of autoimmune reactions and/or modifying the effector cells that can accumulate in the target organ(s). These mechanisms might include: 1) production of cytokines that alter DC or T cell activation and/or proliferation; 2) direct interactions with APC that could alter Ag presentation; and 3) alteration of regulatory cells by direct or indirect mechanisms. The primary EAE mouse model provides an in vivo system to examine the regulatory role of NK cells because they may provide either protection from or exacerbation of the clinical course of autoimmune disease(s). In addition, the ability to selectively deplete NK cell subsets in a primary model allows for a more specific interpretation of the role of NK cells in the initiation of autoimmune disease. In this study, we examined whether NK cells could serve as a regulatory element in primary, peptide-induced EAE.

Cells were isolated from spleen, liver, lymph nodes (LN; axial, inguinal, mesenteric, and cervical), and perfused brain of C57BL/6 mice to examine cell phenotype and cell functions. Liver cells were obtained as previously described (29). LN cells were obtained by gentle dissociation of the organ through a plastic mesh screen in ice-cold medium containing 10% FCS. Brain lymphocytes were obtained by anesthetizing the animal and perfusing saline into the left ventricle of the heart using a peristaltic pump. The right atrium is clipped to allow release of vascular volume. Following complete vascular perfusion and euthanasia, brains were harvested and lymphocytes were isolated following the methodologies used to isolate lymphocytes from the liver tissue. The use of animals for this study was in compliance with policies regarding the humane treatment and care of animals and was approved by the National Cancer Institute-Frederick Animal Care and Use Committee. Animal care was provided in accordance with the procedures outlined in the “Guide for the Care and Use of Laboratory Animals (National Institutes of Health Publication no. 86-23, 1985).

The mAbs 4E5 (Ly49D), 3D10 (Ly49H), and 3A10 (NKG2D) were provided by Dr. W. Yokoyama (Washington University, St. Louis, MO). The mAb 1F8 (Ly49C/I/H) was a gift from Dr. M. Bennett (University of Texas Southwestern Medical Center, Dallas, TX). Fluorochrome-labeled isotype- specific controls were purchased from BD Biosciences and BD Pharmingen along with fluorochrome-labeled Abs to Ly49D, NK1.1, CD4, CD3ε, and CD80. Fluorochrome-labeled Ab to CD86 was purchased from eBioscience. Rabbit F(ab′)2 anti-rat IgG was used as a cross-linking reagent (MP Biomedicals). Anti-asialo GM1 was purchased from Wako Pure Chemical Industries (Japan) and unlabeled anti-NK1.1 and 5E6 (Ly49C/I) were prepared by Hazleton Laboratories (Falls Church, VA). Unlabeled anti-CD3ε was purchased from BD Biosciences and BD Pharmingen. Abs were used for in vivo depletions of cell subsets, flow cytometric, or functional studies. Intracellular IFN-γ detection was performed using Ab kits purchased from BD Biosciences and BD Pharmingen.

Cells were stained as previously described (29) and analyzed on an LSR flow cytometer (BD Biosciences) or a FACSort flow cytometer retrofitted with a solid-state 635-nm laser (BD Biosciences). Data were analyzed using either CellQuest software (BD Bisociences) or FCSExpress2 software (DeNovo Software).

Cytokines were measured using ELISA kits (R&D Systems). Cell stimulations were performed in a 24-well Costar (Corning) plate at a cell concentrations of 1 × 106 cells/ml. Cells were stimulated with MOG peptide (1–10 μg/106 cells) or anti-CD3 (2–5 μg/106 cells; BD Biosciences and BD Pharmingen) plus IL-2 (100 IU/ml; Hoffmann-LaRoche), or MOG peptide (2 μg/106 cells). Unless otherwise stated, samples were collected after overnight or 72-h incubation (37°C, 5% CO2) and were measured in duplicate against the standard curve of the assay and reported as pg/ml. In all assays, the SD was <5 pg/ml.

The multiprobe RNase Protection Assay was performed using the mck-1 or mck-5 template set (BD Pharmingen). Total cellular RNA was extracted using TRIzol (Invitrogen Life Technologies) and 1–5 μg of total mRNA was hybridized with a [33P]UTP-labeled RNA probe (1 × 106 cpm/sample) prepared according to the manufacturer’s directions using the BD Pharmingen RiboQuant In Vitro Transcription Kit. Following hybridization, the samples were treated with RNase A and T1 according to the procedure provided by BD Pharmingen. The RNase was inactivated and precipitated using a master mixture containing 200 μl of Ambion RNase inactivation reagent, 50 μl of ethanol, 5 μg of yeast tRNA, and 1 μl of Ambion GycoBlue coprecipitate per RNA sample. The samples were mixed well, incubated at −70°C for 15 min and centrifuged at 20,800 × g for 15 min at room temperature. The pellets were suspended in 3 μl of BD Pharmingen sample buffer and subjected to PAGE as recommended by the manufacturer (BD Pharmingen).

The MOG35–55 (MEVGWYRSPFSRVVHLYRNGK) was commercially synthesized to 95% purity by HPLC (Invitrogen Life Technologies and Advanced ChemTech). CFA was purchased from MP Biomedicals. Heat-killed Mycobacterium tuberculosis H37Ra was purchased from VWR. Pertussis toxin (PT) was purchased from List Biological Laboratories. IL-2 manufactured by Hoffmann-LaRoche was received from the BRB Preclinical Repository at the National Cancer Institute (NCI-Frederick, Frederick, MD).

RPMI 1640 was purchased from Invitrogen Life Technologies and supplemented with l-glutamine, penicillin-streptomycin, and 10% FCS. Saline was purchased from Baxter Healthcare.

For induction of active EAE, mice were injected s.c. in one flank with 200 μl of an emulsion containing 100 μl of CFA, 200 μg of MOG35–55, and 300 μg of pulverized heat-killed M. tuberculosis in saline solution. On the same day, the mice were injected i.p. with 400 ng of PT in 200 μl of saline and this treatment was repeated 48 h later. A booster immunization with an identical emulsion was given 1 wk later in the opposite flank with no PT. Onset of paralysis was anticipated to occur 3–7 days after the booster immunization.

Following immunization, mice were monitored daily for clinical signs of EAE. The clinical grade was a modification of that which was previously reported: 0, no clinical signs; 1, complete loss of tail tonicity; 2, flaccid tail and abnormal gait ataxia and/or paresis of hind limb; 3, hind limb paralysis; 4, hind limb paralysis with foreleg involvement; and 5, death (21).

