Abstract
Background
Circulating tumor cells (CTCs) play a crucial role in the early diagnosis and prognosis of lung cancer. Identification of a more suitable sample source could be a breakthrough towards enhancing CTC detectability in early‐stage lung cancer. We investigated the differences in detectable CTCs between peripheral arterial and venous blood in early‐ and mid‐stage lung cancer patients undergoing surgery and analyzed the association between clinicopathological factors and detectable CTCs in peripheral arterial and venous blood.
Methods
Peripheral arterial and venous blood was collected in 5‐mL samples from 56 patients with surgically resected and pathologically clear at early‐ or mid‐stage lung cancer. Blood specimens were enriched for CTCs based on isolation by size of epithelial tumor cells. The CTCs were identified using Swiss Giemsa staining and immunohistochemistry for CD45/CD31.
Results
In stage I lung cancer, CTC‐positive rate was significantly higher in peripheral arterial than in venous blood (45.45% vs. 17.39%). There was no significant difference in the number of detectable CTCs between peripheral arterial and venous blood. A low degree of differentiation was associated with a high positive rate of CTCs in peripheral venous blood. The number of circulating tumor microemboli was significantly higher in patients with tumor size >3 cm compared with ≤3 cm.
Conclusion
CTC levels in peripheral arterial and venous blood differed little in lung cancer patients.Compared to peripheral venous blood, peripheral arterial blood had a higher CTC positivity rate in early‐stage lung cancer.This study was favorable for early detection and monitoring of lung cancer.
Keywords: circulating tumor cells, lung cancer, peripheral arterial blood, peripheral venous blood, positivity rate
More CTCs could be detected in peripheral arterial than venous blood in early‐stage lung cancer patients.
INTRODUCTION
Lung cancer has the highest morbidity and mortality rates globally, which are still on the increase. 1 Although early‐stage lung cancer has a good prognosis, it is frequently not identified timeously owing to the absence of apparent symptoms. As a result, the majority of lung cancer patients who display symptoms are already at an advanced stage when they visit the clinic, leading to an overall 5‐year survival rate of <20%. 2 Over the last 10 years, low‐dose spiral computed tomography (LDCT) screening of the lungs, which has gained popularity in Europe and the USA, has reduced the mortality rate of lung cancer by ~20%. 3 , 4 This indicates the significance of early screening for lung cancer. In addition to conventional imaging and tumor marker screening, liquid biopsy techniques, including circulating tumor cells (CTCs) and circulating tumor DNA, have yielded highly promising results in the early detection of cancer. 5 , 6
CTCs are released from solid tumors into the bloodstream. Compared with imaging, CTCs can indicate the presence of early tumors or the recurrence and metastasis of tumors after treatment, months or even years in advance. 7 Despite this, accurately capturing CTCs is vital for their widespread clinical application due to their low number and significant heterogeneity. 8 In addition to maximizing the efficiency and stability of the testing technology or equipment, the time or location of blood sample collection may also affect the efficiency of the testing. For example, recent studies have shown that CTC release is influenced by circadian changes in melatonin levels, which lead to the highest CTC counts during sleep. 9 , 10 Specific to the influence of different blood collection sites on the detection of CTCs, several studies have confirmed that the number of CTCs in pulmonary venous blood is significantly higher than that in peripheral venous blood. 11 , 12 , 13 These studies focused on the impact of lung cancer surgical methods (open or minimally invasive) or procedures (such as the sequence of pulmonary artery and vein removal) on the spread of CTCs. However, this is a technically complex, traumatic, and high‐risk unconventional blood collection method, which is not suitable for early screening of lung cancer and follow‐up of disease outcomes after treatment. CTCs are segregated, filtered, destroyed or cleared in the circulatory system; therefore, the role of the capillary network (including the pulmonary and systemic networks) cannot be ignored. In the present study, we investigated for the first time whether there was a difference in the number of CTCs between peripheral arterial blood and peripheral venous blood, and explored strategies to improve detection of CTCs in peripheral blood in early‐stage lung cancer.
