Skip Navigation



Rheumatology Advance Access published online on March 17, 2008
Rheumatology, doi:10.1093/rheumatology/ken028
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow All Versions of this Article:
47/5/609    most recent
ken028v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrowRequest Permissions
Right arrow Disclaimer
Google Scholar
Right arrow Articles by Iking-Konert, C.
Right arrow Articles by Hänsch, G. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Iking-Konert, C.
Right arrow Articles by Hänsch, G. M.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

© The Author 2008. Published by Oxford University Press on behalf of the British Society for Rheumatology. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

T lymphocytes in patients with primary vasculitis: expansion of CD8 + T cells with the propensity to activate polymorphonuclear neutrophils

C. Iking-Konert1, T. Vogl1, B. Prior2, C. Wagner3, O. Sander1, E. Bleck1, B. Ostendorf1, M. Schneider1, K. Andrassy4 and G. M. Hänsch2

1Rheumatology, Department of Endocrinology, Diabetology and Rheumatology, Heinrich-Heine-University of Duesseldorf, 2Institute for Immunology, Ruprecht-Karls University of Heidelberg, 3Klinik für Unfallchirurgie und Orthopädie, Berufsgenossenschaftliche Unfallklinik Ludwigshafen and 4Medizinische Klinik, Ruprecht-Karls University of Heidelberg, Germany.

Correspondence to: C. Iking-Konert, Rheumatology, Department of Endocrinology, Diabetology and Rheumatology, Heinrich-Heine-University of Duesseldorf, Moorenstr 5, 40225 Düsseldorf, Germany. E-mail: Iking-konert{at}med.uni-duesseldorf.de


   Abstract
Top
 Abstract
Introduction
 Patients and methods
 Results
 Discussion
 Conclusion
 Acknowledgements
 References
 
Objectives. To gain insight into the immune pathogenesis of primary ANCA-associated vasculitides, the prevalence of circulating T lymphocytes expressing CD11b as a marker for activation was analysed in patients with WG or microscopic polyangiitis.

Methods. Receptor expression and IFN{gamma} synthesis were measured in T cells of patients with active disease by cytofluorometry and compared with expression in patients in remission and in healthy donors.

Results. During active disease, a small but conspicuous population of CD8+CD28+CD11b+ was found which produced IFN{gamma}. In healthy donors and in patients in remission or undergoing immunosuppressive therapy, CD11b was exclusively associated with CD8+CD28– cells, the latter being more frequent in patients with long-lasting or severe disease. In vitro experiments confirmed that CD11b is up-regulated when T cells are activated. After multiple rounds of restimulation, the CD11b expression persists whereas CD28 expression is lost, compatible with the notion that CD8+CD28+CD11b+ represents a transient phenotype in the course of T-cell activation. The IFN{gamma}-producing T cells activated polymorphonuclear neutrophils (PMN) to express MHC class II, thus generating the same PMN phenotype as in patients with active ANCA-associated vasculitis. A similar PMN phenotype could be generated by cultivation with supernatants of activated T cells or by IFN{gamma} alone, but not by antibodies to proteinase 3.

Conclusions. In active primary vasculitis, a small population of CD8+ T cells, identified by the expression of CD11b, expands, producing IFN{gamma}. These T cells could activate PMN, thus generating a long-living and potentially destructive PMN phenotype.

KEY WORDS: PMN, ANCA, Vasculitis, T cells, CD11b, CD28, CD8, WG, Microscopic polyangiitis


   Introduction
Top
 Abstract
 Introduction
Patients and methods
 Results
 Discussion
 Conclusion
 Acknowledgements
 References
 
The aetiology and pathogenesis of primary vasculitides such as WG or microscopic polyangiitis (MPA) are still elusive. Although, autoantibodies to neutrophil-derived antigens—collectively referred to as ANCA—are found in the majority of patients, the causal relationship between these autoantibodies and the initiation or progression of the disease is still a matter of controversy [1–6].

From in vitro data it was deduced that antibodies to PR3 stimulate neutrophils (polymorphonuclear neutrophils; PMN) [7–11; reviewed in 3, 6]. Whether that occurs in vivo is questionable, because ANCA are of low affinity, and even in patients with antibody high titres, the PMN are negative for surface-bound antibodies [2]. ANCA are not detected on PMN that have infiltrated the tissue—e.g. in the kidney—despite the fact that those PMN express PR3 [12]. Moreover, ANCA are not deposited at inflamed sites, hence WG and MPA are described as ‘pauci-immune’. There is no doubt, however, that PMN are activated in acute disease: transfer of the ‘target’ antigens for ANCA, particularly of PR3 and of elastase, to the cell surface has been observed [13–15], as has up-regulation of activation-associated surface receptors, including integrins, and the Fc receptor CD64 [12, 16, 17]. Moreover, activation, predominantly of PMN, has been shown by gene array [18]. In addition to the expected activation-associated receptor up-regulation, a small but conspicuous number of PMN differentiated to cells with dendritic-like characteristics: expression of MHC class II antigens and of the co-stimulatory receptors CD80 and CD86 has been observed, as has the propensity to present antigen to T cells [19, 20]. In that context, possible interactions of PMN with activated T lymphocytes are of particular interest, because T cell-derived cytokines, especially IFN{gamma}, are potent activators of numerous PMN functions, including up-regulation of MHC class II antigens, of the co-stimulatory antigens CD80 and CD86, of the Fc receptor CD64 and of integrins [21–23; reviewed in 24].

Participation of T lymphocytes in the pathogenesis of WG or MPA has long been postulated. Evidence for T-cell activation is derived from numerous studies analysing the surface receptor profile (reviewed in [4, 6] and [25–28]) or from therapy studies directed predominantly against T lymphocytes [29]. Moreover, we have provided evidence for replicative senescence of T-cell clones [30], which verifies T-cell activation as opposed to alterations within the T-cell compartment due to immunosuppressive therapy. Despite much effort, the specificity of the activated T cells, the activating events and the role of T cells in the pathogenesis of primary vasculitis are still elusive, although numerous attractive hypotheses have been forwarded [27, 28, 31, 32].