Mice were i.p. injected with 100–200 μg of anti-NK1.1 mAb (PK136), anti-asialo GM1 (dilution 1/20; 0.2 ml), or 100 μg of anti-Ly49 Abs 1–3 days before first immunization with the emulsion containing the MOG35–55 peptide.

To analyze primary T cell responses, 1 × 105 LN cells isolated from mice immunized with MOG35–55 peptide were cultured in 96-well flat-bottom plates with MOG35–55 peptide at a concentration of 2 μg/ml in 0.2 ml of RPMI 1640 medium supplemented with 10% FCS. The plates were incubated for 72 h at 37°C in humidified air containing 5% CO2. Incorporation of [3H]thymidine (1μCi/well) for the final 18 h of the incubation was measured on a Trilux 1450 Microbeta liquid scintillation counter following harvest on a TomTec Harvester 96 Mach 3 (PerkinElmer).

TCR Vβ usage was analyzed following previously published methods (30). Briefly, mRNA from lymphocytes isolated from draining, nondraining, and cervical LN and perfused brain was extracted using TRIzol (Invitrogen Life Technologies) according to the manufacturer’s protocol. TCR Vβ products were synthesized using the Qiagen One-Step RT-PCR kit and primers unique for the 5′ end of Vβ1, 6, 8.1, 8.2, 8.3, 14, and 15 as well as a common 3′ primer (Qiagen Operon). Primer sequences have been previously published (30).

Quantitative PCR analysis was performed using SYBR Green Chemistry (Qiagen) according to the manufacturer’s instructions in 10-μl final volumes in 384-well microtiter plates. Specific primers for detection of TCR Vβ usage of tissue lymphocytes in EAE-immunized mice were defined as described above and were purchased from Qiagen. The endogenous control primer MuGAPDH was purchased from Applied Biosystems and the sequence is proprietary in nature. Thermocycling conditions using an Applied Biosystems 7900 SDS were as follows; 95°C for 15 min and 40 cycles of 95°C for 15 s and 60°C for 1 min. Accurate quantification of each mRNA was achieved using the normalization of the sample ΔCT values to one reference. This value, referred to as the ΔCT − sample value (ΔCT − sample = CT − reference), is derived by taking the result of the expression: if 2(−ΔCT) − 1 > 0, then the result = 2(−ΔCT) − 1 or else the result = −1/2(-ΔCT). This equation changes the range for down-regulation from 0 through 1 to -∞ through 0 and up-regulation from 1 through ∞ to 0 through ∝. The samples that were assayed for expression were normalized to murine GAPDH and amplifications were conducted under identical conditions for each gene of interest. The target mRNA expression was normalized to the GAPDH expression, and the relative expression was calculated back to the EAE controls for each cell type. Raw data from each quantitative PCR run were exported into a comparative CT analysis workbook. CT represents the threshold cycle or the PCR cycle at which an increase in reporter fluorescence above baseline signal can be detected. The comparative CT workbook allows for normalization with different endogenous controls on a number of samples and genes. Each graph displays the analyzed results in a format (both numerically and graphically) showing their expression relative to not only an endogenous control but also a reference sample.

We chose to examine whether NK cells might play an important role in the development of autoimmunity by using a primary EAE model. The C57BL/6 (B6) model is particularly useful, since the model for EAE is well established in this mouse strain (31). Additionally, several Abs are routinely used in this strain for in vivo NK cell depletion because Abs to various NK subsets are available and various gene knockout mice have been generated on the B6 background (32, 33, 34). Given the availability of an autoimmune model and specific NK reagents, we theorized that an alteration in the in vivo NK cell content would alter the adaptive responses to the MOG35–55 peptide responsible for initiation of disease in the B6 mouse. The use of a primary immune model is in contrast to studies that used adoptive transfer models or T cell lines (35, 36) to examine the role of NK cells in the development of EAE. Examination of the LN as the primary T and NK cell interactive site for initiation of an immune response resulting in EAE required verification of the presence of a NK population within the LN. Preliminary phenotyping (Fig. 1) of normal B6 LN indicated the consistent presence of 0.5–1.0% NK1.1+CD3 cells. These cells could be depleted with anti-NK1.1 or asialo GM1 Ab with 90% suppression maintained at day 7 and recovery of the population in the LN by day 14. The spleen and liver demonstrated similar kinetics of suppression (data not shown). In addition to pan-NK-depleting Abs, specific subsets of Ly49D (Fig. 1,G) and Ly49H (Fig. 1,H) NK cells could be deleted efficiently from mice in vivo, relative to control mice (Fig. 1 F).

FIGURE 1.

In vivo depletion of draining LN NK cells. A–D, Mice were treated with control Ig (A and B) or anti-NK1.1 (C and D). Mice were evaluated for the NK cell content of their draining LN on day 7 (A and C) and day 14 (B and D). Subsets of Ly49-activating receptors were evaluated by examination of CD3NK1.1+ cells (E; gate r2) and evaluating the expression of Ly49D and/or Ly49H (F). In vivo depletion of these subsets is shown in LN 3 days after Ab administration of anti-Ly49D (G) or anti-Ly49H (H).

FIGURE 1.

In vivo depletion of draining LN NK cells. A–D, Mice were treated with control Ig (A and B) or anti-NK1.1 (C and D). Mice were evaluated for the NK cell content of their draining LN on day 7 (A and C) and day 14 (B and D). Subsets of Ly49-activating receptors were evaluated by examination of CD3NK1.1+ cells (E; gate r2) and evaluating the expression of Ly49D and/or Ly49H (F). In vivo depletion of these subsets is shown in LN 3 days after Ab administration of anti-Ly49D (G) or anti-Ly49H (H).

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To examine the clinical impact of a change in NK cell content during the EAE immunization/induction schedule, NK depletions were done either before initial immunization of MOG peptide or just before secondary immunization (at day 6). Fig. 2 shows the clinical score (Fig. 2), survival (Fig. 2), and frequency of disease (Fig. 2) for mice depleted of NK cells before primary immunization. EAE control mice that received either nothing, mIgG2a (Cntl), or normal rabbit serum (NRS; data not shown) generated a rapid clinical disease, starting about day 15 that peaked at days 22–25, and was generally maintained for 30 additional days. Mice depleted of NK cells with either NK1.1 or asialo GM1 Abs exhibited significantly (p < 0.05) reduced average clinical scores; however, the time to onset of clinical disease was consistently delayed by 2–5 days in the non-NK-depleted mice. Clinical manifestations of EAE in the mice tended to fade slightly after 30–35 days, with variability from experiment to experiment. Survival in both NK-depleted groups was significantly higher than the EAE control group; however, the frequency of the number of animals displaying disease was not statistically different. Mice depleted of NK cells after primary, but before secondary immunization, failed to demonstrate any consistent differences in parameters shown in Fig. 2, A–C (data not shown). Thus, depletion of NK cells before immunization with MOG peptide decreased both the frequency and extent of disease compared with the EAE control mice. Also, any contribution of NKT cells can be ruled out since similar results were obtained with both anti-NK1.1 and asialo GM1 (Fig. 2 A), as the latter treatment (in normal mice) does not remove NKT cells.