METHODS
Patient eligibility and clinical characteristics
This prospective clinical study gained approval from the Ethics Committee of the Affiliated Cancer Hospital of Shandong First Medical University. We included 56 patients who underwent surgical resection for pathologically confirmed lung cancer. The patients were admitted to the hospital between May and August 2023 and all provided their informed consent before enrolment. The patient data collected included: age and gender; cancer staging (i.e., tumor size and lymph node metastasis) using TNM staging (eighth edition); pathological characteristics including type of pathology, degree of differentiation, lymphatic or venous invasion, pleural invasion, and spread through air spaces; and laboratory tests including platelet count, granulocyte‐to‐lymphocyte count and ratio, D‐dimer, and tumor markers.
We collected 5 mL each of peripheral arterial blood (radial artery) and peripheral venous blood (median elbow vein) from the ipsilateral limb. Blood samples were injected into collection tubes (BD Vacutainer 366 452), gently inverted and mixed eight times, stored at 15–30°C, and subjected to CTC testing within 24 h.
CTC detection by CTCBIOPSY
The fundamental principle of CTCBIOPSY is isolation by size of epithelial tumor cells. This technique, reported by Vona et al., 14 has revolutionized and streamlined the filtration membranes and staining, as well as the entire separation and staining process. The specific detection steps were as follows: 5 mL whole blood was mixed with buffer containing 0.2% formaldehyde to a total volume of 8 mL, and filtered using an 8‐mm‐pore membrane. The captured CTCs and circulating tumor microemboli (CTMs) were stained with Romanowsky stain and air‐dried at room temperature. Based on our own experience and the criteria proposed by other research groups, 15 , 16 the isolated cells were identified as tumor cells only if they exhibited the following morphological characteristics: atypical nuclei (irregular shape and presence of nodular, lobulated contours); nuclear–cytoplasmic ratio >0.8; hyperchromatic nuclei and nonhomogeneous staining; thickened, sunken, wrinkled, and jagged nuclear membranes; nuclear long diameter >18 mm; presence of nuclear chromatin side‐shift, large nucleoli, or abnormal mitotic figures; and presence of tumor cell aggregates, or CTMs. If ≥4 features were seen, the cells were considered to be malignant tumor cells (CTCs). All potential CTCs/CTMs were evaluated in a blind review by three senior cytopathologists to ensure independent identification.
CTC identification (CD45/CD31 immunohistochemistry)
For the captured CTCs/CTMs, immunohistochemical staining of CD45 and CD31 was used to exclude blood intrinsic cells for further identification. The procedure was as follows: 100 μL 0.1% Triton X‐100 was added dropwise, incubated at 18–26°C for 15 min, and washed with deionized water for 2 min, three times. Then, 100 μL of 0.3% H2O2 was added dropwise to the sample slides, which were incubated at 18–26°C for 10 min, and washed three times with PBS for 2 min each time. Next, 100 μL primary antibody working solution of CD45 + CD31 was added dropwise. After 2 h incubation at 18–26°C or overnight at 4°C, the samples were rewarmed for 30 min, and washed three times with PBS for 2 min each time.
After color development, the DAB chromogenic solution was discarded. The samples were rinsed in running water for 5 min, stained with hematoxylin for 5 min, and rinsed in water for 1 s. This was followed by hydrochloric acid alcohol differentiation for 3–8 s before returning to blue in tap water for 15 min with a subsequent rinse under running water. Graded ethanol dehydration consisted of 75%, 95% and 100% ethanol for 1 min each. The samples were air‐dried and the film was sealed with neutral resin. The samples were analyzed by light microscopy. If CD45 or CD31 tested positive, blood origin was confirmed, while CTCs/CTMs were verified if both tested negative.
Statistical analysis
All statistical evaluations were performed using SPSS version 28.0. The χ 2 or Fisher's exact test was used to explore the correlation between CTCs and clinicopathological characteristics. Student's t‐test or Mann–Whitney test was used for continuous variables. Differences were considered statistically significant when p was <0.05.
