The host cellular immune response to cytomegalovirus targets the endothelium and is associated with increased arterial stiffness in ANCA-associated vasculitis

Fifty-three CMV-seropositive patients with AAV in stable remission were recruited from the vasculitis clinic at University Hospitals Birmingham NHS Foundation Trust (Birmingham, UK), and 30 age-matched CMV-seropositive healthy volunteers (HVs) were enrolled from the 1000 Elders Cohort (courtesy of Professor Janet Lord, University of Birmingham, UK) and patient household contacts. CD4⁺CD28^null T-cell percentage and phenotype were assessed in all participants. Arterial stiffness was measured in patients with AAV.

Patients were eligible for inclusion if they had a documented diagnosis of AAV and were in stable remission for at least 6 months, on maintenance immunosuppression with a maximum of two agents, and seropositive for CMV (anti-CMV IgG detected in peripheral blood). Exclusion criteria were estimated glomerular filtration rate of less than 15 mL/minute per 1.73 m², B cell–depleting therapy within 12 months or T cell–depleting therapy within 6 months, presence of other chronic infection (HIV, hepatitis B, hepatitis C, or tuberculosis) and treatment with anti-CMV therapies within the previous month.

Thirty-eight of 53 patients with AAV were participants in the ‘Cytomegalovirus modulation of the immune system in ANca-associated VASculitis’ (CANVAS) clinical trial, a proof-of-concept open-label randomised trial of valaciclovir, or no additional treatment, in CMV-seropositive AAV patients in remission [32]. All immune and arterial stiffness assessments reported here were conducted at baseline prior to commencement of valaciclovir.

The study was approved by the Research Ethics Committee of Yorkshire and the Humber (UK). Written informed consent was obtained from all participants.

Up to 50 mL of peripheral blood was obtained and processed within a maximum of 5 hours following venepuncture. Plasma was isolated by centrifugation and cryopreserved at −80 °C. Peripheral blood mononuclear cells (PBMCs) were isolated from heparinised blood by density gradient centrifugation, used immediately in stimulation experiments with CMV lysate to identify cytokine-producing T cells, or cryopreserved in liquid nitrogen.

Cells for flow cytometry experiments were acquired on a BD LSR II Flow Cytometer and analysed by using FACS DIVA Software Version 8.0 (BD, Franklin Lakes, NJ, USA). Monoclonal antibodies used for flow cytometry experiments are listed in Table S1 of Additional file 1. Gating strategies are shown in Figure S1 and S2 of Additional file 1.

Whole blood was stained with anti-CD3, anti-CD4 and anti-CD28 monoclonal antibodies to determine CD4⁺CD28^null T-cell percentage. Quality control was achieved by using a positive control (Cytofix CD4 Normal Range Positive Control; Cytomark, Caltag Medsystems, Buckingham, UK) with a validated acceptance range for CD3⁺CD4⁺ percentage and a fluorescence minus one control to aid CD28 gating. These were assayed with every analytical run.

To identify CMV lysate-stimulated cytokine-expressing T cells, 0.5–1 × 10⁶ PBMCs were re-suspended in supplemented medium—(RPMI), 10% foetal calf serum; sterile filtered and heat inactivated (Sigma-Aldrich, St. Louis, MO, USA), 1% penicillin/streptomycin (P/S) (Gibco, Thermo Fisher Scientific, Waltham, MA, USA—overnight for 16 (±2) hours at 37 °C, 5% CO₂, in the presence of monensin (2 μmol/L) and phycoerythrin-conjugated anti-CD154 monoclonal antibody as previously described [33]. Cells were stimulated with CMV lysate (1:100) prepared from CMV strain AD169-infected human foetal foreskin fibroblasts. Unstimulated cells served as controls. Following overnight incubation, cells were stained with eFluor-506 viability dye (eBioscience, Thermo Fisher Scientific) for 30 min at 4 °C, washed with phosphate-buffered saline and flow cytometry buffer, and co-stained with saturating amounts of anti-CD3, anti-CD4 and anti-CD28 antibodies for 30 min at 4 °C before washing with flow cytometry buffer. Cells were fixed and permeabilised with an intracellular flow cytometry staining kit (eBioscience) and stained for 30 min at 4 °C with saturating amounts of anti-interferon-gamma (anti-IFN-γ), anti-tumour necrosis factor-alpha (anti-TNF-α), anti-interleukin-2 (anti-IL-2), anti-IL-5, anti-IL-10 and anti-T-bet monoclonal antibodies before washing with flow cytometry buffer.