In the present study, the prevalence of circulating T lymphocytes expressing CD11b as a marker for activation was analysed in patients with active WG or MPA, and was compared with patients with inactive disease, or to healthy donors, respectively. CD11b, the {alpha}-chain of the β2 integrin Mac-1, also known as complement receptor 3 (CR3), is expressed on all leucocytes, including T cells. CD11b is up-regulated in the course of T-cell activation in both, CD4+ and CD8+ cells, and most probably participates in migration and extravasation of the cells [33]. From studies with virus-infected mice, it was concluded that CD11b might be the best single marker available for discriminating between naive and memory CD8+ cells [34], or effector and memory cells, respectively [35]. A study with patients suffering from acute viral infections supported this conclusion: during acute virus infection, a population of CD8+CD28+ lymphocytes expressing CD11b expands with functional characteristics of memory/effector cells [36]. Moreover, our own studies of patients with bacterial infections revealed up-regulation of CD11b not only on CD8+ cells, but also on CD4+ cells.

In the current study, we found a small population of CD8+CD28+CD11b+ cells which declined upon immunosuppressive therapy in patients with active disease. The CD8+ cell produced IFN{gamma}, and we propose that these T cells activate PMN, thus generating the PMN phenotype prevailing in active disease.


   Patients and methods
Top
 Abstract
 Introduction
 Patients and methods
Results
 Discussion
 Conclusion
 Acknowledgements
 References
 
Patient characteristics
The study was approved by the ethic committees of the University of Heidelberg and of the University of Düsseldorf. Between 1999 and 2006, after informed consent was obtained, 90 patients were recruited by the renal unit of the University of Heidelberg University Hospital and the Department of Rheumatology/University of Duesseldorf. Ten patients had active disease, with a BVAS (Birmingham vasculitis activity score) >5 [37]. Of these patients, four had untreated, newly diagnosed WG, one had a relapse after remission for 2 yrs, and five had untreated, newly diagnosed MPA. Eighty patients in remission (BVAS < 1) (47 patients with WG and 33 with MPA) were also included. WG and MPA were classified according to the definition of the Chapel Hill conference [38] and to the ACR criteria for the classification of WG [39]. The patients in remission were divided into two groups: those with long-standing disease (>5 yrs) and those with a disease duration of ≤5 yrs.

‘Severe disease’ was defined as follows: systemic disease including at least one major organ (e.g. kidney, lung, central nervous system), and history of more than one relapse. Furthermore the cumulative cyclophosphamide dose was calculated for each patient. Patients were classified by the number of organs involved, as described by Reinhold-Keller et al. [40] and de Groot et al. [41]. For comparison, blood was drawn from healthy donors, matched with regard to age, after having obtained informed consent and observing the institutional guidelines.

Cytofluorometry
The expression of CD8, CD28 and CD11b was measured in whole blood by FACScan using standard procedures. Cells were triple-stained using phycoerythrin (PE) or APC-labeled antibodies to CD8 or CD4 (Becton Dickinson, Heidelberg, Germany), and FITC-labelled antibodies to CD28 (Serotec Düsseldorf, Germany; YTH 913.12) or CD11b (Biozol; Eching, Germany, ICRF44). A PE-labelled anti-CD11b was also used (BD Biosciences, Heidelberg). PMN were double-stained with the specific marker CD66b FITC (clone 80 H3) and with CD14 PE (clone UCHM1), CD64 PE (clone 10.1) or MHC Class II PE (clone WR18) (all these antibodies were purchased from Serotec). The antibody to PR3 (‘Pelicluster ANCA’) produced by Sanquin Amsterdam, was obtained from HISS Diagnostics Freiburg, Germany. The final concentration of antibodies varied between 1 and 10 µg/test. The isotype controls IgG1, IgG2a and IgG2b were used in the same final concentration. To determine the CD4/CD8 ratio, a mixture of anti-CD4 FITC- anti-CD8-PE was used with IgG FITC/IgG PE as control (antibodies obtained from Beckman Coulter, Marseille, France).

Production of IFN{gamma}
Accumulation of IFN{gamma} was measured intracellulary by cytofluorometry. The cells were treated with brefeldin A (10 µg/ml; Sigma, Munich, Germany) for 4–12 h, then incubated with 100 µl Fix and Perm solution (Becton Dickinson) and FITC-labelled antibody to IFN{gamma}. Two different antibody clones were used: clone 25 723.11 (BD Bioscience) and clone D9D10 (Serotec) in a final concentration of 10 µg/ml.

Analysis of the data
For technical reasons not all parameters were measured in all patients or healthy donors. The number of individuals that compared with regard to one particular parameter is given in the figure or the figure legend, respectively. The results of the FACScan data are expressed as percent positive cells in the respective gate in relation to CD8+ cells. Differences between groups were calculated by one-way analysis of variance (ANOVA) and Bonferroni test at a significance level of P < 0.05. The chi-square test was used to analyse the coincidence of low CD4/CD8 ratio and the expression of CD11b or CD28. The threshold for low CD4/CD8 ratio was set as <1.5; the threshold for the other parameters was set using the respective mean values ± 1 S.D. obtained for healthy donors.

Isolation of T cells and PMN and in vitro activation
PMN
Whole blood was centrifuged on Polymorphprep® (Nycomed, Oslo, Norway) according to the supplier's protocol. From the PMN layer, cells were further purified by adsorption to anti-CD15 magnetic beads (AutoMACS) (Miltenyi Biotech, Bergisch Gladbach, Germany).

T cells
From the mononuclear cell layer, CD8+ cells were isolated by adsorption to anti-CD8 magnetic beads.

Co-cultivation experiments: T-cell line
To generate a T-cell line expressing CD11b, the CD8+ cells were stimulated with phytohaemagglutinin (PHA) (1 µg/ml) and placed on irradiated autologous mononuclear cells as feeder cells (3 x 106 CD8+ per 3 x 106 feeder cells). After 24 h, IL-2 was added (2.5 ng/ml). After 16 days in culture, the T cells were harvested and stimulated again with PHA. After the second round of cultivation, 72% of the cells were positive for CD11b, after the 4th round 95% were positive. Of these, 83% were negative for CD28. For the co-culture experiments, equal numbers of T cells and of PMN (2 x 106 in a total volume of 1 ml) were incubated for 24–48 h.

