Gunnar Sturfelt, Department of Rheumatology, University Hospital of Lund, S-221 85 Lund, Sweden. E-mail: firstname.lastname@example.org
The most likely source of autoantigens in systemic lupus erythematosus (SLE) is apoptotic material. Because increased levels of circulating apoptotic cells are found in SLE we wanted to investigate the capacity of serum from patients with SLE or other autoimmune or infectious diseases and normal healthy donors (NHD) to induce apoptosis in normal monocytes, lymphocytes and corresponding cell lines, in relation to clinical and immunological data. Monocytes and lymphocytes from healthy donors were incubated with sera from 37 SLE patients, 37 sex- and age-matched NHD and sera from patients with rheumatoid arthritis, vasculitis, sepsis and mononucleosis. Sera from SLE patients were sampled at both active and inactive disease. The apoptosis-inducing effect (AIE) of these sera was monitored with flow cytometry using annexin V and propidium iodide (PI) binding. The AIE in monocytes and lymphocytes was significantly higher in sera from SLE patients than in other patient groups and NHD (P < 0·001) and was also higher when cell lines were used. Level of C5a in cell culture supernatant correlated with AIE in monocytes (r = 0·451, P = 0·005), suggesting involvement of complement. Heat-inactivation of sera did not affect the AIE, nor did depletion of IgG by protein G absorption of serum. Kinetic analyses showed a peak in apoptosis induction at 12–16 h, with a delayed PI positivity. AIE was equally high using sera from active and inactive SLE cases, and did not correlate with the SLE Disease Activity Index (SLEDAI). Thus, SLE serum has a strong and apparently disease-specific apoptosis-inducing capacity, which could contribute to a high load of potential autoantigen.
Systemic lupus erythematosus (SLE) is an autoimmune disease characterized by typical involvement of many different organ systems and by immunological abnormalities, notably hyperactive B cells producing various autoantibodies. The aetiopathogenesis of SLE includes both genetic and environmental factors, but is largely unknown. However, the role of apoptosis in the pathogenesis of this disease has been increasingly acknowledged.
In human SLE there are data suggesting both increased apoptosis and defective clearance of apoptotic cell material. It has been demonstrated that mononuclear cells from SLE patients display an accelerated rate of apoptosis and altered expression of apoptosis related proteins in vitro[1–3], and that serum from SLE patients contain soluble circulating factors that induce apoptosis . In addition, some of the exogenous factors that are known to induce apoptosis can also induce SLE or exacerbate the disease, including ultraviolet radiation (UVR) , certain drugs [6,7] and virus infections .
Monocyte-derived macrophages from SLE patients also show impaired phagocytosis of apoptotic bodies and cellular debris when compared to macrophage engulfment capacity in normal individuals . With a reduced phagocytic function the apoptotic cell material is not removed properly and the ineffective scavenging process produces a proinflammatory environment. A clustering and concentration of lupus autoantigens (nucleosomal DNA, SS-A/B, snRNP) in the surface blebs of UVR-induced apoptotic keratinocytes have been demonstrated . During the apoptotic process these nuclear autoantigens may be cleaved and processed in such a way that cryptic or neo-epitopes emerge to which the immune system is not tolerant . The apoptotic cellular material is thus present for a prolonged time in the circulation and exposed to the immune system, which would contribute to the development of autoimmunity.
Mechanisms explaining this impaired phagocytosis of apoptotic material involve the complement system, and especially C1q [12,13]. However, less is known about the increased apoptosis rate of circulating karyotic cells in SLE and soluble factors with apoptosis-inducing capacity  have not been identified. The main objectives of this investigation were to confirm and extend these previous observations by comparing the capacity of serum from patients with SLE, rheumatoid arthritis (RA), systemic vasculitis, infectious diseases and normal control subjects to induce apoptosis in monocytes and lymphocytes, and to relate this to clinical and immunological data.
PATIENTS AND METHODS
Preparation of monocytes and lymphocytes
Peripheral blood mononuclear cells were obtained from fresh heparinated blood samples from three different donors (healthy laboratory personnel). The mononuclear cells were isolated by density gradient centrifugation on Lymphoprep™ (Axis Shield Poc AS, Oslo, Norway) at 605 g for 30 min. The lymphocytes and monocytes thus obtained were washed three times in RPMI-1640 medium with l-glutamine (PAA Laboratories GmbH, Linz, Austria) and 0·1% human serum albumin (Sigma, St Louis, MO, USA) (medium) and centrifuged each time at 605 g for 5 min.
