Self-Sterilizing and Regeneratable Microchip for the Precise Capture

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Self-Sterilizing and Regeneratable Microchip for the Precise Capture and Recovery of Viable Circulating Tumor Cells from Patients with Cancer Lanlan Hui, Yi Su, Tingting Ye, Zhao Liu, Qingchang Tian, Chuanjiang He, Yueqi Zhao, Pu Chen, Xiaojia Wang, Weidong Han, Yan Luo, and Ben Wang ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b15406 • Publication Date (Web): 13 Dec 2017 Downloaded from http://pubs.acs.org on December 15, 2017

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ACS Applied Materials & Interfaces

Self-Sterilizing and Regeneratable Microchip for the Precise Capture and Recovery of Viable Circulating Tumor Cells from Patients with Cancer Lanlan Hui,†§ Yi Su,†§ Tingting Ye,†§ Zhao Liu,# Qingchang Tian,†§ Chuanjiang He,†§ Yueqi Zhao,‡ Pu Chen,¶ Xiaojia Wang,& Weidong Han,$ Yan Luo♯ & Ben Wang†§* †

Cancer Institute (Key Laboratory of Cancer Prevention and Intervention, National Ministry

of Education & Key Laboratory of Molecular Biology in Medical Sciences, Zhejiang Province), The Second Affiliated Hospital, School of Medicine, Zhejiang University, Hangzhou, 310009 China §

Institute of Translational Medicine, School of Medicine, Zhejiang University, Hangzhou,

310029 China #

College of Computer Science and Technology, Zhejiang University, Hangzhou, 310027,

China ‡

Department of Chemistry, Zhejiang University, Hangzhou, 310027 China

¶

&

$

School of Medicine, Wuhan University, Wuhan, 430071 China

Department of Medical Oncology, Zhejiang Cancer Hospital, Hangzhou, 310022 China

Department of Medical Oncology, Sir Run Run Shaw Hospital, School of Medicine,

Zhejiang University, Hangzhou, 310016 China ♯

College of Basic Medical Sciences, School of Medicine, Zhejiang University, Hangzhou,

310058 China 1 ACS Paragon Plus Environment

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KEYWORDS: circulating tumor cells (CTCs), polydimethylsiloxane (PDMS), zinc oxide, antibacterial, recyclable, epithelial to mesenchymal transition (EMT).

ABSTRACT: Cancer cells metastasize and are transported in the bloodstream, easily reaching any site in the body through the blood circulation. A method designed to assess the number of circulating tumor cells (CTCs) should be validated as a clinical tool for predicting the response to therapy and monitoring disease progression in patients with cancer. Although CTCs are detectable in many cases, they remain unavailable for clinic usage due to their high testing cost, tedious operation and poor clinical relevance. Herein, we developed a regeneratable microchip for isolating CTCs that is available for robust cell heterogeneity assays on-site without the need for a sterile environment. The ivy-like hierarchical roughened zinc oxide (ZnO) nanograss interface was synthesized and directly integrated into the microfluidic devices and enables effective CTC capture and flexible, nontoxic CTC release during incubation in a mildly acidic solution, thus enabling cellular and molecular analyses. The microchip can be regenerated and recycled to capture CTCs with the remaining ZnO without affecting the efficiency, even after countless cycles of cell release. Moreover, microbial infection is avoided during its storage, distribution and even in open-space usage, which ideally appeals to the demands of point-of-care (POC) and home testing and meets to the requirements for blood examinations in undeveloped or resource-limited settings. Furthermore, the findings generated using this platform based on the cocktail of anti-epithelial cell adhesion molecule (EpCAM) and anti-vimentin antibodies indicate that CTC capture was more precise and reasonable for patients with advanced cancer.

1. INTRODUCTION Approximately 90% of patients with cancer die from metastatic tumors.1,2 Subgroups of cancer cells, known as circulating tumor cells (CTCs), are shed from the primary tumor and 2 ACS Paragon Plus Environment

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transported through the bloodstream to distant sites in the body, establishing lesions in the lung, brain, liver, or bone due to blood circulation and tissue tropism and ultimately leading to the death of the patient by damaging the CTC-metastasized organs.3,4 In primary, nonmetastatic cancers, CTCs are thought to be released from localized tumors, but in advanced cancers, CTCs might also emanate from metastatic lesions.5,6 The assessment of the CTC number should be validated as a clinical tool for predicting the response to therapy and monitoring disease progression in patients with cancer.7,8 Additionally, enriched CTCs can be employed for cellular and molecular studies, which might provide vital clues for improving our understanding of the mechanisms underlying metastatic dissemination of cancer and facilitating the discovery of novel prognostic biomarkers or demonstrate therapeutic targets.9,10 Several excellent pioneer studies have examined cancer cell capture and adhesion11–16. However, a significant challenge to the clinical application of CTCs is the extraordinary rarity and low purity (10-9) of CTCs in the blood due to the vast number of blood cells present in the circulation and tumor cell heterogeneity, which have resulted in their increased technical difficulty of methods related to CTC use and poor clinical relevance.17,18 Many techniques have been employed, including filtration and density-gradient centrifugation based on physical properties, such as size,19–21 deformability,22 density23 and charge24. More importantly, immunoassay-based detection specifically captures targeted cancer cells by primarily relying on the antigen affinity of antibodies that recognizes tumorspecific biomarkers, particularly epithelial cell adhesion molecules (EpCAM)25,26 . A series of devices for CTC enrichment that are composed of various organic and inorganic materials have been developed. Devices for CTC isolation must exhibit high capture efficiency, sensitivity, and release flexibility to successfully sort CTCs. Generally, CTC tests should be performed in a sterile environment, and a clean room is essential for maintaining aseptic conditions, particularly for the ex vivo culture of CTCs for individualized testing of 3 ACS Paragon Plus Environment