Isolated T cells
CD8+ cells derived from healthy donors were suspended in RPMI supplemented with fetal calf serum (10%), penicillin/streptomycin (1%), L-glutamine (1%), Hepes (1%) and IL2 (2.5 ng/ml) and were placed into multi-well culture dishes (24 wells) coated with anti-CD3 (clone UCHT-1; Immunotec, Marseille, France 100 ng/well) in a concentration of 106 per well. An equal number of autologous PMN was added. After 48 h in culture, MHC class II expression on PMN was measured by cytofluorometry. For comparison, PMN were cultivated without T cells, or with T cells without stimulation.

Production of T-cell supernatants
CD8+ T cells derived from healthy donors were activated with PMA (20 ng/ml) and ionomycin [1 µg/ml for different time intervals (1, 4, 12, 24 and 48 h)]. Supernatants were harvested, IFN{gamma} was measured by a commercially available ELISA (R&D Systems, Minneapolis, MN, USA), and 200 µl of the supernatant was added to PMN 2 106 suspended in 800 µl AIM-V (Gibco, Eggenstein, Germany). MHC class II expression on the PMN was measured after 1, 24 and 48 h by cytofluorometry using an antibody to CD66b to identify the PMN.

Stimulation of PMN with anti-PR3
1–5 µg of anti-PR3, and for comparison, mouse IgG, were added to isolated PMN or to whole blood. After 24 and 48 h, MHC class II expression was measured by cytofluorometry. Death of PMN was determined by propidium iodide staining.


   Results
Top
 Abstract
 Introduction
 Patients and methods
 Results
Discussion
 Conclusion
 Acknowledgements
 References
 
Analysis of PMN and of T cells in patients with active disease
Peripheral blood cells of patients with active WG (n = 5) or active MPA (n = 5) were analysed by cytofluorometry. On PMN, the following activation-associated receptors were up-regulated: CD66b; CD64 and MHC class II, the latter particularly on PMN of patients with WG. Moreover, surface expression of PR3 was detected in all patients, including the patients with MPA. Expression of MPO was seen in one of the MPA patients. After onset of the immunosuppressive therapy, the receptor expression declined within days (examples are shown in Fig. 1A; data of all parameters and patients are summarized in Fig. 1B).


Figure 1
View larger version (35K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		FIG. 1. Expression of CD14, CD64 and MHC class II antigens on PMN (identified by expression of CD66b) was analysed by FACScan. When the disease was active, expression was seen, and declined after onset of immunosuppressive therapy. As an example, data obtained for Patient #8 are shown: (A) upper panel, active disease BVAS 9; lower panel, 12 days after starting the immunosuppressive therapy; BVAS 1. (B) Data of 10 patients are summarized (receptor expression is given as percentage positive PMN). For each receptor the mean values obtained when disease was active differed from the values obtained after therapy, as calculated by ANOVA (P < 0.05).

 
Among the receptors tested on T lymphocytes, marked differences were seen for CD11b expression: on T cells of healthy donors CD11b expression was mainly associated with CD8+CD28– cells. In patients with active disease, a population of CD8+CD28+CD11b+ cells appeared, up to 18% of the CD8+ cells [mean 8.9 ± 4.7% vs 1.2 ± 2.0% in healthy donors (n = 20); P < 0.001]. In remission, the population of CD8+CD28+CD11b+ cells disappeared (an example is shown in Fig. 2A). On CD4+ cells, expression of CD11b was well below 1%.


Figure 2
View larger version (31K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		FIG. 2. Expression of CD11b and synthesis of IFN{gamma}. (A) The CD8+ cells were gated and the expression of CD28 and of CD11b was measured. In patients with active disease (Patient #7, BVAS 4, is shown as an example in the left panel), four populations of cells were identified: CD28+CD11b–; CD28+CD11b+; CD28–CD11b–; CD28–CD11b+. Under therapy, the CD28+CD11b+ cells disappeared. In healthy donors, there is no population of CD28+CD11b+ cells (right panel). (B) When T cells of the patients were kept ex vivo for 4 h in the presence of brefeldin A, IFN{gamma} could be visualized, almost exclusively in the CD11b+ cells. (C) Expression of CD28 and CD11b on CD8+ T cells of patients in remission: both CD8+ cells expressing CD28 and CD8+ cells negative for CD28 are found in the patients; CD11b was almost exclusively expressed by the CD28– cells. An example is shown in Fig. 2C in the left panels; the markers M1 are set according to the isotype control. Data of all patients are summarized in the right panel.

 
The CD8+CD11b+ cells produced IFN{gamma} ex vivo: when kept for 4 h at 37°C in the presence of brefeldin A, IFN{gamma} could be visualized intracellularly, predominantly in the CD11b+ cells (one of five patients is shown in Fig. 2B). Within days after onset of therapy with immunosuppressants, the IFN{gamma}-producing cells were no longer detectable.

Analysis of CD8 T cells of patients in remission: loss of CD28 and expression of CD11b
As described above, CD8+CD28+CD11b+ cells were not detectable in patients in remission and in those patients CD11b was expressed exclusively on the CD28– cells (an example and the summary of all data are shown in Fig. 2C). In agreement with data published by us and others [30, 42, 43], CD8+CD28– cells were seen more frequently in the patients with ANCA-associated vasculitis (46.7 ± 19.3% in the patients compared with 22.4 ± 8.5% in healthy donors; P < 0.01), particularly in those with disease for ≥5 yrs (38.1 ± 14.4% in patients with disease for ≤5 yrs; 49.7 ± 18.1% in patients with disease for >5 yrs; P < 0.05).

CD11b expression did not correlate with reduced CD28 expression, or with clinical parameters such as severity of disease, organ involvement or cumulative cyclophosphamide dosage (data not shown). With regard to CD11b expression, there was no difference between patients with WG and patients with MPA.