RESULTS
There were 56 patients (33 [58.9%] male and 23 [41.1%] female), with a mean age of 61.1 (range 32–79) years. There were four cases (7.1%) at stage 0 (carcinoma in situ), 24 (42.9%) at stage I, 16 (28.6%) at stage II, and 12 (21.4%) at stage III. According to pathological type, 44 cases (78.6%) were diagnosed as adenocarcinoma, five (8.9%) as squamous carcinoma, and seven (12.5%) as other types. Other clinical information such as tumor size, lymph node metastasis, platelets, neutrophil/lymphocyte ratio, and tumor markers is summarized in Table 1.
TABLE 1.
Patient characteristics.
| Characteristics | Patients (N = 56) |
|---|---|
| Age, year | |
| Median | 61.1 |
| Range | 32–79 |
| Sex, n (%) | |
| Female | 23 (41.1) |
| Male | 33 (58.9) |
| Histological cell type, n (%) | |
| Adenocarcinoma | 44 (78.6) |
| Squamous cell carcinoma | 5 (8.9) |
| Others | 7 (12.5) |
| pTNM stage, n (%) | |
| Stage 0 | 4 (7.1) |
| Stage I | 24 (42.9) |
| Stage II | 16 (28.6) |
| Stage III | 12 (21.4) |
| Tumor size, n (%) | |
| >3 cm | 27 (48.2) |
| ≤3 cm | 29 (51.8) |
| Lymph node metastasis, n(%) | |
| Negative | 39 (69.6) |
| Positive | 17 (30.4) |
| Histology differentiation, n(%) | |
| Poor | 25 (44.6) |
| Middle | 19 (33.9) |
| Well | 5 (8.9) |
| Uncertain | 7 (12.5) |
| Lymphatic or venous invasion, n(%) | |
| No | 42 (75.0) |
| Yes | 14 (25.0) |
| Pleural invasion, n(%) | |
| No | 40 (71.4) |
| Yes | 16 (28.6) |
| Spread through air spaces, n(%) | |
| No | 32 (57.1) |
| Yes | 24 (42.9) |
| CEA (ng/mL), median ± SD | 6.87 ± 13.70 |
| Cyfra21‐1 (ng/mL), median ± SD | 2.59 ± 2.12 |
| Platelet (109/L), median ± SD | 248.82 ± 83.23 |
| Neutrophil (109/L), median ± SD | 3.72 ± 1.51 |
| Lymphocyte (109/L), median ± SD | 1.75 ± 0.57 |
| N/L, median ± SD | 2.41 ± 1.35 |
| D‐Dimer, median ± SD | 0.46 ± 0.68 |
Abbreviations: CEA, carcinoembryonic antigen; Cyfra21‐1, cytokeratin 19 fragment; N/L, neutrophil‐to‐lymphocyte ratio; TNM, tumor‐node‐metastasis.
There was little significant difference between the average number of CTCs in peripheral arterial blood (2.94 ± 6.54) and peripheral venous blood (1.47 ± 3.37) (t = 1.588, p = 0.118). There was no evidence of a significant difference in the average number of CTMs in peripheral arterial blood compared with venous blood (2.98 ± 10.98 vs. 3.83 ± 11.03) (t = 0.377, p = 0.708). There was no significant difference in the detection of arterial blood CTCs (28/53, 52.83%) and venous blood CTCs (23/55, 41.82%) with respect to detection rate (CTC/CTM ≥1). Similarly, the CTM detection rate (18/53, 33.96% vs. 17/55, 30.91%) did not show any significant difference (χ 2 = 1.313, p = 0.252 and χ 2 = 0.115, p = 0.735, respectively) (Figure 1). If the threshold for determining CTC positivity was set at ≥2/5 mL, both the arterial blood group (18/53, 33.96%) and venous blood group (13/55, 23.64%) had similar rates of positivity with no significant difference (χ 2 = 1.406, p = 0.236). In terms of overall positivity rate of CTC ≥2 or CTM ≥1, there was no significant difference between the arterial blood group (26/53, 49.06%) and venous blood group (22/55, 40.0%) (Table 2).