drug susceptibility. The capability to maintaining CTC tests under sterile conditions is difficult and expensive, particularly in developing countries where researchers lacking extensive and reliable sterilization infrastructures to store and deliver sterile products, which severely increases the testing cost and the risk of contamination and limits the coverage of current medical examination programs. A useful technology must be able to perform complete CTC analyses at a low cost using simple to operation procedures, an infrastructure that is independent as possible, and producing results that are more relevant to clinical outcomes to enable the robust performance of CTC tests in any setting that can be translated to clinical settings, including those with limited resources. Herein, we developed a self-sterilizing and regeneratable polydimethylsiloxane (PDMS)-based microfluidic chip that directly synthesizes and integrates with a zinc oxide (ZnO) nanostructure coating for CTC capture to overcome these limitations. The ZnO nanograss combined with a cylinder array microstructure of PDMS enhances the interaction with cells, improving the efficiency and selectivity of target cell capture. The regeneratable microchip enables effective CTC capture and flexible, nontoxic CTC release during an incubation in a mildly acidic solution, and it can be regenerated at a low cost and recycled for countless cycles to capture and release CTCs without affecting efficiency. The self-sterilizing microchip also allows CTCs to be robustly analyzed in an aseptic and environmentally friendly manner, even in nonsterile environments due to the antibacterial activity of the ZnO nanograss27–29. Thus, this new device holds immense potential for use in the clinic and enabling point-of-care (POC) and home testing, particularly in undeveloped countries and resource-limited settings. Furthermore, the molecular recognition-dependent CTC-capture platform engrafted with a cocktail of antiEpCAM and anti-vimentin antibodies provides more promising and relevant results for the CTC profile regarding the clinical outcomes and allows the CTC-based liquid biopsy to accurately reflect the disease status, particularly for patients with advanced cancer, compared

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with techniques exclusively relying on the anti-EpCAM antibody only in current clinical practices. 2. EXPERIMENTAL SECTION 2.1. Construction of the PDMS substrate with polymer pillars and the integrated ZnO nanostructure. The substrate of the CTC microchip was designed and fabricated at the Microelectromechanical System (MEMS) Technology Center of Zhejiang University using soft lithography and PDMS. First, the structure of the device was designed with AutoCAD (Autodesk, US) followed by the fabrication of the plastic pattern and silicon wafer. The patterned silicon mold was produced as described below. A layer of negative photoresist (SU8-2150) was spin-coated at 500 rpm for 10 s and 2800 rpm for 50 s. After curing in an oven at 65°C for 5 min and at 95°C for 20 min, the wafer was exposed to UV light for 18 s and cured in an oven under the same conditions. Next, the wafer was washed three times with developing liquid, and the mold was exposed to trimethylchlorosilane (TMSC) vapor for 2 min, followed by transfer to a Petri dish. The well-mixed PDMS pre-polymer was poured onto the surface of the mold and vacuumed with a vacuum pump to replicate the pattern and produce a substrate. After being cured in a 90°C oven for 2 hours, the PDMS substrate was ready for use after being peeled off of the mold. The ZnO seed layer (approximately 500 nm) was synthesized on the surface of the clean and dry PDMS substrate with the cylinder array by sputter coating at 120°C for 30 min. Thereafter, a growth-promoting mixture of zinc nitrate hexahydrate (10 mM, Zn(NO3)2·6H2O) and hexamethylenetetramine (10 mM) solution was prepared to promote ZnO nanocrystal growth. Finally, the PDMS plates were immersed into the ready growth-promoting mixture on a heater at 90°C for 48 hours. 2.2. Preparation of the functionalized substrate. Cell binding molecules were introduced onto the nanomaterial-coated PDMS substrate through a sequential chemical covalent coupling method to confer a selective cell-capture capability. Briefly, nanometer-scale ZnO 5 ACS Paragon Plus Environment


crystals were securely grown on the surface of the PDMS substrate, followed by plasma etching for 1 min. Next, the substrate was soaked in 4% (v/v) (3-aminopropyl) trimethoxysilane in ethanol at room temperature for 1 hour, washed with ddH2O and dried with nitrogen gas. The modified surface was then immersed into a 10 mM N, N’disuccinimidyl carbonate (DSC) solution in acetonitrile for 10 min at room temperature, leading to DSC attachment to the surface. After the solution was removed, the surface was rinsed with ddH2O and dried with nitrogen gas again. The substrate was subsequently incubated with 10 µg/mL NeutrAvidin in phosphate-buffered saline (PBS; 0.01 M, pH 7.2) at room temperature for 60 min, and biotinylated anti-EpCAM or anti-vimentin antibodies were diluted to 10 µg/mL in a 1% BSA solution in PBS for 60 min at room temperature. After each solution was removed, the device was soaked in PBS buffer to remove noncovalently attached molecules. Finally, the device was incubated with 1 mL of a 5% BSA solution for 60 min at room temperature to reduce nonspecific cell capture and stored at 4°C until use. 2.3. Cell capture. Mimetic samples were generated by spiking from 101 to 105 4,6-diamidino2-phenylindole (DAPI)-stained cancer cells into 1 mL of processed blood drawn from healthy donors to simulate clinical samples. The healthy blood was processed to obtain partial blood using the following steps: first, diluted blood, which was a 1:1 mixture of peripheral blood and PBS, was discretely added into a 15 mL centrifuge tube containing the same volume of lymphocyte separation medium. After centrifugation at 650 g for 25 min at 4°C, the cells in the middle layer were carefully collected into a new 15 mL centrifuge tube containing 4-5 mL of PBS followed by centrifugation at 400 g for 20 min at 4°C. Finally, the rinsing process was conducted once more before a volume of PBS equivalent to the volume of whole blood was added to simulate clinical samples. Additionally, the substrate was functionalized through a series of chemical covalent coupling reactions, blocked with blocking buffer and integrated into a silver paper to form a groove for containing liquid. Next, the partial blood sample 6 ACS Paragon Plus Environment

