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CD20+inflammatory T-cells are present in blood and brain of multiple sclerosis patients and can be selectively targeted for apoptotic elimination

Multiple Sclerosis and Related Disorders

Abstract

Background

A subset of T-cells expresses the B-cell marker CD20 and in rheumatoid arthritis secretes Interleukin (IL)-17. IL-17 secreting T-cells (Th17) have also been implicated in the inflammatory response in the central nervous system in multiple sclerosis (MS) and may be a potential target for elimination by biologic therapeutics. ScFvRit:sFasL comprises of a rituximab-derived antibody fragment scFvRit genetically fused to human soluble FasL that specifically eliminated T-cells.

Objective

To determine the presence and phenotype of CD20+T-cells in blood and brain of MS patients. Second, to determine whether scFvRit:sFasL can selectively eliminate CD20+T-cells. After CD20-selective binding, scFvRit:sFasL is designed to trigger FasL-mediated activation-induced cell death of T-cells, but not B-cells.

Methods

Flow cytometry and immunohistochemistry were used to screen for CD20+inflammatory T-cells in MS blood and brain tissue. ScFvRit:sFasL pro-apoptotic activity was evaluated by Annexin-V/PI staining followed by flow cytometry assessment.

Results

Peripheral blood (n=11) and chronic but not active lesions of MS patient brains (n=5) contained CD20+inflammatory T-cells. Activated CD20+T-cells were predominantly CD4+and secreted both IL-17 and INF-γ. ScFvRit:sFasL triggered CD20-restricted FasL-mediated activation-induced cell death in peripheral blood CD20+T-cells, but not CD20+B-cells.

Conclusion

CD20+inflammatory T-cells are present in blood and chronic brain lesions of MS patients. ScFvRit:sFasL selectively eliminated CD20+T-cells and may eliminate pathogenic T-cells without B-cell depletion.

Graphical abstract

 

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Highlights

 

  • We detected CD20+T-cells in the blood and brains of MS patients.
  • Some CD20+T cells in the brains were positive for IL-17.
  • Blood CD20+ and CD20- T cells both produced INF-γ and IL-4.
  • Rituximab alone is a poor therapeutic at eliminating T-cells.
  • An engineered scFvRit:sFasL molecule specifically kills CD20+T-cells.

Keywords: Biologics, Chronic lesions, Human tissue, Interleukin-17, T-cells.

1. Introduction

Multiple sclerosis (MS) is a demyelinating autoimmune disease of the central nervous system (CNS) that is characterised by T-cell mediated nerve damage and immune infiltrates in the brain. These infiltrates predominantly contain CD3+/CD8+T-cells ( Lassmann, 2011 ) and macrophages and induce microglial activation with subsequent microglial and macrophage-mediated phagocytosis of myelin leading to the pathologic hallmarks characterised by T-cell mediated demylination and axonal damage ( Huizinga et al., 2012 ).

T-cells are considered to play a major role in MS pathology, with myelin-specific T-cells triggering the hallmark demyelination of axons. Indeed, various T-cell subtypes have been implicated in MS, including CD8+cytotoxic T-cells, CD4+T helper type-1 (Th1) and, more recently, T helper type 17 (Th17) cells. Mounting evidence supports a role for Th17 cells in MS autoimmunity, relapse, and neuronal and blood-brain-barrier (BBB) damage in patients and murine MS models (Harrington et al, 2005 and Kebir et al, 2007).

However, although considered T-cell mediated, other cells may also contribute to MS pathology. For instance, resident microgliatrigger myelin damage in the absence of T-cell infiltration ( van Horssen et al., 2012 ). Interestingly, B-cells have also recently been implicated in MS pathology, with B-cell activation consistently being detected in MS patients, as evidenced by oligoclonal bands (OCB). A significant role for B-cells is apparently supported by the therapeutic effect of the B-cell depleting anti-CD20 antibody rituximab (RTX). When RTX binds to cell surface CD20, the Fc region of RTX is recognised predominantly by natural killer cells, γδ T-cells, dendritic cells and macrophages leading to cell lysis via perforin and/or phagocytosis in an antibody dependent cell cytotoxicity (ADCC) process. RTX upon binding to CD20 also recruits complement components, resulting in the target cells being lysed via complement dependent cytotoxicity (CDC). Bar-Or and co-workers recently suggested that such depletion of B-cells, results in a reduction in proinflammatory cytokine release by these cellsin vivo, which in turn diminished proinflammatory (Th1 and Th17) responses in CD4 and CD8 lymphocytes ( Bar-Or et al., 2010 ). RTX has been evaluated in a number of phase II clinical trials for MS with reports of beneficial activity ( Castillo-Trivino et al., 2013 ), although its use for treating relapsing-remitting MS is still inconclusive ( He et al., 2013 ).

RTX treatment has beneficial effects in MS very early after administration; however, this is not by depletion of antibody-producing plasma cells ( Franciotta et al., 2008 ). This implies that RTX activity is not directly dependent on reducing antibody levels, but may instead be dependent on RTX-mediated effects on e.g. T-cells ( Barun and Bar-Or, 2011 ). In line with this hypothesis, RTX treatment has been shown to trigger T-cell eliminationin vivo( Cross et al., 2006 ). In RA, depletion of B-cells was suggested to reduce T-cell numbers by reducing levels of T-cell promoting chemokines and cytokines normally released by B-cells ( Melet et al., 2013 ). Recently, we hypothesised that a small but likely pathogenic subset of T-cells that expresses low levels of CD20 (CD20dim) are targeted by RTX treatment ( Eggleton and Bremer, 2014 ) and may partly account for the beneficial effects of RTX in autoimmune diseases.