The fraction of lymphocytes and monocytes obtained according to this procedure was resuspended in medium with 15% normal human serum (NHS) added to a concentration of 4 × 106 cells/ml. Flow cytometry (Epics XL-MCL Beckman-Coulter, FL, USA) analysis on these cells by detection of cell surface CD14 and CD45 showed that approximately 10% of the cells were monocytes. Eight hundred µl of this cell-suspension was plated on a chamber slide 4 well glass slide (Nalge Nunc International, IL, USA) at 37°C in an atmosphere containing 5% CO2 and 96% humidity for 1 h in order for the monocytes to adhere. Non-adherent cells were removed by washing three times with medium. Flow cytometry analysis of these non-adherent cells showed that at least 90% were lymphocytes, and these cells were therefore used as source of lymphocytes.
Incubation of monocytes and lymphocytes with patient or normal sera
The cells adherent to the glass slides (>80% monocytes as assessed by flow cytometry) were incubated for 16 h at 37°C, 5% CO2, 96% humidity, with serum from patients or from normal human donors (NHD) (20% serum in medium, 0·6 ml/chamber). As a positive control, staurosporine 5 µm (Sigma) in medium and 20% NHS was used, and as a negative control serum from one healthy donor was used. After incubation the spontaneously detached cells were removed. The cells remaining adherent were also detached by first adding phosphate buffered saline (PBS) pH 7·2 (0·0062 m sodium phosphate, 0·15 m NaCl) and then 0·5 mm EDTA-PBS for 3 min at room temperature. The cells were washed twice in PBS pH 7·2 containing 0·1% HSA. All cells from each well were subsequently analysed by flow cytometry for annexin V and propidium iodide (PI) binding. The cell supernatant was used for further analyses (see below). Lymphocytes were purified as described above and 1·5 × 106 cells/ml were incubated for 16 h together with medium and 20% test serum in a solution with a total volume of 0·2 ml. The cells were washed twice in PBS pH 7·2 containing 0·1% HSA, and then analysed by flow cytometry for annexin V and PI binding.
Binding of annexinV–fluoroisothiocyanate (FITC) and propidium iodide (PI) (PharMingen, Becton Dickinson, San Diego, CA, USA) to the cells was used to detect viable, early apoptotic and late apoptotic or necrotic cells by flow cytometry. As shown by kinetic studies (see Results), cells first entered the early apoptotic stage, followed by the late apoptotic or necrotic stage. There were no indications that any serum could induce necrosis directly. Therefore, cells positive for annexin V and a combination of annexin V and PI were scored as apoptotic. Unless stated otherwise, the apoptosis-inducing effect (AIE) is defined as the percentage of apoptotic cells induced by the test serum minus the percentage of apoptotic cells induced by the negative control (serum from one NHD). All sera were analysed at least twice. The mean percentage of apoptotic monocytes induced by the reference serum was 11·2% in 25 experiments (s.d. 3·8) and in lymphocytes 6·6% (s.d. 2·4). The mean AIE of the positive control was 51·4% in monocytes (s.d. 9·8), and 58·2% in lymphocytes (s.d. 13·5), yielding coefficient of variation (CV) values for monocytes and lymphocytes of 19% and 23%, respectively. One SLE serum with high AIE was tested simultaneously on the three different cell donors yielding mean AIE in monocytes 68·5% (s.d. 8·6) and mean AIE in lymphocytes 34·1% (s.d. 3·8). The same experiment was repeated with a SLE serum with low AIE; mean monocyte AIE was 1·4% (s.d. 3·2), and mean lymphocyte AIE was − 2·6% (s.d. 1·0).
To confirm that cells were apoptotic, monocytes on glass slides were stained in May–Grünwald–Giemsa (Merck, Darmstadt, Germany) and their morphology was studied by light microscopy (see Results). Forward and side scatter plots were also used to confirm that cells were apoptotic, as well as Caspase 3 activity as detected by flow cytometry (Intergene, Oxford, UK) (see Results).