FIGURE 1.
Circulating tumor cells/circulating tumor microemboli (CTC/CTM) detected in peripheral arterial blood (black arrows indicate CTC/CTM, white arrows indicate white blood cells). (a) Romanowsky staining of morphological characteristics of CTC: nuclear cytoplasmic ratio >0.8; nucleoli are abnormally enlarged; nuclear membranes appear thickened, sunken, wrinkled, and jagged. (b) CD45/CD31 immunohistochemical staining: CTC negative, white blood cells positive. (c) CTM: presence of tumor cells (≥3) aggregation. (d) CD45/CD31 immunohistochemical staining: CTM negative, white blood cells positive. The cells were analyzed under 40x magnification. Scale bar 20 mm.
TABLE 2.
Detection of CTCs and CTMs in arterial and venous blood of lung cancer patients.
| Group | CTCs number (per 5 mL) | CTM number (per 5 mL) | Detectable CTCs number (≥1) | CTM positive rate* | CTCs positive rate** | Overall positivity rate*** |
|---|---|---|---|---|---|---|
| Arterial | 2.94 ± 6.54 | 2.98 ± 10.98 | 28/53 (52.83) | 18/53 (33.96) | 18/53 (33.96) | 26/53 (49.06) |
| Venous | 1.47 ± 3.37 | 3.83 ± 11.03 | 23/55 (41.82) | 17/55 (30.91) | 13/55 (23.64) | 22/55 (40.00) |
| T/X2 value | 1.588 | 0.377 | 1.313 | 0.115 | 1.406 | 0.897 |
| p‐value | 0.118 | 0.708 | 0.252 | 0.735 | 0.236 | 0.344 |
Abbreviations: CTC, circulating tumor cells; CTM, circulating tumor microemboli.
Detectable CTMs ≥1.
Detectable CTCs ≥2.
Detectable CTCs ≥2 or detectable CTMs ≥1.
The disparity in the positivity rates of CTCs in peripheral arterial and venous blood in varying TNM stages of lung cancer was most prominent in stage I. The positivity rate of CTCs in peripheral arterial blood was 45% (10/22) while that in peripheral venous blood was 17.39% (4/23), which was a significant difference (χ 2 = 4.132, p = 0.042). However, there was little significant difference in the positive rate of CTCs between arterial and venous blood in other stages (Table 3). In addition, patients with a tumor size >3 cm displayed a significantly higher number of CTMs in venous blood than patients with tumor size ≤3 cm (p = 0.03) (Figure 2). In venous blood, the degree of differentiation was associated with CTC positive rate, suggesting that CTC positivity increased with a lower degree of differentiation (p = 0.020; Figure 3). Pathological factors, including age, gender, pathological type, tumor markers, platelet count, and neutrophil/lymphocyte count or ratio were not associated with the numbers of CTCs or CTMs detected.
TABLE 3.
CTC positive rate of arteriovenous blood in patients with different TNM stages of lung cancer.
| TNM | CTC positive rate of arterial blood (≥2) | CTC positive rate of venous blood (≥2) | X 2 value | p‐value |
|---|---|---|---|---|
| Stage I (n = 24) | 10/2 (45.45) | 4/23 (17.39) | 4.132 | 0.042 |
| Stage II (n = 16) | 6/15 (40.00) | 5/16 (31.25) | 0.259 | 0.611 |
| Stage III (n = 12) | 2/12 (16.67) | 2/12 (16.67) | 0.000 | 1.000 |
Note: CTC positive rate (cutoff 2): detectable CTCs ≥2.
Abbreviations: CTC, circulating tumor cells; TNM, tumor‐node‐metastasis.
FIGURE 2.
The number of circulating tumor microemboli (CTM) was significantly higher in patients with tumor size >3 cm compared with ≤3 cm.
FIGURE 3.
Degree of differentiation was associated with circulating tumor cells (CTCs)positive rate: the lower the degree of differentiation, the higher the positive rate of CTCs.
