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A Microfluidic Chip Integrated with Hyaluronic Acid-Functionalized Electrospun Chitosan Nanofibers for Specific Capture and Nondestructive Release of CD44-Overexpressing Circulating Tumor Cells Mengyuan Wang, Yunchao Xiao, Lizhou Lin, Xiaoyue Zhu, Lianfang Du, and Xiangyang Shi Bioconjugate Chem., Just Accepted Manuscript • DOI: 10.1021/acs.bioconjchem.7b00747 • Publication Date (Web): 07 Feb 2018 Downloaded from http://pubs.acs.org on February 11, 2018

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Bioconjugate Chemistry is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

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Bioconjugate Chemistry

A Microfluidic Chip Integrated with Hyaluronic Acid-Functionalized Electrospun Chitosan Nanofibers for Specific Capture and Nondestructive Release of CD44-Overexpressing Circulating Tumor Cells

Mengyuan Wang, † Yunchao Xiao, † Lizhou Lin, ‡ Xiaoyue Zhu, † Lianfang Du, ‡ Xiangyang Shi*†§

†

State Key Laboratory for Modification of Chemical Fibers and Polymer Materials, College of

Chemistry, Chemical Engineering and Biotechnology, Donghua University, Shanghai 201620, People’s Republic of China. ‡

Department of Ultrasound, Shanghai General Hospital, Shanghai Jiaotong University School of

Medicine, Shanghai 200080, People’s Republic of China. §

Key Laboratory of Textile Science and Technology, Ministry of Education, Donghua University,

Shanghai 201620, People’s Republic of China.

________________________________________________________ *Corresponding author. E-mail: [email protected]

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Bioconjugate Chemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract Detection of circulating tumor cells (CTCs) in peripheral blood is of paramount significance for early-stage cancer diagnosis, estimation of cancer development, and individualized cancer therapy. Herein, we report the development of hyaluronic acid (HA)-functionalized electrospun chitosan nanofiber (CNF)-integrated microfludic platform for highly specific capture and nondestructive release of CTCs. First, electrospun CNFs were formed and modified with zwitterion of carboxyl betaine acrylamide (CBAA) via Michael addition reaction and then targeted ligand HA through a disulfide bond. We show that the formed nanofibers still maintain the smooth fibrous morphology after sequential surface modifications, have a good hemocompatibility, and exhibit an excellent antifouling property due to the CBAA modification. After embedded within a microfluidic chip, the fibrous mat can capture cancer cells (A549, a human lung cancer cell line) with an efficiency of 91% at a flow rate of 1.0 mL/h. Additionally, intact release of cancer cells is able to be achieved after treatment with glutathione for 40 min to have a release efficiency of 90%. Clinical applications show that 9 of 10 non-small-cell lung cancer patients and 5 of 5 breast cancer patients are diagnosed to have CTCs (1 to 18 CTCs per mL of blood). Our results suggest that the developed microfluidic system integrated with functionalized CNF mats may be employed for effective CTCs capture for clinical diagnosis of cancer.

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INTRODUCTION According to the cancer report from American Association for Cancer Research in 2017, the number of cancer cases and deaths from cancer in a global level is continuously increasing every year, and the estimated cancer incidence is expected to exceed 24 million in 2035. Over 90% of cancer mortality is caused by cancer metastasis, which is closely associated with the circulating tumor cells (CTCs) in human blood.1 Briefly, CTCs are tumor cells disseminated from primary tumors to the bloodstream and circulate in the blood circulation system.2 Many studies suggest that CTCs are responsible for metastasis and recurrence of tumors, and the number of CTCs is closely connected with neoplasm stage of cancer.3 Therefore, isolation and analysis of CTCs in human blood is of paramount importance for cancer diagnosis, tumor stage and grade identification, prognosis evaluation and personalized therapy of cancer.4-6 Although CTCs have attracted a great deal of attention in clinics,7-12 the progress of CTC separation technologies has been slow due to the extraordinary rareness of CTCs (approximately one CTC among 106-107 white blood cells (WBCs)).13, 14 Different methods have been developed to detect CTCs. For instance, physical separation method has been used to isolate CTCs depending on the different densities and sizes between CTCs and WBCs. Another notable system, CellSearchTM, an FDA-approved commercial system, is based on the magnetic separation of CTCs using antibody-modified particles. However, these conventional methods generally have low capture efficiencies owing to the similar size of CTCs to WBCs and cancer cell heterogeneity.15,