Determination of CD4:CD8 ratio in the patients
The determination of the ratio of CD4+ to CD8+ cells is widely used to monitor alterations within the T-cell compartment. As illustrated by histogram plotting, in the majority of patients in remission (58 of 80) the CD4:CD8 ratio was well below 1.5 (normal range 1.5–2.5) (Fig. 3A). A median value of 1.0 was determined for the patients and 2.05 for healthy donors (n = 50). The mean values (1.12 ± 0.56 for the patients and 1.81 ± 0.44 for the healthy donors) were significantly different as calculated by ANOVA and the Bonferroni test (significance level 10–5). There was no significant difference between the WG and the MPA patients (WG: median 1.13; mean ± S.D. 1.16 ± 0.65; MPA: median 1.16; mean 1.33 ± 0.5).


Figure 3
View larger version (20K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		FIG. 3. Determination of the CD4:CD8 ratio in patients with WG or MPA. (A) The CD4:CD8 ratio was determined by cytofluorometry in 80 patients, and is depicted here as a histogram. The x-axis shows the CD4:CD8 ratio (range 1–4; increment 0.2); the y-axis shows the number of patients (black) or of age-matched healthy donors (red), respectively per increment. The lower panels (B) and (C) show the relationship between the CD4:CD8 ratio and the percentage of CD8+CD28– T cells, or CD8+CD11b+ T cells. Patients with a CD4:C8 ratio <1.5 had a higher proportion of CD8+CD28– cells ({chi}2 6.29; P < 0.01).

 
To search for a relationship between the shift of the CD4:CD8 and clinical parameters, patients were grouped according to the following criteria: duration of the disease, severity, organ involvement, systemic vs localized disease, renal involvement and accumulated cyclophosphamide dose. The general tendency was that patients with long-lasting, severe disease or renal involvement had a lower CD4:CD8 ratio. For a limited number of patients (n = 18), we had the opportunity to measure the CD4:CD8 ratio over an extended period of time (6–24 months): in patients with stable, well-controlled disease the ratio of CD8:CD4 remained stable. In some patients, even a shift back towards CD4 was seen. In six patients, the CD4:CD8 ratio shifted further towards CD8; these patients suffered from progressive smoldering disease (data not shown).

Since Marinaki et al. [44] described a decline in the absolute number of CD4+ cells in the course of the disease, CD4+ and CD8+ cells were counted in patients with a CD4:CD8 ratio of ≤1. Here we found that indeed the number of CD4+ was decreased (range 64–392/µl, mean ± S.D. 187 ± 95; normal range 320–1473/µl; n = 10), while CD8+ increased or stayed within normal limits.

Notably, a low CD4:CD8 ratio correlated with a high percentage of CD8+CD28– cells (Fig. 3B), while there was no correlation regarding CD11b expression (Fig. 3C).

In vitro experiments linking IFN{gamma}-producing T cells and PMN
So far, our data indicated that in patients with active WG CD8+CD28+CD11b+ cells expand, which ex vivo produced IFN{gamma} without further activation. Because IFN{gamma} is among the few cytokines able to induce the MHC class II expression on PMN, we attempted to test directly the interaction of CD8+CD28+CD11b+ cells with PMN in vitro. Since induction of MHC class II antigens requires 24–48 h autologous cells, these experiments could not be performed with CD8+CD28+CD11b+ cells from the patients, because during active disease, PMN are already activated. Therefore, we generated CD8+ expressing CD11b of healthy donors on up to 90% of the cells by in vitro activation. These cells also produced IFN{gamma} (Fig. 4A–D, upper panel). When these cells were co-cultivated with autologous PMN, an up-regulation of MHC class II antigens on the PMN was observed. By 24 h, 12.7 ± 8.0% of PMN had acquired MHC class II; by 48 h it was 24.9 ± 7.5%. Expression of MHC class II on freshly isolated PMN was <1% (Fig. 4E–H). In the absence of the T cells, the PMN did not acquire MHC class II, and the majority of PMN became apoptotic. In the presence of T cells, however, up to 80% of PMN survived (data not shown).


Figure 4
View larger version (26K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		FIG. 4. Generation of CD8+CD11b+ cells in vitro and co-culture with PMN. Upper panel: peripheral CD8+ isolated from a healthy donor (A) were stimulated repeatedly with PHA. Expression of CD11b was determined after the second (B) and the fourth round (C) of re-stimulation. The majority of these CD8+CD11b+ cells synthesized IFN{gamma} (D). Lower panel: autologous PMN were added to CD8+ T cells derived from a healthy donor and activated by immobilized anti-CD3. After 48 h, expression of MHC class II on the PMN was measured. In the forward-side scatter image (E), the two cell populations can be seen. The gate was set around the PMN population (R1), and class II expression was measured (F) showing the isotype control. For comparison, PMN cultivated in the absence of T cells are shown (G), as are the PMN before culture (H). PMN acquire MHC class II only in the presence of T cells.

 
In another set of experiments, freshly isolated CD8+ cells were used in place of the T-cell line. When these cells were co-cultivated with PMN, the MHC class II induction was marginal. When, however, the T cells were placed on immobilized anti-CD3, and the PMN were added, MHC class II expression on the PMN was induced. On average, by 48 h, 10.5 ± 7.5% (mean ± S.D. of five independent experiments) of PMN acquired MHC class II. The induction of MHC class II on PMN paralleled the up-regulation of CD11b on the CD8+CD28+ cells (data summarized in Table 1).


View this table:
[in this window]
[in a new window]

  		TABLE 1. Induction of MHC class II expression on PMN of healthy donors by CD8+ T cells

 
In a further set of experiments, supernatants of PMA/ionomycin-activated CD8+T cells were used. With supernatants of cells having been activated for 24–48 h, MHC class II expression could be up-regulated. At these time points, IFN{gamma} was maximally accumulated in the cell supernatant (depending on the individual donors, the IFN{gamma} concentration amounted to 600–800 units/106 cells after 24–48 h). Supernatants harvested after 1 h, which did not yet contain IFN{gamma}, had no effect on MHC class II expression, ruling out direct effects of PMA/ionomycin on PMN. The up-regulation of MHC class II antigens on PMN could be inhibited by antibodies to the IFN{gamma} receptor (data are summarized in Fig. 5).