DISCUSSION
CTCs are rare in peripheral blood, with usually 1 per 109 (billion) blood cells or 1 per 106–107 leukocytes. 17 CTCs have a high degree of heterogeneity, including morphological and molecular information heterogeneity. In terms of morphological heterogeneity, CTCs exist in various forms such as single cells, cell clusters and platelet‐encapsulated cells, and mixed clusters with leukocytes and neutrophils, in addition to different cell volume and nuclear morphology. 18 , 19 , 20 In terms of molecular information heterogeneity, the nucleic acid information and protein expression carried by different patients, different periods and even each CTCs may be different. 21 , 22 These biological characteristics pose challenges to the capture, identification, and downstream analysis of CTCs, so the overall detection rate of lung cancer CTCs is not high, even in advanced cancer. Allard et al. 23 reported that 2183 peripheral venous blood specimens from 964 distant metastatic carcinomas were detected with the CellSearch system, and the positive rate of CTCs was only 36%. Tanaka et al. 24 used a semiautomated system to detect CTCs in peripheral venous blood of 125 diagnosed primary lung cancers (31 of which had distant metastases) with a sample size of 7.5 mL, and the detection rate was only 30.6%.
Improving the detection rate and content of CTCs as much as possible is a prerequisite to guarantee their widespread clinical application. Peripheral venous blood collection is simple, minimally invasive, and is the main sample source for CTC detection. Some studies have investigated the effect of other blood sample sources on the detection of CTCs. Terai et al. 25 obtained blood samples from the common femoral artery and the anterior elbow vein of 17 patients with multiple hepatic metastases of uveal melanoma for CTC measurements. They found that CTCs were detected more frequently (100%) and in greater numbers (median 5, range 1–168) (venous 52.9%; median 1, range 0–8). Murlidhar et al. 12 drew pulmonary and peripheral venous blood specimens from patients with lung cancer in the perioperative period and assessed the CTC load using a microfluidic device, from 108 blood samples analyzed from 36 patients, Pulmonary venous blood had a significantly higher number of CTCs than preoperative (p < 0.0001) and intraoperative peripheral venous blood (p < 0.001). Similarly, Tamminga et al. 13 showed a 70% CTC detection rate in the pulmonary veins of 31 patients undergoing surgery, which was significantly higher than the count in the peripheral radial artery (22%).
Patients with lung cancer exhibit notably higher levels of CTCs in the pulmonary vein or common femoral artery compared to peripheral venous blood, and this disparity may be attributed to differential CTC attrition within the circulatory system. CTCs, as they traverse the bloodstream, are susceptible to various processes including isolation, filtration, degradation, clearance, and even programmed cell death, with a mere 0.01% of CTCs contributing to the formation of metastatic lesions. 26 , 27 The pivotal role played by the capillary network in this context cannot be underestimated. In a theoretical framework, if lung cancer CTCs enter the bloodstream through the bronchial vein, those found in peripheral venous blood must navigate a complex route involving the right cardiac system, pulmonary artery, pulmonary capillary network, pulmonary vein, left cardiac system, aorta, branching networks, systemic capillary networks, and the systemic venous system. This involves traversal through two distinct capillary networks. Conversely, lung cancer CTCs within peripheral arterial blood samples effectively bypass the systemic capillary network and systemic venous system. Should lung cancer CTCs enter the bloodstream via the pulmonary vein, although CTCs in peripheral venous blood samples avoid the pulmonary capillary network, they still necessitate passage through the left cardiac system, aorta, branching networks, systemic capillary networks, and the systemic venous system. This route includes traversal through an additional capillary network. In contrast, lung cancer CTCs in peripheral arterial blood samples undergo a more straightforward course, involving only the left cardiac system and the aorta, completely avoiding both capillary networks and the systemic venous system. As a result, lung cancer CTCs in peripheral arterial blood endure significantly less attrition than their counterparts in peripheral venous blood. Given this mechanism, the collection of pulmonary venous blood for CTC analysis emerges as a potentially highly effective approach to enhance the detection and positivity rates of lung cancer CTCs. This is primarily because the pulmonary vein serves as the exclusive conduit for tumor cells from an entire lung lobe directly into the bloodstream, thereby bypassing the need for protracted blood circulation and exposure to multiple capillary networks. However, it is crucial to acknowledge that collecting pulmonary vein blood necessitates open‐heart surgery and is primarily a disposable procedure. This method is characterized by its technical complexity, invasiveness, and elevated associated risks, rendering it impractical for early lung cancer diagnosis, screening, or post‐treatment monitoring. Conversely, the collection of peripheral arterial blood, particularly radial arterial blood, presents a considerably more straightforward, less invasive, and generally well‐accepted alternative for patients. Nonetheless, it is essential to explore whether this approach offers advantages over peripheral venous blood collection in the context of lung cancer diagnosis and monitoring.