FIGURE 2.

In vivo EAE after NK cell or subset depletion. EAE was measured by clinical score (A and D), number of mice surviving (B and E), or disease incidence (C and F) in C57BL/6 mice. A–C, The mice were treated with control IgG (▴), depleted of NK cells with anti-NK1.1 (PK136; •), or anti-asialo GM1 (▾), whereas in D–F, mice were depleted of NK subsets with anti-Ly49D (4E5; ▵), anti-Ly49C/I/H (1F8; ♦), or anti-Ly49C/I (5E6; □). Values represent mean and SD of 10 mice/group and are representative of more than three experiments. Abs were given to mice i.p. 1–3 days before immunization.

FIGURE 2.

In vivo EAE after NK cell or subset depletion. EAE was measured by clinical score (A and D), number of mice surviving (B and E), or disease incidence (C and F) in C57BL/6 mice. A–C, The mice were treated with control IgG (▴), depleted of NK cells with anti-NK1.1 (PK136; •), or anti-asialo GM1 (▾), whereas in D–F, mice were depleted of NK subsets with anti-Ly49D (4E5; ▵), anti-Ly49C/I/H (1F8; ♦), or anti-Ly49C/I (5E6; □). Values represent mean and SD of 10 mice/group and are representative of more than three experiments. Abs were given to mice i.p. 1–3 days before immunization.

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Because total NK depletion altered the course of disease, we sought to determine whether Ly49 NKR-bearing subsets of NK cells might demonstrate a similar alteration. Thus, Abs to Ly49C/I (present on ∼50% of NK cells), Ly49D (present on 25–30% of NK cells), and Ly49H (present on 25–30% of NK cells) were used to deplete NK subsets. Preliminary phenotypic screening verified that the LN NK cells expressed normal patterns of activating and inhibitory NKRs (Fig. 1) as compared with spleen and liver (data not shown). The preliminary screening confirmed that a selective loss of the Ly49D NKR was observed when mice were treated with Abs to Ly49D or Ly49H (3D10). Thus, our NKR depletion can effectively alter NKR- bearing subsets in the LN, the site where immunity to MOG is being induced. Elimination of activating NKRs Ly49D and Ly49H resulted in a more efficient alteration of clinical score (Fig. 2,D) that was comparable to elimination of the entire NK population or the inhibitory NKRs Ly49C/I and Ly49G2 (data not shown; but similar to NK1.1). These activating NKR-depleting Abs not only diminished the clinical score but altered the frequency of disease (Fig. 2,F) and improved mouse survival (Fig. 2 E). These data suggested that more effective disease alteration was coincident with the activating Ly49-bearing NK subsets. In addition, it is unlikely that these Ab treatments might be altering other leukocyte subsets, since these activating NKRs are not expressed on T cells or monocytes/macrophages (37, 38).

Data in Fig. 2 suggest that Ly49D- and Ly49H-bearing subsets might be critical for EAE development. To further evaluate this result, we used a specific anti-Ly49H Ab (3D10) for depletion. Due to the major overlap between Ly49D and Ly49H (coexpressed by ∼75–80% of NK cells), we examined how critical this Ly49 NKR was in the development of EAE. Table I compares clinical scores, percent survival, and percentage of mice with EAE between B6 control mice and mice that lack the Ly49H NK cell subset. We found that the removal of the Ly49H-expressing cells with Ab resulted in reduced clinical scores. This alteration of autoimmunity was also observed in the frequency of disease, where only 40–50% of mice depleted with Ly49H Abs developed disease and had a 90% survival outcome compared with 100% of the EAE control mice developing disease with significantly diminished survival. Thus, collectively, these data are highly indicative of the specificity of NK cells for altering the clinical course of disease with Ly49 receptors playing a role.

Table I.

Role of Ly49H in NK-mediated modulation of EAE

StrainMean Clinical Score at Day 24 (SE)% SurvivalNo. Surviving% with EAENo. with EAE
Expt. 1      
C57BL/6 2.4 (0.4) 93 13/14 93 13/14 
C57BL/6 anti-Ly49H 1.5 (0.5)a 100 10/10 90 9/10 
C57BL/6 anti-NK1.1 1.4 (0.3)a 100a 10/10a 90a 8/10a 
StrainMean Clinical Score at Day 24 (SE)% SurvivalNo. Surviving% with EAENo. with EAE
Expt. 1      
C57BL/6 2.4 (0.4) 93 13/14 93 13/14 
C57BL/6 anti-Ly49H 1.5 (0.5)a 100 10/10 90 9/10 
C57BL/6 anti-NK1.1 1.4 (0.3)a 100a 10/10a 90a 8/10a 
a

Values are significantly different from those of C57BL/6 control.

To further evaluate the alterations in adaptive immunity in NK-depleted mice, draining LN cells from EAE or control mice were evaluated for a T cell proliferative response (Fig. 3,a) and for production of either Th1/2 (IL-2, IL-4) or inflammatory cytokines (IFN-γ, TNF-α, IL-6, IL-10) 10 days after immunization (Table II). Cells were unstimulated or stimulated in vitro with 2 μg/ml MOG peptide or anti-CD3 plus IL-2 (as a positive control). Table II shows that although each group of mice had intact proliferative responses to CD3 and IL-2, mice depleted of NK cells with Ab treatment with anti-NK1.1, anti-Ly49H, anti-Ly49D, or anti-asialo GM1 (Fig. 3 a) demonstrated a profound deficit in proliferative capacity in response to MOG peptide. Mice depleted of other NKR-bearing subsets (Abs to Ly49C/I or Ly49G2) also had diminished proliferative responses but exhibited more variable results (data not shown).

FIGURE 3.