Two SLE sera with a high AIE and two normal sera were selected for IgG absorption on protein G sepharose (Amersham Pharmacia Biotech, Uppsala, Sweden). The concentration of IgG was measured both before and after the protein G sepharose absorption by electroimmunoassy using rabbit antihuman IgG and the human serum protein calibrator no. X 0908 (Dako A/S, Glostrup, Denmark) was used as a standard.
The untreated SLE sera contained 19·5 and 16·0 g/l, respectively, and the normal sera 9·6 and 5·0 g/l of IgG, respectively. After protein G sepharose treatment, IgG concentrations in all the sera were below 0·1 g/l. AIE of treated and untreated sera was measured as described above.
In order to test the effect of heat inactivation, serum from two SLE patients with high AIE was incubated for 30 min at 56°C and tested.
Patients and controls
SLE patients taking part in a prospective control programme at Department of Rheumatology, Lund University Hospital, Sweden were studied. Sera sampled at a period of active disease from 37 consecutive patients were investigated. In 35 of these patients, sera sampled from a period of inactive disease were also used. The median number of American College of Rheumatology (ACR) classification criteria  fulfilled was six (four to 10). Median age was 40 years (range 11–72 years) and 34 of the 37 patients were women. Blood sampling was performed in a standardized manner, all samples being stored for 1 h at room temperature before centrifugation. Disease activity was determined according to the SLE Disease Activity Index 2000 (SLEDAI-2K)  and the median SLEDAI score in active disease was 10 points (2–24). Current medication with prednisolone, hydroxychloroquine, other immunosuppressive agents (azathioprine, cyclophosphamide, cyclosporin A, mycophenolate mofetil and methotrexate) was recorded both at the time-points of active and inactive disease. Disease manifestations at flare and medication are presented in Table 1. Sera from 37 age- and sex-matched healthy blood donors, 30 patients with active rheumatoid arthritis (RA), 10 patients with primary systemic vasculitis (eight Wegener's granulomatosis, one Churg–Strauss, one Mb Behçet), 10 previously healthy individuals with infectious mononucleosis (Epstein–Barr virus infection) and 10 previously healthy individuals with Streptococcus pneumoniae septicaemia were used as controls.
Table 1. Clinical manifestations and SLEDAI scores at flare of the 37 SLE patients and their treatment with prednisolone (Predn), hydroxychloroquine, and other immunosuppressive agents at flare are shown
AIE (%) mono
AIE (%) lymph
Clinical manifestations at flare
AZA = azathioprine, CTX = cyclophosphamide, CYA = cyclosporine A, HCG = hydroxychloroquine, MYC = mycofenolatmofetil, IvIg = intravenous immunoglobulin, MTX = methotrexate. Apoptosis-inducing effect (AIE) of SLE serum on monocytes (mono) and lymphocytes (lymph) are also shown.
Routine laboratory tests included CRP, complement components (C1q, C3 and C4), anti-dsDNA antibodies, full blood count and urinary analyses. Low levels of complement components were defined as values below the reference interval at Department of Clinical Immunology, University Hospital, Lund, Sweden; C1q (<76% of the normal), C3 (<0·77 g/l) and C4 (<0·12 g/l). IgG anti-dsDNA antibodies were measured with a commercially available enzyme linked immunosorbent assay (ELISA) (Euroimmun, Lübeck, Germany). Serum concentrations of complement component C1q was determined by electroimmunoassay, C3 and C4 by nephelometry, and C5a with a commercially available ELISA (Pharmingen, San Diego, CA, USA). Tumour necrosis factor-alpha (TNF-α) in serum and cell culture supernatant was determined with Immulite®, an automated chemiluminescent enzyme immunoassay (DPC, Gwynedd, UK). IgG anticardiolipin (aCL) antibodies were measured by ELISA according to methods used routinely at Department of Clinical Immunology, University Hospital, Lund, Sweden. Soluble Fas-ligand (sFasL) was measured in serum with a commercially available ELISA (MBL, Nagoya, Japan).
Monocytoid and lymphocytoid cell lines
The monocytoid U937 cell line and the lymphocytoid Jurkat T-cell line were used.
The cell lines were cultured at 37°C, 96% humidity and 5% CO2 using RPMI-1640 medium supplemented with 10% fetal calf serum, Na-pyruvate and non-essential amino acids as a growth medium. The medium was exchanged every third day and the cells were subcultured every seventh day.