The expression of chemokine receptors on unstimulated CD4⁺CD28^null T cells from 17 patients with AAV was defined by staining whole blood with anti-CXCR3, anti-CCR4, anti-CCR6, anti-CD3, anti-CD4 and anti-CD28 monoclonal antibodies. T helper 1 (Th1)-skewed CD4 T cells were identified as CXCR3⁺CCR6⁻, Th2-skewed as CCR4⁺CCR6⁻ and Th17-skewed as CCR4⁺CCR6⁺ [34].

In order to phenotype unstimulated CD4⁺CD28^null T cells with respect to their expression of endothelial homing receptors and cytotoxic molecules, cryopreserved PBMCs from 10 patients with AAV were stained with Fixable Viability dye eFluor-506 as described above; co-stained with anti-CD3, anti-CD4, anti-CD28, anti-CX3CR1, anti-CD49d and anti-CD11b antibodies; and fixed and permeabilised as already described, followed by intracellular staining with anti-perforin and anti-granzyme B antibodies.

Soluble markers of inflammation (IL-2, TNF-α, IFN-γ, IL-10, IL-17A, IL-6 and highly sensitive C-reactive protein) were measured in plasma by Luminex array (ProcartaPlex, eBioscience) in accordance with the instructions of the manufacturer, read on a Bio-Rad Luminex 200 instrument (Bio-Rad, Hercules, CA, USA) and analysed by using ProcartaPlex Analyst 1.0 Software (eBioscience).

Plasma anti-CMV IgG titre was assayed by using an enzyme-linked immunosorbent assay as previously described [30]. CMV seropositivity was defined as an anti-CMV IgG titre of more than 10 units.

Arterial stiffness was estimated by measuring carotid-to-femoral pulse wave velocity (PWV) using the non-invasive, non-operator-dependent Vicorder system (Skidmore, Bristol, UK) that employs a volume displacement method, as previously described [32]. Briefly, the patient was allowed to rest for 5 min prior to inflating a 100-mm-wide cuff on the non-dominant arm to measure peripheral blood pressure. A 30-mm-wide cuff was then placed on the neck at the level of the carotid artery and a 100-mm-wide cuff placed around the proximal thigh. The distance between the mid-clavicular point and the mid-point of the thigh cuff, the aortic path length, was measured with the patient supine. With the patient at a supine 30° head-tilt position, the cuffs were inflated to 60 mm Hg. The Vicorder instrument uses the resultant oscillometric signal to extract the pulse waveforms and pulse transit time to calculate carotid-to-femoral PWV. The mean value of three consistent readings was used for subsequent analysis.

We initially undertook a phenotypic analysis of CD4⁺CD28^null T cells from 53 CMV-seropositive AAV patients in stable remission. PBMCs were stimulated in vitro with CMV lysate, and the proportion of CMV-specific cells within the CD4⁺CD28^null and CD4⁺CD28⁺ subfractions was determined. A greater proportion of CMV-specific T cells was observed within the CD4⁺CD28^null population as determined by expression of the activation marker CD154 and cytokine expression: IFN-γ, TNF-α and IL-2 (Fig. 1a). Neither cell type expressed significant amounts of IL-5 or IL-10.

The percentage of CD4⁺CD28^null T cells was strongly correlated with the total CMV-specific CD4⁺ response (rho = 0.846, P <0.001; Fig. 1b), indicating that the size of the CD4⁺CD28^null T-cell accumulation is a good measure of the impact of CMV infection on the CD4 compartment in this population. CD4⁺CD28^null T-cell percentage also correlated with the humoral response to CMV, anti-CMV IgG titre (rho = 0.324, P = 0.018; Fig. 1b).