CD20dimT-cells have been identified in both healthy controls and in rheumatoid arthritis (RA) blood and synovial fluid ( Wilk et al., 2009 ). These CD20+T-cells have undergone a shift from a Th1 sub-type in healthy controls to a Th17 sub-type in RA ( Eggleton et al., 2011 ). Importantly, depletion of CD20dimT-cells by RTX may contribute to reduced T-cell mediated inflammation in RA (Avivi et al, 2013 and Stroopinsky et al, 2012). Of note, Th17 cells have been reported in MS lesions ( Tzartos et al., 2008 ) and are reported to be involved in MS pathology and response to therapy ( Broux et al., 2013 ). Th17 cells secrete IL-17, which upon interaction with other cytokines can exacerbate inflammatory responses in MS.

In light of the apparent effects of RTX on T-cell immunity, the selective elimination of CD20+inflammatory T-cells may yield clinical benefit, in the absence of deleterious B-cell depleting side effects such as opportunistic infections. The promise of T-cell targeted therapy for autoimmune disease has become apparent from Abatacept (CTLA4-Ig), which selectively inhibits T-cell immune responses. Recently, we reported on an approach to selectively trigger activation induced cell death (AICD) in activated (autoreactive) T-cells. In brief, AICD is an immunological safeguard process that ensures timely resolution of T-cell inflammatory responses by crosslinking of the T-cell receptor Fas by cognate membrane-expressed ligand FasL on activated T-cells (Brunner et al, 1995 and Dhein et al, 1995).

Membrane FasL can also be cleaved into a homotrimeric soluble form (sFasL), which is essentially unable to trigger Fas-mediated apoptosis . However, by genetic fusion of sFasL to a T-cell-selective scFv antibody fragment the inactive sFasL domain is converted into a membrane-bound and fully active FasL upon T-cell binding of the antibody fragment. Previously, we have shown that a CD7-targeted scFv:sFasL fusion protein enables induction of apoptosis in activated CD7+T-cells ( Bremer et al., 2006 ).

Here, we hypothesized that a subset of T-cells found in the blood and brain of MS patients would be CD20+ and of Th17 subtype. Moreover, we hypothesized that these CD20dimTh17 cells could be eliminated by recombinant fusion protein scFvRit:sFasL by selectively triggering activation induced cell death.

2. Patients, materials and methods

2.1. MS and control blood samples

Ethical approval was obtained from the National Research Ethics Service reference 06/Q2102/56. All MS cases (n=11; 10 female and 1 male) were relapsing-remitting in type and diagnosed by McDonald criteria (McDonald et al, 2001 and Polman et al, 2005). They were recruited from neurology clinics and had suffered at least one relapse during the preceding year, ages ranged from 22 to 53 years (mean=40 years), Expanded Disability Status Scale (EDSS) range 2–3 (mean=2.2). Exclusion criteria for enrolment included a relapse or corticosteroid therapy within 30 days; cyclophosphamide or mitoxantrone treatment within 12 months; treatment with interferon-β, glatiramer acetate or natalizumab within 60 days. Healthy control samples (n=12; 11 female and 1 male) between the ages of 21–51 (mean age=41) were obtained from the Clinical Research Facility at Exeter.

2.2. Experimental methods

2.2.1. Cell isolation

Peripheral blood mononuclear cells (PBMCs) were isolated from whole blood and subpopulations delineated by flow cytometry as previously described ( Eggleton, and et al., 2011 ).

2.2.1.1. Detection and quantification of CD20+IL-17 secreting T-cells in MS patients and control subjects

The number of CD3+/CD20+T-cells in the peripheral blood of 11 MS patients and 12 age-sex matched control subjects that were also capable of secreting IL-17 was identified using an IL-17 cell detection kit (Miltenyi Biotec), as previously described (Eggleton et al, 2011 and Szabo-Taylor et al, 2012). Blood samples were tested in a double-blinded way to avoid bias. Lymphocyte subsets from isolated PBMCs were phenotypically delineated by staining for CD3 (T-cells), or CD19+(B-cells) employing allophycocyanin (APC)-conjugated anti-CD3 or anti-CD19. The number of CD20+cells in each sample group was determined by co-staining with fluorescein isothiocyanate (FITC)-conjugated anti-CD20 (Biolegend). PBMCs were stimulated with the superantigen –Cytostim™ and IL-17 secretion detected using a phycoerythrin (PE)-conjugated IL-17 capture antibody in accordance with the manufacturers (Miltenyi Biotec).

2.2.1.2. Quantification of serum levels of IL-17, 21 and 23 by enzyme-linked immunosorbance (ELISA) assays

Quantification of the cytokines IL-17, IL-21 and IL-23 in sera from 11 MS patients and 12 age-sex-matched controls subjects were double blinded and examined by commercial sandwich ELISA (R&D Systems), in accordance with the manufacturer׳s instructions.