Before incubation of cell lines with test serum, the cells were washed twice in RPMI-1640 containing 0·1% HSA and the cell concentration was adjusted to 2 × 105 cells/ml. Cells were incubated with 20% test serum for 16 h, in a total volume of 200 µl, at 37°C, 96% humidity and 5% CO2. As controls NHS and NHS containing 5 µm staurosporine were used, in addition to serum from three patients with RA and three NHD. After incubation, the cells were washed twice in PBS pH 7·2 containing 0·1% HSA and analysed by flow cytometry for annexin V and PI binding as described above.
Mann–Whitney U-test was used for group comparisons and Spearman ranks test for correlations. Comparisons of values in active disease with respect to inactive disease were made with Wilcoxon's signed ranks test for two dependent samples.
Apoptosis was induced in monocytes and lymphocytes by sera [at a concentration 20% (v/v)] from two SLE patients (nos 27 and 33) (Fig. 1a). As a positive control 5 µm staurosporine added to NHS was used, and as negative control serum from a NHD. The peak of apoptosis was reached rapidly in lymphocytes after 12 h. With monocytes the highest apoptosis level was reached after 16 h. It could also be demonstrated that annexin-V binding in monocytes precedes PI reactivity, supporting the idea that SLE serum induces apoptosis and not primary necrosis (Fig. 1b). The same pattern was observed in lymphocytes (data not shown). Morphological studies by light microscopy confirmed that monocytes were apoptotic; these cells were shrunken and displayed nuclear condensation fragmentation (Fig. 2a). Apoptosis was also confirmed by forward and side scatter distribution (Fig. 2b). In addition, Caspase 3 activity was demonstrated in monocytes incubated with sera from two patients with SLE (78 and 61%, respectively). In contrast, sera from two NHD induced Caspase 3 activity in 5% and 1% of monocytes, respectively.
The percentage of apoptotic cells induced by different serum concentrations (2·5, 5, 10, 15 and 20%) was also investigated using serum from two SLE patients and one normal healthy donor. The lowest concentration of SLE serum, 2·5%, was sufficient to induce a high degree of apoptosis in both monocytes and lymphocytes. However, this serum concentration was also associated with the highest standard deviation regarding monocytes (data not shown). No clear-cut dose–response effect could be seen within this concentration interval.
The results from the analyses of apoptosis-inducing effect (AIE) of serum from patients with SLE, RA, vasculitis, sepsis, mononucleosis and NHD are shown in Fig. 3. The AIE on monocytes and lymphocytes was significantly higher with serum from SLE patients compared to all other patient groups and the NHD (P < 0·001). Serum from SLE patients sampled at a time-point with active disease did not differ with regard to AIE on monocytes or lymphocytes from serum of the same patients sampled at a time-point of inactive disease. There was no correlation between SLEDAI and AIE in monocytes (r = 0·139, P = 0·414) or lymphocytes (r = 0·223, P = 0·185).
Patients with active glomerulonephritis did not differ in AIE compared to patients lacking this manifestation (data not shown). In Table 1 treatment with prednisolone, hydroxychloroquine and other immunosuppressive agents at flare is shown, as well as SLEDAI scores and AIE. There was no correlation between dosage of prednisolone and AIE of monocytes (r = 0·028, P = 0·872) or lymphocytes (r = 0·149, P = 0·378). Patients treated with hydroxychloroquine did not differ in AIE compared to the patients without such treatment (data not shown). Serum from the nine patients treated with any kind of other immunosuppressive treatment could induce slightly more apoptosis compared to serum from patients without such treatment in lymphocytes (P = 0·033), but not in monocytes (P = 0·392).
In Table 2 the relationship between complement components, autoantibodies, sFasL, TNF-α and AIE of serum from patients with active SLE disease are shown. Patients with low C1q and/or low C4 levels had particularly high AIE for both monocytes and lymphocytes. This was also seen for serum with low C3 levels, but in this case the correlation was less pronounced. Levels of C5a in cell culture supernatant correlated with the AIE using monocytes, but much less for lymphocytes. With regard to the serum concentration of auto-antibodies, weak correlations were observed between IgG antidsDNA antibody levels and AIE on monocytes and between IgG aCL antibody levels and lymphocyte apoptosis. A weak correlation could also be seen between sFasL levels and AIE on lymphocytes. Levels of TNF-α in cell culture supernatants, but not in tested sera, correlated with AIE in both monocytes and lymphocytes.
