Circulating tumour cells as liquid biopsy: technical aspects

Cancer is one of the leading causes of death worldwide, and metastasis is responsible for more than 90% of the mortality of cancer patients. Metastasis occurs when tumour cells leave the primary tumour, travel through the bloodstream as circulating tumour cells (CTCs), and then colonise secondary tumour sites distant from the primary tumour. CTCs, which are cells that have become detached from primary tumours, are the most representative biomarkers of the biological functions of metastatic development and their analysis has the potential to reveal the key mechanisms of tumour progression.

 

The analysis of CTCs in the blood of patients with cancer was termed ‘liquid biopsy’.^1,2 Blood samples can be obtained and analysed at the time of diagnosis and during treatment. The analysis of the liquid biopsy provides important information on the molecular properties of tumour lesions. This information contributes to the early detection of metastatic lesions and aids in the personalised treatment of cancer patients, such as prognostic evaluation, stratification of patients for targeted therapies, real-time monitoring of treatment efficacy, identification of therapeutic targets, and resistance mechanisms. Numerous clinical studies and meta-analyses, including large cohorts of patients, have shown that the number of CTCs is an important indicator of the risk of progression or death in patients with metastatic solid cancer (for example, breast, prostate, colon).^3–7

 

However, in-depth investigation of CTCs remains technically challenging. CTCs occur as very low concentrations of one tumour cell among millions of blood cells. Their identification and characterisation require extremely sensitive and specific analytic methods, which are usually a combination of enrichment and detection procedures.

 

This article focuses on current technologies used for the enrichment and detection of CTCs. A number of innovative technologies to improve methods for CTC detection have been developed, including CTC microchips, filtration devices, quantitative reverse-transcription polymerase chain reaction (PCR) assays, automated microscopy systems and functional assays. Among the considerable number of promising CTC detection techniques that have been developed, the analytical specificity and clinical utility must be demonstrated for their introduction into clinical practice. 

 

CTC enrichment includes a large panel of technologies based on the different properties of CTCs that must differentiate them from the normal haematopoietic cells: (i) physical properties (for example, size, density, electric charges and deformability); and (ii), biological properties (for example, surface protein expression and invasion capacity; see Table 1).

 

 

Biological properties are mainly used in immunological procedures with antibodies against either tumour-associated antigens (positive selection) or common leucocyte antigen CD45 (negative selection). Positive enrichment typically attains high cell purity, which depends on antibody specificity. Negative enrichment technologies evade some of the pitfalls of positive enrichment; for example, CTCs are not tagged with a difficult-to-remove antibody, they are not activated or modified via an antibody–protein interaction and antibody selection does not bias the subpopulation of CTCs captured. However, these advantages come at the cost of purity, as negative enrichment strategies typically have a much lower purity than positive enrichment8–10 and require a suitable CTC detection step.

 

Mostly used, immunomagnetic systems target an antigen with an antibody, which is coupled to a magnetic bead, and the antigen–antibody complex is isolated via exposure to a magnetic field. Positive selection is usually carried out with antibodies against the epithelial cell adhesion molecule (EpCAM). Among the current EpCAM-based technologies, the FDA-approved CellSearch™ system has gained considerable attention over the past ten years and is frequently compared with all new CTC detection methods as the gold standard. 

 

However, capturing CTCs lacking EpCAM has involved the use of antibody cocktails against various other epithelial cell surface antigens (for example, EGFR, MUC1), or against tissue-specific antigens (for example, PSA, HER2) and against mesenchymal or stem cell antigens (Snail, ALDH1).¹¹ Many commercial platforms for CTC detection using positive magnetic enrichment are available, including: 

 

At present, there is a focus on the development of microfluidics devices (‘chips’) such as Ephesia Chip¹⁶ or CTC-iChip.¹⁷ Interestingly, a unique in vivo device, GILUPI CellCollector^®, was developed to capture CTCs directly in the vein of patients, with a structured medical Seldinger guidewire functionalised with an antibody targeting EpCAM to trap CTCs. This wire allows screening of a large volume of blood during the 30-min collection period. 

