Introduction
Patients with the autoimmune disorder systemic lupus erythematosus (SLE) exhibit, for yet non-clarified reasons, a decreased ability to degrade DNA. The phenomenon was first observed in the 1960s [
1], and has recently regained new interest with the discovery of neutrophil extracellular traps (NETs) [
2]. NETs consist of chromatin covered with antimicrobial proteins and constitute a candidate autoantigen target in SLE. SLE is characterized by an autoimmune reaction against many nuclear antigens historically proposed to originate from apoptotic cells that are not properly cleared [
3]. DNA in various forms is degraded by specific nucleases where DNase-I is the main enzyme responsible for degrading DNA and chromatin released into serum. The role of DNase-I in SLE received attention when it was discovered that the presence of a DNase-I inhibitor correlated with levels of nuclear autoantibodies [
1] and that SLE patients had decreased serum nuclease activity [
4]. Attempts were subsequently made to restore the activity by infusion of recombinant DNase-I but without reaching a sufficient serum concentration to lead to clinical improvements [
5]. Recently, it was confirmed that sera from a subgroup of SLE patients do not degrade NETs [
6,
7].
Although NETs pose a highly interesting target in SLE, other chromatin sources like apoptotic and necrotic cells should not be forgotten. The importance of serum nucleases for the degradation of apoptotic and necrotic cells has been studied previously and it appears that this process is dependent on additional cofactors, such as complement C1q [
8], serum amyloid P [
9], factor VII-activating protease [
10] and plasmin [
11] - at least for degradation of necrotic chromatin. These cofactors are thought to open the normally condensed chromatin structure by displacing histone H1. NETs consist of decondensed, open chromatin and therefore degradation is not dependent on such cofactors. In our previous study we instead observed that C1q inhibited degradation of NETs and hypothesized this to be a trade-off for opsonisation [
6], which was later confirmed by another group [
12]. The current literature hence proposes that multiple factors are important in the degradation of DNA depending on the source and nature of the chromatin. Patients with SLE have been described to have a decreased DNase activity [
4] but how that relates to different forms of DNA is still unclear. We therefore set out to investigate how patients with SLE degrade DNA from a range of clinically relevant sources. The aim was to generate a more comprehensive image of DNA degradation in SLE and determine what sources of DNA most likely are involved in disease pathology. In the study, we focused on DNA sources with known serum nuclease-dependant degradation and used DNA in the form of NETs as well as chromatin from both primary and secondary necrotic cells and compared to degradation of purified DNA using a zymographic approach. The results additionally led us into some fundamental characterization of the interactions between DNase-I and serum proteins.
Methods
Patients and sera
A total of 66 SLE patients (5 men and 61 women) with a median age of 39 years (range 18–75), fulfilling at least four American College of Rheumatology (ACR) 1982 classification criteria for SLE [
13] were recruited at the Clinic of Rheumatology, Skåne University Hospital in Lund (Sweden). The distribution of ACR classification criteria for SLE is described in Table
1. Disease activity was recorded using the SLE disease activity index 2000 (SLEDAI-2K) [
14]. Sera from 103 healthy volunteers with matched age and sex were used as controls in the study. All patients and healthy controls gave informed consent to participate in the study, which was approved by the local ethics committee (Lund University) according to the Helsinki declaration.
Table 1
Patient characteristics according to American College of Rheumatology (ACR) criteria
Malar rash | 44 (66.7) |
Discoid rash | 18 (27.3) |
Photosensitivity | 42 (63.6) |
Oral ulcer | 16 (24.2) |
Arthritis | 53 (80.3) |
Serositis | 35 (53.0) |
Nephritis | 32 (48.5) |
Neurological disorder | 3 (4.5) |
Hematological disorder | 37 (56.1) |
Immunological disorder | 51 (77.3) |
Anti-nuclear antibodies | 66 (100) |
Degradation of cell chromatin
Jurkat T-cells (ATCC, Manassas, VA, USA) were cultured in Roswell Park Memorial Institute medium (RPMI) with 10 % foetal calf serum at 37 °C and 5 % CO
2. For experiments, the cells were washed twice, kept in RPMI and either used directly as live cells alternatively rendered apoptotic, primary or secondary necrotic. Apoptosis was induced by incubation with 1 μM staurosporine (Sigma-Aldrich, St Louis, MO, USA) at 37 °C for 3 h. Primary necrosis was induced through incubation in 15 % EtOH at 37 °C for 1 h and secondary necrosis was induced with 20 μM oxaliplatin (Teva, Petach Tikva, Israel) at 37 °C for 48 h as previously described [
15]. The cell states were confirmed with Annexin V (Immunotools, Friesoythe, Germany) and Via-Probe (BD, San Jose, CA, USA) staining. For degradation experiments, cells were washed and incubated with 1.5 % (for degradation of primary necrotic chromatin) or 3 % (for degradation of secondary necrotic chromatin) patient sera, pooled normal human serum (NHS) or fractionated sera for 3 h at 37 °C in 10 mM Tris-HCl, pH 7.5, 50 mM NaCl, 10 mM MgCl
2 and 2 mM CaCl
2 (DNase buffer). To analyze DNA degradation, we used Hoechst (Invitrogen, Waltham, MA, USA) or Via-Probe to detect cellular double stranded DNA (dsDNA). Loss of signal indicates nearly complete degradation to small fragments (below 20 bp). DNA content was analyzed in a CyFlow Space (Partec, Görlitz, Germany) flow cytometer.
