Patients with lung cancer were screened and evaluated by CTC testing between June 2018 and September 2020 at the Department of Thoracic Surgery of Hanzhong 3201 Hospital (Hangzhou, China). The inclusion criteria were as follows: i) Primary lung cancer confirmed by histology or cytology; and ii) Eastern Cooperative Oncology Group Performance Status ≤2. The exclusion criteria were as follows: i) Prior malignancy in the past 5 years, except non-melanoma skin cancer; and ii) life expectancy ≤3 months. Peripheral blood samples were shipped to and processed at Biotecan Medical Laboratory within 24 h of collection. Clinical data, including age, body mass index, histological subtype, smoking status, sites of metastasis, treatment received and clinical outcome, were collected. All patients provided informed consent before sample and data collection. The present study was approved by the Ethics Committee of Hanzhong 3201 Hospital (approval no. 20180603A). The flowchart of the study design is shown in Fig. 1. The control subjects consisted of 20 healthy individuals recruited from the Physical Examination Center of 3201 Hospital during the same study period. To minimize selection bias, participants were selected based on a systematic random sampling strategy from a pool of eligible individuals who visited the center for routine health check-ups. Inclusion criteria were strictly applied to ensure a comparable baseline demographic profile. Participants were required to have: i) No history of malignant tumors, autoimmune diseases or active infections within the preceding 3 months; ii) normal hepatic and renal function tests; and iii) no history of benign pulmonary diseases. The sample size of 20 was determined based on the availability of eligible subjects during the recruitment period and to match the statistical power required for comparative analysis with the patient cohort, ensuring adequate representation of the background population.

Peripheral blood (7.5 ml) was collected using an acid citrate dextrose anticoagulant tube (Becton, Dickinson and Company) and using a Human Circulating Rare Cell Subtraction Enrichment kit (cat. no. SE-001; Cytelligen, Inc.) according to the manufacturer's instructions. In brief, each tube of peripheral blood was centrifuged at 800 × g for 8 min at room temperature, followed by transfer of the lower layer of cells into centrifuge tubes that contained 3 ml of human CTC separation matrix. All probes (CEP8) and antibodies (anti-CD45, anti-CD31) used in the iFISH procedure were components of the above commercial kit. After centrifugation at 450 × g for 8 min at room temperature, the red blood cells were discarded. Subsequently, the buffy coat cells that included white blood cells (WBCs) and tumor cells were transferred to new tubes and incubated with an immunomagnetic particle-conjugated anti-CD45 antibody for 20 min. WBCs were then depleted by applying a magnetic bead sorting system and the magnetic beads-free solution was centrifuged at 450 × g for 8 min (4°C), followed by two rinses with human CTC buffer at room temperature. Finally, the cell pellet was resuspended and mixed with cell fixative solution and was coated on CTC slides (cat. no. CS-01; Cytelligen, Inc.) and dried overnight at 30°C, which were to be examined by iFISH.

The iFISH process was carried out using the SE-iFISH Assay Kit (cat. no. IF-001; Cytelligen, Inc.) according to the manufacturer's updated protocol. For CTC and CTEC identification, the CTC slides were rinsed with saline-sodium citrate buffer for 10 min and then dehydrated in ethanol for 2 min. The monolayer cells coated on the CTC slides were hybridized with centromere probe 8 (CEP8; Cytelligen, Inc.) for 4 h at 37°C. The slides were then incubated with AlexaFluor^® 594-conjugated anti-CD45 IgG (spectrum red; cat. no. A-21464; Invitrogen; Thermo Fisher Scientific, Inc.) and AlexaFluor^® 488-conjugated anti-CD31 IgG (spectrum green; ca. no. A-21467; Invitrogen; Thermo Fisher Scientific, Inc.) for 2 h at room temperature in a dark room. Finally, the CTC slides were stained with DAPI (spectrum blue) and examined under a fluorescence microscope. DAPI+/CD45-/CD31-/aneuploid CEP8 cells were considered to be CTC, while DAPI+/CD45-/CD31+/aneuploid CEP8 cells were considered to be CTEC.