2.2.1.3. Quantification of intracellular IFN-γ and IL-4 and IL-17 in CD20+ and CD20- T-cells by flow cytometry

To examine the cytokine profile of CD20+T-cells, PBMCs from healthy controls were stimulated with phorbol-12-myristate 13-acetate (PMA) for 4 h, permeabilised with Fix/PERM kit (Caltag/An der Grub) and triple-stained with anti-CD3-APC, anti-CD20-FITC and interferon gamma (IFN-γ)-PERCPcy5.5, or IL-4-PERCPcy5.5, or IL-17-PERCPcy5.5, and analysed using an Accuri flow cytometer (BD Biosciences, CA, USA). To exclude the possibility of detecting false-positive T-cells expressing CD20 whilst actually associated with B cells in the form of T/B-cell doublets, a Quanta SC flow cytometer (Beckman Coulter) which analyses cells on the basis of accurate volume/diameter measurements was used to exclude this possibility.

2.2.1.4. Examination of brain tissues for CD20+inflammatory T-cells

To determine the predominance of IL-17 secreting CD20+inflammatory T-cells in MS brain tissue with varying degrees of lesion inflammation compared with normal control human brain we performed triple layer enzyme immunohistochemistry using a Vectastain®ABC system (Vector Laboratories, Peterborough, UK). We chose this system as it provides high sensitivity and minimal background staining and is less susceptible to non-specific staining/fluorochrome interference as sometimes observed in fluorescent staining and confocal imaging of similar tissue. Snap-frozen blocks of post-mortem normal control (NC) (n=5) or MS cerebral sub-ventricular deep white matter samples (n=15) were obtained from the NeuroResource Tissue Bank, UCL Institute of Neurology, London, with next-of-kin informed consent for tissue donation and ethical approval from Central London REC 1 (I.D.08/H0718/62) and approval for the study from the Local Research Ethics Committee (I.D.04/Q2102/111).

Tissue sections (10 μm) from each snap-frozen block were cut in a cryostat and mounted onto Vectabond-coated slides (Vector Laboratories). The NC cases had an average age at death of 53.8 years (range 28–84) with a time between death and snap freezing of 19.8 h (range 9–30). A total of 15 blocks were examined from 12 cases of MS containing either normal appearing white matter (NAWM) (n=2) or lesions (n=13). The average age of the MS patients at death was 52 years (range 26–67), the disease duration 19 years (range 2–33) and the death-to snap-freezing time 30.5 h (range 9–66). As previously described, lesions were classified into acute (n=4), sub acute (n=4) and chronic (n=5) on the basis of the number and distribution of oil red-O positive macrophages, the extent of demyelination, cellularity in the borders and parenchyma of lesions, and perivascular cuffing ( Li et al., 1993 ). Briefly, the features of acute lesions were patchy demyelination, many phagocytic macrophages containing oil red-O stained myelin debris, and hyper cellularity at the lesion border and cuffing around blood vessels. Sub acute lesions consisted of a demyelinated plaque with fewer macrophages, which were predominately located at the lesion border, and less perivascular cuffing. A chronic lesion consisted of a hypocellular demyelinated plaque where there were no oil red-O stained macrophages present either within or surrounding the plaque.

Immunohistochemistry was performed using a modification of a well established method ( Gutowski et al., 1999 ). Consecutive sections were fixed in acetone and examined by sequential triple label enzyme immunohistochemistry ( Holley et al., 2010 ) to identify CD8+or CD4+T-cells that were also CD20+and IL-17+. Staining of T-cells was visualized using 0.05% (w/v) 3,3′-diaminobenzidine tetrahydrochloride (DAB) to give brown colouration. T-cells that were also CD20+were visualized with Vector SG (blue/grey) (Vector Laboratories) and T-cells that were also IL-17+were visualized with Vector VIP kit (pink) (Vector Laboratories). The sections were dehydrated and cleared in ethanol–xylene and mounted in DPX.

IgG antibodies were titrated to achieve optimum and specific staining by single layer immunohistochemistry. Rabbit anti-human IL-17 and mouse anti-human CD8 (IgG1 - Abcam) were used at 1:200 and 1:10 dilution, respectively and mouse anti-human CD4 (IgG1) and mouse anti-human CD20 (IgG2a) (Vector Laboratories) were used at 1:40 and 1:200 dilution, respectively. Isotype controls confirmed specific staining and that blocking was successful. Images were viewed using an Olympus BX60 microscope with C-mount and Nikon S10 digital camera. Ten fields of view or more were examined for each panel.

2.2.1.4.1. Production of scFvRit:sFasL fusion protein, CD20 transfection and apoptosis assay

Fusion protein scFvRit:sFasL was constructed and produced as previously reported ( Bremer et al., 2008 ). To confirm CD20-restricted activity, human CD20-negative Jurkat cells were transfected by electroporation with human CD20 plasmid pB1-21, as previously described elsewhere ( Bubien et al., 1993 ). Successful transfection of CD20 on Jurkat cells was assessed by fluorescent staining of cells treated with anti-CD20-FITC (Biolegend) antibody and determined by flow cytometry. Parental CD20-negative Jurkat cells and Jurkat-CD20+cells (3×104/well in 48-well plate) were treated with scFvRit:sFasL for 24 h, after which cell death was determined by Annexin-V/PI staining on an Accuri C6 flow cytometer (BD biosciences). To determine apoptotic activity of scFvRit:sFasL towards CD20+primary T-cells, the CD20+ and CD20- T-cell populations were isolated by staining T-cells with CD3-PERCPcy5 and anti-CD20-FITC and subsequent cell sorting on a MoFlo high-speed cell sorter (Cytomation, CO, USA). The isolated populations were treated with 400 ng/ml scFvRit:sFasL for 48 h (3×104/well in 48-well plate), after which apoptosis was determined using annexin V-FITC/PI staining. As a control, Jurkat cells were treated with a range of RTX concentrations (100 ng/ml–10 µg/ml).