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addition, these methods generally suffer problems including low sensitivity, time consuming, and damage of CTCs,17, 18 which are difficult to be further analyzed for isolated CTCs.19 The testing platform associated with nanomaterial-based microfluidic chips has attracted considerable attention in the past few years. The introduction of microfluidic system to sort CTCs is 3



quite promising because of its outstanding advantages in accurate operation of liquid samples with a small volume, low cost, high throughput, and high sensitivity of detection.20, 21 For better separation of CTCs, the microfluidic channel is generally integrated with a substrate, such as glass slide,22 silion nanopillars,23 polymer brushes,24 nanowire,23 nanofibrous mat,25, 26 and so on. Among them, electrospun nanofibrous mat owns advantages of simple preparation process and easy surface functionalization,27, 28 ability to mimic native extracellular matrix29 because of its large specific surface area, high porosity, and biocompatibility, and micro-topographical features to regulate cell behaviors in vitro.30-33 In order to achieve efficient and sensitive isolation of CTCs, a variety of molecules such as antibodies,34 aptamers,35 polypeptide,36 or E-selectin37 have been modified on the microfluidic channel or nanofibers to serve as targeting ligands. However, antibody and polypeptide molecules are expensive and are easy to be inactivated.38 Hence, it is extremely important to replace antibodies or peptides with cheaper and more easily available targeting molecules. For instance, our prior work reported that hyaluronic acid (HA)-modified nanofibers could effectively and specifically capture cancer cells overexpressing-CD44 receptors.25, 39 To achieve deep analysis of CTCs in a cellular and molecular level, it is essential to gently release CTCs from nanofibers or microfluidic channel surface.40 Although many attempts, such as enzymatic treatment,41 laser microdissection,26 voltage application,42 or variation of temperature,43, 44 have been used to release CTCs from the capture matrix, it is still a great challenge to release cells in a nondestructive manner. Thus, it is imperative to design a rational linker between capture matrix and targeting ligands for intact release of the cancer cells. Disulfide bond has been reported to be a redox-sensitive linker, which can be rapidly broken via treatment with glutathione (GSH), dithiothretiol, or other reducing reagents.45, 46 Previously, researchers have demonstrated that the 4


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introduction of zwitterions (e.g., carboxybetaine, sulfobetaine, phosphobetiane etc.)47-49 can effectively enhance the antifouling properties of the nanomaterials. So we speculate that it may be feasible to decrease the nonspecific cell adhesion to further enhance the purity of captured CTCs via modification of zwitterion molecules onto the nanofibrous mats or other substrates.

Figure 1. Schematic representation of the fabrication of CNF mats and microfluidic system for capture and release of CTCs. CNFs, chitosan nanofibers; CBAA, carboxyl betaine acrylamide; MPTMS, 3-mercaptopropyltrimethoxysilane; GSH, glutathione.

In this study, a facile approach to fabricate HA-functionalized electrospun chitosan nanofibers (CNFs)-embedded microfluidic chip was developed for highly efficient capture and intact release of cancer cells or CTCs. Specifically, electrospun CNFs were modified with zwitterion of carboxyl betaine acrylamide (CBAA) via Michael addition reaction to decrease nonspecific cell adhesion. Then, HA was conjugated to the surface of CNFs via a disulfide bond (Figure 1). The functionalized 5



CNFs were systematically characterized in terms of structure, morphology, hemocompatibility, and antifouling property. Next, we performed cancer cell capture and release assays under both a static condition using the functionalized CNFs and a dynamic condition using a microfluidic system integrated with the functionalized CNFs. CD44-overexpressing A549 cells (a human lung cancer cell line) were used as a model cell type, and the captured cancer cells were released by GSH treatment. Finally, the clinical utility of the fiber-integrated microfluidic system was tested using cancer patient blood samples and three-color immunocytochemistry was utilized to identify and count the separated CTCs. According to a thorough literature investigation, this is the first example related to the development of chitosan nanofiber-embedded microfluidic system for isolation and detection of CTCs.