Responses of draining LN to MOG peptide. A, Proliferation of draining LN cells from MOG-immunized mice (day 15 following first immunization) were cultured with 2–25 μg/ml MOG peptide (), anti-CD3 (2 μg/ml), and IL-2 (100 U/ml) (▩) or nothing (control (Cntl); ▪). Normal mice, EAE control mice, and EAE mice treated with anti-NK1.1, anti-asialo GM1, anti-Ly49H, and anti-Ly49D were evaluated. Cells were stimulated for 48 h, then pulsed with [3H]TdR (1 μCi/well) for 18 h. B, Frequency of cytokine production in CD4+ T cells from draining LN cells. Cells used in B were evaluated for intracellular production of TNF-α and IFN-γ after 8 h of stimulation with MOG peptide. Cells were stimulated with 2 μg/ml MOG peptide for 2 h, then brefeldin A was added for the remaining 6 h. Values are pools of three to five mice per treatment. C, Cytokine production in CNS leukocytes and peripheral LN. Panel shows the production of IL-2 after 18 h in either brain lymphocytes (Lymphs) or draining LN cells after stimulation with 2 μg/ml MOG peptide in naive C57BL/6 mice (▪), EAE control mice (), and EAE NK1.1-depleted (Dep., dep., Depl.) mice (). D, IFN-γ mRNA expression at 6 h in cells isolated from draining, nondraining, or cervical lymph nodes. Values are pools of three to five mice per treatment. The autoradiograph is a representative example of the RNase protection results. Lymphs, Lymphocytes.

FIGURE 3.

Responses of draining LN to MOG peptide. A, Proliferation of draining LN cells from MOG-immunized mice (day 15 following first immunization) were cultured with 2–25 μg/ml MOG peptide (), anti-CD3 (2 μg/ml), and IL-2 (100 U/ml) (▩) or nothing (control (Cntl); ▪). Normal mice, EAE control mice, and EAE mice treated with anti-NK1.1, anti-asialo GM1, anti-Ly49H, and anti-Ly49D were evaluated. Cells were stimulated for 48 h, then pulsed with [3H]TdR (1 μCi/well) for 18 h. B, Frequency of cytokine production in CD4+ T cells from draining LN cells. Cells used in B were evaluated for intracellular production of TNF-α and IFN-γ after 8 h of stimulation with MOG peptide. Cells were stimulated with 2 μg/ml MOG peptide for 2 h, then brefeldin A was added for the remaining 6 h. Values are pools of three to five mice per treatment. C, Cytokine production in CNS leukocytes and peripheral LN. Panel shows the production of IL-2 after 18 h in either brain lymphocytes (Lymphs) or draining LN cells after stimulation with 2 μg/ml MOG peptide in naive C57BL/6 mice (▪), EAE control mice (), and EAE NK1.1-depleted (Dep., dep., Depl.) mice (). D, IFN-γ mRNA expression at 6 h in cells isolated from draining, nondraining, or cervical lymph nodes. Values are pools of three to five mice per treatment. The autoradiograph is a representative example of the RNase protection results. Lymphs, Lymphocytes.

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Table II.

Cytokine release after in vitro stimulation

Immunization CytokineaNormal ControlFirst ImmunizationFirst and NK Depleted
TNF-α 899 189 
IFN-γ 2203 468 
IL-6 229 82 
IL-12p70 29 21 
IL-10 16 32 44 
IL-2 100 23 
IL-4 <10 <10 <10 
IL-5 <10 <10 <10 
Immunization CytokineaNormal ControlFirst ImmunizationFirst and NK Depleted
TNF-α 899 189 
IFN-γ 2203 468 
IL-6 229 82 
IL-12p70 29 21 
IL-10 16 32 44 
IL-2 100 23 
IL-4 <10 <10 <10 
IL-5 <10 <10 <10 
a

Cytokine production from draining LN cells. LN cells from normal mice, EAE control mice, and EAE mice treated with anti-NK1.1 were cultured with 2 μg/ml MOG peptide. Cells were stimulated for 24 h, then supernatants were collected and evaluated for cytokine release. EAE mice LN were harvested after the second immunization at day 10. Supernatants were measured using CBA bead technology (BD Biosciences). All values are pools of three to five mice per treatment.

Because the NK or NK subset depletion altered the proliferation of LN T cells, we evaluated the cytokine production in these draining LN leukocytes. As shown in Table II, there were strong levels of MOG-induced TNF-α and IFN-γ production; modest levels of IL-6, IL-12 p70, IL-10, and IL-2 while IL-4 and IL-5 were undetectable. MOG-induced TNF-α and IFN-γ production was strongly diminished with NK depletion. When the frequency of CD4+ cells expressing these cytokines (Fig. 3,B) was evaluated, a similar pattern of reduction was observed. Other cytokines evaluated (IL-4, IL-13, and IL-10) were either not produced or were not altered by NK depletion. Thus, the MOG-induced production of cytokines by CD4+ cells was dramatically reduced when NK cells were depleted before immunization. Recent studies have demonstrated that the classical Th1/Th2 CD4+ T cells have been joined by an IL-17-producer T cell, termed Th17 (39, 40, 41). This Th17 cell has a defined cytokine profile; it has its own set of lineage-specific developmental genes and is the main proinflammatory CD4+ effector T cell involved in murine models of CIA and EAE. To evaluate the potential role for NK cells to modulate IL-17 and/or IL-23 production, draining node lymphocytes were evaluated for their ability to produce IFN-γ, IL-17, and IL-23 in either control or NK-depleted mice 10 days after immunization. Fig. 4,a demonstrates the result of one typical experiment examining individual mice, where the production of IFN-γ was significantly diminished. When IL-17 or IL-23 production was examined, no consistent increase was observed in the levels of either cytokine, although the ratio of IFN-γ and IL-17 is dramatically altered by NK depletion. Fig. 4 b shows the number of CD4- or CD8-positive cells producing either IFN-γ or IL-17 from LN. A similar pattern of IFN-γ reduction was seen, whereas IL-17-producing T cell subsets were not changed upon NK depletion.

FIGURE 4.

Production of Th17 cytokines after NK1.1 depletions. A, Cytokine production in draining LN for IFN-γ (□ and ▪), IL-17 (○ and •), and IL-23 (⋄ and ♦). Panel shows the production after 18 h in draining LN cells after stimulation with 1 μg/ml MOG peptide in individual mice from EAE control mice (○, □, and ⋄) and EAE NK1.1-depleted mice (•, ▪, and ♦). B, Intracellular staining of CD4 (▪) and CD8 (▩) T cells. Cytokine production was analyzed in draining LN cells 6 h after stimulation with 1 μg/ml MOG peptide (three pooled mice). Values express the number of IFN-γ- or IL-17-positive cells from the EAE- immunized mice with or without NK depletion.

