Technology in Cancer Research and Treatment ISSN 1533-0346 2014 June 30. Epub ahead of print.
A Combined Negative and Positive Enrichment Assay for Cancer Cells Isolation and Purification www.tcrt.org DOI: 10.7785/tcrt.2012.500447 Cancer cells that detach from solid tumor and circulate in the peripheral blood (CTCs) have been considered as a new “biomarker” for the detection and characterization of cancers. However, isolating and detecting cancer cells from the cancer patient peripheral blood have been technically challenging, owing to the small sub-population of CTCs (a few to hundreds per milliliter). Here we demonstrate a simple and efficient cancer cells isolation and purification method. A biocompatible and surface roughness controllable TiO2 nanofilm was deposited onto a glass slide to achieve enhanced topographic interactions with nanoscale cellular surface components, again, anti-CD45 (a leukocyte common antigen) and anti-EpCAM (epithelial cell adhesion molecule) were then coated onto the surface of the nanofilm for advance depletion of white blood cells (WBCs) and specific isolation of CTCs, respectively. Comparing to the conventional positive enrichment technology, this method exhibited excellent biocompatibility and equally high capture efficiency. Moreover, the maximum number of background cells (WBCs) was removed, and viable and functional cancer cells were isolated with high purity. Utilizing the horizontally packed TiO2 nanofilm improved pure CTC-capture through combining cell-capture-agent and cancer cell-preferred nanoscale topography, which represented a new method capable of obtaining biologically functional CTCs for subsequent molecular analysis.
Boran Cheng, M.D.‡ Shuyi Wang, M.D.‡ Yuanyuan Chen, M.D. Yuan Fang, M.D. Fangfang Chen, M.D. Zhenmeng Wang, M.D. Bin Xiong, M.D., Ph.D.* Department of Oncology, Zhongnan Hospital of Wuhan University, Hubei Key Laboratory of Tumor Biological Behaviors, Hubei Cancer Clinical Study Center, Wuhan, Hubei 430071, P. R. China These authors contributed equally
‡
to this work.
Key words: Cancer cells; Cell capture; Purity; TiO2 nanoparticle chip.
Introduction Tumor metastasis, the process by which cancer cells leave primary tumor and colonize distant sites, is the common cause of mortality for cancer patients with solid malignant tumors (1). Being observed since 1869, circulating tumor cells (CTCs) (2), which are defined as cancer cells detaching from the solid tumor and circulating within peripheral blood (3), have been considered to play a crucial role in tumor metastasis and disease progression. In the past decade, abundant evidence has been provided to support the idea that CTC counts could serve as prognostic indicators for predicting clinic outcomes and monitoring treatment responses (4-7). For example, metastatic breast cancer patients with 5 or more CTCs counts (enumerated by CellSearch-Veridex) in 7.5 mL whole blood before
Abbreviations: CTCs: Circulating Tumor Cells; EpCAM: Epithelial Cell Adhesion Molecule; WBCs: White Blood Cells; TIP: Titanium (IV) Isopropoxide; GMBS: 4-Maleimidobutyric Acid N-hydrosuccinimide; MTPMS: 3-Mercaptopropyl Trimethoxysilane; PFA: Paraformaldehyde; BSA: Bovine Serum Albumin; DAPI: 49, 6-Diamidino-2-phenylindole Dihydrochloride; SA: Streptavidin; PBS: Phosphate-buffered Saline; PDMS: Polydimethylsiloxane; TiSP: TiO2 Spheres; DMEM: Dulbecco’s Modified Eagle Medium.
*Corresponding author: Bin Xiong, M.D., Ph.D. Phone: 186 186-0711-5962 E-mail: [email protected]
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treatments are reported to suffer poorer progression-free survival and overall survival (8). More importantly, CTCs, regarded as “liquid biopsy”, provide an alternative access to solid tumor, which enables serial sampling for the study of tumor genomic variations (9) throughout the whole treatment and gives guidance to new drug discovery and personalized therapy. However, in absence of effective enrichment, the analysis of CTCs is extremely challenging as they are such a small sub-population (a few to hundreds per mL) among a large number of hematologic cells (109 cells mL–1) (8, 10) in the peripheral blood. Thus, to fully explore the clinic value of CTCs, a robust platform with sufficient sensitivity, excellent biocompatibility but low cost is required for rare CTCs detections. Previously, we have demonstrated a highly efficient CTCs capture assay using a transparent nanostructured substrates made of TiO2 nanoparticles (11). Enhanced local topographic interactions (12, 13) between cancer cells surface (e.g., microvilli) and the antibody coated nanoparticles lead to improved CTC capture efficiency. However, further applications of this platform were constrained by the contamination of the non-specifically captured white blood cells (WBCs), which is incompatible with many subsequent studies such as gene expression, gene mutation, polymorphism and genetic sequence test. It’s urgent to develop a new assay to get high purity cancer cells for molecular and functional analysis. Herein, on the basis of our previously reported nanostructured TiO2 nanoparticles substrate, we develop a new CTC detection strategy by combining negative depletion and positive enrichment, which is capable of efficiently capturing high purity and viable cancer cells in patient blood samples. A biocompatible and surface roughness controllable TiO2 nanofilm was deposited onto a glass slide to achieve enhanced topographic interactions with nanoscale cellular surface components (14, 15) (e.g., filopodia), again, anti-CD45 (16) (a leukocyte common antigen) and anti-EpCAM (17) (epithelial cell adhesion molecule) were then coated onto the surface of nanosubstrate for advance depletion of WBCs and specific isolation of CTCs, respectively. This method also exhibited high capture efficiency comparing to the traditional positive enrichment technologies based on single capture agent labeled nanosubstrate, such as our previously reported horizontally packed TiO2 nanofibers (18) and TiO2 nanoparticles substrate (11). More importantly, the maximum number of background cells (WBCs) was removed and viable and functional cancer cells were isolated with high purity, which made these cancer cells available for subsequent studies. Finally, rare CTCs captured from colon cancer patient’s peripheral blood sample using our nanostructured chip were identified by three-color immunocytochemistry. We conclude that this improved method is capable of isolating pure CTCs for subsequent molecular analysis.