2.3. Statistical analysis

Between groups analysis of cytokine protein profiles and differences in T-cell phenotypes were tested using a Mann–WhitneyU-test. The effect of anti-CD20 agents on T-cell killing was analysed using a 2-sided unpaired studentTtest. Probability values ofP< 0.05 were considered to be statistically significant.

3. Results

3.1. CD20+inflammatory T-cells containing pro-inflammatory cytokines are present in the peripheral blood of MS patients

Blood CD20+T-cells were detected using flow cytometry for CD3/CD20 or CD19/CD20 on peripheral blood mononuclear cells (PBMCs) isolated from MS patients and healthy control blood ( Fig. 1 A). Typically 60% of CD19+B-cells immunostained for CD20 with intermediate intensity, 20% with high intensity and 20% with low intensity, in both MS and control subjects ( Fig. 1 A (ii)) and these were excluded from further analysis. The remaining less intensely positive CD20 cells consisted of CD3+T-cells ( Fig. 1 A (iii)). CD3/CD20 double-positive cells were single cells as determined by strict gating on fsc/ssc scatter plot ( Fig. 1 B (i)) and further analysed ( Fig. 1 B(ii)). To confirm different location of doublets on fsc/ssc scatter plot, T-cell/B-cell doublets were experimentally induced using bi-specific anti-CD3/anti-CD19 antibody. Treatment with this bispecific antibody triggered formation of second double CD3/CD20 double positive population but at distinct fsc/ssc scatter ( Fig. 1 B (iii)). As a further precaution, T-cell subpopulations were assessed with a Quanta SC flow cytometer and Cell Lab Quanta™ software (Beckman Coulter). This particular flow cytometer analyses cells on the basis of accurate volume/diameter measurements (not forward side scatter, which is an arbitrary measurement of cell size) and side scatter analysis. Therefore, it is possible to gate cells of a specific cell diameter, eliminating the possibility of analysing T-cell-B-cell aggregates ( Fig. 1 C). Blood of both MS and control subjects contained a median percentage of 2–6% of CD3+/CD20+T-cells. The CD4:CD8 cell ratio for CD20 was similar for both groups ( Fig. 2 A). Employing triple-colour flow cytometry the percentage of T-cells (CD4 or CD8) that secreted IL-17 was assessed. In peripheral blood of MS and control groups, the median percentage (interquartile range; IQR) of IL-17 secreting CD4 T-cells was similar ( Fig. 2 B). Flow cytometry analysis, indicated the proportion of IL-17 secreting cells that were CD20+were similar in both groups ( Fig. 2 C). Both CD20- and CD20+T-cells stained positive for both IFN-γ and IL-4 ( Fig. 2 D), but a greater proportion of CD20+T-cells were capable of secreting IFN-γ (97.9%) compared to CD20-T-cells (11.8%).

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Fig. 1 Gating, selection and demonstration of CD20+T cell single cell populations. (A) (i) Typical flow cytometry dot plot of the forward and side scatter of peripheral blood mononuclear cells (PBMCs). (ii) Dot plot of P1 gated cells as indicated in (A(i)) stained with anti-CD20-FITC and anti-CD19-PE to detect and exclude CD20+B-cells. (iii) Dot plot of P1 gated cells, except CD20+B-cells, stained with antiCD20-FITC and anti-CD3-CyQ. (B) (i) Dot plot of PBLs stained with bio-specific antibody for anti-CD3/anti-CD19. (ii) Dot plot of low forward scatter (single cell) R1 gated cells (as indicated in B(i), stained with anti-CD20-FITC and anti-CD3-CyQ. (iii) Dot plot of higher forward scatter (doublets) R2 gated cells, stained as above. (C) Lymphocytes were stained with anti-CD3-APC/anti-II-17-PE/CD20-FITC, whereupon (i) single cells (7–9 μm in diameter) positive for anti-CD3 were gated, sorted and evaluated for (ii) expression of CD20 and IL-17. The photomicrograph shows permeabilised CD3+isolated lymphocytes positive for both CD20 and IL-17.

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Fig. 2 Quantification and phenotype analysis of subsets of CD20+inflammatory T-cells in 11 MS patents and 12 healthy subjects peripheral blood. (A) PBMCs were isolated MS and healthy control subjects and tripled stained for CD20, CD19, CD4 or CD8. Cells were gated for CD20 and the percentage of the CD4+, CD8+T-cells and CD19+B-cells determined. (B) MS and control PBLs were stimulated with ‘cytostim’ and the number of CD3+T-cells actively secreting IL-17 determined by flow cytometry and was 0.75% (IQR, 1.15) and 0.49% (IQR, 0.69), respectively and for CD8 cells 1.8 % (IQR, 1.85) and 1.7 (IQR, 1.6), respectively. (C) The proportion of gated IL-17-secreting lymphocytes in MS and healthy subjects that were either CD4+ or CD8+ and CD20+was evaluated by triple-stained flow cytometry. In MS patients peripheral blood, the median percentage (and IQR) of IL-17 secreting CD4 or CD8 cells that were also CD20+were 14.8% (IQR, 34.5) and 7.8 (IQR, 14.6), respectively, compared to control subjects IL-17 secreting CD4 and CD8 cells in which 14.8% (IQR, 24.5) and 8.2 % (IQR, 19.6) expressed surface CD20, respectively. (D) Typical comparison of intracellular INF-γ production in total CD3+, CD20- and in CD20+T-cells (Top panels) and IL-4 production in total CD3+, CD20- and CD20+T-cells (bottom panels) (N=3).