FIGURE 4.

Production of Th17 cytokines after NK1.1 depletions. A, Cytokine production in draining LN for IFN-γ (□ and ▪), IL-17 (○ and •), and IL-23 (⋄ and ♦). Panel shows the production after 18 h in draining LN cells after stimulation with 1 μg/ml MOG peptide in individual mice from EAE control mice (○, □, and ⋄) and EAE NK1.1-depleted mice (•, ▪, and ♦). B, Intracellular staining of CD4 (▪) and CD8 (▩) T cells. Cytokine production was analyzed in draining LN cells 6 h after stimulation with 1 μg/ml MOG peptide (three pooled mice). Values express the number of IFN-γ- or IL-17-positive cells from the EAE- immunized mice with or without NK depletion.

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Since autoimmunity in the EAE model is known to be caused by an infiltration of T cells that induces a demyelination of neural tissue as well as inflammation of the CNS (31), we further evaluated the brain lymphocytes that were found in the CNS (after saline perfusion) using this primary immunization model system. IL-2 production was examined in either brain or draining LN lymphocytes after MOG stimulation in vitro (Fig. 3,C). Ag-induced IL-2 production was observed in lymphocytes from both brain and LN lymphocytes of EAE-induced mice. Although naive mice lacked any significant IL-2 production, NK depletion of EAE-induced animals resulted in a significant reduction of IL-2 production by draining LN cells but not those cells isolated from the brain. Evaluation of T cell subsets in brain lymphocytes revealed no change in CD4 or CD8 frequency after NK1.1 depletion (20.2 and 23.1% for CD4 and 19.3 and 18.8% for CD8). In addition, CNS lymphocytes were evaluated for the presence of T regulatory cells (CD3+CD4+ CD25+) and no differences in frequency were observed upon comparison of control or NK- depleted mice (data not shown). When peripheral nodes (draining, nondraining, and cervical) were examined from the same mice for IFN-γ mRNA (Fig. 3,D), NK depletion resulted in decreased IFN-γ mRNA levels. This is consistent with the lower levels of MOG-induced T cell activation that were seen as measured by IL-2 production (Fig. 3,C). Measurement of intracellular IFN-γ and TNF-α from the brain lymphocytes of EAE-induced mice demonstrated that these lymphocytes were quite capable of producing both cytokines (Fig. 5). The frequency of T cells producing cytokines after MOG peptide immunization was not significantly different (data not shown), but the total number of TNF-α (Fig. 5,a)- and IFN-γ (Fig. 5 b)-producing cells was significantly lower after NK cell depletion. Thus, these data suggested that although NK depletion can significantly alter the number and quality of responses in the LN, the MOG-reactive cells that reach the CNS are capable of making cytokines at a similar frequency as compared to the non-NK-depleted mice. Therefore, it appears that the decrease in the number of lymphocytes reaching the CNS in NK-depleted mice is responsible for the overall diminution of clinical disease that we observed.

FIGURE 5.

Cytokine production from CNS leukocytes. CNS was perfused with saline and lymphocytes were isolated as described in Materials and Methods. Bar graphs show the CD4+ or CD8+ T cell cytokine production in normal mice (▦), MOG-immunized mice (▪), or MOG-immunized and NK1.1-depleted mice (▨). Values represent the total number of CD4+ or CD8+ T cells in the CNS isolate that are producing either TNF-α (A) or IFN-γ (B). Values are pools of three to five mice per treatment.

FIGURE 5.

Cytokine production from CNS leukocytes. CNS was perfused with saline and lymphocytes were isolated as described in Materials and Methods. Bar graphs show the CD4+ or CD8+ T cell cytokine production in normal mice (▦), MOG-immunized mice (▪), or MOG-immunized and NK1.1-depleted mice (▨). Values represent the total number of CD4+ or CD8+ T cells in the CNS isolate that are producing either TNF-α (A) or IFN-γ (B). Values are pools of three to five mice per treatment.

Close modal

Because the results above suggested that the CNS cells were identical in the EAE and the NK-depleted EAE mice, only differing in numbers, while the draining LN T cells from the depleted mice were functionally deficient, we examined this site to determine whether an innate interaction between NK and DC might be altered in NK-depleted mice. When LN and brain of EAE-NK-depleted mice were examined for the presence of DC (expression of CD11c and MHC class II, I-Ab; Table III) and compared with the EAE control mice on day 7 following immunization, there were significantly fewer DC present in the NK-depleted EAE brain and cervical LN. Although the draining LN did not show significant changes in the actual number of DC, the percentage of DC was increased. The number of brain DC was diminished by 59% while the percentage was almost identical. In addition, the frequency and intensity of expression of CD80 and CD86 on DC (CD11c+class II+) was not altered by NK cell depletion (data not shown). As expected after NK1.1 depletion, a dramatic decrease was observed in the NK cell numbers in the LN that began to return by day 14 (data not shown). It should be noted that immunization with MOG peptide increased the number of draining LN NK cells compared with naive mice, a finding recently reported in another adaptive model (42). These data suggest the potential for tissue-specific interactions between DC and NK cells.

Table III.

Evaluation of DC and NK cells in EAE brain and LN

Organ SiteGroupTotal Cells (×106)% DCaNo. of DCa (×105)% NKaNo. of NKa (×104)
Brain EAE 4.8 26.1 (100) 13.0 (100) 1.5 (100) 3.70 (100) 
Brain NK deplb 2.2 24.6 (94) 5.4 (41)c 0.2 (13)c 0.18 (5)c 
Brain Control 0.1 9.4 (36) 0.1 (0.7) 1.1 (73) 0.00 (0) 
Draining LN EAE 14.0 13.9 (100) 20.0 (100) 2.1 (100) 23.00 (100) 
Draining LN NK depl 8.0 21.9 (158)c 18.0 (90) 0.2 (10)c 12.00 (52)c 
Draining LN Control 18.0 3.8 (27) 6.8 (34) 1.1 (52) 6.00 (26) 
Cervical LN EAE 8.2 15.0 (100) 12.0 (100) 1.5 (100) 7.40 (100) 
Cervical LN NK depl 3.0 11.1 (74) 3.3 (27) 0.2 (13)c 0.48 (7)c 
Cervical LN Control 1.6 2.3 (15) 1.6 (13) 0.4 (26) 0.63 (6) 
Organ SiteGroupTotal Cells (×106)% DCaNo. of DCa (×105)% NKaNo. of NKa (×104)
Brain EAE 4.8 26.1 (100) 13.0 (100) 1.5 (100) 3.70 (100) 
Brain NK deplb 2.2 24.6 (94) 5.4 (41)c 0.2 (13)c 0.18 (5)c 
Brain Control 0.1 9.4 (36) 0.1 (0.7) 1.1 (73) 0.00 (0) 
Draining LN EAE 14.0 13.9 (100) 20.0 (100) 2.1 (100) 23.00 (100) 
Draining LN NK depl 8.0 21.9 (158)c 18.0 (90) 0.2 (10)c 12.00 (52)c 
Draining LN Control 18.0 3.8 (27) 6.8 (34) 1.1 (52) 6.00 (26) 
Cervical LN EAE 8.2 15.0 (100) 12.0 (100) 1.5 (100) 7.40 (100) 
Cervical LN NK depl 3.0 11.1 (74) 3.3 (27) 0.2 (13)c 0.48 (7)c 
Cervical LN Control 1.6 2.3 (15) 1.6 (13) 0.4 (26) 0.63 (6) 
a