Methods Preparation of TiO2 Nanoparticles Chip (19-21) The TiO2 nanoparticles chip was fabricated in polydimethylsiloxane (PDMS) using a standard soft lithography and replica molding method. The process has been reported previously (11). The width and height of micro-channel were 1 mm and 1 mm, respectively. The PDMS with microchannel was reversible bonding with glass substrate. 1.0 g synthesized TiO2 spheres (TiSP) (Sigma-Aldrich, St Louis, MO, USA) was dispersed into a mixture of 0.05 g lauric acid, 0.2 g ethyl cellulose and 10.0 mL terpinol and ethanol mixture solution (v/v 1:1) to form slurry. The TiSP slurry can be diluted with ethanol to a concentration of 5 mg mL–1. Then the capillary force can drive the TiO2 nanoparticles slurry filled the micro-channel. After baking at 708C for half hour, the PDMS was carefully peeled off from the glass substrate. The substrate with TiO2 nanoparticles on the surface substrate was annealed at 5008C for 15 min. Finally the PDMS was aligned onto the patterned TiO2 nanoparticles substrate. Surface Modification with Antibody (22) According to the previously established method (23), the TiO2 nanoparticles chip was modified with 4% (v/v) 3-mercaptopropyl trimethoxysilane (MPTMS) (Sigma-Aldrich) in ethanol at room temperature for 1 h, then, the substrate was treated with the coupling agent N-y-maleimidobutyryloxy succinimide ester (GMBS, 0.25 mM) (Sigma-Aldrich) for 1 h make for GBMS attachment to the substrate, next, the substrate was treated with 20 μg mL–1 of streptavidin (SA) (Invitrogen, Carlsbad, CA, USA) at room temperature overnight, r esulting in immobilization onto GMBS, and washed with 1X PBS to remove excess streptavidin before adding 10 μL biotinylated anti-human CD45 (Mouse IgG1, 2.5 μg mL–1 in PBS) (R&D Systems, Minneapolis, MN, USA) onto Negative depletion portion and 10 μL biotinylated anti-EpCAM (10 μg mL–1 in PBS) (R&D Systems) onto Positive enrichment portion, respectively. Cells and Blood Samples Colorectal cancer cell line (HCT116), gastric carcinoma cell line (MGC803) and cervical cancer cell line (HeLa) were harvested from Hubei Key Laboratory of Tumor Biological Behaviors. Cells were cultured in Dulbecco’s modified eagle medium (DMEM, BD Biosciences, San Jose, CA, USA) supplemented with 10% fetal bovine serum (SigmaAldrich) and 1% penicillin/streptomycin solution at 378C under a humidified 5% CO2 atmosphere. Whole blood samples from healthy donors were obtained from Department of Clinical Laboratory, Zhongnan Hospital of Wuhan University (Wuhan, Hubei, P. R. China) according to a protocol by
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the Institutional Review Board (IRB) and cancer patients’ blood samples were obtained from Department of Oncology, Zhongnan Hospital of Wuhan University (Wuhan, Hubei, P. R. China) under a separate IRB-approved protocol. All blood specimens were collected into anticoagulant tubes (EDTA K2, 2 mL, violet cap) (Wuhan Zhiyuan Medical Technology Co., Ltd., Wuhan, Hubei, China).
(DAPI, 0.33 μg mL–1 in DI water) was used for staining nuclear for 10 min. Finally, detecting and counting of targeted cells using the fluorescence microscope (IX81, Olympus, Tokyo, Japan). Captured CTCs on chip were photographed by using IPP software (Media Cybernetics Inc., Silver Spring, MD, USA).
Cell Capture and Identification
Cell Capture from the Artificial CTC Blood Samples and Patient Blood Samples
1 mL cell suspension or patient peripheral blood samples were introduced onto TiO2 nanoparticles chips at a flow rate of 8 μL min–1. Then, the chip was rinsed with PBS for 10 min. The captured cells on the chip were fixed with 4% paraformaldehyde (PFA) (Sigma-Aldrich) in PBS for 10 min. Then, the chip was loaded with 0.2% Triton X-100 (Sigma-Aldrich) in PBS and incubated for 10 min in order to improve the permeability of the cell and to allow for intracellular staining, 25 μL blocking solution was then loaded (2% donkey serum in PBS/0.2% triton X-100) onto the chip and kept for 30 min at room temperature, followed by staining with anti-cytokeratin-PE (CAM5.2, conjugated with phycoerythrin) (BD Biosciences) and anti-human CD45-FITC (Ms IgG1, clone H130) (BD Biosciences, San Jose, CA, USA) overnight. 49, 6-diamidino-2-phenylindole dihydrochloride
The cell capture efficiency of the optimal capture conditions was validated with artificial CTC samples containing HCT116 cells: 1) 1 mL DMEM media containing HCT116 cells and 2) 1 mL human blood (healthy donor) containing HCT116 cells. After rinsing, followed by staining of antiCK, anti-CD45 and DAPI, the specifically captured HCT116 cells were identified and counted on the substrates. 1.0 mL patient peripheral blood sample was introduced onto our chip according to the procedure described above. Results and Discussion The TiO2 nanoparticles chip was composed of two functional components (Figure 1A and 1B): 1) the biotinylated antiCD45 bioconjugated on TiO2 nanofilm depleted WBCs and
Figure 1: Schematic of the CTC-chip consist of two microfluidic components. (A) The biotinylated anti-CD45 bioconjugated on TiO2 nanoparticles depleted WBCs and (B) the horizontally packed TiO2 nanofilm improved CTC-capture through combining cell-capture-agent and cancer cell-preferred nanoscale topography. SEM images of (C) TiO2 nanofilm and (D) HCT116 cell being captured on TiO2 nanofilm.