 

Positive selection of CTCs requires an assumption about the unknown nature of CTCs in an individual blood sample. This bias is avoided by negative selection in which the blood sample is depleted of unwanted cells.

 

Negative enrichment uses an indirect method to isolate CTCs: they target and remove background cells, such as leucocytes, using antibodies against CD45 (which is not expressed on carcinomas or other solid tumours) and other leukocyte antigens, to achieve a CTC-enriched sample. 

 

The RosetteSepTM CTC Enrichment Cocktail (STEMCELL Technologie) offers a unique method for further depletion of unwanted cells by integrating immunoaffinity-based enrichment with density centrifugation. RosetteSep™ targets unwanted cells and forms a pellet with red blood cells (RBCs) through tetrameric antibody complexes that target an extensive mixture of specialised antigens. An immunomagnetic version is also available – the EasySep™ system – which contains magnetic nanoparticles and tetrameric antibody complexes targeting CD45, for example. 

 

Some technologies are flexible and can function either as positive or as negative enrichment systems by applying different antibodies (for example, replacing anti-EpCAM, specific for epithelial cells with anti-CD45, specific for leucocytes). For example, the CTC-iChip could use an immunomagnetic selection with functionalised beads against EpCAM or CD45 and CD66b.¹⁷

 

Numerous marker-independent techniques have been developed for CTC isolation and detection. Label-free enrichment processes based on physical properties, such as density, size, deformability, and electric charge, avoid molecular bias induced by variability of cell biomarker expression associated with tumour heterogeneity.

 

Density gradient centrifugation was one of the first methods recorded for CTC isolation.¹⁸ Although not originally developed for CTC isolation, Ficoll-Paque^®, a density gradient medium for the separation and isolation of mononuclear cells, has been used in research for some time. However, after such a non-specific pre-analytical process, CTCs are still present in a large number of leucocytes; indeed, only erythrocytes and polynuclear cells are depleted using Ficoll-Paque centrifugation. To improve this enrichment, a subsequent positive or negative step is usually required and performed.

 

Designed for CTC isolation, OncoQuick (Greiner Bio-One) employs a liquid separation medium that has been optimised for the specific enrichment of CTCs only, based on their buoyant density under appropriate conditions and no additional step is required because even the leucocytes are eliminated from the cell monolayer between the plasma and the Ficoll. 

 

Microfiltration technologies, based on the precedent that CTCs generally exhibit a larger morphology than leukocytes, such as ScreenCell^®,¹⁹ ISET^®,²⁰ CellSieve™,^21,22 or Parsortix™²³ involve flowing blood through pores or microfluidic steps of calibrated size to trap larger cells (the CTCs) while smaller cells pass through. 

 

Some other size-based microfluidic devices use inertial focusing to separate CTCs from blood. Vortex technology relies on inertial microfluidics and laminar microscale vortices to position cells along channel walls upstream of micro-vortices designed to stably trap CTCs.²⁴ In the same way, the ClearCell FX technology uses Dean Flow and hydrodynamic focusing within a curved channel for differential collection of CTCs and WBC streams.²⁵ However, this method includes lysing the RBCs prior to isolation of CTCs.

 

Such technologies or approaches have the advantages of being less complicated, sometimes rapid, and requiring minimal equipment. However, some of these approaches may be prone to clogging and the release of the CTCs into suspension for further analysis is challenging.

 

An innovative approach for cell separation, dielectrophoresis (DEP), exploits the distinct electrical fingerprints of different cells, which depend on their composition (for example, cell membrane, nucleus, organelles), morphology (for example, size, shape), and phenotype. ApoStream^®, a commercial system for CTC enrichment, applies the first strategy through the use of a microfluidic device using DEP field-flow assist cell separation technique.²⁶

 

Few methods have been developed for specialised detection of CTC clusters. In most cases, CTC clusters were observed when detecting individual CTCs. Multiple studies using microfiltration for CTC isolation have reported capturing CTC clusters (microemboli).^19,27–29 These reports indicate that CTC clusters stay intact when isolated with microfiltration methods, whereas other enrichment strategies might either fail to capture clusters or break them apart. Sarioglu et al reported the development of a novel three-dimensional microfiltration system (the Cluster-Chip) specifically designed to capture CTC clusters.³⁰ The simple, but sophisticated, design of the Cluster-Chip captures these circulating tumour microemboli using multiple rows of shifted triangular pillars. 