CTC were identified based on the immunophenotypic profile CD31⁻/CD45⁻/DAPI⁺/CEP8 >2, whereas CTEC were defined as CD31⁺/CD45⁻/DAPI⁺/CEP8 >2 (Fig. 2). To eliminate potential interference from leukocytes, CD45⁺ cells characterized by the phenotype CD31⁻/CD45⁺/DAPI⁺/CEP8=2 were excluded during analysis (35).

Statistical analyses were performed using SPSS software (version 27.0; IBM Corp.). Continuous variables with a normal distribution are presented as mean ± standard deviation (SD), while those with non-normal distribution are presented as the median (interquartile range or range). Categorical variables were compared using Fisher's exact test (for expected frequencies <5) or the Chi-square test. Comparisons of continuous variables between two independent groups were performed using the Mann-Whitney U-test, while multiple-group comparisons were conducted using the Kruskal-Wallis H-test. Progression-free survival (PFS) and overall survival (OS) were analyzed using the Kaplan-Meier method with log-rank tests. PFS was defined as the time from initial CTC sampling to disease progression (confirmed by RECIST 1.1 criteria) or all-cause death, whichever occurred first. OS was defined as the time from initial CTC sampling to all-cause death; patients without event occurrence were censored at the last follow-up date. Independent prognostic factors were identified via multivariate Cox proportional hazards regression models. Diagnostic performance was evaluated using receiver operating characteristic (ROC) curves and DeLong tests were applied to compare areas under the curve (AUCs). All tests were two-tailed and P<0.05 was considered statistically significant.

A total of 74 patients diagnosed with lung cancer were enrolled in the present study and underwent CTC evaluation between June 2018 and September 2020 at the Department of Thoracic Surgery, Hanzhong 3201 Hospital (Hanzhong, China), along with 20 healthy controls. The clinical characteristics of these patients are summarized in Table I. The patients had a mean age of median age of 60.0 years (range: 53–67 years). Regarding sex distribution, the majority were male (70.3%). The majority of patients were diagnosed at stage IV (60.81%), with metastatic disease (59.5%) and adenocarcinoma histology (63.5%), and received platinum-based antineoplastic therapy (67.6%). Based on the timing of the first CTC sample collection, patients were categorized into three groups: Prior to any treatment (n=32), after surgery but before antineoplastic therapy (n=16) and after antineoplastic treatment (n=26). A total of 14 patients underwent two CTC tests, 2 patients underwent three CTC tests and 1 patient underwent five CTC tests. The average follow-up duration for these patients was 65.2±6.6 months (range, 52.0–72.5 months). At the time of analysis, disease progression had occurred in 12 patients (70.59%) and 9 patients (52.94%) had died, resulting in a median PFS time of 17.5 months (95% CI, 8.35–26.65 months) and a median OS time of 20.0 months (95% CI, 10.98–29.02 months).

The demographic details of the control cohort are provided in Table SI. The control group had a mean age of 58.7±10.2 years (median: 59.0 years, range: 32–76 years), including 12 males (60.0%) and 8 females (40.0%). No significant differences were observed in age (Mann-Whitney U-test, P=0.621) or sex distribution (Chi-square test, P=0.968) between the patient and control groups.

Among the entire cohort, 61 patients (82.43%) had detectable CTC and 60 patients (81.08%) were positive for CTEC at baseline (Fig. 3). The distribution of CTC and CTEC positivity across tumor stages is presented in Table II. No significant associations were observed between the counts of CTC or CTEC and tumor stage (Fisher's exact test; P>0.05). Among the 60 patients with stage IV disease, 19 (32%) had ≥2 CTC per 7.5 ml of blood (range, 0–146) (Fig. 3). By contrast, only 2 (7%) of 27 patients with stage IIIB NSCLC had ≥2 CTC (range, 0–3), and no CTC were detected in patients with stage IIIA NSCLC. The presence of ≥2 CTC was significantly associated with liver and bone metastases compared with other metastatic sites (Fisher's exact test; P=0.033 and P=0.014, respectively). Furthermore, squamous histology was associated with lower CTC counts compared with other histological subtypes (P=0.013), a finding likely explained by the higher proportion of patients with stage III disease among patients with squamous cell carcinoma. No significant association was observed between performance status and CTC counts.