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containing cancer cells was added to the groove and shaken at 80 rpm on a shaker for 30 min. Finally, the suspension was removed, the plate was washed three times with PBS, and the substrate without silver paper was observed under a fluorescence microscope. 2.4. Cell recovery. First, the acidic release solution was prepared by adding 0.1 M hydrochloric acid to PBS to generate a pH of approximately 5.6. Next, the device was washed with PBS to remove nonspecific captured blood cells and was then immersed in the acid solution for 2-3 min. The last step was to terminate the release by adding PBS and collecting the targeted cells through centrifugation. The number of released cells was determined by fluorescence microscopy and the procedural counting method. The LIVE/DEAD viability assay was performed to examine the viability of released cells that had been incubated with the weak acid solution for 1-6 min. Cell viability was detected according to by the manufacturer’s protocols. Afterwards, the cells were washed with PBS on a shaker, collected by centrifugation and observed under a fluorescence microscope, followed by counting cells using procedural counting method (Dataset S2). 2.5. Acquisition of environmental bacteria. A 6-well plate containing 3 mL of complete cell culture medium per well was placed in an open space at room temperature to simulate the open environment of device storage, distribution and the procedure of cell-capture procedure. After 18 hours, the medium was concentrated, and the suspension was removed. Next, 10 mL of Luria-Bertani (LB) medium was added and cultured overnight at 37°C and 200 rpm. Finally, the environmental bacteria were obtained and were ready to be employed to test the antibacterial activity of the ZnO nanocrystals. 2.6. Antimicrobial activity of the ZnO coating and the CTC-capture microchip. The antibacterial activity of the ZnO coating was examined by testing the medium turbidity as a qualitative measure of cell growth and by plating assays to compare the cell viability. The environmental bacteria from each experiment were freshly prepared by incubating a miniscule 7 ACS Paragon Plus Environment


amount (0.5 µL of freshly prepared environmental bacterial culture using a pipette tip with a volume of 2.5 µL) into 10 mL of the liquid LB media in a microchip with the ZnO nanocoating as the experimental group and without the nanocoating as the control group to examine the bacterial growth rate in the presence of ZnO nanocoating. The bacteria were grown in an incubator maintained at 37°C and shaken at 200 rpm. The optical density of 200 µL of bacteria medium was measured at 600 nm every two hours, and the background (turbidity due to medium) was eliminated by collecting blank readings. During the optical density measurement, plating assays were performed after an 8 hours incubation. Next, 200 µL of freshly cultured environmental bacteria were diluted 1×105-fold times to adjust the bacterial density, spread on the plates, covered with discs, and incubated for 18 hours at 37°C, after which the number of clone was counted. Culture medium without the ZnO coating served as the control group. MCF7 cells were captured and released with our platform as described above, and then cultured in a 6-well plate to examine the antibacterial activity of the capture device coated with the ZnO nanograss. The enriched cells were proliferated in cell culture flasks for the next 5 days. The optical density of the medium was measured at 600 nm, and the background was corrected by collecting blank readings and comparing them with the original cells that did not undergo the procedure of cell capture and release procedure. 2.7. Assaying the toxicity of the ZnO coating. Cell Counting Kit-8 was employed to precisely investigate whether the ZnO nanomaterial harms the targeted cells, and we simulated the procedure of the cell enrichment and cell release. First, MCF7 cells in good condition were evenly divided into two parts, one of which was added to the silver papermade groove, whose bottom was the PDMS substrate coated with ZnO nanocrystals and was incubated at 37°C in a 5% CO2 atmosphere for 30 min. The other part, which served as the normal control, was maintained in complete culture medium at 37°C in a 5% CO2 atmosphere for 30 min. After the incubation with the acid solution for 2-3 min and centrifugation of cell 8 ACS Paragon Plus Environment

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solution, complete culture medium was added to adjust both cell densities to 2×104/mL, and the cell suspension was plated into wells of 96-well plates. Then, the cells were cultured at 37°C in a 5% CO2 atmosphere for 24 hours, and the culture medium was replaced with 200 µL of complete culture medium mixed with Cell Counting Kit-8 detection solution at the ratio of 10:1. A blank control group was created by plating 200 µL of complete culture medium mixed with Cell Counting Kit-8 detection solution. The plate was then incubated at 37°C in a 5% CO2 atmosphere until the color of the culture solution changed. Finally, the optical density was measured at 450 nm using a SpectraMax M5 microplate spectrophotometer, and the background (turbidity from the complete medium and Cell Counting Kit-8 detection solution) was eliminated by collecting blank readings. 2.8. Clinical trials. Thirty-six samples were collected from 12 patients with primary (n = 6) or advanced (n = 6) breast cancer recruited from three hospitals (Zhejiang Cancer Hospital, Sir Run Run Shaw Hospital and The Second Affiliated Hospital of Zhejiang University, Dataset S1) were recruited. First, 3 mL of the blood samples from patients with cancer was processed as described above. Three functionalized devices were used for each sample based on the anti-EpCAM antibodies, anti-vimentin antibodies and cocktail, respectively. Next, 3 mL of processed blood was added to three devices. After a 30 min incubation on a shaker, the device was washed with PBS. The device was incubated with 1 mL of blocking buffer containing 5% BSA (w./v.) for 1 hour. Both the anti-PanCK detection antibody (conjugated to fluorescein isothiocyanate, FITC) and anti-CD45 detection antibody (phycoerythrin (PE)) were diluted twenty-fold by adding 0.5% BSA in PBS to acquire the best staining results. The captured CTCs were incubated with these antibodies overnight at 4°C, followed by washed with PBS. DAPI (1:1000 diluted in PBS) was incubated with captured cells for 15 min to stain their nuclei and cells were then washed with PBS. Finally, the captured cells were observed under a fluorescence microscope. 9 ACS Paragon Plus Environment


2.9. Statistical analysis. All data were analyzed using Student’s t test and are presented as the means ± SEM of at least triplicate samples from independent analyses, as indicated. P < 0.05 was considered statistically significant. * represented P < 0.05, ** represented P < 0.01, and *** represented P < 0.001. Statistical tests were performed with GraphPad Prism software (La Jolla, CA, USA).