RESULTS AND DISCUSSION Synthesis and Characterization of CBAA-CNFs and CBAA-CNFs-HA. Electrospun CNF mats were prepared according to the literature47 using the chitosan/polyethylene oxide (PEO) mass ratio of 9:1 (total polymer concentration of 3.0 wt%) and characterized through scanning electron microscopy (SEM, Figure 2a). The formed CNFs display a smooth fibrous morphology with a uniform fiber diameter distribution (average fiber diameter = 151 ± 30 nm) and the nanofibrous mat exhibits a porous structure with a random orientation. To render the CNFs with water stability, the CNFs were crosslinked with glutaraldehyde (GA) and then observed by SEM (Figure 2b). Crosslinking treatment does not seem to cause any significantly changes to the nanofiber morphology, while the average fiber diameter increases to 175 ± 41 nm after crosslinking. This may be due to the crosslinking process-induced fiber swelling. Besides, we note that the appearance of

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nanofirous mat changes from white to light yellow after crosslinking, further indicating the success of GA crosslinking.50

Figure 2. SEM images and fiber diameter distribution histogram of electrospun CNFs before (a) and after (b) GA vapor crosslinking.

The formed CNFs were modified with CBAA via Michael addition with the chitosan primary amines. In order to link the targeting ligand HA onto the surface of CNFs via a redox-responsive linker,

HA-Cys-3-mercaptopropyltrimethoxysilane

(HA-Cys-MPTMS)

segments

were

first

synthesized (Figure S1, Supporting Information) and characterized by attenuated total reflection-Fourier transform infrared (ATR-FTIR) spectroscopy (Figure S2, Supporting Information). As can be seen in curve 1, the peak at 2960 cm-1 is attributed to the stretching vibrations of -CH3, and the -NH- characteristic peak of L-cysteine ethyl ester hydrochloride (H-Cys-OEt.HCl) appears at 1580 cm-1 and 1541 cm-1. For curve 2, the -OH characteristic peaks of HA at 1043 cm-1 and 3292 cm-1 can be detected. Compared with free HA (curve 2), a new broad peak appearing at 1739 cm-1 in 7



curve 3 can be attributed to the amide group, suggesting that HA-Cys conjugate has been successfully produced via an amide bond. In curve 4, the peak at 2562 cm-1 is assigned to the -SH vibrations of MPTMS, and the methylene characteristic peaks of MPTMS appear at 2942 cm-1 and 2840 cm-1. In contrast with curve 3 (HA-Cys), an enhanced broad peak appearing at 2922 cm-1 in curve 5 can be associated to the -CH2 stretching vibrations, and a new peak at 564 cm-1 can be assigned to the absorption peak of disulfide bond. This suggests that MPTMS has been successfully grafted with the HA-Cys conjugate via a redox-sensitive disulfide bond. We used 1H NMR to further characterize HA-Cys and HA-Cys-MPTMS (Figure S3, Supporting Information). Clearly, the peaks between 3.18 ppm and 3.79 ppm (H2-H11) and the characteristic peak at 1.87 ppm (H12) can be assigned to the HA protons. The methyl protons of H-Cys-Oet.HCl at 1.19 ppm (Hd) can also be clearly observed (Figure S3a, Supporting Information). This suggests the successful modification of H-Cys-Oet.HCl onto the surface of HA. After conjugation of MPTMS, the characteristic peaks at 2.55 ppm (He) and 2.29 ppm (Hf) assigned to the methylene protons of MPTMS appear, suggesting the successful formation of the HA-Cys-MPTMS segments (Figure S3b, Supporting Information). Through NMR peak integration, the number of H-Cys-Oet.HCl molecules attached to each HA and the number of MPTMS molecules conjugated onto each HA-Cys were calculated to be 2.0 and 1.8, respectively. The modification of CBAA or HA-Cys-MPTMS onto the CNFs to form CBAA-CNFs or CBAA-CNFs-HA was also characterized by ATR-FTIR (Figure S4, Supporting Information). In curve 1, the peaks at 3024 cm-1 and 1673 cm-1 are attributed to the C=C stretching vibrations of CBAA monomer, and the peak at 1365 cm-1 corresponds to the methyl groups of CBAA. For curve 2, the peaks at 3366 cm-1 and 3280 cm-1 correspond to the chitosan amine groups and hydroxyl groups of chitosan or PEO for the formed CNFs, respectively. As compared to the pristine CNFs, the new 8