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Figure 2: (A) Surface modification procedure of the TiO2 nanoparticles chip with antibody for cancer cell capture. The cell capture yields of the TiO2 nanoparticle chip with (B) different surface roughness of TiO2 nanofilms ranging from 20 to 100 nm and (C) at different flow rate (with flow rate of 8, 16, 24, 32, 40 and 48 μL min–1). (D) Capture yields at different spiking cell numbers ranging from 50 to 1000 mL–1. (E) Capture yields of negative depletion and positive enrichment methods using suspension of cervical (HeLa), colon (HCT116), gastric (MGC803) cell lines and WBC. Error bars show standard deviations (n 5 3).
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A Combined Assay for Cancer Cells Isolation and Purification 2) horizontally packed TiO2 nanofilm improved CTC-capture through combining cell-capture-agent (anti-EpCAM) and cancer cell-preferred nanoscale topography (Figure 1C). The captured cancer cells on the nanofilm substrate appeared as many extended pseudopodia (Figure 1D). This finding provided further evidence supporting the working mechanism of our nanostructure-based cell-capture approach. These two components were fabricated according to the previously established procedures (details in Methods). The roughness of the nanofilm substrates were controlled by changing the temperature and reaction time. As illustrated in Figure 2A, the biotin-TiO2 nanoparticles chips were prepared for cell capture. To assess how the surface roughness of TiO2 nanoparticles chip impacts the capture yields, we prepared an EpCAM-positive cancer cell line (HCT116) in Dulbecco’s modified Eagle medium (DMEM) at a concentration of 105 mL–1 and captured the spiked cells with anti-EpCAM-grafted substrates prepared with different surface roughness (ranging from 20 to 100 nm). As shown in Figure 2B, when the surface roughness of the substrates were around 85 nm, maximum cell capture yields were achieved for HCT116 cell line (approximately 76%). Therefore this optimal substrate was selected for subsequent studies. The effect of roughness on the cell capture yields can be attributed to the local topographic interactions between nanosubstrate and cancer cells. The capture performance of TiO2 nanoparticles chip was assessed with a 1.0 mL suspension of HCT116 cells (1000 cells mL–1 in DMED medium) at flow rate of 8, 16, 24, 32, 40 and 48 μL min–1. As observed in Figure 2C, the highest cellcapture efficiency (76 6 2%) was accomplished at flow rate of 8 μL min–1, the recovery efficiency of cell capture decreased significantly with increasing flow rate (up to 24 μL min–1), we ascertain 8 μL min–1 to be the maximal flow rate for subsequent studies to decrease shearing force in order to preserve potentially fragile CTCs. We then validated the cell capture efficiency of the anti-CD45 and anti-EpCAM coated TiO2 nanoparticles chip by using a series of artificial blood samples prepared by spiking HCT116 cells into DMEM and healthy donor’s blood at concentrations of approximately 50, 100, 250, 500, 1000 cells mL–1. The results in Figure 2D indicated that the recovered cell numbers ranging from a few to hundreds CTCs mL–1. Spiked HCT116 cells were captured from DMEM and healthy donor’s blood with a yield of 76.1% and 63.6%, respectively. Finally, to illustrate the cell capture specificity and optimized cell capture performance of TiO2 nanoparticles chip, we used two EpCAM-positive cancer cell lines-HCT116 (colorectal cancer cell line) and MGC803 (gastric carcinoma cell line) for specific cell capture, a non-specific cells line-HeLa (cervical cancer cell line) as an adherent epithelial cell control, and CD45-positive cell-WBCs as suspension lymphocyte control were used for
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a specific cell capture contrast experiment. For the EpCAMpositive cell and CD45-positive cell, the maximal cell capture yields were achieved, whereas only relatively low yields of non-specific cells were captured. Summarized results in Figure 2E supported the idea that the CD45 and EpCAMcoated substrates were capable of depleting WBCs, capturing HCT116, MGC803 cells, but not HeLa cells (optimal capture condition of TiO2 nanofilm substrate with 8 μL min–1). During the capability test of purity cancer cells capture using TiO2 nanoparticle chips, we performed cell capture efficiency and purity studies using artificial blood samples which were prepared by spiking healthy donor’s blood (containing ca. 4-10 3 106 WBCs mL–1) with HCT116 cells at a concentration of approximately 1000 cells mL–1. 1 mL artificial blood samples were introduced onto TiO2 nanoparticles chips at a flow rate of 8 μL min–1. In parallel, positive enrichment without negative depletion method was also examined as a control. After rinsing, fixation, permeabilization, blocking, followed by staining with anti-CK, anti-CD45 and DAPI, the specifically captured HCT116 cells and non-specifically immobilized WBCs were identified on the Chips. The fluorescence stain results in Figure 3A showed that the Positive enrichment without Negative depletion method captured many more WBCs than Negative depletion with Positive enrichment method. Cell capture assay results in Figure 3B and 3C showed that Negative depletion with Positive enrichment method exhibited equally high capture yields but higher purity comparing to Positive enrichment w ithout Negative depletion. 626 6 49 HCT116 and 2000-5000 WBCs were identified on the Negative depletion with Positive enrichment platform, and 644 6 32 HCT116 and 5000-10000 WBCs were identified on the Positive enrichment without Negative depletion platform. Our results in Figure 3 showed that there was significant difference in levels of preference between these two methods, which suggested that higher purity cancer cells in cancer patients’ peripheral blood could potentially be captured by our platform for subsequent studies. Further, Cell viability was tested by FDA-PI assay at two stages: before capture and after capture. The results indicated that the cell viability reaches 84% (after capture), making these cells suitable for subsequent studies (Figure 3D-3F). Because the captured cancer cells are alive, they could be captured from cancer patients and potentially be physically isolated for proteomic and genomic analyses. In order to validate the clinical utility of our platform for rare cancer cells capture from cancer patient peripheral blood samples, we applied the optimized cell capture conditions to study colon cancer patient peripheral blood samples (preserved in EDTA-K2 tubes). 1.0 mL peripheral blood sample was introduced onto our chip, after negative depletion and positive enrichment, captured cells then rinsed with PBS, fixed with 4% PFA and then permeabilized with