3. RESULTS AND DISCUSSION

3.1. Construction and optimization of the microfluidic chip with hierarchically fractal interface for CTC capture. PDMS is widely applied for CTC enrichment, with sources ranging from blood samples in vitro30 to blood vessels in vivo,31 due to its good biocompatibility, high flexibility, low toxicity, fine optical transparency and easy fabrication with the designed microstructure. The construction of the hierarchically fractal interface for CTC capture is schematically illustrated in Figure 1a. The microfluidic device used in the subsequent experiments was a simple 3 cm × 2 cm PDMS substrate with a micropillar surface composed of circular columns arranged in an equilateral triangular format (Figure 1b and c). When incubated with a cell suspension, the substrate formed in a vessel made of silver paper (Figure S1a and b). The silver paper groove resembles a cup. The activated PDMS substrate was placed on the bottom, and the cell solution was added into the groove and incubated for 30 minutes. We choose silver paper for four reasons. First, cells flowed through the device once, however, cells in the silver paper groove remained in the groove for 30 minutes during an incubation on a shaker and thus had more chances to interact with antibodies, yielding a high capture rate. Second, we can generate the desired shape with silver paper that is maintained for a long time without alterations. Next, we can easily make a groove with silver paper for capturing cells and remove the silver paper when observing the results since the PDMS readily transmits of light. Finally, we can generate markers with the silver paper to 10 ACS Paragon Plus Environment

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identify different samples. The size of the PDMS substrate on the bottom was invariant, but the height of the groove gradually changed. The multilevel structure combined with the micropillar enhanced the interaction between the surface and the cells, improving its capture efficiency for target cells. After we covered the circular columns with ivy-like ZnO nanograss to improve the specific surface area, CTCs had more opportunities to bind to the device (Figure 1d and e). We explored an improved method that would easily and rapidly synthesize the ZnO coating by firmly fixing it to the PDMS substrate using magnetron sputtering to initially deposit a ZnO seed layer of approximately 500 nm. Typical hexagonal ZnO prisms were grown on the seed layer after an incubation with a growth-promoting medium at 90°C in the first round (Figure 1d). The X-ray diffraction (XRD) analysis confirmed that the ZnO coating was pure and had one crystal phase. The XRD pattern was perfectly coincident with the pattern of wurtzite not only in the first growth step but also in the second growth step after the first round was dissolved by the acidic incubation solution (Figure 1e and f). The ZnO nanograss was firmly attached to the PDMS substrate, and the device was functionalized through sequential chemical covalent coupling;32 and thus, the device was sufficiently robust to capture targeted cells. We performed a theoretical investigation of fluidic dynamics to guide the rational design of the microfluidic device for cell capture using the microfluidic module in the software of Comsol MultiphysicsTM (COMSOL, US). Five different arrangements of arrays of 80 µm tall pillars were designed on the PDMS substrate (Figure S1c-g). The first array is hereafter denoted by d100s130, indicating pillars of 100 µm in diameter with a 130 µm distance between the centers of two adjacent pillars. The other samples were similarly labeled d160s210, d160s240, d160s260, and d200s250. The physical cell-capture process was simplified as 1000 particles released from the channel inlet at one time. The micropillar size and interpillar distance were optimized to maximize the capture efficiency (Figure 1g, Figure 11 ACS Paragon Plus Environment


S1h-k and Movies S1, S3, S5 and S6). We analyzed the profiles of the cell trajectory, velocity field and the mechanical shear stress of the devices with five different column arrangements. The d100s130 device exhibited higher shear stress and velocity and a lower capture yield than the other groups (Figure 1h and Movie S2). In contrast, the columns of d160s240 had the lowest shear stress and velocity and the highest capture yield among the five different sizes (Figure 1i and Movie S4); thus, d160s240 was chosen as optimal geometric arrangement. 3.2. Cell-capture performance of the CTC microchip. An immunoassay-based CTCcapturing surface primarily relies on antibodies recognizing tumor-specific biomarkers, particularly EpCAM and vimentin. Thus, these antibodies were engrafted onto the hierarchically fractal interfaces. After sequential chemical covalent coupling, biotinylated antibodies were firmly fixed onto the nanomaterial-coated substrate. Because each NeutrAvidin molecule has four epitopes to combine with biotinylated antibodies, the usefulness of NeutrAvidin further improves the enrichment efficiency. First, we evaluated the relationship between the cell concentration and capture efficiency under different physiological conditions. A series of mimetic samples at concentrations ranging from 10 to 10,000 tumor cells per milliliter was prepared by spiking DAPI-stained cancer cells in PBS, processed blood or whole blood drawn from healthy donors. We chose two cancer cell lines: the MCF7 epithelial cell line and the MDA-MB231 mesenchymal cell line. The results showed that the mean capture yields of MCF7 and MDA-MB231 cells were all greater than 80% in both PBS and processed blood but were less than 60% in whole blood under the same conditions (Figure 2a and b). However, no significant differences when the cell density was varied. PDMS is well known to nonspecifically adsorb proteins. Abundant proteins on the cell surface cause cells to randomly adhere to the PDMS substrate (Figure 2c). We investigated whether the ZnO coating would reduce the nonspecific adhesion of white blood cells but 12 ACS Paragon Plus Environment

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maintain the capture yield of targeted cancer cells. A series of experiments was conducted to further investigate the random adhesion of white blood cells and the specific capture of targeted cancer cells. First, some slippery PDMS substrates were activated with antibodies, and others were coupled with the ZnO coating. After an incubation with DAPI-stained white blood cells in processed blood and three washing with PBS, the fluorescent signals were significantly reduced in the device with the ZnO coating (Figure 2c). The quantitative analysis of the random adhesion of white blood cells on the PDMS substrate with and without the ZnO-nanograss coating clearly proved that the ZnO coating reduced the random adhesion of blood cells (P < 0.001) (Figure 2d). The entire microchip with the ZnO-coated PDMS substrate and polymer pillars was activated using the steps described in Figure 1a to further confirm this finding. Accordingly, the PDMS substrate without the ZnO coating was functionalized by randomly adhering EpCAM-biotinylated antibodies. Next, mimetic samples prepared by spiking DAPI-stained MCF7 cells into processed blood drawn from healthy donors that were then incubated with the differently treated devices for 30 min. Finally, after rinses with PBS, the capture efficiency was confirmed by fluorescence microscopy. The targeting cell-capture efficiencies of the ivy-like ZnO-coated device and the PDMS-only device were not significantly different (P > 0.05) (Figure 2e). These results were consistent with the quantitative real-time polymerase chain reaction (qRT-PCR) results and again illustrated that the ivy-like ZnO coating reduced the random adhesion of white blood cells but did not influence the capture yield of targeted cancer cells, resulting in a high selectivity for cell capture. Currently, several questions remain: what is the relationship between the capture efficiency and the percentage of cells expressing biomarkers on the cell surface? Can the device be extensively applied to enrich different cell types using special markers? A series of experiments was conducted to precisely answer these questions. 13 ACS Paragon Plus Environment