As well as expressing IFN-γ and TNF-α, the majority of CD4⁺CD28^null T cells expressed the Th1 lineage transcription factor T-bet (Fig. 1c) and a Th1-skewed chemokine receptor profile (CXCR3⁺CCR6⁻) (Fig. 1d). In contrast, the chemokine receptor staining profile that identifies Th2-skewed (CCR4⁺CCR6⁻) and Th17-skewed (CCR4⁺CCR6⁺) cells was not seen on CD4⁺CD28^null T cells. Indeed, CD4⁺CD28^null T cells comprised the majority of the Th1 compartment (median 51.6%, 30.0–77.3). In addition, the percentage of CD4⁺CD28^null T cells was strongly correlated with the overall size of the Th1 compartment (Fig. 1c), indicating that the size of the CD4⁺CD28^null T-cell expansion in CMV-seropositive AAV patients determines the relative proportion of Th1 cells, a subset known to exert a strong pro-atherosclerotic effect [35].

There was no difference between AAV patients and age-matched CMV-seropositive HVs in the induction of CD154 expression on CD4⁺CD28^null T cells or their cytokine expression profile following CMV lysate stimulation (Fig. 1e). However, patients with AAV had larger expansions of CD4⁺CD28^null T cells compared with HVs (11.3% [3.7–19.7] versus 6.7 [2.4–8.8]; P = 0.022).

IFN-γ and TNF-α activate endothelial cells and increase surface expression of chemokines and adhesion molecules such as IFN-γ–inducible protein-10 (IP-10), fractalkine, vascular adhesion molecule-1 (VCAM-1) and intercellular adhesion molecule-1 (ICAM-1) [36]. Having identified high levels of expression of the IP-10 receptor CXCR3 on the surface of CD4⁺CD28^null T cells, we evaluated the surface expression of the receptors for fractalkine, VCAM-1 and ICAM-1 (CX3CR1, CD49d and CD11b) on unstimulated PBMCs from 10 patients with AAV. Co-expression of all three endothelial homing receptors was significantly more common on CD4⁺CD28^null compared with CD4⁺CD28⁺ T cells (51.6% [42.7–64.4] versus 5.0 [1.7–6.3]; P = 0.006; Fig. 2a). In addition, the majority of unstimulated CD4⁺CD28^null T cells contained intracellular stores of both perforin and granzyme B (74.5%, 63.5–92.7; Fig. 2b), suggesting that they have the capacity to target and lyze endothelial cells.

To determine whether the size of the CD4⁺CD28^null T-cell expansion is associated with arterial stiffness, as a clinical marker of cardiovascular pathology, carotid-to-femoral PWV was measured in the patients with AAV. The CD4⁺CD28^null T-cell percentage was found to be significantly correlated with increased systolic blood pressure (rho = 0.305, P = 0.026), pulse pressure (rho = 0.428, P = 0.001) and PWV (rho = 0.371, P = 0.006; Fig. 3).

On univariable analysis, age, percentage of CD4⁺CD28^null T cells, plasma concentration of TNF-α, and blood pressure parameters (Table 2) were associated with increased PWV. In contrast, we did not observe an association between anti-CMV IgG titre and PWV.

To account for confounding factors, we constructed a multivariable linear regression model including all variables that were associated with PWV on univariable analysis with a P value of less than 0.1 (Tables 2 and 3). This demonstrated that the percentage of CD4⁺CD28^null T cells associated with increased arterial stiffness independently of age, proteinuria, peripheral mean arterial blood pressure, and plasma concentration of TNF-α. There was a 0.66 m/s [95% confidence interval 0.13–1.19] increase in PWV for every 10% increase in CD4⁺CD28^null T cells (P = 0.016; Table 3). This relationship did not change when systolic blood pressure or pulse pressure was substituted for mean arterial blood pressure, and the size of the CD4⁺CD28^null T-cell expansion remained independently associated with increased PWV.