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peak at 1652 cm-1 for the crosslinked CNFs can be assigned to the aldimine linkage between the chitosan amines and GA (curve 3). In addition, a new peak at 1050 cm-1 can be associated to the ether bond formed between the GA and the PEO/chitosan hydroxyl groups. This indicates the success of the crosslinking reaction between GA aldehyde/chitosan amines and the GA aldehyde/chitosan or PEO hydroxyl groups. In contrast with curve 3, a new peak appearing at 1249 cm-1 in curve 4 can be attributed to the stretching vibrations of C-N bond formed between the chitosan amines and CBAA. In addition, the C=C characteristic peaks disappear in curve 4. These spectral changes indicate that CBAA has been modified onto the surface of CNFs via Michael addition successfully. By comparison with CBAA-CNFs (curve 4), the Si-O-C vibrations of MPTMS at 1150 cm-1 can be detected (curve 5), which suggests that HA-Cys-MPTMS has been successfully immobilized onto the surface of CNFs to get the CBAA-CNFs-HA product. The surface hydrophilicity of the fibrous mats was assessed by water contact angle measurements (Figure S5, Supporting Information). Pristine CNFs have a water contact angle of 46.9°. After further CBAA and HA modification, the contact angle of the nanofibrous mats reduces to 35.0° and 24.8°, respectively, at the same time point. The enhanced hydrophilicity of the CNFs further suggests the success of the modification of CBAA and HA onto the surface of CNFs. The fiber surface modifications as characterized by ATR-FTIR and contact angle measurements indirectly confirm that the composition of chitosan is also on the fiber surface. Furthermore, thermal gravimetric analysis (TGA) was carried out to quantitatively analyze the degree of CBAA and HA modification onto the CNFs (Figure S6, Supporting Information). It can be seen that pure CNF mat displays a weight loss of 86% at 900 oC, while CBAA-CNFs and CBAA-CNFs-HA have a weight loss of 83% and 65% at the same temperature, respectively. By subtracting the weight loss of CBAA-CNFs from that of the pure CNF mat, the CBAA and 9