Figure 5
View larger version (11K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		FIG. 5. Induction of MHC class II expression on PMN: PMN were incubated for the times indicated; the percentage of CD66b and MHC class II-positive cells was measured by FACScan. Isolated PMN of healthy donors were incubated with IFN{gamma} (100 U/ml), or supernatants of activated T cells harvested at 1 or 24 h after stimulation of CD8+ cells with PMA/ionomycin, respectively. Depending on the individual donors, the IFN{gamma} concentration amounted to 600–800 units/106 cells after 24–48 h. Pre-incubation of PMN with an antibody to the IFN{gamma} receptor (for 30 min at 30°C) inhibited the MHC class II expression. Antibodies to PR3 did not induce MHC class II expression [data are given as percentage MHC class II-positive PMN (mean plus/minus S.D. of n = 3)] measured either before culture, or 24 h (open bars) or 48 h (filled bars) after culture.

 
Since autoantibodies to PR3 are thought to play a role in PMN activation ([7, 9, 10, 45]; reviewed in [3, 6]), we assessed the effect of PR3 antibodies on the expression of MHC class II by PMN. Neither short-term nor long-term culturing of PMN with anti-PR3 resulted in an up-regulation of MHC class II (Fig. 5).


   Discussion
Top
 Abstract
 Introduction
 Patients and methods
 Results
 Discussion
Conclusion
 Acknowledgements
 References
 
A hallmark of WG or MPA is the activation of PMN. Activated PMN are found in the peripheral blood, but also as infiltrates in affected tissues [12, 16–20]. Although PMN are mainly recognized for their role in host defence, there is strong evidence that they also participate in chronic inflammatory disease as major effectors of tissue destruction [46–49]. Activation of PMN is primarily attributed to bacterial products or pro-inflammatory cytokines (e.g. IL8). Less well-acknowledged is the fact that IFN{gamma}, a typical T-cell-derived cytokine, is a potent activator of PMN. IFN{gamma} induces a rather unique alteration of PMN; namely, the transdifferentiation of PMN to cells with characteristics of dendritic cells, including prolongation of the life span, acquisition of CD64, CD83, MHC class II and the co-stimulatory molecules CD80 and CD86, as well as the ability to function as accessory cells for T-cell activation ([20–23], reviewed in [24] and [50]).

We now propose that it is indeed the T-cell response that leads generation of IFN{gamma} to the activation of PMN in WG or MPA. Our proposition is based on the following facts: (i) during acute disease PMN are activated. Aside from the usual activation-associated receptors, MHC class II antigens and CD64 are up-regulated, both known to be induced by IFN{gamma}, but not by other inflammatory cytokines such as IL-8, or by complement activation products (for review see [50]) or by ANCA. (ii) There is unquestionable evidence that IFN{gamma} is generated during active disease and that it is critically involved in the pathogenesis, particularly of WG [51–54]; (iii) T cells are activated in the course of the primary vasculitides, and T-cell activation progresses over an extended period of time. Evidence includes the replicative senescence within the T-cell compartment [30], the reduced expression of CD28 [30, 42, 43, 51, 55, 56], the acquisition of CD57 [42] and the shift of the CD4:CD8 cell ratio (Fig. 3A).

How and why the T cells are activated has not yet been determined, nor has it been possible to unequivocally establish antigen specificity ([57–60]; reviewed in [28]). According to data in the literature, in WG or MPA both, CD4 and CD8, T cells are activated, and CD4 cells have been identified as a source of IFN{gamma} [51–54].

We now provide evidence for the activation of CD8+ cells during active disease. As a marker for T-cell activation, we determined CD11b expression, because studies with virus-infected mice have indicated that CD11b expression is not only a reliable indicator of T-cell activation but also discriminates between recently activated effector cells and memory cells [34, 35]. Indeed, in our patients with active disease, a small but conspicuous population of CD8+CD28+CD11b+ cells appeared. Expansion of such CD8+CD28+CD11b+ T cells has been reported before in patients infected with a virus [34]. Although, virus-infection (e.g. with CMV or EBV) could not be ruled out in our patients, we think that it is rather unlikely, because there were no clinical indications of virus infection, and 9 of the 10 patients had newly diagnosed disease and were not undergoing immunosuppressive therapy. Moreover, from in vitro experiments it is known that CD11b up-regulation is not specific for virus-infection, but occurs regularly in the course of T-cell activation [61].

CD8+CD28+CD11b+ cells are considered to be a transient ‘intermediate’ phenotype in the process of CD8+ activation, giving rise to the more persistent phenotype CD8+CD28–CD11b+ [36], which is found in patients in remission or under immunosuppressive therapy as well as in healthy donors. The percentage of these cells increases with age, most probably reflecting a history of previous activation, and the fact, that—in contrast to activated CD4+—CD8+ are rather resistant towards apoptosis, and thus accumulate over a lifetime (reviewed in [62]).

Activation and expansion of CD8+ cells in the course of the active disease, combined with their higher resistance to apoptosis, could account for the shift of the CD4:CD8 ratio towards CD8, the more so since CD4+ cells are lost [44].

In combination, our data provide evidence for the activation-induced expansion of CD8+, apparent as CD8+CD28+CD11b+. Ex vivo, these cells produce IFN{gamma}. Following immunosuppressive therapy, IFN{gamma} production is no longer apparent. It is of note, however, that IFN{gamma} production is not limited to CD11b+ cells.

According to our hypothesis, PMN are activated by IFN{gamma} derived most likely from the activated CD8+ cells. To test this more directly, CD8+CD28+CD11b+ cells were generated from healthy donors, because the use of patient-derived cells was precluded by the fact that autologous PMN would be required, which during active disease, however, are already activated. Therefore, T cells of healthy donors were used. With a cell line consisting of >90% CD8+CD28+CD11b+ cells, MHC class II expression on PMN could be induced. Similarly, T cells activated by a one-step exposure to anti-CD3, which up-regulated CD11b expression on the CD8+CD28+ cells, induced MHC class II on the PMN, as did the supernatants of activated T cells.