 

After enrichment, the CTC fraction still contains a substantial number of leucocytes, and CTCs need to be identified specifically at the single-cell level by a robust and reproducible method that can distinguish them from normal blood cells.

 

CTCs can be detected by using a combination of membrane and/or intra-cytoplasmic anti-epithelial, anti-mesenchymal, anti-tissue-specific marker or anti-tumour-associated antibodies.¹¹ However, many CTC assays use the same identification step as the CellSearch^® system: cells are fluorescently stained for cytokeratins (CK), the common leucocyte antigen CD45 and a nuclear dye (DAPI). Through multicolour image analysis with a fluorescence microscope, CTCs are defined as CK+/CD45−/DAPI+ cells. Although some antigens are applicable to various different cancer types (for example, CK for breast, colon and prostate cancer and other epithelial tumours), tissue-specific antigens are also suitable (PSA for prostate cancer).

 

Ultra-high speed automated digital microscopy using fiber optic array scanning technology (FAST) has been developed to detect CTCs that are labelled by antibodies with fluorescent conjugates.^31,32 Other slide-based automated scanning microscopes have been introduced for detecting CTCs, including the Ikoniscope^® imaging system and the Ariol^® system,^33,34 and show promising results that still need to be validated in large clinical studies.

 

Some researchers also use flow cytometry technology, which offers high throughput detection and labeling of several targets at the same time (9 or 12 targets).³⁵ Moreover some cytometers allow cell sorting for post-processing analysis.³⁶ Indeed, immunological detection offers the advantage of allowing isolation of stained CTCs for subsequent molecular characterisation. While manual isolation by micromanipulation of CTCs is possible,³⁷ it is rather arduous and time-consuming; an alternative automated single cell selection device has therefore been developed. 

 

The DEPArray™ technology, based on a dielectrophoresis strategy by trapping single cells in DEP cages,³⁸ is designed for single-cell recovery of CTCs. Multiple clinical studies have used DEPArray™ to detect and recover single CTCs for subsequent genetic analyses following enrichment.^39–41 The use of this technology for CTC recovery will likely be limited to samples with a relatively high number of CTCs due to a cell-loss of approximately 40% during sample loading.⁴²

 

Nucleic acid-based CTC detection methods are the most widely used alternatives to immunological assays. These technologies identify specific tumour DNA or mRNA to confirm the presence of CTCs indirectly.⁴³ Detection involves designing specific primers supposedly associated with CTC-specific genes. These genes either code for tissue-, organ-, or tumour-specific proteins, or, more specifically, contain known mutations, translocations, or methylation patterns found in cancer cells.⁴⁴

 

The nucleic acid-based method has the highest sensitivity but lacks specificity, owing to the potential of captured non-cancerous cells to generate false-positive signals, thus decreasing the overall accuracy. According to previous work, not all the detection targets are tumour specific, because the same targets can be found in blood cells.^45,46 Pantel et al have reported that CK-19, a major marker for CTC detection, is also present in immune cells.⁴⁷ Multiplex PCR, such as the AdnaTest kit (AdnaGen AG), could overcome this limitation.^45,48 Moreover, considering the genetic heterogeneity of CTCs, a multiplex PCR assay might provide improved sensitivity and specificity rates.