Capture yields were compared among the following different cancer cell lines with varying expression of the epithelial marker EpCAM, as confirmed by immunocytochemistry to evaluate the effect of special biomarker expression on the efficiency of CTC enrichment: HeLa cervical cancer cells, Bxpc3 pancreatic cancer cells and MCF7 breast cancer cells (Figure 2f-h). HeLa cells rarely expressed EpCAM, with a positive percentage of 1.8%, as shown by flow cytometry. However, HepG2 cells and MCF7 cells showed relatively higher positive percentages of expression (59.4% and 98.9%, respectively; Figures 2k and S2a). Each cell line was analyzed after being spiked into processed blood at a density of approximately 103 cells per milliliter to investigate the relationship between the EpCAMpositive percentage and cell-capture yield. HeLa cells exhibited the lowest capture yield (11.90 ± 2.73%), a value that was markedly less than the Bxpc3 cell line (66.53 ± 2.11%) and the MCF7 cell line, which exhibited the highest capture rate (85.47 ± 1.88%) (Figure 2m). Similar results were obtained by applying the mesenchymal marker vimentin to serve as the special biomarker for breast cancer cells of MCF7 and MDA-MB231. Specific expression of vimentin was shown by immunocytochemistry (Figure 2i and j) and confirmed by flow cytometry (Figure 2l and Figure S2b). The percentage of vimentin-positive MDA-MB231 cells was 91.6% compared to 0.7% for MCF7 cells. The capture results ranged from 89.57 ± 3.51% for MDA-MB231 cells to 22.10 ± 2.97% for MCF7 cells (Figure 2n). Based on these results, the device was broadly applicable for different types of cells using specific expressed biomarkers, and the capture yield was closely correlated with the percentage of cells expressing the specific markers, indicating that the capture yield corresponded with the percentage of targeted cells positive for surface markers. 3.3. Recovery of targeted cells and recycling of the CTC-capturing microchip. A perfect capture device should not only have a high capture efficiency but also flexible cell release and no toxicity to targeted cells. In our study, the targeted cells were released via the dissolution 14 ACS Paragon Plus Environment

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of nanoscale ZnO layers following treatment with an acidic solution. To the best of our knowledge, almost all CTC enrichment devices are disposable, which is wasteful and not environmentally friendly; but a recyclable CTC-capture device will avoid these disadvantages. Here, we developed the first regeneratable microchip, and the recycling procedure is shown in Figure 3a. The remaining ZnO coating on the device can be activated for use in countless rounds of cell capture and release since the overall thickness of ZnO coating was much greater than the thickness of the dissolved layer, which was extremely thin (Figure S3h and i). As the release time increased, the nanocrystal coating gradually dissolved until the ZnO nanograss was completely removed. The steps listed above represent the first round of recycling. Then, a new layer of ZnO nanograss was grown on the remaining seed layer, and numerous rounds of cell capture and release were conducted repeatedly. Thus, the second round included the new growth of ZnO crystals on the remaining seed layer and further recycling as in the first round. The morphology and crystal phase of ZnO nanograss grown in the second round changed, as shown by scanning electron microscopy (SEM) (Figure 1e and f), but the function and efficiency remained the same. These results illustrated the possibility of repeating the cell capture and recovery procedure for countless times as needed. During the cycle, safe and extensive release of the captured cells is of pivotal importance. When treated with an acidic solution with a pH less than or equal to 6.0, the majority of ZnO nanocrystals dissociated into Zn2+ divalent cations.33 Given the desired balance between cell viability and the corrosivity of a strong acid, we finally chose a solution with a pH of 5.6 as the release solution. We then wanted to determine the appropriate release time to obtain extensive recovery and high cell viability. We incubated the device with the release solution for 1-5 min after cell capture. The recovery efficiency was calculated as the number of released cells divided by the total number of captured cells; meanwhile, a LIVE/DEAD cell assay was employed to examine the cell viability of the released MCF7 cells. Double staining 15 ACS Paragon Plus Environment


with Calcein-AM/EthD-1 revealed that the number of dead cells increased as the incubation time was extended (Figure S3a-e). Based on the results of a quantitative analysis, approximately 90% of cells were recovered in 2 min, and nearly 98% of cells were released at t = 5 min (Figure S3f). The cell viability was greater than 90% at 3 min because of the mild, cell-friendly release conditions, but approximately 20% of the cells were dead at 5 min (Figure S3g). Based on the chart plotting the trends in cell recovery and cell viability, approximately 90-95% of the captured cells were viable and recovered after 2-3 min of treatment (Figure 3b). We achieved a regeneratable CTC-capturing device due to the ZnO coating. Nevertheless, we wanted to determine whether the capture yield was affected during cycling. Therefore, the capture efficiencies of the 1st, 2nd, 3rd, 4th, 5th, 6th, 11th, 16th and 21st cycles were examined with mimetic samples by spiking DAPI-stained MCF7 cells into processed blood drawn from healthy donors. Student’s t test did not reveal significant differences between the 1st cycle and the other 8 cycles in the first- and second-grown layers of the ZnO coating (Figure 3c and d). We therefore developed an unprecedented recyclable CTC-capture device without affecting the capture yield that exhibited easy, reliable and highly efficient recycling by engrafting PDMS with an ivy-like ZnO coating. 3.4. Sterile-needless usage availability based on the nanometer ZnO coating. Sterility is of extraordinary importance during device storage, distribution and rounds of cell capture and recovery to analyze the cellular and molecular characteristics of CTCs. The antibacterial activity was examined by testing culture turbidity as a qualitative measure of bacterial growth and by performing plating assays to compare bacterial viability in the absence and presence of the ZnO nanocoating. Bacteria in the environment were first obtained to simulate the experimental conditions as closely as possible, enabling us to draw scientific conclusions. The culture turbidity was examined by measuring the optical density of the LB medium at 600 nm. The bacteria from the control environment showed a normal growth curve, including a slow 16 ACS Paragon Plus Environment

