Immune senescence and immune activation in elderly colorectal cancer patients

Characteristics of the study population

Eighty-one patients with colorectal cancer were enrolled in this study. Their clinical and biological characteristics at baseline are reported in Table 1. Median age was 76 years; 60.5% were male. According to TNM staging system, 3 patients (3.7%) had stage I, 22 (27.2%) stage II, 36 (44.4%) stage III and 20 (24.7%) stage IV. A comprehensive geriatric assessment (CGA) was performed in all patients; 55 were classified as fit (67.9%), 20 were vulnerable (24.7%) and 6 were frail (7.4%). Seventy patients (86.4%) received a systemic treatment in an adjuvant (63.0%) and/or metastatic setting (37.0%), while 11 patients (13.6%) were not treated, due to frailty recognized at CGA or early stage of disease with no indication to adjuvant treatment.

Immune senescence markers

TREC levels decreased significantly with increasing age in patients (r_s=−0.403; p<0.001) (Figure 1A), while telomere length did not correlate with age (r_s=−0.128, p=0.314) (Figure 1B). These findings are in agreement with our first exploratory study [7]. Consistently with the knowledge that TREC is a marker for thymic output, a positive correlation was found between TREC and CD4 recent thymic emigrant (RTE) cells (r_s=0.270, p=0.026), and CD8 RTE cells (r_s=0.290, p=0.022); of interest, there was a trend to negative correlation between TREC and activated CD8 cells (r_s=-0.230, p=0.054). Levels of TREC and length of telomeres were not prognostic of disease relapse in the disease-free-survival (DFS) conducted in the 61 I-III staged patients (Table 2), neither of a negative outcome event (relapse/progression or death) in the event-free-survival (EFS) performed in all 81 patients included in the study (Table 3).

Among the immunological cellular subtypes, high percentage (up to the best cut-off value) of senescent CD8 cells, but not CD4 cells, was significantly associated with a higher risk of disease relapse in the DFS (Hazard Ratio (HR) [95% Confidence Interval]= 3.5 [1.0-11.6], p=0.0434) (Table 2 and Figure 2A), and higher risk of a negative event in the EFS (HR=3.0 [1.4-6.3], p=0.0051) (Table 3). Notably, senescent CD8 cells were highly expressed in IV staged patients (Table 1) and significantly associated with disease stage (I-III vs IV p=0.0043); when adjusted for stage (I-III vs IV), this marker missed the association with EFS (HR adjusted =1.8 [0.8-4.0], p=0.1470) (not shown).

Low percentage of CD4 RTE cells was not prognostic of DFS (Table 2). Its prognostic value of EFS (HR=2.8 [1.2-6.9], p=0.0203) (Table 3) was lost when adjusted for stage (HR adjusted=1.6 [0.6-4.1], p=0.2956). Interestingly, high percentages of CD8 RTE cells were prognostic of both DFS (HR= 8.4 [1.1-64.7], p=0.0410) (Table 2) and EFS (HR= 2.4 [1.1-5.7], p=0.0369) (Table 3), and remained independently associated to EFS, after adjusting for stage (HR adjusted= 2.4 [1.0-5.7], p=0.0442].

The CD4/CD8 ratio was investigated as a surrogate marker of immune senescence [11]. While low CD4/CD8 ratio (<0.68) lacked of prognostic role in the EFS (Table 3), I-III staged patients with low CD4/CD8 ratio were at higher risk of disease relapse (HR= 7.0 [2.1-23.3], p=0.0014) (Table 2 and Figure 2B).

Immune activation markers

High percentages (up to the best cut-off value) of immune activated CD8 cells were significantly associated with higher risk of disease relapse in the DFS (HR= 4.1 [1.1-15.0], p=0.0297) (Table 2 and Figure 2C), and negative event in the EFS (HR= 4.4 [2.1-9.1]; p<0.0001) (Table 3). As pointed out for senescent CD8 cells, the activated CD8 cells were highly expressed in IV staged patients and significantly associated to disease stage (I-III vs IV p=0.0003); when adjusted for stage (I-III vs IV), this marker missed the association with EFS (HR adjusted =1.2 [0.5-2.7], p=0.7218) (not shown).

CRC patients with high levels of circulating pro-inflammatory cytokines IL-6 and C reactive protein (CRP) showed higher risk of disease relapse in the DFS (HR= 5.6 [1.1-28.7], p=0.0368 for IL-6 and 2.2 [0.7-7.0], p=0.1682 for CRP, Table 2) and negative event in the EFS (HR= 5.4 [2.5-11.8], p<0.0001 for IL-6 and 4.6 [2.3-9.1], p<0.0001 for CRP, Table 3).