Percent or number has been normalized to EAE group (100%) within each specific tissue examined.

b

depl, Depleted.

c

Values are indicators of significant differences between the EAE group and the NK-depleted group.

We next examined the maturational status (based on expression of CD80 and CD86) of draining and cervical LN DC from EAE and NK-depleted EAE mice evaluated at day 7 postimmunization (Fig. 6). NK depletion modified the DC composition as found in draining LN (site of immunization); there was an increase in the percentage of CD80+CD86+ DC in EAE NK-depleted mice, suggesting a more mature DC population as compared with the EAE control mice. This change was not seen in the cervical LN and these DC effects were lost by day 14 of treatment. By comparing the ratio of bright (mature) to dim (immature) CD80+CD86+ DC during treatment (Fig. 6, B and C), we found that the NK depletion resulted in an increased frequency of CD80+CD86+ (mature) DC in the draining nodes, whereas immature DC are favored in the cervical nodes.

FIGURE 6.

Effects of NK depletion on DC. DC were enumerated from LN at specified times after immunization. A, Results of flow cytometric analysis of DC (examining CD11c+class II+ cells) evaluating the expression of CD80 and CD86 from cervical (left dot plots) or draining LN (right dot plots) at the immunization site at days 7 and 14, with or without anti-NK1.1 treatment (NK depl.). B (draining) and C (cervical) nodes were evaluated for relative expression of CD80/86bright–dim DC in mice immunized with MOG peptide with (EAE NK Dep.; ▴) or without NK depletion (EAE; ▵). n, Values are pools of three to five mice per treatment.

FIGURE 6.

Effects of NK depletion on DC. DC were enumerated from LN at specified times after immunization. A, Results of flow cytometric analysis of DC (examining CD11c+class II+ cells) evaluating the expression of CD80 and CD86 from cervical (left dot plots) or draining LN (right dot plots) at the immunization site at days 7 and 14, with or without anti-NK1.1 treatment (NK depl.). B (draining) and C (cervical) nodes were evaluated for relative expression of CD80/86bright–dim DC in mice immunized with MOG peptide with (EAE NK Dep.; ▴) or without NK depletion (EAE; ▵). n, Values are pools of three to five mice per treatment.

Close modal

Previous studies using MOG35–55 in various H-2 mice have indicated that TCR Vβ1, 6, 8,14, and 15 (16, 43, 44, 45) are used in C57BL/6 mice (46, 47) with the major TCR βs being 1 and 8. Since our studies have demonstrated an alteration in T cell responses that is characterized by decreased clinical disease, loss of Ag responsiveness, and reduced cytokine production, especially in the peripheral lymphoid tissue, we examined the TCR Vβ usage in the NK-depleted EAE mice by quantitative real-time PCR (30, 47) primer sequences. The data from a representative experiment (n = 3) are shown in Fig. 7. TCR Vβ quantitative usage in EAE-immunized mice was compared with NK-depleted, immunized mice. All TCR Vβs were normalized to a housekeeping gene and compared with the EAE controls. At the site of clinical disease in the brain, NK depletion resulted in a decrease in TCR Vβ8.1 usage, by 100–200%. Decreases were also observed in TCR Vβ14 and 15. However, when either cervical or draining LN were examined, an overall reduction in the measured TCR Vβs was seen across the analysis, consistent with the lack of reactivity to in vitro MOG peptide seen in these same tissues.

FIGURE 7.

Analysis of TCR usage in EAE mice depleted of NK cells. Lymphocytes from perfused brains (A) or cervical (B) or draining (C) nodes were removed and analyzed for TCR Vβ usage by real-time quantitative PCR using primers specific for the Vβs specified. Mice were immunized with MOG without NK1.1 depletion and values were normalized to this treatment (error bar). Values represent change in EAE NK-depleted mice relative to EAE autoimmune mice (with SE). The NK1.1 depletion values (▪) are shown for selected Vβs.

FIGURE 7.

Analysis of TCR usage in EAE mice depleted of NK cells. Lymphocytes from perfused brains (A) or cervical (B) or draining (C) nodes were removed and analyzed for TCR Vβ usage by real-time quantitative PCR using primers specific for the Vβs specified. Mice were immunized with MOG without NK1.1 depletion and values were normalized to this treatment (error bar). Values represent change in EAE NK-depleted mice relative to EAE autoimmune mice (with SE). The NK1.1 depletion values (▪) are shown for selected Vβs.

Close modal

Numerous recent reviews (48, 49, 50) have postulated the concept that NK cells can play an important role in the regulation of adaptive immunity. However, to date, there are limited definitive reports that define where and how NK cells can modulate their complex interactions. Perhaps the most studied and well-defined reports are the studies in CMV infection and the development of CD8 effector T cells (1, 2). Although encephalitogenic peptides (20) or TCR peptides (19, 51) in association with MHC molecules are recognized as the receptor ligands for some regulatory T cells, less is known about how other regulatory cells are triggered.

In fact, early studies (44, 52) used anti-asialo GM1 polyclonal sera for NK cell deletion, which can damage macrophages (53) or T cells (54). Other studies (45, 55, 56, 57) did not distinguish NK cells from NKT cells (58, 59, 60), a novel regulatory lymphocyte population that produces a large amount of IL-4 after TCR ligation (61).