 

Functional assays exploit aspects of live cellular activity for CTC detection. Interestingly, these assays have the particularity to focus on the discovery of ‘metastasis competent cells’. In order to detect only viable CTCs, the functional Epithelial ImmunoSPOT (EPISPOT) assay, was introduced for in vitro CTC detection.⁴⁹ This technology assesses the presence of CTCs based on secretion, shed or release of specific proteins during 24–48 hours of short-term culture.⁵⁰ EPISPOT has been applied to blood and bone marrow samples, and validated in several different cancers, for example, breast and colon with the CK19 EPISPOT assay, or prostate with the PSA-EPISPOT assay.^51–54

 

This test is currently being further developed into a liquid format with micro-droplets (called EPIDROP) that allows capture of single viable CTCs and subsequent molecular characterisation. Another in vitro functional assay, Vita-Assay™ (Vitatex), exploits the preferential adhesion of invasive rare blood cells to a specialised matrix to enrich viable CTCs from blood up to one million-fold.⁵⁵ This method has also been tested in metastatic prostate⁵⁶ and breast cancer.⁵⁷

 

In vivo, important information can be obtained by transplantation of patient-derived CTCs into immunodeficient mice: metastases that were grown after xenotransplantation of enriched CTCs have the most characteristics of metastasis-initiator cells.⁸ A report on patients with small-cell lung cancer showed that CTCs from patients with either chemosensitive or chemorefractory tumours are tumourigenic in immunocompromised mice, and the resultant CTC-derived explants mirrored the response of the donor patient to platinum and etoposide chemotherapy.⁵⁸ However, at present, these in vivo assays require very high CTC yields in the transplanted blood sample, which have so far only been achieved in a few patients.

 

CTCs hold the key to understand the biology of metastasis and provide a biomarker to non-invasively measure the evolution of tumour subclone during treatment and disease progression. Improvements in technologies to yield purer CTC populations might now enable better cellular and molecular investigations. Characterisation of CTCs allows better insights into tumour heterogeneity, within most assays, including immunofluorescence, array CGH, next generation sequencing of both DNA and RNA, and fluorescence in situ hybridisation (FISH).

 

Protein analyses on single CTCs are currently performed by immunostaining with antibodies directed against protein of interest. Multiple labelling is possible but usually restricted to a few proteins of interest for tumour cell biology and cancer therapy. This may help to identify signaling pathways relevant to metastasis development and treatment responses. In breast cancer patients, the HER2 status of CTCs could be assessed and show discrepancies with primary tumour status.^59,60 More recently, immune checkpoint regulators such as programmed death-ligand 1 have become exciting new therapeutic targets and could be used for liquid biopsy in future clinical trials on patients undergoing immune checkpoint blockage.^61,62

 

For single-cell sequencing to identify genomic and transcriptomic characteristics of CTCs, most studies have focused on genomic analyses and carried out whole genome amplifications (WGAs) to increase the amount of DNA, which is subsequently subjected to the analyses of specific mutations and copy number variations using conventional and next-generation sequencing technologies.^37,63,64 As an example, CTCs with mutated KRAS genes will escape anti-EGFR therapy and their early detection might help to guide therapy in individual patients, although it noteworthy that WGA has the inherent risk of inducing a bias and the results need to be therefore carefully validated. Besides isolation of single CTCs, enrichment by 3–4 log units might be sufficient to obtain a concentration of one CTC in 1000 blood cells, which is in the range that is suitable for highly sensitive mutation analyses technologies such as digital PCR.⁶⁵ 

 

Another approach is FISH analysis of single CTCs identified by immunocytochemistry.^66,67 Such an immuno-FISH approach can be combined with automated detection of CTCs and might be easier to implement in future clinical diagnostics.

 

The recent explosion in the field of CTC biology is reflected in the myriad of CTC technologies developed within the last decades. New technologies have arisen to address new challenges as our understanding of CTC biology evolves permanently. Recent focuses on the epithelial-to-mesenchymal transition and stem cell markers in CTCs illuminated the potential values of new biomarkers on CTCs and they may provide information of clinical interest.

 

While clinical studies using CellSearch™ and other CTC technologies have affirmed that CTC enumeration provides relevant prognostic information and clinical validity, the potential for liquid biopsy to address clinical utility is still under investigation. In conclusion, liquid biopsy diagnostics might help to focus the current cancer screening modalities, which could potentially reduce health care costs. 