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Figure 3: Capture yields and purity of cancer cells with different methods (Negative depletion and Positive enrichment, Positive enrichment without Negative depletion). (A) The fluorescent micrographs of cancer cells captured from the artificial blood samples. Three-color immunocytochemistry method based on FITC-labeled anti-CD45, PE-labeled anti-CK, and DAPI nuclear staining was applied to identify and enumerate CTCs from non-specifically trapped WBCs. Scale bars are 50 μm. (B) Capture cell number. (C) Capture yields and purity against two different cell-capture methods. (D) Plotted the viability of HCT116 cancer cells at different stages. Error bars show standard deviations (n 5 3). The fluorescent micrographs of FDA-PI stain HCT-116 cancer cells before capture (E), after capture (F). Viable (green) and non-viable (red) cancer cells were distinguishable.
Figure 4: Identification of CTCs based on a 3-color immunocytochemistry staining using PE-conjugated anti-CK, FITC-conjugated anti-CD45 (a marker for WBC) and DAPI (nuclear specific) was validated on a colon cancer patient.
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A Combined Assay for Cancer Cells Isolation and Purification 0.02% Triton-X100. After that, captured cells were stained by PE-labeled anti-CK, FITC-labeled anti-CD45 and DAPI (14, 24). Specifically captured CTCs (CK1/CD45-/DAPI1, 10 μm cell sizes 30 μm) were identified from WBCs (CK-/CD451/DAPI1, sizes 15 μm) using fluorescence microscopy (Figure 4). In general, these data demonstrated that our platform for follow-up studies (molecular analysis of purity CTCs captured from patients) was possible, but at the same time, there was not enough CTCs number in further analysis from this colon cancer patient. Conclusion In conclusion, CTCs isolation based on negative depletion and positive enrichment allowed the detection of cancer cells with high purity, exhibited similar capture yield comparing to traditional method. Regarding the strong desire of obtaining pure CTCs from patients for study in genetic analysis, we expect the improved method can be used for the potential clinical research in future (treatment responses monitor and patient prognosis indication). Conflict of Interest We certify that this manuscript has not been published in whole or in part nor it being considered for publication e lsewhere. The authors have no conflicts of interest to declare. Acknowledgements The work was supported by National High Technology Research and Development Program of China (Grant No. 2012AA02A502, 2012AA02A506). References 1. Steeg, P. S., Bevilacqua, G., Kopper, L., Thorgeirsson, U. P., Talmadge, J. E., Liotta, L. A., Sobel, M. E. Evidence for a novel gene associated with low tumor metastatic potential. J Natl Cancer Inst 80, 200-204 (1988). DOI: 10.1093/jnci/80.3.200 2. Ashworth, T. A case of cancer in which cells similar to those in the tumours were seen in the blood after death. Aust Med J 14, 146-149 (1869). 3. Steeg, P. S. Tumor metastasis: mechanistic insights and clinical challenges. Nat Med 12, 895-904 (2006). DOI: 10.1038/nm1469 4. Schuster, R., Bechrakis, N. E., Stroux, A., Busse, A., Schmittel, A., Scheibenbogen, C., Thiel, E., Foerster, M. H., Keilholz, U. Circulating tumor cells as prognostic factor for distant metastases and survival in patients with primary uveal melanoma. Clinical Cancer Research 13, 1171-1178 (2007). DOI: 10.1158/1078-0432.CCR-06-2329 5. Taback, B., Chan, A. D., Kuo, C. T., Bostick, P. J., Wang, H. J., Giuliano, A. E., Hoon, D. S. Detection of occult metastatic breast cancer cells in blood by a multimolecular marker assay: correlation with clinical stage of disease. Cancer Res 61, 8845-8850 (2001). 6. Keilholz, U., Goldin-Lang, P., Bechrakis, N. E., Max, N., Letsch, A., Schmittel, A., Scheibenbogen, C., Heufelder, K., Eggermont, A.,
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23. He, R., Zhao, L., Liu, Y., Zhang, N., Cheng, B., He, Z., Cai, B., Li, S., Liu, W., Guo, S. Biocompatible TiO2 nanoparticle-based cell immunoassay for circulating tumor cells capture and identification from cancer patients. Biomedical Microdevices, 1-10. DOI: 10.1007/ s10544-013-9781-9 24. Zhang, P., Chen, L., Xu, T., Liu, H., Liu, X., Meng, J., Yang, G., Jiang, L., Wang, S. Programmable fractal nanostructured interfaces for specific recognition and electrochemical release of cancer cells. Advanced Materials 25, 7603-7609 (2013). DOI: 10.1002/ adma.201300888 Received: November 10, 2013; Revised: April 8, 2014; Accepted: April 21, 2014
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A Combined Negative and Positive Enrichment Assay for Cancer Cells Isolation and Purification www.tcrt.org DOI: 10.7785/tcrt.2012.500447 Cancer cells that detach from solid tumor and circulate in the peripheral blood (CTCs) have been considered as a new “biomarker” for the detection and characterization of cancers. However, isolating and detecting cancer cells from the cancer patient peripheral blood have been technically challenging, owing to the small sub-population of CTCs (a few to hundreds per milliliter). Here we demonstrate a simple and efficient cancer cells isolation and purification method. A biocompatible and surface roughness controllable TiO2 nanofilm was deposited onto a glass slide to achieve enhanced topographic interactions with nanoscale cellular surface components, again, anti-CD45 (a leukocyte common antigen) and anti-EpCAM (epithelial cell adhesion molecule) were then coated onto the surface of the nanofilm for advance depletion of white blood cells (WBCs) and specific isolation of CTCs, respectively. Comparing to the conventional positive enrichment technology, this method exhibited excellent biocompatibility and equally high capture efficiency. Moreover, the maximum number of background cells (WBCs) was removed, and viable and functional cancer cells were isolated with high purity. Utilizing the horizontally packed TiO2 nanofilm improved pure CTC-capture through combining cell-capture-agent and cancer cell-preferred nanoscale topography, which represented a new method capable of obtaining biologically functional CTCs for subsequent molecular analysis.