To overcome the problems inherent in the previous studies, we selected a model of primary EAE induced in B6 with MOG35–55 peptide (31). Our initial studies evaluated the effects of depleting NK cells with anti-NK1.1, anti-asialo GM1, and select anti-Ly49s to alter populations of NK cells in adaptive tissue and draining LN and evaluate the effects of these treatments on the clinical course of EAE in vivo. Our studies clearly demonstrated that NK depletion before immunization diminished the onset of EAE and extent of paralysis. The use of anti-asialo GM1 and select anti-Ly49s ruled out a role for NKT cells since these cells remained unaffected by these treatments. In addition, the depletion of NK cells after immunization failed to alter clinical disease in vivo, thus supporting a regulatory role for NK cells early in immune development but not in the effector phase. Our findings that elimination of NK cells expressing Ly49D and Ly49H, a highly overlapping but minor subset of NK cells, had an effect similar to that of total NK depletion implicated this subset of the NK cells in the clinical response. Furthermore, when our studies were translated into an in vitro evaluation of T cell responses to cognate MOG Ag, NK-depleted lymphoid tissues demonstrated a diminished T cell response as measured by proliferation and cytokine production (IFN-γ and TNF-α).

Recent studies evaluating IL-17 production by T cells has implicated this cytokine as a major autoimmune regulator. Komiyama et al. (39) demonstrated that IL-17−/− mice had suppressed development of EAE. These animals exhibited delayed onset, reduced clinical scores, altered histological changes, and early recovery. The major producer of IL-17 was CD4+ T cells and their data suggested that IL-17 and IFN-γ mutually cross-regulate expression of each cytokine. These observations indicate that IL-17 plays a crucial role in the development of EAE. Furthermore, Carballeda et al. (40) demonstrated that mice lacking IL-23 (p19−/−) did not develop EAE. In addition, disease resistance by IL-23 knockout mice was associated with loss of IL-17-producing CD4+ T lymphocytes. These studies demonstrated that the IL-17/23 inflammation and related molecules were critical initiators of EAE and perhaps other autoimmune diseases. Suryani et al. (41) demonstrated that MOG peptide induced an IL-17+IFN-γ+ population of CD4+ CNS-infiltrating MOG35–55-specific T cells, which is the majority population relative to IL-17+IFN-γ cells. These studies implied that the Th1 lineage is more encephalitogenic than is suggested by adoptive transfer of Th1 (IL-17/IFN-γ+) cells since these cells may be terminally differentiated. In the present study, the removal of NK cells did not consistently alter the IL-17 or IL-23 levels induced by MOG stimulation, although a dramatic decrease in the ratio of IL-17:IFN-γ production was observed. These findings suggest that the target of NK effects is not within the IL-17/23 regulation of effector T cells.

Evaluation of the CNS leukocytes, unlike the draining LN, indicated both a qualitative alteration (TCR Vβ) and a quantitative change, since a reduced frequency of MOG-reactive T cells that emigrate to the CNS and mediate autoimmune reaction was observed. Analysis of TCR Vβ usage after NK cell depletion revealed a qualitative shift in the T cell population because T cells containing Vβ8, 14, and 15 were decreased. Because these T cells have been shown to be the major effector repertoire in C57BL/6 mice (31), loss of this population as a result of NK cell depletion would explain the altered clinical outcome observed in our study. Finally, the evaluation of LN DC demonstrated an alteration in mature DC levels (based on CD80 and CD86 expression), indicating that NK cells may prevent optimal secondary T cell responses and Ag presentation. Thus, the role of NK cells may be nonlytic since studies with perforin knockout mice (data not shown) demonstrated no regulatory role in primary EAE induction and disease. Collectively, these data support the recent hypothesis (62) that in addition to being antitumor and antiviral effector cells, NK cells mediate a critical link between NK cells and adaptive T and B cell responses as indicated in previous reports in the CMV infection model system (1, 63) and in the multiple sclerosis model (25, 28, 64).

The mechanisms by which NK cells regulate these adaptive responses are not yet fully elucidated, but their site of regulation has been clearly shown to involve the draining LN after MOG peptide immunization, an immunoregulatory site not generally thought to involve NK cells until recently. The cytokines that are critical for the maturation of EAE responses and that determine the differentiation of T cells into MOG- specific Th1 (IL-2, IFN-γ, TNF-α) T cells would be expected to have a regulatory impact on numerous other adaptive responses. Our findings are consistent with the proposed hypothesis that through cytokine production, NK cells can regulate the ability of intracellular bacteria and viruses to induce and regulate Ag-specific T cells (65, 66).

Our data suggest an important LN interaction among NK cells, DC, and the development of Ag-specific T and perhaps B cells. NK cells have been proposed to promote Th1 responses in part by early production of IFN (67) and regulation of DC (6). These reported effects of NK cells influencing the Ag-specific autoimmune response brings together a number of these regulatory circuits. However, such observations do not exclude the possibility that other cell types or other signals from NK cells might be involved in NK-DC-T cell interactions. For example, we have already shown that cytokines (IL-12 and IL-18) induced by other cell types can significantly costimulate NK cells in an in vivo environment and alter the inhibitory circuit that is mediated by NKRs (68). Thus, further experimentation is required to more fully define all of the signals that regulate NK-DC interactions.

The study of NK cell function in vivo has been challenging and has resulted in contradictory results as reviewed by Shi (25). These previous studies have used anti-NK1.1 or anti-asialo GM1 (not present on NKT cells) to selectively examine NK cells in a variety of autoimmune models and our findings are not inconsistent with the data reported in those model systems.

Shi (27) demonstrated in IL-18-deficient mice that immunization with MOG35–55 failed to induced Th1 and autoantibody responses and subsequent clinical EAE. This lack of autoimmunity could be restored by IL-18 but was dependent on the presence of NK cells. In addition, these studies suggested a role for NK-induced IFN-γ for CNS pathology. Our studies are quite consistent with this report for a central role of NK cells in the development of Th1 cells and MOG-induced EAE.

In another study, Shi (26) reported that NK cells can determine the outcome of B cell-mediated autoimmunity. Using this myasthenia gravis model, mice depleted of NK cells or NK cell-deficient IL-18−/− mice failed to develop Th1 responses and Abs to the acetylcholine receptor. These results established an important link between NK cells and autoreactive T and B cells. Our studies are quite consistent with this report for a central role of NK cells in the development of this B cell-mediated autoimmunity.

Other reports of mice lacking IL-15 or T-bet demonstrated both defective Th1 responses and autoimmunity. However, these deficiencies target multiple immune effectors and are a bit more difficult to apply directly to NK cells (reviewed in Ref. 64).