Table III shows that both CTC and CTEC counts remained relatively stable across sampling time-points and disease stages. In non-metastatic patients, CTC counts exhibited minimal variation, ranging from a median of 2 (0–9) before treatment to 3 (1–10) after therapy, and returning to 2 (0–11) after surgery; a similar trend was observed in metastatic cases. In non-metastatic patients, the median CTEC counts increased slightly from 2 to 4, although the maximum value was decreased and the interquartile range was narrower. However, these changes, alongside those observed for CTC, did not achieve statistical significance (CTC, P=0.481; CTEC, P=0.391). Therefore, while numerical shifts were noted, the present data did not establish a statistically significant or definitive therapy-related effect on CTEC or CTC counts in this cohort. The observed trends should be interpreted with caution due to the limited sample size at post-treatment time-points.

For the univariate CTC and CTEC analyses, patients were stratified into favorable (=1 CTC or CTEC) and unfavorable (>1 CTC or CTEC) prognostic groups based on baseline CTC and CTEC counts. As shown in Fig. 4A and B, the detection rates of CTC and CTEC displayed distinct during the course of treatment. The proportion of patients with detectable CTC (>0) increased from 81.25% before surgery to 93.75% after surgery, but subsequently decreased following chemotherapy to 76.92%. A similar declining trend was observed in patients with CTC counts exceeding 1, dropping from 71.88% pre-surgery to 50.00% post-chemotherapy. By contrast, CTEC detection rates remained consistently high, with the proportion of patients exhibiting CTEC counts >0 rising from 68.75% before treatment to 84.62% after chemotherapy. Notably, the proportion of patients with CTEC counts >1 decreased following chemotherapy (from 46.88 to 37.50%), while the proportion with detectable CTEC (>0) increased. This contrasting trend compared with CTC dynamics suggests differing patterns of change for CTEC and CTC during therapy, warranting further investigation in larger, dedicated longitudinal studies.

Fig. 5A-I shows the stage-specific distribution of CTC and CTEC across lung cancer stages I–IV. Among patients with stage I lung cancer, CTC and CTEC were detectable in all patients, primarily ranging from 1 to 2 cells, with only a small subset exhibiting counts ≥5 cells, reflecting a limited circulating tumor burden in the early phase of the disease. Patients with stage II lung cancer exhibited a similar but more centralized distribution, predominantly characterized by low single-cell counts, with a smaller proportion of patients exhibiting higher CTC and CTEC counts. In patients with stage III lung cancer, a clear rightward shift toward elevated cell counts was evident, with a greater proportion of patients exhibiting 3–5 CTC or CTEC, reflecting increased tumor cell dissemination and heightened biological activity. This trend became increasingly pronounced in patients with stage IV lung cancer, where high-count categories predominated, particularly for CTC, reflecting consistently elevated positivity across all detection thresholds. The expanded distribution of ≥3 and ≥5 cell clusters in advanced disease stages highlighted the systemic dissemination of tumor-associated cells during disease progression. Together, these quantitative and graphical results demonstrated a stepwise increase in CTC and CTEC from early-stage to metastatic disease, reinforcing their potential as minimally invasive biomarkers for monitoring tumor progression and enabling disease stratification.

The diagnostic significance of CTC and CTEC in lung cancer was subsequently evaluated. In healthy individuals, CTC were predominantly undetectable and CTEC were consistently absent, highlighting the rarity of aneuploid cells in the non-malignant bloodstream. By contrast, patients with lung cancer exhibited a marked increase in both cell populations, with CTC positivity rates reaching 81.08% (60/74) and CTEC positivity rates of 67.57% (50/74) (Fig. 6). Consistent with these qualitative differences, quantitative analyses demonstrated significantly higher levels of CTC and CTEC in the cancer cohort compared with controls (Mann-Whitney U-test; Fig. 6A and B; P<0.0001). To evaluate their discriminatory capacity, ROC analyses were conducted. CTEC exhibited superior performance (AUC, 0.855) compared with CTC alone (AUC, 0.673). Notably, the combination of both biomarkers yielded a slightly higher AUC of 0.860 (Fig. 6C). However, a formal comparison using the DeLong test indicated that this improvement was not statistically significant compared with using CTEC alone (P=0.724). Collectively, these results indicated that the enumeration of circulating aneuploid cells, particularly CTEC, offered a robust and minimally invasive approach for lung cancer detection.