Boran Cheng, M.D.‡ Shuyi Wang, M.D.‡ Yuanyuan Chen, M.D. Yuan Fang, M.D. Fangfang Chen, M.D. Zhenmeng Wang, M.D. Bin Xiong, M.D., Ph.D.* Department of Oncology, Zhongnan Hospital of Wuhan University, Hubei Key Laboratory of Tumor Biological Behaviors, Hubei Cancer Clinical Study Center, Wuhan, Hubei 430071, P. R. China These authors contributed equally
‡
to this work.
Key words: Cancer cells; Cell capture; Purity; TiO2 nanoparticle chip.
Introduction Tumor metastasis, the process by which cancer cells leave primary tumor and colonize distant sites, is the common cause of mortality for cancer patients with solid malignant tumors (1). Being observed since 1869, circulating tumor cells (CTCs) (2), which are defined as cancer cells detaching from the solid tumor and circulating within peripheral blood (3), have been considered to play a crucial role in tumor metastasis and disease progression. In the past decade, abundant evidence has been provided to support the idea that CTC counts could serve as prognostic indicators for predicting clinic outcomes and monitoring treatment responses (4-7). For example, metastatic breast cancer patients with 5 or more CTCs counts (enumerated by CellSearch-Veridex) in 7.5 mL whole blood before
Abbreviations: CTCs: Circulating Tumor Cells; EpCAM: Epithelial Cell Adhesion Molecule; WBCs: White Blood Cells; TIP: Titanium (IV) Isopropoxide; GMBS: 4-Maleimidobutyric Acid N-hydrosuccinimide; MTPMS: 3-Mercaptopropyl Trimethoxysilane; PFA: Paraformaldehyde; BSA: Bovine Serum Albumin; DAPI: 49, 6-Diamidino-2-phenylindole Dihydrochloride; SA: Streptavidin; PBS: Phosphate-buffered Saline; PDMS: Polydimethylsiloxane; TiSP: TiO2 Spheres; DMEM: Dulbecco’s Modified Eagle Medium.
*Corresponding author: Bin Xiong, M.D., Ph.D. Phone: 186 186-0711-5962 E-mail: [email protected]
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treatments are reported to suffer poorer progression-free survival and overall survival (8). More importantly, CTCs, regarded as “liquid biopsy”, provide an alternative access to solid tumor, which enables serial sampling for the study of tumor genomic variations (9) throughout the whole treatment and gives guidance to new drug discovery and personalized therapy. However, in absence of effective enrichment, the analysis of CTCs is extremely challenging as they are such a small sub-population (a few to hundreds per mL) among a large number of hematologic cells (109 cells mL–1) (8, 10) in the peripheral blood. Thus, to fully explore the clinic value of CTCs, a robust platform with sufficient sensitivity, excellent biocompatibility but low cost is required for rare CTCs detections. Previously, we have demonstrated a highly efficient CTCs capture assay using a transparent nanostructured substrates made of TiO2 nanoparticles (11). Enhanced local topographic interactions (12, 13) between cancer cells surface (e.g., microvilli) and the antibody coated nanoparticles lead to improved CTC capture efficiency. However, further applications of this platform were constrained by the contamination of the non-specifically captured white blood cells (WBCs), which is incompatible with many subsequent studies such as gene expression, gene mutation, polymorphism and genetic sequence test. It’s urgent to develop a new assay to get high purity cancer cells for molecular and functional analysis. Herein, on the basis of our previously reported nanostructured TiO2 nanoparticles substrate, we develop a new CTC detection strategy by combining negative depletion and positive enrichment, which is capable of efficiently capturing high purity and viable cancer cells in patient blood samples. A biocompatible and surface roughness controllable TiO2 nanofilm was deposited onto a glass slide to achieve enhanced topographic interactions with nanoscale cellular surface components (14, 15) (e.g., filopodia), again, anti-CD45 (16) (a leukocyte common antigen) and anti-EpCAM (17) (epithelial cell adhesion molecule) were then coated onto the surface of nanosubstrate for advance depletion of WBCs and specific isolation of CTCs, respectively. This method also exhibited high capture efficiency comparing to the traditional positive enrichment technologies based on single capture agent labeled nanosubstrate, such as our previously reported horizontally packed TiO2 nanofibers (18) and TiO2 nanoparticles substrate (11). More importantly, the maximum number of background cells (WBCs) was removed and viable and functional cancer cells were isolated with high purity, which made these cancer cells available for subsequent studies. Finally, rare CTCs captured from colon cancer patient’s peripheral blood sample using our nanostructured chip were identified by three-color immunocytochemistry. We conclude that this improved method is capable of isolating pure CTCs for subsequent molecular analysis.