Median [interquartile-IQR] levels of sCD14 and LPS were higher in patients who will experience a negative event than in patients who will not (2268 [2050-2549] vs 2062 [1870-2249] ng/ml; p=0.005, for sCD14; 47 [31-70] vs 38 [19-54] pg/ml; p=0.054, for LPS) (Figure 3A and 3B). In I-III staged patients, levels of sCD14 tended to be higher in patients who will have a disease relapse than in those who will not (2224 [1906-2581] vs 2062 [1850-2253] ng/ml; p=0.063), while no differences were found for LPS levels (45 [31-87] vs 38 [18–55] pg/ml; p=0.189) (not shown).

Circulating levels of sCD14 and LPS were significantly increased with increasing percentages of activated CD8 cells in all patients (regression coefficient β=0.007, p=0.0001 for sCD14, and β=0.168, p<0.0001 for LPS) (Figure 3C and D). Notably, both sCD14 and LPS were strongly associated with activated CD8 in the group of patients who will experience a negative event (β= 0.013, p=0.0032 for sCD14, and β= 0.225, p=0.0003 for LPS), but not in the group of patients who will not (β=0.0004, p=0.5514 for sCD14, and β=0.020, p=0.2133 for LPS) (Figure 3C and 3D). Moreover, also in 61 I-III staged patients, levels of sCD14 and LPS were significantly increased with increasing immune activation. In particular, circulating levels of sCD14 and LPS were associated with increased percentages of activated CD8 in patients who will have a disease relapse (β= 0.022, p=0.0045 for sCD14, and β= 0.284, p=0.038 for LPS). (Figure 3E and 3F).

Study population

A total of 81 patients aged ≥70 years with new diagnosis of stage I-IV colorectal cancer, consecutively admitted to Unit of Medical Oncology 1, Istituto Oncologico Veneto-IRCCS, between October 2010 and May 2017, were enrolled into the study. Clinical, biomolecular and pathological features for each patient were collected in a specific database. Treatments for limited and advanced disease were collected. Each patient performed a comprehensive geriatric assessment (CGA) and a CT scan at baseline to define the stage of the disease and subsequently as for clinical practice. A blood sample was collected at tumor diagnosis for each patient. Data about white blood cells count, IL-6 and C-Reactive Protein (CRP) were registered.

The present study was approved by the Institutional Ethics Committee (CE IOV 2010/18) and conducted according to the Declaration of Helsinki and the guidelines of Good Clinical Practice. Written informed consent was obtained from all patients.

Biomarkers analyses

Sample collection

Peripheral blood mononuclear cells (PBMC) were isolated from peripheral blood by centrifugation on a Ficoll-Paque (Pharmacia, Uppsala, Sweden) gradient. PBMC and plasma samples were cryopreserved and stored, respectively, in liquid nitrogen and at -80°C until they were employed.

Flow cytometric analysis

Approximately 250,000 PBMC were stained for 15 min in the dark using the Live/Dead Fixable Near-IR Dead Cell Stain Kit (Life Technologies, Carlsbad, CA, USA) and the following labelled monoclonal antibodies (mAbs): anti-CD3 [fluorescein isothiocyanate (FITC)], anti-CD4 [peridinin chlorophyll protein (PerCP)], anti-CD8 (PerCP), anti-CD31 [phycoerythrin (PE)], anti-CD45RA [allophycocyanin (APC)], anti-CD38 PE, anti-HLA-DR APC, anti-CD57 PE and anti-CD28 APC. All samples were analysed by four-color flow cytometer FACSCalibur (Becton Dickinson) equipped with a 488 nm argon-ion laser and a 635 nm red diode laser. A total of 50,000 events was collected in the lymphocyte gate using morphological parameters (forward- and side-scatter). Data were processed using CellQuest Pro Software (Becton Dickinson) and analysed using Kaluza^® Analysis Software v.1.2 (Beckman Coulter, Inc, Fullerton, CA, USA). The gating strategy for flow cytometry analysis to identify CD4+ and CD8+ cell subsets is described in Supplementary Figure 1.

Quantification of T-cell receptor rearrangement excision circle (TREC)

Thymic output in PBMC was studied by measuring TREC levels by real-time polymerase chain reaction (PCR), as previously described [26]. TREC levels were expressed as TREC copy number/10⁵ PBMC [26,27].

Determination of telomere length

Relative telomere length (T/S) was determined by monochrome quantitative multiplex PCR assay as previously described [28].

Quantification of microbial translocation markers

Plasma levels of lipopolysaccharide (LPS) levels were quantified with the Human Lipopolysaccharides ELISA Kit (Cusabio, Wuhan, China), and results were expressed as LPS ng/ml plasma. Plasma levels of human sCD14 were determined with the Quantikine Human sCD14 Immunoassay (R&D Systems Inc. Minneapolis, MN, USA), according to manufacturer’s instructions, and results were expressed as sCD14 ng/ml plasma.