Recent studies by Huang (28) demonstrated a link between CX3CR1 expression and brain pathology in EAE, showing that NK cells were markedly reduced in the CNS of CX3CR1-deficient mice. However, the recruitment of other leukocytes, e.g., T cells, NKT cells, and monocytes/macrophages, were unchanged. The lack of NK cells was associated with increase clinical EAE mortality and hemorrhagic inflammatory lesions. These changes were not observed in CD1d-deficient mice, effectively ruling out a role for NKT cells. These results strongly suggested a critical role for CX3CR1+ NK cells in CNS pathology. Although our studies did not examine subsets of NK cells, we did observe primarily changes in T cell numbers with intact activation status in the CNS. However, increased NK cell numbers in the CNS compared with control mice were observed in our studies (Table III). Thus, our studies are consistent with the report by Huang et al. (28) that NK cells do increase in the EAE CNS.

Overall and in the context of the studies discussed above, our results are consistent with recent postulations (25, 47, 48, 49) that NK cells serve as an important potential link between innate and adaptive immunity. However, this link represents a very complex interaction and may be highly dependent upon the model system used. Thus, further studies are required to elucidate the role of NK cells in innate-adaptive interactions.

Our findings are in contrast to those reported by Zhang et al. (21). In that study, in vivo NK cell depletion increased EAE clinical pathology (Fig. 1 in Ref. 21) and increased MOG-induced proliferation and IFN-γ production from T cells (Fig. 5 in Ref. 21). In addition, NK depletion did not alter monophasic EAE (Fig. 4 in Ref. 21). In contrast to these results, our studies using anti-NK1.1, anti-asialo GM1, and anti-Ly49 reagents all demonstrated a decrease in clinical pathology and a decreased frequency of disease (current study, Fig. 1) as well as decreased MOG-induced proliferation and IFN-γ production (current study, Fig. 3). In contrast to our study which only evaluated primary EAE generation, Zhang et al. used adoptive transfer models of either primary T cells or T cell lines (Figs. 7 and 8 in Ref. 21) that mediated an EAE autoimmune response. A possible explanation for the differences between our report and that of Zhang et al. is that in transfer studies it is well known that NK cells rapidly eliminate transplanted cells in the circulation. Thus, some of the conclusions reached by Zhang and coworkers could have been due to the measured alterations in transfer efficiency. The reasons for the discrepancies in the in vivo depletions are not apparent; however, there are several notable differences between our report and that of Zhang and coworkers. First, the clinical EAE observed in the control mice in the studies by Zhang et al., where NK depletion was performed, was <1, which represents a very low level of disease. Our studies generally had control EAE that was 2–3 in magnitude (equivalent to 4–5 based on the scale used by Zhang et al.). Second, the anti-NK1.1 was used at a much higher dose by Zhang et al. (500 μg/mouse), whereas our studies used a split dose of 100 μg/mouse. These split doses deplete NK cells more efficiently in liver and LN. Both doses removed detectable NK cells as reported but the large excess of anti-NK1.1 might have other consequences not observed with administration of the lower dose. Third, Zhang et al. used footpad injection, followed by a booster immunization after 1 wk with PT being given i.v. Our studies used s.c. immunization followed by a 48-h boost with the PT given i.p. Fourth, our PT source was different from that of Zhang et al., a variable that might change the clinical score in this murine EAE model (our unpublished data). Fifth, our in vitro studies emphasized the LN (draining, nondraining, and cervical) and brain lymphocyte responses to MOG peptide, whereas studies by Zhang et al. evaluated the spleen. Sixth, our studies evaluated the brain immunology, DC maturation status, and TCR Vβ repertoire, issues that were not examined by Zhang et al. Seventh, both studies found that NK depletion after immunization failed to have any effect on clinical disease, supporting a role for NK cells early in the generation of an adaptive T cell response. Thus, the differences in the timing of immunization and PT routes as well as the cells evaluated may play an important role in the discrepancies in our results with those previously reported by Zhang et al.

Studies by Bakker et al. (69) demonstrated an important role for DAP12 in autoimmune disease and concluded that an impaired Ag priming was the underlying mechanism responsible for the resistance of DAP12-deficient mice to the development of EAE. Since DAP12 is present in both APC and NK cells, these conclusions are not in conflict with the data that we have presented here because our studies clearly demonstrated that primary T cell responses are minimized and that DC are altered when NK cells are depleted. In addition, the lack of DAP12 in NK cells, and as shown by the role of activating Ly49 cells, would significantly alter their ability to secrete cytokines, since many cytokines are expressed upon triggering activating receptors (67, 68, 70). Furthermore, our data directly implicate Ly49D and Ly49H NKR subsets, one of which is a potent source of NK cytokine production (71, 72, 73). Consistent with regulation of early Ag presentation by DC, our data also implicate an early involvement of NK cells since their depletion after the primary immunization (after day 7) did not modulate EAE clinically or the T cell responses.

In this current study, we demonstrate that NK cells, a major arm of innate immunity, participate in the development of primary MOG-induced T cell autoimmune disease. The requirement of NK cells was demonstrated in vivo by the reduced ability of NK or NKR subset-depleted mice to exhibit potent clinical disease. When studies were translated in vitro from mice depleted of their NK cells or a NKR subset, T lymphocytes demonstrated both a qualitative and quantitative failure to exhibit a T cell anti-MOG response as compared with EAE non-NK-depleted mice. NK cells were shown to regulate in the LN the immune responses that initiate the autoimmune disease, since the absence of the appropriate NK cells resulted in decreased production of inflammatory cytokines (TNF-α and IL-6) and a decrease in the growth-promoting cytokine IL-2.

In summary, the current data indicate that NK cells function in autoimmune pathology and that the alteration of the immune status of NK cells that either activate or inhibit their function would promote or interfere with normal innate homeostasis. Just as NK cells have been proposed to be a first line of defense against tumor cell progression, this study indicates that NK cells can contribute to the quality and quantity of adaptive T and B cell responses.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported in whole or in part with federal funds from the National Cancer Institute, National Institutes of Health, under Contract N01-CO-12400. This work was also supported in part by the Intramural Research Program of the National Institutes of Health, National Cancer Institute, Center for Cancer Research.

2

The publisher or recipient acknowledges the right of the U.S. government to retain a nonexclusive, royalty-free license in and to any copyright covering this article. The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. government.

4

Abbreviations used in this paper: DC, dendritic cell; EAE, experimental autoimmune encephalomyelitis; MOG, myelin oligodendrocyte glycoprotein; PT, pertussis toxin; LN, lymph node; CT, threshold cycle.

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