Monocytes show immunoregulatory capacity on CD4+ T cells in a human in-vitro model of extracorporeal photopheresis

Abstract

Extracorporeal photopheresis (ECP) is a widely used immunomodulatory therapy for the treatment of various T cell-mediated disorders such as cutaneous T cell lymphoma (CTCL), graft-versus-host disease (GvHD) or systemic sclerosis. Although clinical benefits of ECP are already well described, the underlying mechanism of action of ECP is not yet fully understood. Knowledge on the fate of CD14⁺ monocytes in the context of ECP is particularly limited and controversial. Here, we investigated the immunoregulatory function of ECP treated monocytes on T cells in an in-vitro ECP model. We show that ECP-treated monocytes significantly induce proinflammatory T cell types in co-cultured T cells, while anti-inflammatory T cells remain unaffected. Furthermore, we found significantly reduced proliferation rates of T cells after co-culture with ECP-treated monocytes. Both changes in interleukin secretion and proliferation were dependent on cell-contact between monocytes and T cells. Interestingly, blocking interactions of programmed death ligand 1 (PD-L1) to programmed death 1 (PD-1) in the in-vitro model led to a significant recovery of T cell proliferation. These results set the base for further studies on the mechanism of ECP, especially the regulatory role of ECP-treated monocytes.

Keywords: PD-L1/2; Th17 cells; extracorporeal photopheresis; monocytes; proliferation.

Figure 1

Percentages of regulatory T cells (T_regs), pro‐ and anti‐inflammatory T cell subsets after co‐culture with ECP‐treated monocytes of healthy donors. (a) In‐vitro ECP‐treated or untreated monocytes of healthy donors were co‐cultured with autologous CD4⁺ T cells with anti‐CD3 antibody (100 ng/ml) at a monocyte : T cell ratio of 1 : 4 for 5 days. Induction of CD3⁺CD4⁺CD25⁺forkhead box protein 3 (FoxP3)⁺ T_regs (n = 8), proinflammatory CD3⁺CD4⁺interferon (IFN)‐γ⁺ T helper type 1 (Th1) cells (n = 27), CD3⁺CD4⁺interleukin (IL)‐17A⁺ Th17 cells (n = 27), CD3⁺CD4⁺IL‐17A⁺IFN‐γ⁺ Th17/Th1 cells (n = 27), CD3⁺CD4⁺IL‐2⁺ T cells (n = 19) and anti‐inflammatory CD3⁺CD4⁺IL4⁺ Th2 cells (n = 18) was assessed by intracellular flow cytometry analysis. (b–f) Necessity of cell‐contact or cytokine environment for changes in T cell subsets was analysed by incubation of ECP untreated or treated monocytes for 24 h and subsequent addition of either monocytes and 100 ng/ml anti‐CD3 antibody (cell‐contact) or supernatant of monocytes, 100 ng/ml anti‐CD3 antibody and 1 μg/ml anti‐CD28 antibody (supernatant) to CD4⁺ T cells (n = 7 or n = 5). Co‐cultures were investigated after 5 days. ECP untreated or treated monocytes directly co‐cultured with anti‐CD3 antibody and CD4⁺ T cells were used as reference (reference). ****P  < 0·0001, **P  < 0·01, *P  < 0·05.

Figure 2

Influence of in‐vitro ECP‐treated monocytes on autologous T cell proliferation. ECP untreated or treated monocytes were co‐cultured with CD4⁺ T cells from the same donor with anti‐CD3 antibody (100 ng/ml) for 5 days after in‐vitro ECP. (a,b) Proliferation rates of CFSE‐labelled CD3⁺CD4⁺ T cells were assessed via flow cytometry (histogram shows one donor of 31, n = 31). (c) Necessity of cell‐contact or cytokine environment for changes in proliferation was analysed (n = 4). Therefore, ECP untreated or treated monocytes were cultured for 24 h exclusively. In cell‐contact‐dependent approaches freshly isolated CD4⁺ T cells were cultured with precultured monocytes at a monocyte : T cell ratio of 1 : 4 in the presence of 100 ng/ml anti‐CD3 antibody for 5 days (cell‐contact). For cytokine‐mediated approaches supernatant of precultured monocytes were added to freshly isolated CD4⁺ T cells with 100 ng/ml anti‐CD3 antibody and 1 μg/ml anti‐CD28 antibody (supernatant). Approaches where ECP untreated or treated monocytes were directly co‐cultured with 100 ng/ml soluble anti‐CD3 antibody and CD4⁺ T cells were used as reference (reference). Co‐cultures were investigated after 5 days. ****P  < 0·0001, **P  < 0·01.

