The pro-inflammatory potential of T cells in juvenile-onset systemic lupus erythematosus

The Liverpool Paediatric Research Ethics Committee approved this study. Written informed consent was obtained from all participating subjects or their parent/guardian. JSLE patients fulfilled the revised American College of Rheumatology (ACR) criteria for the diagnosis of SLE[14] before the age of 17 years. Paediatric non-inflammatory controls were children without inter-current infection or non-inflammatory musculoskeletal symptoms and had undergone elective surgery. All paediatric patients were recruited at Alder Hey Children’s National Health Service (NHS) Foundation Trust. JSLE disease activity data was collected using a paediatric adaptation of the British Isles Lupus Assessment Group (BILAG) 2004 (pBILAG2004) disease activity score[15].

Peripheral blood mononuclear cells (PBMCs) were isolated from fresh heparinised blood within 1 hour of collection by 1-step centrifugation through Polymorph Prep (Axis-shield, UK) following manufacturer’s instructions.

PBMCs were re-suspended at a concentration of 1 × 10⁶ cells/ml in RPMI 1640 medium (Sigma-Aldrich) supplemented with 10% fetal calf serum (Sigma-Aldrich) and 1% penicillin/Streptomycin (Sigma-Aldrich). PBMCs were seeded into a 96 well culture plate (Nunc) at 5 × 10⁵/well in triplicate and stimulated with beads coated with antibodies to CD3/CD28 (2.5 × 10⁵ beads/well; Miltenyi Biotech) producing a final bead-to-cell ratio of 1:2. The PBMC were incubated at 37°C in an atmosphere containing 5% CO₂. After 2 days cell free supernatants were collected and the concentration of IL-17A in supernatants was assessed using an enzyme-linked immunosorbant assay (ELISA) kit according to the manufacturer’s instructions (ebioscience, San Diego, US), the detection limit was 7.8 pg/ml.

IL-17A concentration in plasma samples was measured using a commercially available Human IL-17A Immunoassay Quantikine® ELISA kit (R + D Systems Inc., Minneapolis, MN, USA) following manufacturer’s instructions.

Levels of IL-6, IL-21, IL-22 and IL-17 F in cell culture supernatant were measured using Bio-plex Pro Human Th17 cytokine assay and levels of IL-12, IFNγ, IL-4 and IL-13 were measured in serum samples using a Bio-Plex Pro Human Cytokine 27-Plex Panel (Bio-Rad Laboratories) following the manufacturer’s instructions.

RNA was extracted from PBMCs using the RNeasy Mini kit (Qiagen Inc.). The concentration and purity of RNA was confirmed by the relative absorbance at 260 nm and measuring the 260/280 nm ratio using an ND-1000 NanoDrop spectrometer (Thermo Scientific). First strand cDNA synthesis was initiated from 100 ng total RNA using random hexamers (Promega) and avian myeloblastosis virus reverse transcriptase (Promega) using conditions described by the manufacturer in a final volume of 25 μl. The primers used were as follows: IL17A forward 5′ GAA TCT CCA CCG CAA TGA GGA CCC 3′; reverse 5′ GTT GAT GCA GCC CAA GTG GCG 3′; RORC forward 5′ GTC CCG AGA TGC TGT CAA GT 3′; reverse 5′ TGA GGG TAT CTG CTC CTT GG 3′; IL23R forward 5′ ACA GGG CAC CTT ACT TCT GAC AA 3′; reverse 5′ AGC AAA GAC GAT CAT TCC CAA T 3′; IL23 forward 5′ AGA GGG CAC CTT ACT TCT GAC AA 3′; reverse 5′ AGC AAA GAC GAT CAT TCC CAA T 3′; RPL13A forward 5′ TTT CCA AGC GGC TGA CGA AG 3′; reverse 5′ AGC AAA GAC GAT CAT TCC CAA T 3′. All quantitative real-time PCR took place using the SYBR green fluorescence method with SYBR green qPCR mastermix (Stratagene) as specified by the manufacturer. Real-time PCR reactions took place in triplicate on a MX4000® Multiplex Quantitative QPCR system (Stratagene) using standard thermal cycling conditions. Non-template controls were prepared by replacing the cDNA fraction of the PCR reaction with an equivalent volume of nuclease free water (Promega). RPL13A was monitored as an internal standard for the PBMCs. mRNA expression for each gene was normalised to these internal standards. Absolute quantification was obtained using the standard curve method of analysis.