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growth period, a logarithmic phase and a stationary phase, after incubation in liquid LB medium for 10 hours. Nevertheless, in the presence of the nanoscale ZnO coating, the same environmental bacteria grew extremely slowly in liquid LB media (Figure 4a). Plating assays were conducted to test the viability of bacteria grown in 200 µL of bacterial medium after an 8 hours incubation. A significantly higher clone count was recorded in the absence of the nanocrystal coating, which served as the control group, than that in the presence of the ZnO nanocoating (Figure 4b-d). Furthermore, no clones were observed in the experimental group (Figure 4b and d). The clone counts of the two groups indicated that the ZnO nanocoating exhibited a robust antibacterial ability (Figure 4b). We then captured and released MCF7 cells with our platform in the open space and then cultured the collected cells in 6-well plates for 5 days. Notably, those enriched cells adhered well to the plate and proliferated in cell culture flasks during the subsequent 5 days. Compared with the original cells that did not undergo the procedure of cell capture and release procedure, the optical density at 600 nm of the medium from 5 day cultures showed almost the same value, which indicated the antibacterial function during the capture and release process (Figure 4e). Based on these findings, the microchip engrafted with the ZnO coating killed bacteria and served as a robust CTC analysis platform without the need for a sterile environment. However, the antibacterial activity raised concerns about the toxicity to the targeted cells when the ZnO nanocoating is used as a component of the capture device. The toxicity of the ZnO nanomaterial was measured using the Cell Counting Kit-8 assay. The optical density at 450 nm was not significantly different between cells grown in the absence and presence of the ZnO nanocoating, indicating the material had no effect on the cell viability (Figure 4f). Thus, the ZnO coating alleviated concerns about the microbial contamination and toxicity of the cell-capture microchip to targeted cells.



3.5. CTC capture from the blood samples of patients with breast cancer. The platform was applied to blood samples donated by patients who provided informed consent according to an Institutional Review Board-approved protocol at Zhejiang University after having optimized the performance of the CTC-capture device. Thirty-six samples from 12 patients with primary (n = 6) and advanced (n = 6) breast cancer were tested (dataset S1). Most of the patients had been treated with chemotherapy; however, the untreated specimens collected before excision ranged from 3 of 6 in patients with primary cancer to 1 of 6 in patients with advanced cancers. Next, 3 mL of blood was collected from each patient, and 1 mL of blood from each sample was analyzed. In this study, we attempted to capture CTCs using antibodies against the epithelial marker EpCAM, the mesenchymal marker vimentin, and a cocktail comprising a 1:1 mixture of antiEpCAM and anti-vimentin antibodies. CTCs enriched from a series of patient blood samples were identified using a comprehensive image analysis, consisting of staining with DAPI for DNA content, FITC-conjugated anti-PanCK antibodies against CK4, CK5, CK6, CK8, CK10, CK13, CK18, CK19 and CK20 for cancer cells, and PE-conjugated anti-CD45 antibodies for leukocytes. DAPI+, PanCK+, and CD45- cancer cells were distinct from DAPI+, PanCK-, and CD45+ normal hematologic cells (Figure 5a and b). We analyzed the capture results obtained EpCAM antibodies, vimentin antibodies and the cocktail (1:1 mixture of EpCAM and vimentin antibodies) from both patients with primary cancer and advanced cancer (Figure 5c and d). CTCs were identified in 4 of 6 (66.7%) clinically localized non-metastatic cancers, and the results obtained using EpCAM antibodies roughly reflected the CTC count in the blood specimens (Figure S4a). CTCs were identified in 6 of 6 (100%) advanced cancer samples using either EpCAM antibodies or vimentin antibodies. Surprisingly, the number of captured cells obtained using EpCAM antibodies was lower than the total number of captured cells obtained using the cocktail (Figure S4b). Cells from patients with advanced cancer had 18 ACS Paragon Plus Environment

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undergone the EMT, which induced the loss of epithelial characteristics and acquisition of the mesenchymal phenotype and cell heterogeneity. Therefore, CTC capture using the cocktail is more reasonable and precise. In addition, the CTC count of advanced cancer specimens was much higher than those of non-metastatic cancer samples and all values were ≥ 5 CTCs because CTCs were thought to be released from localized tumors in primary, non-metastatic cancers; however, in advanced cancers, CTCs may also emanate from metastatic lesions. The only specimen from a patient with advanced cancer who had not received treatment contained a significantly higher CTCs count, which illustrated the positive effect of chemotherapy on other groups by successfully killing cancer cells. In summary, simply engrafting a ZnO coating onto the PDMS pattern endows the CTCcapturing microchip with efficient and sensitive cell enrichment, flexible and harmless targeted cell release, and infinite recycling potential, without microbial infection in an open environment or the need for sterilization. PDMS has been widely employed to isolate and enrich CTCs due to its good biocompatibility, high flexibility, low toxicity, fine optical transparency and easy fabrication of the designed structure.31,34 However, PDMS nonspecifically adsorbs large amounts of serum proteins and white blood cells due to the large number of proteins on their surfaces. The engraftment of an ivy-like ZnO coating onto the PDMS surface enables the devices to reduce the nonspecific adhesion of blood cells. Because the ZnO coating was firmly attached to the surface of the PDMS substrate, it would not detach or affect the capture efficiency and purity. Increasingly, nanomaterials have been applied to enrich CTCs, and low atomic number inorganic materials are believed to be nontoxic and biocompatible and have also been used as drug carriers, cosmetics, and fillings in medical materials.3,35–37 We selected the ZnO coating as a component of the CTC enrichment device for several reasons.