Figure 3

Comparison of T cell subsets and proliferation rates after ex‐vivo and in‐vitro ECP of patient samples. The samples were collected from the apheresis bag during ECP procedure before addition of UVADEX^® (ECP^–, ex vivo) and after ultraviolet A (UVA) irradiation (ECP⁺, ex vivo). Monocytes from patients were isolated, co‐cultured with freshly isolated CD4⁺ T cells from healthy donors at a monocyte : T cell ratio of 1 : 4 with 100 ng/ml anti‐CD3 antibody and analysed 5 days after ECP. Furthermore, monocytes from patient samples before addition of UVADEX^® were isolated and treated with the ECP in‐vitro model (ECP^– and ECP⁺, in vitro). Afterwards, monocytes were co‐cultured with CD4⁺ T cells from healthy donors and anti‐CD3 antibody (100 ng/ml) and investigated after 5 days. Induction of CD3⁺CD4⁺CD25⁺forkhead box protein 3 (FoxP3)⁺ regulatory T cells (T_regs) (n = 4*), proinflammatory CD3⁺CD4⁺interferon (IFN)‐γ⁺ T helper type 1 (Th1) cells (n = 6*), CD3⁺CD4⁺interleukin (IL)‐17A⁺ Th17 cells (n = 6*), CD3⁺CD4⁺IL‐17A⁺IFN‐γ⁺ Th17/Th1 cells (n = 6*), CD3⁺CD4⁺IL‐2⁺ T cells (n = 5*) and anti‐inflammatory CD3⁺CD4⁺IL4⁺ Th2 cells (n = 5*) in approaches with (a) ex‐vivo‐ and (b) in‐vitro‐treated monocytes was assessed by intracellular flow cytometry analysis. (c) Determination of T cell proliferation rates after co‐culture with ex‐vivo‐ and in‐vitro‐treated monocytes via CFSE labelling and flow cytometry analysis (n = 3). *One of 4 or 6 data points after ex‐vivo ECP treatment is missing due to coagulation of sample material. **P  < 0·01, *P  < 0·05.

Figure 4

Characterization of in‐vitro ECP‐treated monocytes. ECP untreated or treated monocytes were cultured after in‐vitro ECP for 2 days and (a) absolute cell counts referred to originally seeded cell numbers directly after ECP treatment (day 0) were determined (n = 9). (b) Determination of proportions of living, early apoptotic, late apoptotic and dead cells was performed by calcium‐dependent staining of ECP untreated or treated monocytes with annexin V after 2 days and successive flow cytometry analysis (n = 6). (c) Moreover, ECP untreated or treated monocytes were co‐cultured with CFSE‐labelled CD4⁺ T cells from the same donor in different monocyte : T cell ratios with anti‐CD3 antibody (100 ng/ml) for 5 days after in‐vitro ECP and proliferation rates of CD3⁺CD4⁺ T cells were assessed via flow cytometry (n = 3–5). Induced percentages of (d) CD14⁺CD209‐human leucocyte antigen D‐related (HLA‐DR)⁺CD86⁺ monocytes and (e) CD14⁺CD209⁺HLA‐DR⁺CD86⁺ macrophages (n = 15) as well as (f) HLA‐DR‐CD33⁺CD14⁺monocytic myeloid derived suppressor cells (M‐MDSCs) (n = 16) were assessed via flow cytometry analysis 2 days after in‐vitro ECP. ****P  < 0·0001, **P  < 0·01, *P  < 0·05.

Figure 5

Investigation of the connection between programmed cell death ligand 1 (PD‐L1) and PD‐L2 and T cell proliferation. ECP untreated or treated monocytes were cultured for 2 days and proportions of living (a) CD14⁺PD‐L1⁺ and (b) CD14⁺PD‐L2⁺ monocytes were determined via flow cytometry. Furthermore, ECP untreated or treated monocytes were co‐cultured with CD4⁺ T cells from the same donor with anti‐CD3 antibody (100 ng/ml) for 5 days after in‐vitro ECP and proportions of living (c) CD3⁺CD4⁺PD‐L1⁺ and (d) CD3⁺CD4⁺PD‐2⁺ monocytes were determined via flow cytometry. (e) To block PD‐L1‐PD‐1 and PD‐L2‐PD‐1 interactions 25 µg/ml of pembrolizumab, anti‐PD‐L1 or anti‐PD‐L2 antibody was added to co‐cultures of either untreated or ECP‐treated monocytes with CFSE‐labelled CD4⁺ T cells from the same donor with anti‐CD3 antibody (100 ng/ml) stimulation for 5 days after in‐vitro ECP. Proliferation rates of CD3⁺CD4⁺ T cells were assessed via flow cytometry. **P  < 0·01, *P  < 0·05.