Table 2. The relationship between levels of complement components C1q, C4, C3, the complement split product C5a, IgG antidsDNA and anticardiolipin antibodies, soluble Fas-ligand (sFasL) and TNF-α in serum or cell-culture supernatant and apoptosis-inducing effect (AIE) in lymphocytes and monocytes of serum from patients with active SLE disease. Spearman's rank correlation test was used to calculate correlation coefficient (r) and P-values. The variables in the left column refer to serum if not stated otherwise
AIE (%) monocytes
AIE (%) lymphocytes
The AIE of SLE sera (three sera from patients with active SLE disease and three from inactive SLE) and controls was also tested on the monocytoid cell line U937 and the T cell line Jurkat in order to strengthen the findings from normal monocytes and lymphocytes. None of the sera from three RA patients or from the NHD induced apoptosis in U937 cells or Jurkat cells, as shown in Fig. 4. In contrast, serum from SLE patients had a marked AIE on both these cell lines.
IgG absorption of SLE sera by protein G did not decrease their AIE. The AIE of one untreated SLE serum on monocytes was 60·6%versus 78·7% when IgG was removed, and the other serum gave 57·8% before and 75·3% after treatment. Similar figures were obtained for AIE on lymphocytes (data not shown). AIE on monocytes with NHS was – 0·3 before treatment and 7·7 after treatment. Heat inactivation (HI) of two SLE sera did not influence the AIE; AIE on monocytes was 66·8% before and 56·5% after HI in one serum and 32·7% before and 32·4% after HI in the other SLE serum. Similar figures were obtained from lymphocyte apoptosis studies (data not shown).
Apoptotic cells have been suggested to be a major source of autoantigens in SLE since clustering and concentration of lupus autoantigens in the surface blebs of apoptotic cells have been demonstrated . These autoantigens are apparently processed further in such a way that neo-epitopes will emerge to which the immune system is not tolerant. It has become increasingly acknowledged that impaired clearance of such apoptotic material is of great importance in the pathogenesis of SLE .
There are few reports of an increased rate of apoptosis in human SLE. In the present investigation we demonstrate that serum from SLE patients induces apoptosis in monocytes and lymphocytes from normal healthy donors, as well as in the monocytoid cell line U937 and the T cell line Jurkat. The induction of apoptosis could not be observed with sera from patients with other autoimmune diseases such as RA or systemic vasculitis, or infectious diseases such as mononucleosis or sepsis, or healthy individuals. Thus the AIE observed is not related to non-specific inflammatory events, and seems to be specific for SLE.
The results of the present investigation extend and confirm previous findings from our laboratory . The possibility of pro-apoptotic mechanisms operating in human SLE is supported further by findings of Fas-dependent accelerated apoptosis in vitro of monocytes/macrophages from SLE patients  and similar findings in lymphocytes [1,2], T cell lines  and in CD34+ stem cells exposed to SLE serum . Increased levels of circulating apoptotic monocytes , lymphocytes [3,20] and neutrophil granulocytes  in SLE patients have been reported which might also reflect pro-apoptotic events, although impaired clearance of such apoptotic cells would yield the same outcome. Thus, our current data demonstrating apoptosis inducing factors in serum of patients with SLE are in accordance with the findings of others. Together with impaired clearance of apoptotic material, a high rate of apoptosis would result in an increased load of circulating autoantigen, which could lead ultimately to immune response and the formation of immune complexes deposited in tissues causing disease.
Even though the cell cultures of monocytes and lymphocytes have high purity, it could be speculated whether interactions of, for example, a small lymphocyte subpopulation and monocytes could influence the apoptosis induction. The marked apoptosis induction by SLE sera in both Jurkat T cell line and U937 monocyte cell line indicate strongly, however, that the apoptosis seen in monocytes and lymphocytes is not mediated by cell–cell contact.