First, the flexible release of targeted cells was achieved with the ZnO coating. Solutions with a pH less than or equal to 6.0 have a marked effect on the stability of ZnO nanomaterials, and most of the components are dissolved and exist as Zn2+ ions.33 A highly acidic solution with a lower pH completely and rapidly releases targeted cells conjugated to ZnO nanograss, but they also harm cells and affect cell viability. Thus, we chose a solution with a pH less than but close to 6.0 as the release solution. Next, antibacterial function is important for the storage and distribution of the medical device and cellular and molecular analyses. Antibacterial agents are broadly classified as organic or inorganic. Organic antimicrobial agents are always less stable at a high temperature or pressure; more importantly, they influence the capture efficiency and purity during CTC capture. Inorganic antibacterial materials, such as metal oxides, are robust and durable and thus have a clear advantage over organic agents. The presence of the ZnO coating enables the long-term storage of the enrichment device without bacterial infection and ensures that captured CTCs are sterile and can be harvested for subsequent cellular and molecular analyses, even in an open environment. Notably, the antibacterial function is affected by the different morphologies of ZnO, by increasing levels of reactive oxygen species produced by the aqueous suspension of ZnO or directly by electrostatic forces of ZnO nanoparticles binding to the bacterial surface.38 Finally, after 48 hours of growth with a firmly settled seed layer of approximately 500 nm, the thickness of the ZnO coating was much larger than the dissolved layer, as the dissolving ability is limited after the device is incubated with a weak acid solution for several minutes.39 The release treatment only dissolves a nanoscale-thin layer attached to antibodies and cells. After cell release, a new functionalized substrate appeared on the remaining ZnO nanocrystals through a sequential covalent coupling method. When needed, the regenerative functionalized device can be applied to a new round of cells for capture and release, showing that cell 20 ACS Paragon Plus Environment

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capture and release could be repeated. After all the initially grown ZnO nanocrystals were dissolved, a new layer of ZnO nanograss can be grown on the seed layer. Next, the newly grown layer of ZnO nanocrystals could be applied to a series of cell capture and release cycles, indicating that the substrate could be continuously recycled. Given the commercial potential of the device, the regeneration capability is of high importance and offers significant advantages. CTCs emanating from primary tumors with phenotypic changes allow the cancer cells to penetrate blood vessels, accompanied by the EMT.40 The EMT plays a vital role in the loss of epithelial characteristics from epithelial cells and the acquisition of the mesenchymal phenotype. Additionally, the EMT causes the downregulation of epithelial markers and upregulation of mesenchymal markers. Many CTC-capture techniques that rely on the epithelial marker EpCAM have been reported and have enriched a considerable number of cancer cells, such as FDA-cleared CELLSEARCH® Circulating Tumor Cell (CTC) Test.41 However, the EMT and heterogeneity of CTCs limited their broad and precise clinical applications. This study included 6 patients with primary breast cancer and 6 patients with advanced breast cancer. Based on our findings, the CTC-capture device based on the antibody cocktail of EpCAM and vimentin antibodies, rather than the EpCAM antibody alone, is more precise and reasonable for disease monitoring and evaluating patients with an advanced stage of cancer. In this case, the numbers and molecular profiles of CTCs from a large cohort of patients with different metastatic cancers undergoing systemic treatment must be investigated thoroughly and compared with standard radiographic methods to provide more clinically relevant information for personized cancer management and precise treatment. Because the CTC-capturing microchip developed in the present study is a versatile engrafting platform, a series of combination cell capture agents, such as multiple antibodies, targeting peptides or aptamers, should be exploited and used for basic cancer biology research and clinical cancer 21 ACS Paragon Plus Environment


management in the future, and may also provide a potential platform for sorting other rare cell types, such as stem cells. 4. CONCLUSIONS In conclusion, the CTC microchip successfully isolated cancer cells from not only spiked blood mixtures but also the clinical samples of patients with cancer with a high capture efficiency, sensitivity and specificity, making the CTC heterogeneity analysis simpler and more convenient. In addition, targeted cells were released flexibly and innocuously by incubating the device with an acid solution. The engraftment of the ZnO coating enabled an unprecedented antimicrobial and recyclable CTC analysis platform. These considerable advantages enable the microchip to be stored for long periods and used in the open environment without concerns of bacterial infection, avoiding the need for disinfection processes in the manufacturing, distribution, storage and even on-site usage of the CTC microchip. The simple ZnO coating made the CTC analysis more convenient and robust in the clinic, and in the future, it may satisfy the need for POC and home testing. The ability of the device to be regenerated may offer a large benefit at a low cost, extending the use of CTC biopsies to the clinic. The clinical application of this versatile CTC platform based on the cocktail of anti-EpCAM and anti-vimentin antibodies will make precise and significant strides in cancer diagnosis and personalized therapy management. The quantification of CTCs could be considered a “liquid biopsy” to predict the response to therapy and monitor disease progression in patients with cancer. Moreover, enriched CTCs may be employed for cellular and molecular studies, providing vital clues to improve our understanding of the mechanisms underlying the metastatic dissemination of cancer and to apply to the discovery of novel prognostic biomarkers or therapeutic targets. Thus, the clinical application of this versatile CTC platform will make significant contributions to personalized cancer therapy.


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ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: www.acs.org. Reagents, computational fluid dynamics analysis method, cell line experiments, collection and processing of blood specimens, microstructure characterization and computational fluid dynamics (CFD) analysis of the CTC microchip, surface marker analysis of cancer cells by flow cytometry, recovery and viability profiles of released cells, CTC quantification and profiles of patients with breast cancer, procedural cell counting method, and C++ code for CTC analysis (file type, PDF) are included. Cell capture simulation of different dimensional microchips using the CFD analysis (file type, AVI) is also included. Movies S1-6 describe the procedure for capturing 1000 cells with the device without columns and with five different sizes of columns in one experiment. In the absence of columns, the device rarely captures cells, and the device with d160s240 (160 µm in diameter × 80 µm tall, 240 µm space between the centers of the two columns) captures the greatest number of cells. AUTHOR INFORMATION Corresponding author *Email: [email protected] (Ben Wang) Authors’ contributions L.H. and B.W. designed the project. L.H., Y.S., T.Y., Q.T and Y.Z. conducted the experiments. L.H., C.H., Y.L. and B.W. analyzed the data. Z.L. performed the procedural counting method. P.C. performed the CFD analysis. X.W. and W.H. provided the clinical 23 ACS Paragon Plus Environment


samples. The manuscript was written with contributions from all authors. All authors have approved the final version of the manuscript. Notes The authors have no competing financial interests to declare. ACKNOWLEDGMENTS We thank Dr. L. Lei & Dr. Z. Hong from the Department of Medical Oncology, Dr. L. Fang from the Department of Pharmacy at Zhejiang Cancer Hospital, and Dr. W. Tian & X. Wang for collecting the clinical samples and participating in constructive discussions. We are grateful to the staff of the Core Facilities of the Institute of Translational Medicine of Zhejiang University for assisting with the flow cytometry, confocal laser scanning microscopy, microplate spectrophotometry and fluorescence microscopy. This study was supported by grant from the Natural Science Foundation of China (81401541 and 81570168), the National Key R&D Program of China (2016YFC1100800), the Distinguished Young Scientist Award of Natural Science Foundation of Zhejiang Province (LR16H180001), the Major Project in Science and Technology of Zhejiang Province (2014C03048-2, 2012C13019-1), and the Medical Development Plan of Zhejiang Province (2015PYA001).