As we did not detect significantly different numbers of Th17 cells in MS and control subjects׳ blood, we measured IL-17 in the serum of MS (n=11) and control subjects (n=12), together with IL-21 and IL-23 concentrations (these latter cytokines are implicated in Th17 cell development) (Elyaman et al, 2009 and Shahrara et al, 2008). We did not detect IL-17 in serum from our control or MS subjects. IL-21 was detectable in all serum samples of MS and control subjects with median concentrations of 25.80 pg/ml (IQR, 25.93) and 24.45 pg/ml (IQR, 23.07) respectively. IL-23 was detectable in 7/12 control serum samples and 4/11 MS samples, but there was no significant difference in concentrations.

3.2. Detection of CD20+inflammatory T-cells in the brains of MS patients

Since T-cells have a role in the pathogenesis of MS ( Tzartos et al., 2008 ), we evaluated whether CD4/8+cells in human MS brain expressed CD20. As anticipated, no CD4 or CD8 cells were detected in normal control (NC) brain In macroscopically normal-appearing white matter (NAWM) there was weak non-specific background interspersed with undefined cells (data not shown). In chronic lesions we observed distinct staining of individual cells that were either CD20+cells ( Fig. 3 A and C), CD4+T-cells ( Fig. 3 A) CD8+T-cells ( Fig. 3 C) or IL-17+cells ( Fig. 3 C) were observed. Some CD4+IL-17+T-cells were identified as well as CD4+CD20+T-cells ( Fig. 3 B). CD8+CD20+T-cells were also observed (brown–blue–black) ( Fig. 3 D). In acute MS lesions moderate CD4+and CD8+cell staining was seen. Stronger staining of CD4+and CD8+T-cells was seen in sub-acute lesions but there was no evidence of CD20+or IL-17 staining on CD4 or CD8 T-cells in sub-acute lesions (not shown).

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Fig. 3 Identification of CD20+ve T-cells in chronic white matter lesions of MS brain. Chronic lesion (CL) brain sections were analysed at x 400 magnification for CD4 T-cells (panel A and B) or CD8 T-cells (panel C) and 100 x magnification (Panel D). Panel A shows both CD4+(brown) and CD20+(blue–grey) lymphocytes. Panel B depicts CD4 (brown) / IL-17+(pink) lymphocytes and a single CD4+CD20 (blue–black) lymphocyte. Panel C shows CD8+cells (brown), CD20+cells (blue–grey) and IL-17+(pink) lymphocytes. Panel D shows a number of lymphocytes at the periphery of a blood vessel double stained for CD8+and CD20+, which appear blue–black. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

3.3. An anti-CD20 (scFvRit:sFasL) molecule enhances killing of CD20+T-cells by apoptosis

Recent studies have highlighted the promise of anti-CD20 therapeutics for treatment of MS, (Kitsos et al, 2012 and Stuve et al, 2009), but the available data suggest that T-cell elimination may actually be a major contributor to their efficacy. Our study demonstrates the presence of CD20+T-cells in the brain and blood of MS patients that in the brain are IL17-producing and may be pathogenic. Therefore, we assessed whether the fusion protein scFvRit:sFasL could selectively eliminate CD20+T-cells. In brief, scFvRit:sFasL is essentially inactive, but once bound to CD20 is converted into membrane-bound and pro-apoptotic FasL. Of note, normal human B-cells are resistant to FasL-mediated killing and scFvRit:sFasL ( Bremer et al., 2008 ). In line with this, parental CD20-negative Jurkat T-cells cells were resistant to treatment with scFvRit:sFasL ( Fig. 4 ). In contrast, treatment of Jurkat.CD20 with scFvRit:sFasL triggered apoptosis in >90% of cells ( Fig. 4 ). Furthermore, apoptosis induction by scFvRit:sFasL was competitively inhibited by co-treatment with epitope competing antibody rituximab. Thus, scFvRit:sFasL has CD20-restricted apoptotic activity.

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Fig. 4 Ability of Rituximab (RTX) and scFv-CD20-FasL antibody to target and kill T-cells by apoptosis. (A) RTX and an engineered single chain anti-CD20 human antibody linked to FasL were used to kill CD20+T-cells and Jurkat cells by apoptosis. (B) CD20+ transfectant Jurkat T-cells and parental CD20- Jurkat cells were treated with scFvRit:sFasL, with and without epitope blocking anti-CD20 (RTX). The antibody fragment scFvRit:sFasL specifically killed CD20+Jurkat cell (N=4; mean +SD). (C) RTX alone does not kill 48 h cultured unselected CD3+T-cells isolated from the peripheral blood of healthy controls by apoptosis, as assessed by annexin-V and 7-AAD staining (mean+SD of triplicate experiments/individual). (D) scFvRit:sFasL antibody specifically targets 48 h cultured FACs sorted CD20+but not CD20- T-cells from healthy subjects for killing by apoptosis (N=4; mean±SD,p<0.05).