Methods Preparation of TiO2 Nanoparticles Chip (19-21) The TiO2 nanoparticles chip was fabricated in polydimethylsiloxane (PDMS) using a standard soft lithography and replica molding method. The process has been reported previously (11). The width and height of micro-channel were 1 mm and 1 mm, respectively. The PDMS with microchannel was reversible bonding with glass substrate. 1.0 g synthesized TiO2 spheres (TiSP) (Sigma-Aldrich, St Louis, MO, USA) was dispersed into a mixture of 0.05 g lauric acid, 0.2 g ethyl cellulose and 10.0 mL terpinol and ethanol mixture solution (v/v 1:1) to form slurry. The TiSP slurry can be diluted with ethanol to a concentration of 5 mg mL–1. Then the capillary force can drive the TiO2 nanoparticles slurry filled the micro-channel. After baking at 708C for half hour, the PDMS was carefully peeled off from the glass substrate. The substrate with TiO2 nanoparticles on the surface substrate was annealed at 5008C for 15 min. Finally the PDMS was aligned onto the patterned TiO2 nanoparticles substrate. Surface Modification with Antibody (22) According to the previously established method (23), the TiO2 nanoparticles chip was modified with 4% (v/v) 3-mercaptopropyl trimethoxysilane (MPTMS) (Sigma-Aldrich) in ethanol at room temperature for 1 h, then, the substrate was treated with the coupling agent N-y-maleimidobutyryloxy succinimide ester (GMBS, 0.25 mM) (Sigma-Aldrich) for 1 h make for GBMS attachment to the substrate, next, the substrate was treated with 20 μg mL–1 of streptavidin (SA) (Invitrogen, Carlsbad, CA, USA) at room temperature overnight, r esulting in immobilization onto GMBS, and washed with 1X PBS to remove excess streptavidin before adding 10 μL biotinylated anti-human CD45 (Mouse IgG1, 2.5 μg mL–1 in PBS) (R&D Systems, Minneapolis, MN, USA) onto Negative depletion portion and 10 μL biotinylated anti-EpCAM (10 μg mL–1 in PBS) (R&D Systems) onto Positive enrichment portion, respectively. Cells and Blood Samples Colorectal cancer cell line (HCT116), gastric carcinoma cell line (MGC803) and cervical cancer cell line (HeLa) were harvested from Hubei Key Laboratory of Tumor Biological Behaviors. Cells were cultured in Dulbecco’s modified eagle medium (DMEM, BD Biosciences, San Jose, CA, USA) supplemented with 10% fetal bovine serum (SigmaAldrich) and 1% penicillin/streptomycin solution at 378C under a humidified 5% CO2 atmosphere. Whole blood samples from healthy donors were obtained from Department of Clinical Laboratory, Zhongnan Hospital of Wuhan University (Wuhan, Hubei, P. R. China) according to a protocol by
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the Institutional Review Board (IRB) and cancer patients’ blood samples were obtained from Department of Oncology, Zhongnan Hospital of Wuhan University (Wuhan, Hubei, P. R. China) under a separate IRB-approved protocol. All blood specimens were collected into anticoagulant tubes (EDTA K2, 2 mL, violet cap) (Wuhan Zhiyuan Medical Technology Co., Ltd., Wuhan, Hubei, China).
(DAPI, 0.33 μg mL–1 in DI water) was used for staining nuclear for 10 min. Finally, detecting and counting of targeted cells using the fluorescence microscope (IX81, Olympus, Tokyo, Japan). Captured CTCs on chip were photographed by using IPP software (Media Cybernetics Inc., Silver Spring, MD, USA).
Cell Capture and Identification
Cell Capture from the Artificial CTC Blood Samples and Patient Blood Samples
1 mL cell suspension or patient peripheral blood samples were introduced onto TiO2 nanoparticles chips at a flow rate of 8 μL min–1. Then, the chip was rinsed with PBS for 10 min. The captured cells on the chip were fixed with 4% paraformaldehyde (PFA) (Sigma-Aldrich) in PBS for 10 min. Then, the chip was loaded with 0.2% Triton X-100 (Sigma-Aldrich) in PBS and incubated for 10 min in order to improve the permeability of the cell and to allow for intracellular staining, 25 μL blocking solution was then loaded (2% donkey serum in PBS/0.2% triton X-100) onto the chip and kept for 30 min at room temperature, followed by staining with anti-cytokeratin-PE (CAM5.2, conjugated with phycoerythrin) (BD Biosciences) and anti-human CD45-FITC (Ms IgG1, clone H130) (BD Biosciences, San Jose, CA, USA) overnight. 49, 6-diamidino-2-phenylindole dihydrochloride
The cell capture efficiency of the optimal capture conditions was validated with artificial CTC samples containing HCT116 cells: 1) 1 mL DMEM media containing HCT116 cells and 2) 1 mL human blood (healthy donor) containing HCT116 cells. After rinsing, followed by staining of antiCK, anti-CD45 and DAPI, the specifically captured HCT116 cells were identified and counted on the substrates. 1.0 mL patient peripheral blood sample was introduced onto our chip according to the procedure described above. Results and Discussion The TiO2 nanoparticles chip was composed of two functional components (Figure 1A and 1B): 1) the biotinylated antiCD45 bioconjugated on TiO2 nanofilm depleted WBCs and
Figure 1: Schematic of the CTC-chip consist of two microfluidic components. (A) The biotinylated anti-CD45 bioconjugated on TiO2 nanoparticles depleted WBCs and (B) the horizontally packed TiO2 nanofilm improved CTC-capture through combining cell-capture-agent and cancer cell-preferred nanoscale topography. SEM images of (C) TiO2 nanofilm and (D) HCT116 cell being captured on TiO2 nanofilm.