 

The authors received support from DGOS, the National Institute of Cancer (INCA), ARC Foundation and CANCER-ID, an Innovative Medicines Initiative Joint Undertaking under grant agreement no. 115749, based on financial contributions from the European Union’s Seventh Framework Program (FP7/2007-2013), and EFPIA companies’ in-kind contribution.

 

1 Pantel K, Alix-Panabieres C. Circulating tumour cells in cancer patients: challenges and perspectives. Trends Molec Med 2010;16(9):398–406.

2 Merker JD et al. Circulating tumor DNA analysis in patients with cancer: American Society of Clinical Oncology and College of American Pathologists Joint Review. J Clin Oncol 2018:Jco2017768671.

3 Cristofanilli M et al. Circulating tumor cells, disease progression, and survival in metastatic breast cancer. N Engl J Med 2004;351(8):781–91.

4 Cohen SJ et al. Relationship of circulating tumor cells to tumor response, progression-free survival, and overall survival in patients with metastatic colorectal cancer. J Clin Oncol 2008;26(19):3213–21.

5 de Bono JS et al. Circulating tumor cells predict survival benefit from treatment in metastatic castration-resistant prostate cancer. Clin Cancer Res 2008;14(19):6302–9.

6 Zhang L et al. Meta-analysis of the prognostic value of circulating tumor cells in breast cancer. Clin Cancer Res 2012;18(20):5701–10.

7 Bidard FC et al. Clinical validity of circulating tumour cells in patients with metastatic breast cancer: a pooled analysis of individual patient data. Lancet Oncol 2014;15(4):406–14.

8 Baccelli I et al. Identification of a population of blood circulating tumor cells from breast cancer patients that initiates metastasis in a xenograft assay. Nat Biotechnol 2013;31(6):539–44.

9 Yang L et al. Optimization of an enrichment process for circulating tumor cells from the blood of head and neck cancer patients through depletion of normal cells. Biotechnol Bioeng 2009;102(2):521–34.

10 Lara O et al. Enrichment of rare cancer cells through depletion of normal cells using density and flow-through, immunomagnetic cell separation. Exp Hemetol 2004;32(10):891–904.

11 Alix-Panabieres C, Mader S, Pantel K. Epithelial-mesenchymal plasticity in circulating tumor cells. J Molec Med (Berlin, Germany) 2017;95(2):133–42.

12 Powell AA et al. Single cell profiling of circulating tumor cells: transcriptional heterogeneity and diversity from breast cancer cell lines. PLoS One 2012;7(5):e33788.

13 Giordano A et al. Epithelial-mesenchymal transition and stem cell markers in patients with HER2-positive metastatic breast cancer. Mol Cancer Ther 2012;11(11):2526–34.

14 Ramirez P et al. Oncological evaluation by positron-emission tomography, circulating tumor cells and alpha fetoprotein in patients with hepatocellular carcinoma on the waiting list for liver transplantation. Transplant Proc 2016;48(9):2962–5.

15 Bredemeier M et al. Establishment of a multimarker qPCR panel for the molecular characterization of circulating tumor cells in blood samples of metastatic breast cancer patients during the course of palliative treatment. Oncotarget 2016;7(27):41677–90.

16 Autebert J et al. High purity microfluidic sorting and analysis of circulating tumor cells: towards routine mutation detection. Lab Chip 2015;15(9):2090–101.

17 Karabacak NM et al. Microfluidic, marker-free isolation of circulating tumor cells from blood samples. Nat Protoc 2014;9(3):694–710.

18 Seal SH. Silicone flotation: a simple quantitative method for the isolation of free-floating cancer cells from the blood. Cancer 1959;12(3):590–5.

19 Desitter I et al. A new device for rapid isolation by size and characterization of rare circulating tumor cells. Anticancer Res 2011;31(2):427–41.

20 Farace F et al. A direct comparison of CellSearch and ISET for circulating tumour-cell detection in patients with metastatic carcinomas. Br J Cancer 2011;105(6):847–53.

21 Adams DL et al. The systematic study of circulating tumor cell isolation using lithographic microfilters. RSC Adv 2014;9:4334–42.