Next, the apoptotic activity of scFvRit:sFasL and the antibody RTX towards CD20+T-cells were evaluated using isolated CD3+T-cells from healthy subjects. Treatment with RTX (0–100 µg/ml) for 48 h had little effect on the percentage of apoptotic cells compared to untreated cells ( Fig. 4 B and C). This is consistent with the finding that ADCC and CDC are the main effector mechanisms of RTX. Treatment of sorted CD20+T-cells treated with 400 ng/ml scFvRit:sFasL led to a significant increase in apoptosis compared to CD20- T-cells vs. their background (medium alone) values ( Fig. 4 D). When background apoptosis was subtracted from treatments of cells with scFvRit:sFasL, 2-fold more CD20+T-cells (22.7%) were killed compared to 10.8% of CD20- T-cells. Thus, scFvRit:sFasL can selectively eliminate CD20+ T-cells in the absence of toxicity towards normal CD20+B-cells.

4. Discussion

In the current study we identified the presence of CD20+T-cells in the peripheral blood of MS patients and in chronic lesions of MS brain, where these cells produced the pro-inflammatory cytokine IL-17. Further, we demonstrated that CD20+T-cells can be selectively targeted for apoptotic elimination using fusion protein scFvRit:sFasL comprised of a rituximab-derived antibody fragment and the pro-apoptotic effector molecule sFasL.

B-cell depleting drugs such as RTX have been trialled to treat MS (Hauser et al, 2008 and Hawker et al, 2009). However, the mechanism by which elimination of CD20+B-cells is effective in MS is not fully understood. The reduction of peripheral B-cells may reduce the generation of CNS-autoreactive antibodies against myelin proteins. However, the immediate beneficial effects of RTX in MS patients argue against autoantibodies being the cause. Activated B-cells can also play a role in autoimmunity beyond autoantibody production, by for instance acting as antigen presenting cells and by secreting cytokines and chemokines that activate T-cells ( Yanaba et al., 2008 ). In addition, our own work and the work of others have shown that there is a small population of CD20+T-cells in the blood of healthy subjects and autoimmune patients, (Eggleton et al, 2011, Szabo-Taylor et al, 2012, and Wilk et al, 2009). Treatment of RA patients with RTX also eliminates CD20+T-cells from the blood ( Wilk et al., 2009 ).

CD20+T-cells were here also found in blood of healthy controls and MS patients and stained positive by flow cytometry for INF-γ, indicating that CD20+T-cells possess a key pro-inflammatory cytokine. However, since the potential pathological effects of Th1/Th17 cells occur in the CNS, we examined the cerebral sub-ventricular white matter of MS and control subjects for CD20+T-cells. We observed the presence of CD8 and CD4 Th17 cells in the lesions of MS patients, in agreement with previous observations ( Tzartos et al., 2008 ). In chronic lesions, a proportion of T-cells not associated with perivascular cuffs stained positive for CD20. This shows for the first time that CD20+inflammatory T-cells are present in chronic lesions. Suggesting CD20+inflammatory T-cells in the brain could potentially be targeted by anti-CD20 (e.g. RTX).

Within peripheral blood there are potentially three mechanisms by which anti-CD20 therapeutics can kill cells. The main killing mechanisms are via ADCC and CDC and both processes are pro-inflammatory in nature and occur when RTX, along with complement components and immune cells cross the BBB ( Stuhler et al., 2006 ). The third mechanism, apoptosis, is non-inflammatory and previously we have generated antibody fragments linked to death receptor ligands that can eliminate cells by apoptosis ( Wiersma et al., 2014 ), but in our study the ability of RTX to kill T-cells directly by apoptosis was poor ( Fig. 4 C). We showed it is possible to target CD20 and increased expression of Fas on autoreactive T-cells to promote apoptotic signalling upon binding to scFvRit:sFasL. A distinguishing feature of scFvRit:sFasL is that it does not induce apoptosis in CD20-ve T-cells or Jurkat cells, suggesting it specifically targets CD20+ve T-cells. Further, it does not trigger apoptosis in normal B-cells as these are insensitive to FasL/Fas-mediated apoptosis. Thus, scFvRit:sFasL can potentially be used to selectively deplete CD20+T-cells in the absence of B-cell depletion.

5. Conclusion

In conclusion, we have identified a subset of CD20+T-cells in the brain of MS patients that may respond to anti-CD20 therapeutics and explain in part why anti-CD20 may be beneficial in MS. In addition, we have demonstratedin vitrothat an scFvRit:sFasL molecule can eliminate selective subsets of CD20+T-cells, which we have observed in MS patients׳ blood and brain tissue. This raises the possibility for anti-CD20 therapeutics selectively targeting pathogenic CD20+T-cells without B-cell depletion.

Conflict of interest

The authors declare no conflicts of interest and have not received any funding from any non-charitable sources.

Funding

Funding for this study was kindly provided by grants from The Northcott Devon Medical Foundation (JEH), and a Royal Devon and Exeter Foundation Trust Hospital research grant (PE, JEH & NJG) and Alberta Innovates Health Solution Visiting Scientist Fellowships (PE & EB).

Acknowledgement

We acknowledge support from Dr Gillian Baker and colleagues at the NIHR Clinical Research Facility, Exeter, for patient selection, sample collection and blinding of the study and to Prof. Marek Michalak of the University of Alberta for access to research facilities.