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Figure 2: (A) Surface modification procedure of the TiO2 nanoparticles chip with antibody for cancer cell capture. The cell capture yields of the TiO2 nanoparticle chip with (B) different surface roughness of TiO2 nanofilms ranging from 20 to 100 nm and (C) at different flow rate (with flow rate of 8, 16, 24, 32, 40 and 48 μL min–1). (D) Capture yields at different spiking cell numbers ranging from 50 to 1000 mL–1. (E) Capture yields of negative depletion and positive enrichment methods using suspension of cervical (HeLa), colon (HCT116), gastric (MGC803) cell lines and WBC. Error bars show standard deviations (n 5 3).
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A Combined Assay for Cancer Cells Isolation and Purification 2) horizontally packed TiO2 nanofilm improved CTC-capture through combining cell-capture-agent (anti-EpCAM) and cancer cell-preferred nanoscale topography (Figure 1C). The captured cancer cells on the nanofilm substrate appeared as many extended pseudopodia (Figure 1D). This finding provided further evidence supporting the working mechanism of our nanostructure-based cell-capture approach. These two components were fabricated according to the previously established procedures (details in Methods). The roughness of the nanofilm substrates were controlled by changing the temperature and reaction time. As illustrated in Figure 2A, the biotin-TiO2 nanoparticles chips were prepared for cell capture. To assess how the surface roughness of TiO2 nanoparticles chip impacts the capture yields, we prepared an EpCAM-positive cancer cell line (HCT116) in Dulbecco’s modified Eagle medium (DMEM) at a concentration of 105 mL–1 and captured the spiked cells with anti-EpCAM-grafted substrates prepared with different surface roughness (ranging from 20 to 100 nm). As shown in Figure 2B, when the surface roughness of the substrates were around 85 nm, maximum cell capture yields were achieved for HCT116 cell line (approximately 76%). Therefore this optimal substrate was selected for subsequent studies. The effect of roughness on the cell capture yields can be attributed to the local topographic interactions between nanosubstrate and cancer cells. The capture performance of TiO2 nanoparticles chip was assessed with a 1.0 mL suspension of HCT116 cells (1000 cells mL–1 in DMED medium) at flow rate of 8, 16, 24, 32, 40 and 48 μL min–1. As observed in Figure 2C, the highest cellcapture efficiency (76 6 2%) was accomplished at flow rate of 8 μL min–1, the recovery efficiency of cell capture decreased significantly with increasing flow rate (up to 24 μL min–1), we ascertain 8 μL min–1 to be the maximal flow rate for subsequent studies to decrease shearing force in order to preserve potentially fragile CTCs. We then validated the cell capture efficiency of the anti-CD45 and anti-EpCAM coated TiO2 nanoparticles chip by using a series of artificial blood samples prepared by spiking HCT116 cells into DMEM and healthy donor’s blood at concentrations of approximately 50, 100, 250, 500, 1000 cells mL–1. The results in Figure 2D indicated that the recovered cell numbers ranging from a few to hundreds CTCs mL–1. Spiked HCT116 cells were captured from DMEM and healthy donor’s blood with a yield of 76.1% and 63.6%, respectively. Finally, to illustrate the cell capture specificity and optimized cell capture performance of TiO2 nanoparticles chip, we used two EpCAM-positive cancer cell lines-HCT116 (colorectal cancer cell line) and MGC803 (gastric carcinoma cell line) for specific cell capture, a non-specific cells line-HeLa (cervical cancer cell line) as an adherent epithelial cell control, and CD45-positive cell-WBCs as suspension lymphocyte control were used for
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a specific cell capture contrast experiment. For the EpCAMpositive cell and CD45-positive cell, the maximal cell capture yields were achieved, whereas only relatively low yields of non-specific cells were captured. Summarized results in Figure 2E supported the idea that the CD45 and EpCAMcoated substrates were capable of depleting WBCs, capturing HCT116, MGC803 cells, but not HeLa cells (optimal capture condition of TiO2 nanofilm substrate with 8 μL min–1). During the capability test of purity cancer cells capture using TiO2 nanoparticle chips, we performed cell capture efficiency and purity studies using artificial blood samples which were prepared by spiking healthy donor’s blood (containing ca. 4-10 3 106 WBCs mL–1) with HCT116 cells at a concentration of approximately 1000 cells mL–1. 1 mL artificial blood samples were introduced onto TiO2 nanoparticles chips at a flow rate of 8 μL min–1. In parallel, positive enrichment without negative depletion method was also examined as a control. After rinsing, fixation, permeabilization, blocking, followed by staining with anti-CK, anti-CD45 and DAPI, the specifically captured HCT116 cells and non-specifically immobilized WBCs were identified on the Chips. The fluorescence stain results in Figure 3A showed that the Positive enrichment without Negative depletion method captured many more WBCs than Negative depletion with Positive enrichment method. Cell capture assay results in Figure 3B and 3C showed that Negative depletion with Positive enrichment method exhibited equally high capture yields but higher purity comparing to Positive enrichment w ithout Negative depletion. 