HA-Cys-MPTMS modification percentages can be calculated to be 3% and 21%, respectively. Hemocompatibility Assay. Hemolysis and dynamic anticoagulant assays were employed to assess the hemocompatibility of functional CNFs. As shown in the inset of Figure S7 (Supporting Information), human red blood cells (HRBCs) incubated with three kinds of nanofibers (CNFs, CBAA-CNFs, or CBAA-CNFs-HA) display no obvious hemolytic phenomena, similar to those treated with PBS. The hemolysis rate of HRBCs treated with the CNFs, CBAA-CNFs, and CBAA-CNFs-HA were calculated to be 0.9%, 0.8% and 0.6%, respectively, which are all much less than the standard threshold value (5%),51 suggesting that the developed CNFs and functional CNFs do not seem to cause hemolysis. Anticoagulant assay was also performed to investigate the hemocompatibility of the CNFs, CBAA-CNFs, and CBAA-CNFs-HA (Figure S8, Supporting Information). Compared with the control group (cover slip), the OD values for all three nanofibrous mat groups are higher at the same incubation time, suggesting that functional CNFs have good anticoagulant properties. These results suggest that the developed CNFs and functional CNFs display a good hemocompatibility, which is essential for further CTC capture applications using blood samples. Antifouling Assay. The antifouling property of CNFs and functional CNFs was verified through fibrinogen adsorption and blood-cell attachment assays (Figure 3). In Figure 3a, we can see that the fibrinogen adsorption onto CBAA-CNFs and CBAA-CNFs-HA is much lower than that onto the pristine CNFs (p < 0.001) under the same experimental conditions, suggesting that the nanofibrous mats functionalized with CBAA have excellent resistance to protein adsorption.

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Figure 3. (a) Fibrinogen adsorption rate onto different fibrous mats after incubation at 37 oC for 1 h. (b) The density of WBCs trapped onto the nanofibrous mats. (c) Fluorescence microscopic images of WBCs trapped onto three kinds of fibrous mats after incubation at 37 oC for 1 h. For each sample, 1 mL of WBC suspension (1 × 106 cells/mL) was seeded into each well of the 24-well cell culture plate. Level of significance is ***: p < 0.001.

The excellent antifouling property of the CBAA-modified fibrous mats was further confirmed by blood cell resistance assay (Figure 3b-c). In Figure 3b, the density of WBCs attached on the pristine CNFs is 308 cells/mm2, while the cell density on the CBAA-CNFs and CBAA-CNFs-HA is 5 cells/mm2 and 9 cells/mm2, respectively at the same incubation time point. These results indicate that zwitterion CBAA modification could significantly reduce the blood-cell attachment onto the fiber surface. The excellent antifouling property was also qualitatively validated by fluorescence microscopic images of the WBC attachment onto the fibrous mats (Figure 3c), where much less 11



WBCs are attached onto the CBAA-functionalized CNFs when compared to the pristine CNFs without CBAA. The excellent antifouling property of the CNFs should be due to the modification of zwitterion CBAA molecules consisting of equal molar amount of positive and negative charges in close neighbourhood, thereby holding many water molecules around and form a protection layer of water on the surface of materials. This is likely to shield the interactions between the proteins/WBCs and the fiber surface, in agreement with the literature.52, 53 Capture and Release of Cancer Cells Using the Functionalized CNFs under Static Conditions. We next used the CBAA-CNFs-HA for specific cancer cell capture under static conditions (Figure 4). As shown in fluorescence microscopic images, mixed WBCs and A549 cells are able to be non-specifically captured by the pristine CNFs (Figure 4a), and few WBCs are captured onto the surface of CBAA-CNFs (Figure 4b). In contrast, more A549 cells (red cells) can be captured by the CBAA-CNF-HA mats by virtue of the modified targeting ligand HA (Figure 4c). Quantitative results (Figure 4d) show that the capture efficiency of A549 cells onto different nanofibers increases with the incubation time. Compared with CBAA-CNFs and CNFs, the capture efficiency of CBAA-CNFs-HA is significantly higher (p < 0.001) at the same incubation time points. For instance, the A549 cell capture efficiency on the CBAA-CNFs-HA is 81% (number of captured A549 cells: 244 ± 36) with nonspecifically captured WBCs (1700 ± 500) at 60 min, while only 38% (number of captured A549 cells: 114 ± 30; and number of non-specifically captured WBCs: 36000 ± 2000) and 15% (number of captured A549 cells: 46 ± 20; and number of non-specifically captured WBCs: 1200 ± 300) of A549 cells are captured onto the CNFs and CBAA-CNFs at 60 min, respectively. These results indicate that the developed CBAA-CNFs-HA have an ability to efficiently and specifically capture A549 cells due to the presence of the targeting ligand HA.