Triggering PMN with antibodies to PR3 in concentrations reported to affect the oxidative burst [8] had no effect at all on MHC class II expression, implying that the up-regulation of MHC class II antigens on PMN, as it is seen in vivo in patients with active disease, is not due to stimulation with anti-PR3, but rather to the action of activated T cells.

Whether or not the acquisition by PMN of MHC class II is of any relevance for the progress of the disease cannot be decided as yet. Other IFN{gamma}-induced alterations of PMN, such as prolonged lifespan or enhanced cytotoxicity, are easier to reconcile with a pathogenetic role of PMN [46–49]. On the other hand, we and others [20, 23, 63] have shown, that MHC class II-positive PMN also serve as accessory cells for T-cell activation, which in turn could sustain the T-cell activation and thereby perpetuating the immune response.


   Conclusion
Top
 Abstract
 Introduction
 Patients and methods
 Results
 Discussion
 Conclusion
Acknowledgements
 References
 
In summary, this study provides evidence that T lymphocytes, particularly IFN{gamma}-producing CD8+CD28+CD11b+ cells, expand during active disease. We propose that these T cells—as they do it in vitro—induce the differentiation of PMN which is typically seen in active disease, and that these altered PMN participate in the chronic-destructive inflammatory process.

Formula


   Acknowledgements
Top
 Abstract
 Introduction
 Patients and methods
 Results
 Discussion
 Conclusion
 Acknowledgements
References
 
The authors would like to thank Ms Jeannie Wurz for editing the manuscript.

Disclosure statement: The authors have declared no conflicts of interest.


   References
Top
 Abstract
 Introduction
 Patients and methods
 Results
 Discussion
 Conclusion
 Acknowledgements
 References
 