626 6 49 HCT116 and 2000-5000 WBCs were identified on the Negative depletion with Positive enrichment platform, and 644 6 32 HCT116 and 5000-10000 WBCs were identified on the Positive enrichment without Negative depletion platform. Our results in Figure 3 showed that there was significant difference in levels of preference between these two methods, which suggested that higher purity cancer cells in cancer patients’ peripheral blood could potentially be captured by our platform for subsequent studies. Further, Cell viability was tested by FDA-PI assay at two stages: before capture and after capture. The results indicated that the cell viability reaches 84% (after capture), making these cells suitable for subsequent studies (Figure 3D-3F). Because the captured cancer cells are alive, they could be captured from cancer patients and potentially be physically isolated for proteomic and genomic analyses. In order to validate the clinical utility of our platform for rare cancer cells capture from cancer patient peripheral blood samples, we applied the optimized cell capture conditions to study colon cancer patient peripheral blood samples (preserved in EDTA-K2 tubes). 1.0 mL peripheral blood sample was introduced onto our chip, after negative depletion and positive enrichment, captured cells then rinsed with PBS, fixed with 4% PFA and then permeabilized with
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Figure 3: Capture yields and purity of cancer cells with different methods (Negative depletion and Positive enrichment, Positive enrichment without Negative depletion). (A) The fluorescent micrographs of cancer cells captured from the artificial blood samples. Three-color immunocytochemistry method based on FITC-labeled anti-CD45, PE-labeled anti-CK, and DAPI nuclear staining was applied to identify and enumerate CTCs from non-specifically trapped WBCs. Scale bars are 50 μm. (B) Capture cell number. (C) Capture yields and purity against two different cell-capture methods. (D) Plotted the viability of HCT116 cancer cells at different stages. Error bars show standard deviations (n 5 3). The fluorescent micrographs of FDA-PI stain HCT-116 cancer cells before capture (E), after capture (F). Viable (green) and non-viable (red) cancer cells were distinguishable.
Figure 4: Identification of CTCs based on a 3-color immunocytochemistry staining using PE-conjugated anti-CK, FITC-conjugated anti-CD45 (a marker for WBC) and DAPI (nuclear specific) was validated on a colon cancer patient.
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A Combined Assay for Cancer Cells Isolation and Purification 0.02% Triton-X100. After that, captured cells were stained by PE-labeled anti-CK, FITC-labeled anti-CD45 and DAPI (14, 24). Specifically captured CTCs (CK1/CD45-/DAPI1, 10 μm cell sizes 30 μm) were identified from WBCs (CK-/CD451/DAPI1, sizes 15 μm) using fluorescence microscopy (Figure 4). In general, these data demonstrated that our platform for follow-up studies (molecular analysis of purity CTCs captured from patients) was possible, but at the same time, there was not enough CTCs number in further analysis from this colon cancer patient. Conclusion In conclusion, CTCs isolation based on negative depletion and positive enrichment allowed the detection of cancer cells with high purity, exhibited similar capture yield comparing to traditional method. Regarding the strong desire of obtaining pure CTCs from patients for study in genetic analysis, we expect the improved method can be used for the potential clinical research in future (treatment responses monitor and patient prognosis indication). Conflict of Interest We certify that this manuscript has not been published in whole or in part nor it being considered for publication e lsewhere. The authors have no conflicts of interest to declare. Acknowledgements The work was supported by National High Technology Research and Development Program of China (Grant No. 2012AA02A502, 2012AA02A506). References 1. Steeg, P. S., Bevilacqua, G., Kopper, L., Thorgeirsson, U. P., Talmadge, J. E., Liotta, L. A., Sobel, M. E. Evidence for a novel gene associated with low tumor metastatic potential. J Natl Cancer Inst 80, 200-204 (1988). DOI: 10.1093/jnci/80.3.200 2. Ashworth, T. A case of cancer in which cells similar to those in the tumours were seen in the blood after death. Aust Med J 14, 146-149 (1869). 3. Steeg, P. S. Tumor metastasis: mechanistic insights and clinical challenges. Nat Med 12, 895-904 (2006). DOI: 10.1038/nm1469 4. Schuster, R., Bechrakis, N. E., Stroux, A., Busse, A., Schmittel, A., Scheibenbogen, C., Thiel, E., Foerster, M. H., Keilholz, U. Circulating tumor cells as prognostic factor for distant metastases and survival in patients with primary uveal melanoma. Clinical Cancer Research 13, 1171-1178 (2007). DOI: 10.1158/1078-0432.CCR-06-2329 5. Taback, B., Chan, A. D., Kuo, C. T., Bostick, P. J., Wang, H. J., Giuliano, A. E., Hoon, D. S. Detection of occult metastatic breast cancer cells in blood by a multimolecular marker assay: correlation with clinical stage of disease. Cancer Res 61, 8845-8850 (2001). 6. Keilholz, U., Goldin-Lang, P., Bechrakis, N. E., Max, N., Letsch, A., Schmittel, A., Scheibenbogen, C., Heufelder, K., Eggermont, A.,
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23. He, R., Zhao, L., Liu, Y., Zhang, N., Cheng, B., He, Z., Cai, B., Li, S., Liu, W., Guo, S. Biocompatible TiO2 nanoparticle-based cell immunoassay for circulating tumor cells capture and identification from cancer patients. Biomedical Microdevices, 1-10. DOI: 10.1007/ s10544-013-9781-9 24. Zhang, P., Chen, L., Xu, T., Liu, H., Liu, X., Meng, J., Yang, G., Jiang, L., Wang, S. Programmable fractal nanostructured interfaces for specific recognition and electrochemical release of cancer cells. Advanced Materials 25, 7603-7609 (2013). DOI: 10.1002/ adma.201300888 Received: November 10, 2013; Revised: April 8, 2014; Accepted: April 21, 2014
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