All data are presented as the mean ± SEM. Comparisons between JSLE and control patients were made using the Mann–Whitney test. All analyses were performed using GraphPad Prism 4 software (Graph Pad Software, San Diego, CA). p < 0.05 were considered significant.

Thirty JSLE patients were studied; Table 1 presents data for the demographic, clinical biomarker of disease activity, physician’s global assessment of disease activity, pBILAG2004 disease activity index and current medications of the patient group. Disease activity at time of sampling varied from mild to severe, with a range of therapeutic modalities being used. Due to small volumes of blood obtained from the paediatric samples, we were unable to use the same group of JSLE patients for every experiment. Additional file1: Table S1–S4 describe patient subsets used in each part of this study.

Fifty-seven paediatric non-inflammatory healthy patients were included as controls mean age 13.1 years (range 2–17 years); of which 27 were female (47%). All were of white British ethnicity.

To determine which T helper cell phenotype are active in vivo during ongoing JSLE disease, cytokine levels in the blood were measured (Figure 1). JSLE patients (n = 19) were found to have a significantly higher level of IL-17A (21.5 ± 5.2 pg/ml) in comparison to controls (n = 18, 7.2 ± 2.5 pg/ml, p = 0.028; Figure 1A). mRNA levels of IL-17A, RORC (gene encoding Th17 transcription factor (RORɣT), the Th17 stabilising cytokine IL-23 and its receptor IL-23R were measured in JSLE and control PBMCs (Figure 1B). The relative expression of IL-17A and IL-23 were both significantly higher in JSLE PBMCs in comparison to those of healthy control patients (p = 0.018 and p = 0.010 respectively). The levels of RORC and IL-23R were also higher in JSLE PBMCs but did not reach statistical significance (Figure 1B).

Levels of the Th1 cytokines IL-12 and IFNγ and the Th2 cytokines IL-4 and IL-13 were not statistically significantly increased in JSLE patients (n = 11); BILAG -2004, A/B =6 score, mean (range) (3.8 (0–12)) compared to control (n = 7) patient serum (data not shown). These data indicate that Th17 cell may be the dominant Th cell in JSLE patients, reflected by the significantly higher levels of IL-17A in patient plasma.

Comparison of JSLE patients who had active renal disease (pBILAG2004 score of A or B; n = 6) to those who did not (pBILAG2004 score C-E; n = 13) demonstrated that patients with active renal disease at the time of sampling had higher plasma levels of IL-17A but this difference was not significant (23.92 ± 4.5 pg/ml versus 17.46 ± 6.1 pg/ml, p = 0.16). However, it should be noted that renal JSLE patients had statistically significantly higher levels of IL-17A in comparison to healthy control plasma (23.92 ± 4.5 pg/ml versus 7.23 ± 2.5 pg/ml, p = 0.005).

To fully characterise the production of IL-17A and Th17 related cytokines, PBMCs from JSLE (n = 7) and control (n = 6) patients were stimulated with anti-CD3 and anti-CD28 coated microbeads for 2 days in vitro to induce T cell activation and supernatants were analysed by ELISA. Levels of IL-17A secreted from PBMCs of patients with JSLE (297.8 ± 116.3 pg/ml) were significantly higher than from control PBMCs (26.2 ± 7.3 pg.ml) (p = 0.022) (Figure 2A).

To further measure the Th17 profile of the stimulated cells, 9 JSLE PBMC and 10 control PBMC supernatants were analysed using a Th17 cytokine panel bioplex (Figure 2B). JSLE PBMCs secreted increased levels of cytokines known to be produced by Th17 cells which drive Th17 development including IL-17 F, IL-21 and IL-22 (Figure 2B); the level of IL-21 was statistically significantly higher in JSLE compared to control supernatants (p = 0.003). JSLE supernatants also contained significantly increased IL-6 (p = 0.028). These results demonstrate that upon stimulation, JSLE PBMCs have an increased ability to produce Th17 cytokines and IL-6 required for the further generation of Th17 cells.