Isolation of neutrophils
Neutrophils were isolated from healthy volunteers according to a previously published method [
16]. Briefly, blood from healthy volunteers was separated by centrifugation on a Histopaque 1119 column (Sigma-Aldrich), the granulocyte-rich fraction was isolated and washed, and neutrophils were isolated by centrifugation on a Percoll gradient (65 − 80 %) (GE-healthcare, Fairfield, CT, USA) and isolated from the intersection of the 70 % and 75 % layer, washed and resuspended in RPMI with 10 mM Hepes. Purity of neutrophils (>80 %) was determined by surface marker expression for anti-CD14 (BD), anti-CD15 and anti-CD16 (both from Immunotools) and defined as CD16
+/CD15
+/CD14
low.
Generation and degradation of NETs
Freshly isolated neutrophils, 50,000/sample, were seeded onto a 96-well flat-bottom plate (Nunc, Waltham, MA, USA) with 20 nM PMA (Sigma-Aldrich) for 4 h at 37 °C and 5 % CO2 to generate NETs. After incubation, cell medium was removed and 10 % patient sera, control sera or fractionated sera in DNase buffer was added and incubated for 60 minutes at 37 °C. During this time, degraded NETs were released into solution. Aliquots of the solution containing NETs were then transferred to PBS with a final concentration of 2 mM EDTA to stop further degradation and DNA content was quantified using PicoGreen (Invitrogen). As the internal control, pooled NHS was used and all samples were compared to the mean of the internal controls for each individual experiment. All samples were measured twice, first in duplicates followed by once in singles and the mean of the two measurements was used for analysis.
Gel filtration
Serum diluted to 50 % with or without 500,000 cpm = counts per minute 125I DNase-I (Bioworld, Dublin, OH, USA), labeled using the chloramine T method, was diluted in DNase buffer and separated on a Superose 12 column (GE healthcare) using the ÄKTA system (GE healthcare). The column was washed with DNase buffer and 0.1 ml fractions were collected. For assays using radiolabeled DNase-I, radioactivity was measured in fractions before stored at −80 °C until further used.
DNase zymogram
Sera from SLE patients and controls were diluted in DNase buffer to 30 % and loaded onto a native PAGE gel containing 22 μg/ml denatured calf thymus DNA (Sigma-Aldrich). As internal controls 30 % NHS was used. Gels were run at 90 V for 2 h under native non-reducing conditions, washed with dH2O and incubated with 40 mM Tris–HCl pH 7.5, 8 mM MgCl2, 2 mM CaCl2, 0.02 % NaN3 at 37 °C for 18 h. After this, ethidium bromide (EtBr) was added to a final concentration of 1 μg/ml and gels were incubated at 37 °C for an additional 30 minutes and analyzed in a ChemiDoc™ MP Imaging System (BioRad, Hercules, CA, USA). For complex formation assays, 2.3 μg/ml AF488-labeled DNase-I was incubated with 5–200 μg/ml actin from rabbit skeletal muscle (Sigma-Aldrich) or 150–200 μg/ml Gc-globulin (Sigma-Aldrich) in 10 % NHS or heat-inactivated NHS (Hi-NHS) and run as above.
Statistical analyses
For comparison of chromatin degradation between two groups the Mann-Whitney test was used. For comparison between multiple experimental parameters two-way analysis of variance (ANOVA) was used followed by Bonferroni post hoc test in GraphPad Prism 5 (GraphPad, La Jolla, CA, USA). Association with disease manifestation was tested using Pearson’s chi-square (χ2) test. For principal component analysis, cluster analysis and analysis of association with disease manifestation was performed using JMP 11 (SAS, Cary, NC, USA).
Discussion
For unknown reasons the ability to degrade DNA is decreased in a proportion of patients with SLE. In this report we have systematically investigated the ability of sera from SLE patients to degrade DNA from multiple clinically relevant sources of DNA. We have focused the study on degradation of DNA sources that are dependent on serum nucleases as determined in previous reports [
11,
15] and as shown in Fig.