References

  • Avivi et al., 2013 I Avivi, D Stroopinsky, T. Katz. Anti-CD20 monoclonal antibodies: beyond B-cells. Blood Rev. 2013;27:217-223 Crossref
  • Bar-Or et al., 2010 A Bar-Or, L Fawaz, B Fan, PJ Darlington, A Rieger, C Ghorayeb, et al. Abnormal B-cell cytokine responses a trigger of T-cell-mediated disease in MS?. Ann Neurol. 2010;67:452-461 Crossref
  • Barun and Bar-Or, 2011 B Barun, A. Bar-Or. Treatment of multiple sclerosis with anti-CD20 antibodies. Clin Immunol. 2011;142:31-37
  • Bremer et al., 2006 E Bremer, B ten Cate, DF Samplonius, LF de Leij, W. Helfrich. CD7-restricted activation of Fas-mediated apoptosis: a novel therapeutic approach for acute T-cell leukemia. Blood. 2006;107:2863-2870 Crossref
  • Bremer et al., 2008 E Bremer, B ten Cate, DF Samplonius, N Mueller, H Wajant, AJ Stel, et al. Superior activity of fusion protein scFvRit:sFasL over cotreatment with rituximab and Fas agonists. Cancer Res. 2008;68:597-604 Crossref
  • Broux et al., 2013 B Broux, P Stinissen, N. Hellings. Which immune cells matter? the immunopathogenesis of multiple sclerosis. Crit Rev Immunol. 2013;33:283-306 Crossref
  • Brunner et al., 1995 T Brunner, RJ Mogil, D LaFace, NJ Yoo, A Mahboubi, F Echeverri, et al. Cell-autonomous Fas (CD95)/Fas-ligand interaction mediates activation-induced apoptosis in T-cell hybridomas. Nature. 1995;373:441-444 Crossref
  • Bubien et al., 1993 JK Bubien, LJ Zhou, PD Bell, RA Frizzell, TF. Tedder. Transfection of the CD20 cell surface molecule into ectopic cell types generates a Ca2+conductance found constitutively in B lymphocytes. J Cell Biol. 1993;121:1121-1132 Crossref
  • Castillo-Trivino et al., 2013 T Castillo-Trivino, D Braithwaite, P Bacchetti, E. Waubant. Rituximab in relapsing and progressive forms of multiple sclerosis: a systematic review. PLoS One. 2013;8:e66308 Crossref
  • Cross et al., 2006 AH Cross, JL Stark, J Lauber, MJ Ramsbottom, JA. Lyons. Rituximab reduces B cells and T cells in cerebrospinal fluid of multiple sclerosis patients. J Neuroimmunol. 2006;180:63-70 Crossref
  • Dhein et al., 1995 J Dhein, H Walczak, C Baumler, KM Debatin, PH. Krammer. Autocrine T-cell suicide mediated by APO-1/(Fas/CD95). Nature. 1995;373:438-441 Crossref
  • Eggleton and Bremer, 2014 P Eggleton, E. Bremer. Direct and indirect rituximab-induced T-cell depletion: comment on the article by Melet et al. Arthritis Rheum. 2014;66:1053 Crossref
  • Eggleton et al., 2011 P Eggleton, E Bremer, JM Tarr, M de Bruyn, W Helfrich, A Kendall, et al. Frequency of Th17 CD20+cells in the peripheral blood of rheumatoid arthritis patients is higher compared to healthy subjects. Arthritis Res Ther. 2011;13:R208 Crossref
  • Elyaman et al., 2009 W Elyaman, EM Bradshaw, C Uyttenhove, V Dardalhon, A Awasthi, J Imitola, et al. IL-9 induces differentiation of TH17 cells and enhances function of FoxP3+natural regulatory T cells. Proc Natl Acad Sci U S A. 2009;106:12885-12890 Crossref
  • Franciotta et al., 2008 D Franciotta, M Salvetti, F Lolli, B Serafini, F. Aloisi. B cells and multiple sclerosis. Lancet Neurol. 2008;7:852-858 Crossref
  • Gutowski et al., 1999 NJ Gutowski, J Newcombe, ML. Cuzner. Tenascin-R and C in multiple sclerosis lesions: relevance to extracellular matrix remodelling. Neuropathol Appl Neurobiol. 1999;25:207-214 Crossref
  • Harrington et al., 2005 LE Harrington, RD Hatton, PR Mangan, H Turner, TL Murphy, KM Murphy, et al. Interleukin 17-producing CD4+effector T cells develop via a lineage distinct from the T helper type 1 and 2 lineages. Nat Immunol. 2005;6:1123-1132 Crossref
  • Hauser et al., 2008 SL Hauser, E Waubant, DL Arnold, T Vollmer, J Antel, RJ Fox, et al. B-cell depletion with rituximab in relapsing-remitting multiple sclerosis. N Engl J Med. 2008;358:676-688 Crossref
  • Hawker et al., 2009 K Hawker, P O’Connor, MS Freedman, PA Calabresi, J Antel, J Simon, et al. Rituximab in patients with primary progressive multiple sclerosis: results of a randomized double-blind placebo-controlled multicenter trial. Ann Neurol. 2009;66:460-471 Crossref
  • He et al., 2013 D He, R Guo, F Zhang, C Zhang, S Dong, H. Zhou. Rituximab for relapsing-remitting multiple sclerosis. Cochrane DatabaseSyst Rev. 2013;:12
  • Holley et al., 2010 JE Holley, J Newcombe, JL Whatmore, NJ. Gutowski. Increased blood vessel density and endothelial cell proliferation in multiple sclerosis cerebral white matter. Neurosci Lett. 2010;470:65-70 Crossref
  • Huizinga et al., 2012 R Huizinga, BJ van der Star, M Kipp, R Jong, W Gerritsen, T Clarner, et al. Phagocytosis of neuronal debris by microglia is associated with neuronal damage in multiple sclerosis. Glia. 2012;60:422-431 Crossref