22 Adams DL et al. Cytometric characterization of circulating tumor cells captured by microfiltration and their correlation to the CellSearch((R)) CTC test. Cytometry A 2015;87(2):137–44.

23 Xu L et al. Optimization and evaluation of a novel size based circulating tumor cell isolation system. PLoS One 2015;10(9):e0138032.

24 Lemaire CA et al. Fast and label-free isolation of circulating tumor cells from blood: From a research microfluidic platform to an automated fluidic instrument, VTX-1 Liquid Biopsy System. SLAS Technol 2018;23(1):16–29.

25 Hou HW et al. Isolation and retrieval of circulating tumor cells using centrifugal forces. Sci Rep 2013;3:1259.

26 Balasubramanian P et al. Antibody-independent capture of circulating tumor cells of non-epithelial origin with the ApoStream(R) system. PLoS One 2017;12(4):e0175414.

27 Ilie M et al. “Sentinel” circulating tumor cells allow early diagnosis of lung cancer in patients with chronic obstructive pulmonary disease. PloS One 2014;9(10):e111597.

28 Vona G 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.

29 Harouaka RA et al. Flexible micro spring array device for high-throughput enrichment of viable circulating tumor cells. Clin Chem 2014;60(2):323–33.

30 Sarioglu AF et al. A microfluidic device for label-free, physical capture of circulating tumor cell clusters. Nat Methods 2015;12(7):685–91.

31 Krivacic RT et al. A rare-cell detector for cancer. Proc Natl Acad Sci U S A 2004;101(29):10501–4.

32 Somlo G et al. Multiple biomarker expression on circulating tumor cells in comparison to tumor tissues from primary and metastatic sites in patients with locally advanced/inflammatory, and stage IV breast cancer, using a novel detection technology. Breast Cancer Res Treat 2011;128(1):155–63.

33 Ntouroupi TG et al. Detection of circulating tumour cells in peripheral blood with an automated scanning fluorescence microscope. Br J Cancer 2008;99(5):789–95.

34 Pailler E et al. Method for semi-automated microscopy of filtration-enriched circulating tumor cells. BMC Cancer 2016;16:477.

35 Lustberg MB et al. Heterogeneous atypical cell populations are present in blood of metastatic breast cancer patients. Breast Cancer Res 2014;16(2):R23.

36 Zhang L et al. The identification and characterization of breast cancer CTCs competent for brain metastasis. Sci Transl Med 2013;5(180):180ra48.

37 Heitzer E et al. Complex tumor genomes inferred from single circulating tumor cells by array-CGH and next-generation sequencing. Cancer Res 2013;73(10):2965–75.

38 Abonnenc M et al. Programmable interactions of functionalized single bioparticles in a dielectrophoresis-based microarray chip. Anal Chem 2013;85(17):8219–24.

39 Fabbri F et al. Detection and recovery of circulating colon cancer cells using a dielectrophoresis-based device: KRAS mutation status in pure CTCs. Cancer Lett 2013;335(1):225–31.

40 Mesquita B et al. Molecular analysis of single circulating tumour cells following long-term storage of clinical samples. Mol Oncol 2017;11(12):1687–97.

41 Paolillo C et al. Detection of activating estrogen receptor gene (ESR1) mutations in single circulating tumor cells. Clin Cancer Res 2017;23(20):6086–93.

42 Peeters DJ et al. Semiautomated isolation and molecular characterisation of single or highly purified tumour cells from CellSearch enriched blood samples using dielectrophoretic cell sorting. Br J Cancer 2013;108(6):1358–67.

43 Esmaeilsabzali H et al. Detection and isolation of circulating tumor cells: principles and methods. Biotechnol Adv 2013;31(7):1063–84.

44 Riethdorf S, Wikman H, Pantel K. Review: Biological relevance of disseminated tumor cells in cancer patients. Int J Cancer 2008;123(9):1991–2006.

45 Fehm T et al. Detection and characterization of circulating tumor cells in blood of primary breast cancer patients by RT-PCR and comparison to status of bone marrow disseminated cells. Breast Cancer Res 2009;11(4):R59.