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Figure 1. Construction and simulation of the CTC microchip. (a) Sketch of the fabrication process for the fractal and hydrophilic interface based on the recognition of EpCAM or vimentin on the cell surface. (b) Photograph of the 2 × 3 cm enrichment substrate with a series 32 ACS Paragon Plus Environment

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of columns. (c) Image of the CTC-capture device with columns (d160s240), in which any three cylindrical pillars form an equilateral triangle. SEM images of ZnO nanograss coated on the surface of the substrate from (d) the first synthesis cycle and (e) second cycle (scale bar, 2 µm). (f) XRD analysis of ZnO crystals grown on the substrate during the first and second synthesis cycle. (g) CFD readout profiles of the CTC enrichment system for the cell capture yield of the devices with five different sizes after 1,000, 3,000, 6,000 and 10,000 cells flowed through in a single trial. CFD analysis for the cell capture, velocity field and mechanical shear stress of the PDMS substrate with polymer pillars of (h) d100s130 and (i) d160s240 after 1,000 cells flowed through the device once.



Figure 2. Cell-capture performance of the CTC microchip. Regression analysis of the capture efficiency using a series of mimetic samples at cell densities ranging from 10 to 10,000 tumor cells per milliliter that were prepared by spiking DAPI-stained (a) MCF7 cells and (b) MDAMB231 cells into PBS, processed blood or whole blood drawn from healthy donors. (c) 34 ACS Paragon Plus Environment

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Fluorescent images and (d) quantitative analysis of the nonspecific adhesion of white blood cells on the bare and ZnO-nanograss-coated PDMS substrate (scale bar, 100 µm). (e) Capture yield of the devices with EpCAM antibodies that randomly adhered to and activated nanograss PDMS substrates. (f) Percentage of EpCAM-positive HeLa, HepG2 and MCF7 cells, as investigated by flow cytometry. (g) Percentage of vimentin-positive MCF7 and MDA-MB231, as evaluated by flow cytometry. Confocal images of (h) HeLa, (i) HepG2 and (j) MCF7 cells stained with DAPI and anti-EpCAM antibodies on the cell surface (scale bar, 20 µm). Confocal images of (k) MCF7 and (l) MDA-MB231 cells stained for DAPI and antivimentin antibodies on the cell surface (scale bar, 20 µm). (m) Capture yield based on EpCAM expression in the three cell lines. (n) Capture yield based on vimentin expression in the two cell lines. Data are presented as the means ± SEM (n ≥ 3 and 2 independent replicates). * P < 0.05, ** P < 0.01 and *** P < 0.001 (Student’s t test).



Figure 3. Recovery of targeted cells and recycling of the CTC microchip. (a) Schematic illustration of the regeneratable device for cell capture, release and device regeneration. (b) Trends in the cell recovery and viability of released cells after incubation with a mildly acidic solution for 1-5 min. (c) The capture efficiencies of the 1st, 2nd, 3rd, 4th, 5th, 6th, 11th, 16th and 21st rounds of cell capture and release were examined with mimetic samples by spiking DAPI-stained MCF7 cells into processed blood drawn from healthy donors. The capture rate was not significantly different between the first cycle and the other eight rounds. (d) The capture efficiencies of the 1st, 2nd, 3rd, 4th, 5th, 6th, 11th, 16th and 21st rounds of cell capture and release were examined with the newly grown ZnO coating after the first grown nanocrystals were completely dissolved. The capture rate was not significantly different between the first cycle and the other eight rounds. Data are present as the means ± SEM (n ≥ 3 and 2 independent replicates). * P < 0.05, ** P < 0.01 and *** P < 0.001 (Student’s t test).


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Figure 4. Self-sterilization performance based on the antimicrobial function of the nanoscaleZnO crystal-coated CTC microchip. (a) Growth curve of environmental bacteria incubated with liquid LB medium for 10 hours, as determined by measuring the optical density at 600 nm. The control group showed a normal growth curve, including a slow growth period, a logarithmic phase and a stationary phase. However, the ZnO-treated group showed slow growth without a typical growth curve. (b) Quantitative analysis of the bacterial clones of the control group and ZnO-treated group after incubation in medium for 8 hours. (c) and (d) Bacterial clones from the control and nanomaterial-treated groups grown on solid medium. No clones were observed in the ZnO-treated group in six replicates. (e) Optical densities at 600 nm of the complete medium as a blank control, complete medium after cultured with normal MCF7 cells as a control, and MCF7 cells subjected to enrichment, release, and culture for 5 days as the experimental group. (f) Toxicity analysis of ZnO, as measured by recording the optical density at 450 nm. Untreated MCF7 cells and cells incubated on the ZnOnanocrystal-coated PDMS plate were compared. Data are presented as means ± SEM (n ≥ 3 and 2 independent replicates). * P < 0.05, ** P < 0.01 and *** P < 0.001 (Student’s t test).



Figure 5. Clinical trials of the CTC microchip using blood samples from patients with breast cancer. (a) Images of immunofluorescence staining for captured DAPI+, PanCK+ and CD45− cancer cells (scale bar, 5 µm). (b) Images of immunofluorescence staining for randomly adhered DAPI+, PanCK− and CD45+ leukocytes (scale bar, 5 µm). CTCs captured from (c) clinical non-metastatic, localized breast cancer and (d) metastatic, advanced breast cancer samples based on the individual antibodies or cocktail of EpCAM and vimentin antibodies.


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Self-Sterilizing and Regeneratable Microchip for the Precise Capture

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