This study found that there was no obvious difference in the mean number of CTCs and the number of detectable CTCs between peripheral blood from the radial artery and peripheral blood from a vein in the elbow. In addition, there was little difference in the mean number of CTMs and the number of detectable CTMs between the two sample sites. These results demonstrate that using blood collected from the peripheral radial artery does not enhance the detection of lung cancer CTCs when compared to using median elbow veins. Previous research has highlighted a noteworthy variation in patient prognosis when CTC ≥2 is used as the affirmative diagnostic threshold. 28 Our study showed that the positive arterial blood CTCs were higher in those with early‐stage (stage I) lung cancer than in peripheral venous blood. This suggests that detecting CTCs in peripheral arterial blood compared to peripheral venous blood may lead to earlier diagnosis, thus aiding in the early detection and monitoring of lung cancer. Our findings indicate that there is a correlation between the degree of differentiation and the positive rate of CTCs, as well as between the size of the tumor and the number of CTMs in peripheral venous blood. This emphasizes the importance of venous blood collection in our study.
In conclusion, our study revealed no significant difference in the CTC levels between the peripheral radial artery and venous blood in lung cancer patients. Additionally, there was no observable increase in the number of CTCs by puncturing the radial artery, which could potentially lead to additional trauma. It is important to note that venous blood samples for CTC testing are typically collected concurrently with other blood tests. However, in the case of early‐stage lung cancer, particularly for patients scheduled for surgical resection, increasing the CTC positivity rate is possible by collecting peripheral radial artery blood. This method can be synchronized with blood gas analysis to reduce trauma and is more favorable for the early diagnosis and efficacy monitoring of lung cancer.
AUTHOR CONTRIBUTIONS
Conceptualization, Zhen‐dan Wang and Guo‐dong Zhang; Resources, Sheng Li; Methodology, Ying Ma, Ji‐yan Liu and Di‐hua Li; Validation, Guo‐dong Zhang; Investigation, Zhen‐dan Wang; Data curation, Yi‐fei Feng; Writing–original draft preparation, Zhen‐dan Wang and Yu‐shuo Wang; Writing–review and editing, Guo‐dong Zhang and Sheng Li; Funding acquisition, Zhen‐dan Wang and Sheng Li. All authors have read and agreed to the published version of the manuscript.
FUNDING INFORMATION
This project was supported by the Natural Science Foundation of Shandong Province (ZR2020QH205) and Academic Promotion Project of Shandong First Medical University (2019QL004).
CONFLICT OF INTEREST STATEMENT
The authors declare that there is no conflict of interest.
ACKNOWLEDGMENTS
The authors thank assistant doctors Xue Hongwu and Shi Zhigang from the Department of Thoracic Surgery at Shandong Cancer Hospital, for their contribution to patient information collection. The authors appreciate the support of the staff of the Department of Anesthesia and Surgery in arterial blood collection and the nursing staff of the Department of Thoracic Surgery at Shandong Cancer Hospital for their help and assistance with venous blood collection.