1. Interestingly, the study reveals that the majority of patients with SLE (61 %,
n = 40) exhibit decreased ability to degrade chromatin from either NETs, primary or secondary necrotic cells by serum. As there are multiple nucleases expressed in different tissues and cellular locations [
21], it is tempting to speculate that studies of other means of degradation such as endogenous degradation of apoptotic chromatin by caspase-activated DNase [
22] as well as other intracellular DNases such as TREX1 [
23,
24] and DNase-gamma [
25] may expand this group further and should be addressed in future studies.
Although there was relatively poor correlation between decreased degradation of different types of chromatin, we did identify a subset of patients with a distinct clinical phenotype associated with a general decrease in NET/chromatin degradation. Patients in cluster 1 mainly displayed a decreased degradation of primary necrotic chromatin, which points in a direction of a separate mechanism of degradation and aligns with previous reports describing cofactor-dependent degradation of the highly condensed chromatin from primary necrotic cells that require decondensation before efficient degradation. Presence of anti-DNA antibodies appears to prevent degradation of the decondensed NETs [
7] and possibly also chromatin from secondary necrotic cells, which aligns with the clinical associations in cluster 3. Additionally, there are also reports of protein-DNA complexes, originating from NETs, being less efficiently degraded in SLE [
26] which require further investigation in relation to other sources of DNA. To address the decreased ability to degrade primary necrotic chromatin in cluster 1, we used western blot to analyze protein levels of some DNase-I cofactors, such as C1q [
8] and factor VII-activating protease [
10]. We did observe a trend towards lower levels in particular of factor VII-activating protease in cluster 1, however this did not reach statistical significance (data not shown). Most likely, a more systematic approach employing more sensitive methods would be required to properly address this issue. The data in this study do suggest such study should focus on patients in cluster 1.
Mice that lack DNase-I develop SLE-like disease [
27] and nephritis in mice and humans is also associated with a decrease in DNase-I expression [
28]. Multiple studies have found decreased nuclease activity in patients with SLE [
4,
29,
30]. However, these studies measured total nuclease activity using radial diffusion or oligonucleotide immune assays that neither separate forms of serum nucleases nor do they take into account the presence of potential nuclease inhibitors in serum. By using a zymographic approach, we separated serum proteins based on charge and therefore reduced the likelihood of interference with inhibitors that are not strongly bound to DNase-I. It is interesting to note that in this setting there was no difference in DNase-I activity between healthy controls and SLE patients, at least not in the investigated cohort. Even though DNase-I mutations in SLE are rare [
31], investigation of such mutations or transcription forms in individuals with low activity may be of interest. The data do however suggest that it is not the DNase-I activity per se that is affected in these SLE patients but rather DNase-I is prevented from degrading chromatin, potentially by sterically blocking access for DNase-I by dsDNA antibodies generated during the disease. Additionally, we cannot rule out that patients with low ability to degrade primary necrotic chromatin produce lower levels of cofactors. Together, the data hence suggest that the decreased degradation observed in SLE is not a predisposing factor but rather a consequence of the autoimmune response most likely further fuelling disease pathology such as kidney disease. This aligns well with the pathology of the DNase
−/− mouse [
32]. We recently found that an inability of SLE patients to degrade NETs is, in the majority of cases, not permanent but usually recovers within a period of 6 months indicating that this is a dynamic process that correlates with disease activity and levels of nuclear autoantibodies [
17].
A pharmacological study in rats first observed that DNase-I binds a serum protein, which most likely is globular actin [
18]. In the present study we can demonstrate that this complex is surprisingly active and appears to be the major form of DNase-I in the serum of healthy individuals. Inhibition of the actin-DNase-I complex is thought to be dependent on ATP, which was not included in our experimental setup. However when we used an ATP analog to confirm inhibition, disassociation of the complex occurred most likely due to polymerization of actin (data not shown). It is possible that this does not occur in vivo where there are actin depolymerizing factors and ATP stabilizing factors. Recently, this issue was addressed using gelsolin to prevent actin polymerization [
33]. Intriguingly, as extracellular ATP is a marker of inflammation it is tempting to speculate that this may prevent NETs from being degraded at a site of inflammation and infection.
Acknowledgements
The author would like to acknowledge all patients and their physicians as well as the following funding bodies: the Swedish Research Council (K2012-66X-14928-09-5), Foundations of Österlund, Greta and Johan Kock, King Gustav V´s 80th Anniversary, Knut and Alice Wallenberg, Inga-Britt and Arne Lundberg, Torsten Söderberg, Royal Physiographic Society in Lund and grants for clinical research (ALF and from the Skåne University Hospital).
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Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
JL, KC and MM performed the experiments and analyzed the results. BG acquired and analyzed clinical parameters and AAB recruited patients and supervised the study together with AMB, and both also analyzed and interpreted the data. JL drafted the manuscript together with AMB. All authors have read, revised and approved the final manuscript.