  1. Falk RJ, Jennette JC. Are ANCA pathogenic – oh yes they are. J Am Soc Nephrol (2002) 13:1977–9.[Free Full Text]
  2. Abdel-Salam B, Iking-Konert C, Schneider M, Andrassy K, Hänsch GM. Autoantibodies to human cytoplasmic antigens (ANCA) do not bind to polymorphonuclear neutrophils in whole blood. Kidney Int (2004) 66:1009–17.[CrossRef][Web of Science][Medline]
  3. Harper L, Savage COS. Pathogenesis of ANCA-associated systemic vasculitis. J Pathol (2000) 190:349–59.[CrossRef][Web of Science][Medline]
  4. Sarraf P, Sneller MC. Pathogenesis of Wegener's granulomatosis: current concepts. Expert Rev Mol Med (2005) 13:1–19.
  5. Hänsch GM, Iking-Konert C, Andrassy K. The pathogenesis of ANCA-associated vasculitides: old hypotheses and new insights. Clin Nephrol (2005) 64:460–4.[Web of Science][Medline]
  6. Bosch X, Guilabert A, Font J. Antineutrophil cytoplasmic antibodies. Lancet (2006) 368:404–18.[CrossRef][Web of Science][Medline]
  7. Falk R, Terrel R, Charles L, Jennette J. Anti-neutrophil cytoplasmic autoantibodies induce neutrophils to degranulate and produce oxygen radicals in vitro. Proc Natl Acad Sci USA (1990) 87:4115–9.[Abstract/Free Full Text]
  8. Charles LA, Caldas ML, Falk RJ, Terrell RS, Jennette JC. Antibodies against granule proteins activate neutrophils in vitro. J Leukoc Biol (1991) 50:539–46.[Abstract]
  9. Keogan MT, Esnault VLM, Green AJ, Lockwood CM, Brown DL. Activation of normal neutrophils by anti-neutrophil cytoplasm antibodies. Clin Exp Immunol (1992) 90:228–34.[Web of Science][Medline]
  10. Ralston D, Marsh C, Lowe M, Wewers M. Antineutrophil cytoplasmic antibodies induce monocyte IL-8 release. Role of surface proteinase-3, alpha1-antitrypsin, and Fcgamma receptors. J Clin Invest (1997) 100:1416–24.[Web of Science][Medline]
  11. Hattar K, van Burck S, Bickenbach A, et al. Anti-proteinase antibodies (c-ANCA) prime CD14-depedent leukocyte activation. J Leukoc Biol (2005) 78:992–1000.[Abstract/Free Full Text]
  12. Brouwer E, Huitema MG, Mulder AHL, et al. Neutrophil activation in vitro and in vivo in Wegener's granulomatosis. Kidney Int (1994) 45:1120–31.[Web of Science][Medline]
  13. Csernok E, Ernst M, Schmitt W, Bainton DF, Gross WL. Activated neutrophils express proteinase 3 on their plasma membrane in vitro and in vivo. Clin Exp Immunol (1994) 95:244–50.[Web of Science][Medline]
  14. Müller-Kobold AC, Kallenberg CG, Tervaert JW. Leucocyte membrane expression of proteinase 3 correlates with disease activity in patients with Wegener's granulomatosis. Br J Rheumatol (1998) 37:901–7.[Abstract/Free Full Text]
  15. Morcos M, Zimmermann F, Radsak M, et al. Autoantibodies to polymorphonuclear neutrophil elastase do not inhibit but enhance elastase activity. Am J Kidney Dis (1998) 31:978–85.[Web of Science][Medline]
  16. Haller H, Eichhorn J, Pieper K, Göbel U, Luft F. Circulating leukocyte integrin expression in Wegener's granulomatosis. J Am Soc Nephrol (1996) 7:40–8.[Abstract]
  17. Müller-Kobold AC, Mesander G, Stegeman CA, Kallenberg CG, Tervaert JW. Are circulating neutrophils intravascularly activated in patients with anti-neutrophil cytoplasmic antibody (ANCA)-associated vasulitides? Clin Exp Immunol (1998) 114:491–9.[CrossRef][Web of Science][Medline]
  18. Alcorta D, Barbes DA, Dooley M, et al. Microarray expression analysis of systemic lupus erythematodes and ANCA vasculitis patient leukocytes revealed disease specific gene expression signatures. Kid Blood Press Res (2005) 28:171.
  19. Hänsch GM, Radsak M, Wagner C, et al. Expression of major histocompatibility class II antigens on polymorphonuclear neutrophils in patients with Wegener's granulomatosis. Kidney Int (1999) 55:1181–8.
  20. Iking-Konert C, Vogt S, Radsak M, Wagner C, Hansch GM, Andrassy K. Polymorphonuclear neutrophils in Wegener's granulomatosis acquire characteristics of antigen presenting cells. Kidney Int (2001) 60:2247–62.[CrossRef][Web of Science][Medline]
  21. Gosselin EJ, Wardwell K, Rigby WF, Guyre PM. Induction of MHC class II on human polymorphonuclear neutrophils by granulocyte/macrophage colony-stimulating factor, IFN-gamma, and IL-3. J Immunol (1993) 151:1482–90.[Abstract]
  22. Reinisch W, Tillinger W, Lichtenberger C, et al. In vivo induction of HLA-DR on human neutrophils in patients treated with interferon-{gamma}. Blood (1996) 87:3068.[Free Full Text]
  23. Radsak M, Iking-Konert C, Stegmaier S, Andrassy K, Hänsch GM. Polymorphonuclear neutrophils (PMN) as accessory cells for T-cell activation: MHC class II restricted antigen-dependent induction of T-cell proliferation. Immunology (2000) 101:521–30.[CrossRef][Web of Science][Medline]
  24. Ellis TN, Beaman BL. Interferon-gamma activation of polymorphonuclear neutrophil function. Immunology (2004) 112:2–12.[CrossRef][Web of Science][Medline]
  25. Gross WL, Csernok E, Helmchen U. Antineutrophil cytoplasmic autoantibodies, autoantigens, and systemic vasculitis. APMIS (1995) 103:81–97.[Web of Science][Medline]
  26. Clayton AR, Savage CO. What you should know about PR3-ANCA. Evidence for the role of T cells in the pathogenesis of systemic vasculitis. Arthritis Res (2000) 2:260–2.[CrossRef][Web of Science][Medline]
  27. Sanders JS, Stegeman CA, Kallenberg CG. The Th1 and Th2 paradigm in ANCA-associated vasculitis. Kidney Blood Press Res (2003) 26:215–20.[CrossRef][Web of Science][Medline]
  28. Lamprecht P. Off balance: T-cells in antineutrophil cytoplasmic antibody (ANCA)-associated vasculitides. Clin Exp Immunol (2005) 141:201–10.[CrossRef][Web of Science][Medline]
  29. Schmitt WH, Hagen EC, Neumann I, Nowack R, Flores-Suarez LF, van der Woude FJ, European Vasculitis Study Group. Treatment of refractory Wegener's granulomatosis with antithymocyte globulin(ATG): an open study in 15 patients. Kidney Int (2004) 65:1440–8.[CrossRef][Web of Science][Medline]
  30. Vogt S, Iking-Konert C, Hug F, Andrassy K, Hansch GM. Shortening of telomeres: evidence for replicative senescence of T cells derived from patients with Wegener's granulomatosis. Kidney Int (2003) 63:2144–51.[CrossRef][Web of Science][Medline]
  31. Zhou Y, Huang D, Paris PL, Sauter CS, Prock KA, Hoffman GS. An analysis of CTLA-4 and proinflammatory cytokine genes in Wegener's granulomatosis. Arthritis Rheum. (2004) 50:2645–50.[CrossRef][Web of Science][Medline]
  32. Abdulahad WH, van der Geld YM, Stegeman CA, Kallenberg CG. Persistent expansion of CD4+ effector memory T cells in Wegener's granulomatosis. Kidney Int (2006) 70:938–47.[CrossRef][Web of Science][Medline]
  33. Wagner C, Hänsch M. Receptors for complement C3 on T-lymphocytes: relics of evolution or functional molecules? Mol Immunol (2006) 43:22–30.[CrossRef][Web of Science][Medline]
  34. McFarland HI, Nahill SR, Maciaszek JW, Welsh RM. CD11b (Mac-1): a marker for CD8+ cytotoxic T cell activation and memory in virus infection. J Immunol (1992) 149:1326–33.[Abstract]