Wang Z, Feng Y, Wang Y, Ma Y, Liu J, Li D, et al. Peripheral arterial rather than venous blood is a better source of circulating tumor cells in early lung cancer. Thorac Cancer. 2024;15(8):654–660. 10.1111/1759-7714.15236
Zhen‐dan Wang and Yi‐fei Feng contributed equally to this article.
REFERENCES
- 1. Leiter A, Veluswamy RR, Wisnivesky JP. The global burden of lung cancer: current status and future trends. Nat Rev Clin Oncol. 2023;20(9):624–639. [DOI] [PubMed] [Google Scholar]
- 2. Allemani C, Matsuda T, Di Carlo V, Harewood R, Matz M, Nikšić M, et al. Global surveillance of trends in cancer survival 2000‐14 (CONCORD‐3): analysis of individual records for 37 513 025 patients diagnosed with one of 18 cancers from 322 population‐based registries in 71 countries. Lancet. 2018;391(10125):1023–1075. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 3. de Koning HJ, van der Aalst CM, de Jong PA, Scholten ET, Nackaerts K, Heuvelmans MA, et al. Reduced lung‐cancer mortality with volume CT screening in a randomized trial. New Engl J Med. 2020;382(6):503–513. [DOI] [PubMed] [Google Scholar]
- 4. Aberle DR, Adams AM, Berg CD, Black WC, Clapp JD, Fagerstrom RM, et al. Reduced lung‐cancer mortality with low‐dose computed tomographic screening. New Engl J Med. 2011;365(5):395–409. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 5. Lawrence R, Watters M, Davies CR, Pantel K, Lu YJ. Circulating tumour cells for early detection of clinically relevant cancer. Nat Rev Clin Oncol. 2023;20(7):487–500. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6. Wang Z, Xie K, Zhu G, Ma C, Cheng C, Li Y, et al. Early detection and stratification of lung cancer aided by a cost‐effective assay targeting circulating tumor DNA (ctDNA) methylation. Respir Res. 2023;24(1):163. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7. Liu MC, Shields PG, Warren RD, Cohen P, Wilkinson M, Ottaviano YL, et al. Circulating tumor cells: a useful predictor of treatment efficacy in metastatic breast cancer. J Clin Oncol. 2009;27(31):5153–5159. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8. Ring A, Nguyen‐Sträuli BD, Wicki A, Aceto N. Biology, vulnerabilities and clinical applications of circulating tumour cells. Nat Rev Cancer. 2023;23(2):95–111. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9. Diamantopoulou Z, Castro‐Giner F, Schwab FD, Foerster C, Saini M, Budinjas S, et al. The metastatic spread of breast cancer accelerates during sleep. Nature. 2022;607(7917):156–162. [DOI] [PubMed] [Google Scholar]
- 10. Zhu X, Suo Y, Fu Y, Zhang F, Ding N, Pang K, et al. In vivo flow cytometry reveals a circadian rhythm of circulating tumor cells. Light Sci Appl. 2021;10(1):110. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 11. Shima H, Tanabe N, Sato S, Oguma T, Kubo T, Kozawa S, et al. Lobar distribution of non‐emphysematous gas trapping and lung hyperinflation in chronic obstructive pulmonary disease. Respir Investig. 2020;58(4):246–254. [DOI] [PubMed] [Google Scholar]
- 12. Murlidhar V, Reddy RM, Fouladdel S, Zhao L, Ishikawa MK, Grabauskiene S, et al. Poor prognosis indicated by venous circulating tumor cell clusters in early‐stage lung cancers. Cancer Res. 2017;77(18):5194–5206. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 13. Tamminga M, de Wit S, van de Wauwer C, van den Bos H, Swennenhuis JF, Klinkenberg TJ, et al. Analysis of released circulating tumor cells during surgery for non‐small cell lung cancer. Clin Cancer Res. 2020;26(7):1656–1666. [DOI] [PubMed] [Google Scholar]