Complement component C1q has been shown to be of vital importance in clearance of apoptotic material . It is also well established that low levels of C1q are associated with severe SLE flares, especially with kidney involvement, as a result of immune complex deposition . A high apoptosis rate was associated particularly with low C1q and C4 levels, confirming previous data . This could indicate that impaired complement function is due possibly to consumption, and increased amounts of circulating immune complexes are involved in the apoptosis-inducing process. However, the correlation between low C1q levels and high apoptosis rate could be seen only in sera from patients with active disease, and not inactive disease (data not shown). These contradictory findings warrants further studies. Furthermore, serum from patients with active glomerulonephritis did not induce a particularly high apoptosis rate. More importantly, pretreatment of the sera by heat inactivation or protein G absorption did not change the apoptosis rate. Thus, IgG containing immune complexes are obviously not involved directly in the apoptosis induction, and if complement is involved directly a complement split product such as C5a might be a candidate. High levels of C5a in cell culture supernatants were indeed associated with a high apoptosis rate in monocytes. This is of interest because it has been demonstrated that systemic activation of complement induces C5a-dependent apoptosis of thymocytes and that the blockade of C5a during sepsis rescues thymocytes from apoptosis .
Other serum factors of potential interest as apoptosis inducers are autoantibodies, because purified autoantibodies such as anti-dsDNA and anticardiolipin antibodies do induce apoptosis in vitro[25,26]. In the present investigation there were, however, only weak correlations between degree of apoptosis and levels of these autoantibodies. This is in agreement with the lack of effect of IgG removal by protein G absorption in this study. Serum from patients with severe bacterial infection has been shown to contain increased amounts of soluble Fas-ligand (sFasL) which induce apoptosis in neutrophil granulocytes in vitro. However, no relation of sFasL levels in serum and apoptosis could be observed in this study, and none of the sera from patients with sepsis could induce apoptosis in monocytes or lymphocytes. Notably, no relationship between current medication and rate of apoptosis could be seen, suggesting that drugs are not of importance as apoptosis inducers in the present investigation.
There are many cytokines with potential implications in the current test system, acting either directly to induce apoptosis or being produced by the monocytes and lymphocytes exposed to the SLE sera, causing secondary effects on neighbouring cells. In the present investigation a relationship between high amounts of TNF-α in cell culture supernatants and apoptosis in monocytes could indeed be observed. Hypothetically, a soluble factor in SLE serum could activate monocytes to produce TNF-α which could induce apoptosis in neighbouring cells via TNF receptor I. Ongoing experiments will clarify whether or not the apoptosis induction is death-receptor mediated.
Notably, serum from patients with inactive disease could induce apoptosis to the same degree as serum from the same patients with active disease. Furthermore, in patients with active disease there was no correlation with the global disease activity index SLEDAI. The patients also represented a broad spectrum of clinical manifestations and degree of disease severity that can be seen in SLE, but no apparent clinical manifestation could be associated with high apoptosis. The lack of association between apoptosis induction, clinical disease activity and clinical subsets may indicate that there are several mechanisms operating.
Taken together, serum from SLE patients has a strong apoptosis-inducing effect on monocytes, lymphocytes and corresponding cell lines. The mechanisms need, however, to be investigated further and serum factors responsible for these effects need to be identified. Because apoptotic material probably constitutes the major autoantigen source in SLE, the presence of endogenous circulating pro-apoptotic factors could contribute to an increased load of autoantigen. This, together with impaired clearance of such material, could lead to a subsequent immune response and the formation of immune complexes deposited in tissues causing clinically detectable disease. Thus, the apoptosis-inducing effect we describe may contribute to the pathogenesis of the disease and may be a target for future therapeutic agents.
Technical assistance from Gertrud Hellmer and Annica Andreasson, and help with blood sampling by Lotta Larsson, are gratefully acknowledged. Sera from patients with infectious diseases were kindly provided by Dr Göran Jönsson and Dr Jean-Henrik Braconier, Department of Infectious Diseases, University Hospital, Lund, Sweden. This study was supported by grants from the Swedish National Association against Rheumatism, the Swedish Medical Research Council (grant no. 13489), the Medical Faculty of the University of Lund, Alfred Österlunds Stiftelse, Crafoords Stiftelse, Greta och Johan Kocks Stiftelser, Konung Gustaf V 80-års fond, Lunds Sjukvårdsdistrikt and Professor Nanna Svartz Stiftelse.