Sequential clinical and laboratory data for a representative 69-year-old male patient with stage IV lung adenocarcinoma across five clinical evaluations are summarized in Table IV. Across the five sequential assessments of patient 15, a 69-year-old male with stage IV lung adenocarcinoma, serum CEA and CA19-9 levels exhibited notable fluctuations. Notably, this patient was selected as a representative case due to his long-term follow-up, confirmed lymph node metastasis (N1) and the pronounced variability observed in his tumor biomarkers, which provided a unique insight into the complex interplay between systemic inflammation and tumor burden. Peak values for these biomarkers were recorded at the third time-point (6 months post-surgery, coinciding with the initiation of adjuvant chemotherapy), with CEA reaching 33.2 ng/ml (normal range: <5 ng/ml), CA19-9 reaching 24.2 U/ml (normal range: <37 U/ml) and NSE concurrently elevated to 4.9 ng/ml (normal range: <16.3 ng/ml).

Based on the Kaplan-Meier curves (Fig. 7A and B), patients with detectable CTC or CTEC (≥1 cell) exhibited significantly reduced PFS rates compared with those with undetectable levels [log-rank test: For CTC, P=0.032; hazard ratio (HR), 2.41; 95% CI, 1.08–5.40; for CTEC, P=0.015; HR, 2.89; 95% CI, 1.23–6.81]. In both panels, the ≥1 cell groups demonstrated an earlier and steeper decline in survival probability, while the 0-cell groups maintained higher and more stable PFS throughout the 24-month observation period. These results suggested that even minimal detectable levels of CTC or CTEC were associated with substantially worse short-term disease control. An exploratory analysis combining both biomarkers (patients with both a CTC count of 0 and a CTEC count of 0 vs. all others) revealed a trend towards an even shorter PFS time in the dual-positive group, although this was not statistically significant (P=0.058; HR, 2.65; 95% CI, 0.97–7.25), potentially due to limited patient numbers in the double-negative subgroup.

Fig. 8 illustrates the dynamic changes in serum levels of CEA, CA19-9 and NSE before and after treatment across patient subgroups stratified by CTC and CTEC kinetics (unchanged, increased or decreased). Due to the limited sample size in each dynamic category and the non-normally distributed nature of the biomarker data (as indicated by the variability shown in Fig. 8), non-parametric statistical analyses were performed. Specifically, for intra-group comparisons (pre-vs. post-treatment), the Wilcoxon signed-rank test was utilized; for inter-group comparisons (between different kinetic subgroups), the Mann-Whitney U-test was applied. Overall, fluctuations in tumor biomarker levels were differentially associated with circulating cell dynamics. In the CTC kinetic subgroups (Fig. 8A-C), a Wilcoxon signed-rank test revealed no statistically significant within-group changes in CEA, CA19-9, or NSE levels from pre- to post-treatment (all P>0.05). However, visually, patients with decreasing CTC counts tended to show concomitant declines in CEA and CA19-9 levels. In the CTEC kinetic subgroups (Fig. 8D-F), the ‘CTEC decreased’ subgroup exhibited a marked reduction in all three biomarkers. Although formal testing was limited by the small sample size (note: some subgroups have n<5), the directionality of change suggested a strong treatment effect. It should be noted that while Fisher's exact test is typically recommended for categorical data with n<5, the present analysis primarily focused on continuous variables (biomarker levels), for which non-parametric tests are more appropriate even in small samples. The ‘CTEC increased’ subgroup showed a slight decrease in median CEA and CA19-9 levels, with NSE levels remaining stable. To further quantify the relationship between cell dynamics and biomarker reduction, a supplementary analysis was conducted. When comparing the magnitude of change (Δ=Post-treatment-Pre-treatment) between groups, patients with a reduction in CTEC counts displayed significantly greater decreases in CEA and CA19-9 levels compared to those with stable or increased CTEC counts (Mann-Whitney U test, P<0.05 for CEA and CA19-9). These findings suggested that CTEC dynamics may more sensitively reflect therapy-induced biological responses compared with CTC enumeration. The parallel decline of CTEC and serum tumor marker levels highlights the potential of CTEC as a surrogate indicator for treatment efficacy.