- 14. Vona G, Sabile A, Louha M, Sitruk V, Romana S, Schütze K, et al. Isolation by size of epithelial tumor cells: a new method for the immunomorphological and molecular characterization of circulatingtumor cells. Am J Pathol. 2000;156(1):57–63. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15. Hofman VJ, Ilie MI, Bonnetaud C, Selva E, Long E, Molina T, et al. Cytopathologic detection of circulating tumor cells using the isolation by size of epithelial tumor cell method: promises and pitfalls. Am J Clin Pathol. 2011;135(1):146–156. [DOI] [PubMed] [Google Scholar]
- 16. Hofman V, Long E, Ilie M, Bonnetaud C, Vignaud JM, Fléjou JF, et al. Morphological analysis of circulating tumour cells in patients undergoing surgery for non‐small cell lung carcinoma using the isolation by size of epithelial tumour cell (ISET) method. Cytopathology. 2012;23(1):30–38. [DOI] [PubMed] [Google Scholar]
- 17. Deng Z, Wu S, Wang Y, Shi D. Circulating tumor cell isolation for cancer diagnosis and prognosis. EBioMedicine. 2022;83:104237. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18. Aceto N, Toner M, Maheswaran S, Haber DA. En route to metastasis: circulating tumor cell clusters and epithelial‐to‐mesenchymal transition. Trends Cancer. 2015;1(1):44–52. [DOI] [PubMed] [Google Scholar]
- 19. Cools‐Lartigue J, Spicer J, McDonald B, Gowing S, Chow S, Giannias B, et al. Neutrophil extracellular traps sequester circulating tumor cells and promote metastasis. J Clin Invest. 2013;123(8):3446–3458. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20. Liu X, Song J, Zhang H, Liu X, Zuo F, Zhao Y, et al. Immune checkpoint HLA‐E:CD94‐NKG2A mediates evasion of circulating tumor cells from NK cell surveillance. Cancer Cell. 2023;41(2):272‐87.e9. [DOI] [PubMed] [Google Scholar]
- 21. Gkountela S, Castro‐Giner F, Szczerba BM, Vetter M, Landin J, Scherrer R, et al. Circulating tumor cell clustering shapes DNA methylation to enable metastasis seeding. Cell. 2019;176(1–2):98–112.e14. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 22. Giordano A, Gao H, Anfossi S, Cohen E, Mego M, Lee BN, et al. Epithelial‐mesenchymal transition and stem cell markers in patients with HER2‐positive metastatic breast cancer. Mol Cancer Ther. 2012;11(11):2526–2534. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 23. Allard WJ, Matera J, Miller MC, Repollet M, Connelly MC, Rao C, et al. Tumor cells circulate in the peripheral blood of all major carcinomas but not in healthy subjects or patients with nonmalignant diseases. Clin Cancer Res. 2004;10(20):6897–6904. [DOI] [PubMed] [Google Scholar]
- 24. Tanaka F, Yoneda K, Kondo N, Hashimoto M, Takuwa T, Matsumoto S, et al. Circulating tumor cell as a diagnostic marker in primary lung cancer. Clin Cancer Res. 2009;15(22):6980–6986. [DOI] [PubMed] [Google Scholar]
- 25. Terai M, Mu Z, Eschelman DJ, Gonsalves CF, Kageyama K, Chervoneva I, et al. Arterial blood, rather than venous blood, is a better source for circulating melanoma cells. EBioMedicine. 2015;2(11):1821–1826. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 26. Chambers AF, Groom AC, MacDonald IC. Dissemination and growth of cancer cells in metastatic sites. Nat Rev Cancer. 2002;2(8):563–572. [DOI] [PubMed] [Google Scholar]
- 27. Méhes G, Witt A, Kubista E, Ambros PF. Circulating breast cancer cells are frequently apoptotic. Am J Pathol. 2001;159(1):17–20. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 28. Ren L, Zhong X, Liu W, Xu D, Lei Y, Zhou J, et al. Clinical significance of a circulating tumor cell‐based classifier in stage IB lung adenocarcinoma: a multicenter, Cohort Study. Ann Surg. 2023;277(2):e439–e448. [DOI] [PMC free article] [PubMed] [Google Scholar]