  35. Christensen JE, Andreasen SO, Christensen JP, Thomsen AR. CD11b expression as a marker to distinguish between recently activated effector CD8(+) T cells and memory cells. Int Immunol (2001) 13:593–600.[Abstract/Free Full Text]
  36. Fiorentini S, Licenziati S, Alessandri G, et al. CD11b expression identifies CD8+CD28+ T lymphocytes with phenotype and function of both naive/memory and effector cells. J Immunol (2001) 166:900–7.[Abstract/Free Full Text]
  37. Luqmani R, Bacon P, Moots R, et al. Birmingham vasculitis activity score (BVAS) in systemic necrotizing vasculitis. Q J Med (1994) 87:671–8.[Web of Science]
  38. Jennette JC, Falk RJ, Andrassy K, et al. Nomenclature of systemic vasculitis: the proposal of an international consensus conference. Arthritis Rheum (1994) 37:187–92.[Web of Science][Medline]
  39. Leavitt RY, Fauci AS, Bloch DA, et al. The American College of Rheumatology 1990 Criteria for the classification of Wegener's granulomatosis. Arthritis Rheum (1990) 33:1101–7.[Web of Science][Medline]
  40. Reinhold-Keller E, Kekow J, Schnabel A, et al. Influence of disease manifestation and antineutrophil cytoplasmic antibody titer on the response to pulse cyclophosphamide therapy in patients with Wegener's; granulomatosis. Arthritis Rheum (1994) 37:919–24.[Web of Science][Medline]
  41. de Groot K, Gross WL, Herlyn K, Reinhold-Keller E. Development and validation of a disease extent index for Wegener's granulomatosis. Clin Nephrol (2001) 55:31–8.[Web of Science][Medline]
  42. Schlesier M, Kaspar T, Gutfleisch J, Wolff-Vorbeck G, Peter HH. Activated CD4+ and CD8+ T-cell subsets in Wegener's granulomatosis. Rheumatol Int (1995) 145:213–9.
  43. Moosig F, Czernok E, Wang G, Gross WL. Costimulatory molecules in Wegner's granulomatosis (WG): lack of expression of CD28 and preferential up-regulation of its ligands B7-1 (CD80) and B-2 (CD86) on T-cells. Clin Exp Immunol (1998) 114:113–8.[CrossRef][Web of Science][Medline]
  44. Marinaki S, Neumann I, Kälsch AI, et al. Abnormalities of CD4 T cell subpopulations in ANCA-associated vasculitis. Clin Exp Immunol (2005) 140:181–9.[CrossRef][Web of Science][Medline]
  45. Porges A, Redecha P, Kimberly W, et al. Anti-neutrophil cytoplasmic antibodies engage and activate human neutrophils via Fc gamma RIIa. J Immunol (1994) 153:1271–80.[Abstract]
  46. Savill J. Apoptosis in resolution of inflammation. J Leukoc Biol (1997) 61:375–80.[Abstract]
  47. Savill J, Haslett C, Hellewell PG, Williams TJ. Fate of neutrophil. Immunopharmacology of neutrophils. (1994) San Diego: CA Academic. 295–314.
  48. Dallegri F, Ottonello L. Tissue injury in neutrophilic inflammation. InflammRes (1997) 46:382–91.
  49. Edwards SW, Moulding DA, Derouet R, Moots RJ. The neutrophil: an emerging regulator of inflammatory and immune response. Chem Immunol Allergy—Cassatella M, ed. (2003) 83:204–24. Basel: Karger.[Medline]
  50. Hänsch GM, Wagner C. Expression of MHC class II antigen and coreceptor molecules in polymorphonuclear neutrophils. Chem Immunol Allergy (2003) 83:45–63.[Medline]
  51. Giscombe R, Nityanand S, Lewin N, Grunewald J, Lefvert AK. Expanded T cell populations in patients with Wegener's granulomatosis: characteristics and correlates with disease activity. J Clin Immunol (1998) 18:404–13.[CrossRef][Web of Science][Medline]
  52. Ludviksson BR, Sneller MC, Chua KS, et al. Active Wegener's granulomatosis is associated with HLA-DR+ CD4+ T cells exhibiting an unbalanced Th1-type T cell cytokine pattern: reversal with IL-10. J Immunol (1998) 160:3602–9.[Abstract/Free Full Text]
  53. Komocsi A, Lamprecht P, Csernok E, et al. Peripheral blood and granuloma CD4(+)CD28(-) T cells are a major source of interferon-gamma and tumor necrosis factor-alpha in Wegener's granulomatosis. Am J Pathol (2002) 160:1717–24.[Abstract/Free Full Text]
  54. Spriewald BM, Witzke O, Wassmuth R, et al. Distinct tumour necrosis factor alpha, interferon gamma, interleukin 10, and cytotoxic T cell antigen 4 gene polymorphisms in disease occurrence and end stage renal disease in Wegener's granulomatosis. Ann Rheum Dis (2005) 64:457–61.[Abstract/Free Full Text]
  55. Lamprecht P, Moosig F, et al. CD28 negative T-cells are enriched in granulomatous lesions of the respiratory tract in Wegener's granulomatosis. Thorax (2001) 56:751–7.[Abstract/Free Full Text]
  56. Lamprecht P, Bruhl H, Csernok E, et al. Differences in CCR5 expression on peripheral blood CD4+CD28- T-cells and in granulomatous lesions between localized and generalized Wegener's granulomatosis. Clin Immunol (2003) 108:1–7.[CrossRef][Web of Science][Medline]
  57. Brouwer E, Stegeman C, Huitema M, et al. T cell reactivity to proteinase 3 and myeloperoxidase in patients with Wegener's granulomatosis (WG). Clin Exp Immunol (1994) 98:448–53.[Web of Science][Medline]
  58. Popa ER, Franssen CF, Limburg PC, Huitema MG, Kallenberg CG, Tervaert JW. In vitro cytokine production and proliferation of T cells from patients with anti-proteinase 3- and antimyeloperoxidase-associated vasculitis, in response to proteinase 3 and myeloperoxidase. Arthritis Rheum (2002) 46:1894–904.[CrossRef][Web of Science][Medline]
  59. van der Geld YM, Huitema MG, Franssen CF, van der Zee R, Limburg PC, Kallenberg CG. In vitro T lymphocyte responses to proteinase 3 (PR3) and linear peptides of PR3 in patients with Wegener's granulomatosis (WG). Clin Exp Immunol (2000) 122:504–13.[CrossRef][Web of Science][Medline]
  60. Winek J, Mueller A, Csernok E, Gross WL, Lamprecht P. Frequency of proteinase 3 (PR3)-specific autoreactive T cells determined by cytokine flow cytometry in Wegener's granulomatosis. J Autoimmun (2004) 22:79–85.[CrossRef][Web of Science][Medline]
  61. Wagner C, Hänsch GM, Stegmaier S, Denefleh B, Hug F, Schoels M. The complement receptor 3, CR3 (CD11b/CD18) on T lymphocytes: activation-dependent up-regulation and regulatory function. Eur J Immunol (2001) 31:1173–80.[CrossRef][Web of Science][Medline]
  62. Vallejo AN. CD28 extinction in human T cells: altered functions and the program of T-cell senescence. Immunol Rev (2005) 205:158–69.[CrossRef][Web of Science][Medline]
  63. Fanger NA, Liu C, Guyre PM, et al. Activation of human T cells by major histocompatability complex class II expressing neutrophils: proliferation in the presence of superantigen, but not tetanus toxoid. Blood (1997) 89:4128–35.[Abstract/Free Full Text]
Submitted 5 September 2007; revised version accepted 11 January 2008.
Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?



This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow All Versions of this Article:
47/5/609    most recent
ken028v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrowRequest Permissions
Right arrow Disclaimer
Google Scholar
Right arrow Articles by Iking-Konert, C.
Right arrow Articles by Hänsch, G. M.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Iking-Konert, C.
Right arrow Articles by Hänsch, G. M.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?