46 Punnoose EA et al. Molecular biomarker analyses using circulating tumor cells. PLoS One 2010;5(9):e12517.

47 Pantel K, Brakenhoff RH, Brandt B. Detection, clinical relevance and specific biological properties of disseminating tumour cells. Nat Rev Cancer 2008;8(5):329–40.

48 Chebouti I et al. ERCC1-expressing circulating tumor cells as a potential diagnostic tool for monitoring response to platinum-based chemotherapy and for predicting post-therapeutic outcome of ovarian cancer. Oncotarget 2017;8(15):24303–13.

49 Soler A et al. EpCAM-Independent enrichment and detection of viable circulating tumor cells using the EPISPOT assay. Methods Mol Biol 2017;1634:263–76.

50 Cayrefourcq L et al. Establishment and characterization of a cell line from human circulating colon cancer cells. Cancer Res 2015;75(5):892–901.

51 Deneve E et al. Capture of viable circulating tumor cells in the liver of colorectal cancer patients. Clin Chem 2013;59(9):1384–92.

52 Kuske A et al. Improved detection of circulating tumor cells in non-metastatic high-risk prostate cancer patients. Sci Rep 2016;6:39736.

53 Markou A et al. Multiplex gene expression profiling of in vivo isolated circulating tumor cells in high-risk prostate cancer patients. Clin Chem 2018;64(2):297–306.

54 Alix-Panabieres C. EPISPOT assay: detection of viable DTCs/CTCs in solid tumor patients. Recent Results Cancer Res 2012;195:69–76.

55 Tulley S et al. Vita-Assay method of enrichment and identification of circulating cancer cells/circulating tumor cells (CTCs). Methods Mol Biol 2016;1406:107–19.

56 Friedlander TW et al. Detection and characterization of invasive circulating tumor cells derived from men with metastatic castration-resistant prostate cancer. Int J Cancer 2014;134(10):2284–93.

57 Lu J et al. Isolation of circulating epithelial and tumor progenitor cells with an invasive phenotype from breast cancer patients. Int J Cancer 2010;126(3):669–83.

58 Hodgkinson CL et al. Tumorigenicity and genetic profiling of circulating tumor cells in small-cell lung cancer. Nat Med 2014;20(8):897–903.

59 Riethdorf S et al. Detection and HER2 expression of circulating tumor cells: prospective monitoring in breast cancer patients treated in the neoadjuvant GeparQuattro trial. Clin Cancer Res 2010;16(9):2634–45.

60 Jaeger BAS et al. The HER2 phenotype of circulating tumor cells in HER2-positive early breast cancer: A translational research project of a prospective randomized phase III trial. PLoS One 2017;12(6):e0173593.

61 Topalian SL, Drake CG, Pardoll DM. Immune checkpoint blockade: a common denominator approach to cancer therapy. Cancer Cell 2015;27(4):450–61.

62 Mazel M et al. Frequent expression of PD-L1 on circulating breast cancer cells. Mol Oncol 2015;9(9):1773–82.

63 Paoletti C et al. Comprehensive mutation and copy number profiling in archived circulating breast cancer tumor cells documents heterogeneous resistance mechanisms. Cancer Res 2018;78(4):1110–22.

64 Sho S et al. Precision oncology using a limited number of cells: optimization of whole genome amplification products for sequencing applications. BMC Cancer 2017;17(1):457.

65 Denis JA, Lacorte JM. Detection of RAS mutations in circulating tumor cells: applications in colorectal cancer and prospects. Ann Biol Clin (Paris) 2017;75(6):607–18.

66 Obermayr E et al. Circulating tumor cells: potential markers of minimal residual disease in ovarian cancer? a study of the OVCAD consortium. Oncotarget 2017;8(63):106415–28.

67 Podolak J et al. Androgen receptor amplification is concordant between circulating tumor cells and biopsies from men undergoing treatment for metastatic castration resistant prostate cancer. Oncotarget 2017;8(42):71447–55.