  • Kebir et al., 2007 H Kebir, K Kreymborg, I Ifergan, A Dodelet-Devillers, R Cayrol, M Bernard, et al. Human TH17 lymphocytes promote blood-brain barrier disruption and central nervous system inflammation. Nat Med. 2007;13:1173-1175 Crossref
  • Kitsos et al., 2012 DK Kitsos, S Tsiodras, E Stamboulis, KI. Voumvourakis. Rituximab and multiple sclerosis. Clin Neuropharmacol. 2012;35:90-96 Crossref
  • Lassmann, 2011 H. Lassmann. Review: the architecture of inflammatory demyelinating lesions: implications for studies on pathogenesis. Neuropathol Appl Neurobiol. 2011;37:698-710 Crossref
  • Li et al., 1993 H Li, J Newcombe, NP Groome, ML Cuzner. Characterization and distribution of phagocytic macrophages in multiple sclerosis plaques. Neuropathol Appl Neurobiol. 1993;19:214-223 Crossref
  • McDonald et al., 2001 WI McDonald, A Compston, G Edan, D Goodkin, HP Hartung. Lublin FD, et al. Recommended diagnostic criteria for multiple sclerosis: guidelines from the International Panel on the diagnosis of multiple sclerosis. Ann Neurol. 2001;50:121-127 Crossref
  • Melet et al., 2013 J Melet, D Mulleman, P Goupille, B Ribourtout, H Watier, G. Thibault. Rituximab-induced T cell depletion in patients with rheumatoid arthritis: association with clinical response. Arthritis Rheum. 2013;65:2783-2790 Crossref
  • Polman et al., 2005 CH Polman, SC Reingold, G Edan, M Filippi, HP Hartung, L Kappos, et al. Diagnostic criteria for multiple sclerosis: 2005 revisions to the “McDonald Criteria”. Ann Neurol. 2005;58:840-846 Crossref
  • Shahrara et al., 2008 S Shahrara, Q Huang, AM Mandelin 2nd, RM. Pope. TH-17 cells in rheumatoid arthritis. Arthritis Res Ther. 2008;10:R93 Crossref
  • Stroopinsky et al., 2012 D Stroopinsky, T Katz, JM Rowe, D Melamed, I. Avivi. Rituximab-induced direct inhibition of T-cell activation. Cancer Immunol Immunother. 2012;61:1233-1241 Crossref
  • Stuhler et al., 2006 G Stuhler, S Knop, MS Topp, SM Krober, U Ernemann, U Herrlinger, et al. Intravenously administered rituximab induces remission of EBV associated non Hodgkin lymphoma confined to the brain in a patient after allogeneic stem cell transplantation. Haematologica. 2006;91:ECR01
  • Stuve et al., 2009 O Stuve, VI Leussink, R Frohlich, B Hemmer, HP Hartung, T Menge, et al. Long-term B-lymphocyte depletion with rituximab in patients with relapsing-remitting multiple sclerosis. Arch Neurol. 2009;66:259-261
  • Szabo-Taylor et al., 2012 KE Szabo-Taylor, P Eggleton, CA Turner, ML Faro, JM Tarr, S Toth, et al. Lymphocytes from rheumatoid arthritis patients have elevated levels of intracellular peroxiredoxin 2, and a greater frequency of cells with exofacial peroxiredoxin 2, compared with healthy human lymphocytes. Int J Biochem Cell Biol. 2012;44:1223-1231 Crossref
  • Tzartos et al., 2008 JS Tzartos, MA Friese, MJ Craner, J Palace, J Newcombe, MM Esiri, et al. Interleukin-17 production in central nervous system-infiltrating T cells and glial cells is associated with active disease in multiple sclerosis. Am J Pathol. 2008;172:146-155 Crossref
  • van Horssen et al., 2012 J van Horssen, S Singh, S van der Pol, M Kipp, JL Lim, L Peferoen, et al. Clusters of activated microglia in normal-appearing white matter show signs of innate immune activation. J Neuroinflamm. 2012;9:156 Crossref
  • Wiersma et al., 2014 VR Wiersma, Y He, DF Sampionus, RJ van Ginkel, J Gerssen, P Eggleton, et al. A CD47-blocking TRAIL fusion protein with duel pro-phagocytic and pro-apoptotic activity. Br J Haematol. 2014;164:304-307 Crossref
  • Wilk et al., 2009 E Wilk, T Witte, N Marquardt, T Horvath, K Kalippke, K Scholz, et al. Depletion of functionally active CD20+T cells by rituximab treatment. Arthritis Rheum. 2009;60:3563-3571 Crossref
  • Yanaba et al., 2008 K Yanaba, JD Bouaziz, T Matsushita, CM Magro St, EW Clair, TF. Tedder. B-lymphocyte contributions to human autoimmune disease. Immunol Rev. 2008;223:284-299 Crossref

Footnotes

a University of Exeter Medical School & Neurology Department, Royal Devon and Exeter Hospital, University of Exeter, Devon, UK

b Department of Surgery, University Medical Centre, University of Groningen, Groningen, Netherlands

c NeuroResource, UCL Institute of Neurology, London, UK

lowast Correspondence to: University of Exeter Medical, School, University of Exeter, St. Luke’s Campus, Heavitree Road, Exeter. EX1 2LU UK.

1 These authors made an equal contribution to the study.

2 Present address: Manchester Pharmacy School, University of Manchester, Manchester, United Kingdom.