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Figure 4. Fluorescence microscopic images of A549 cells and WBCs captured onto CNFs (a), CBAA-CNFs (b), and CBAA-CNFs-HA (c) under static conditions at an incubation time of 40 min. (d) The capture efficiency of A549 cells on different nanofibrous mats under static conditions at different incubation time periods. For each sample, 1 mL of the mixed cell suspension (106/mL WBCs and 300/mL A549 cells) was added into each well of a 24-well cell cuture plate. Levels of significance are ** for p < 0.01 and *** for p < 0.001, respectively.

The captured cancer cells were next treated with GSH to cleave the disulfide bonds to release the cells from the nanofibrous mats. Firstly, we used GSH at different concentrations to check the release efficiency of the captured cancer cells (Figure S9, Supporting Information). It can be seen that the release efficiency of A549 cells increases with the GSH concentration, and reaches 92% at the GSH concentration of 50 mM. Subsequently, the release efficiency levels off at the GSH concentration over 50 mM. Hence, we chose the GSH concentration at 50 mM for the subsequent experiments.

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Figure 5. (a) Fluorescence microscopic images of A549 cells captured on the CBAA-CNFs-HA before and after release (incubated with GSH for 40 min), and live-dead cell staining of the released A549 cells. (b) The release efficiency of A549 cells after incubated with GSH (50 mM) for different time periods. (c) The viability of the released A549 cells after incubated with GSH (50 mM) for different time periods.

We further performed the release experiments with 50 mM GSH concentration treatment for different time period to obtain the optimum GSH treatment time. Fluorescence microscopic analysis (Figure 5a) reveals that almost all the captured A549 cells can be released and the released A549 cells have a fairly high cell viability. The release efficiency of A549 cells increases with the incubation time within 40 min, and tends to be constant over 40 min (Figure 5b). Quantitative viability assay of the released A549 cells shows that the viability of A549 cells reaches 89%-100% when treated with GSH for 10-60 min (Figure 5c), suggesting that the GSH treatment does not seem to damage cells and the intact cell release can be achieved within 40 min. 14


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Dynamic Capture of Cancer Cells Using a Fiber-Integrated Microfluidic Chip. Next, we tested the fiber-integrated microfluidic chip system to dynamically capture cancer cells. Firstly, under different flow rates we investigated the capture efficiency of A549 cells (Figure 6a). Apparently, the capture efficiency of A549 cells increases with the decrease of flow rate. The capture efficiency of A549 cells can be as high as 91% at a flow rate of 1.0 mL·h-1, which has no statistically significant difference compared with the capture efficiency at 0.5 mL h-1. Thus, we selected the flow rate of 1.0 mL h-1 for the subsequent dynamic cell capture experiments.

Figure 6. (a) The capture efficiency of A549 cells in a microfluidic chip integrated with the CBAA-CNFs-HA at different flow rates. (b) The capture efficiency of A549 cells when spiked with various number of A549 cells using a flow rate of 1.0 mL h-1. (c) The capture efficiency of spiked A549, HeLa, MCF-7, and U87MG cells, and in each case the cancer cells were spiked into RBC-lysed blood with a cancer cell density of 300 cells/mL. Level of significance is *** for p < 0.001.

The detection range of the microfluidic chip system was then tested by spiking A549 cells in the RBC-lysed blood at different densities (Figure 6b). The microfluidic device exhibits an A549 cell capture efficiency in a range of 80% (number of captured A549 cells = 8 ± 2 at a spiking density of 10 cells per mL)-94% (number of captured A549 cells = 187 ± 10 at a spiking density of 200 cells 15



per mL) when 10-300 A549 cells were spiked to the WBCs, which is comparable to that achieved by CellSearch (> 80%).54 These results indicate that our microfluidic device with a detection threshold of at least 10 cancer cells/million WBCs is competent for clinical samples, which usually have a CTC density at a few to hundreds cells per milliliter. Next, the capture performance of different types of cancer cells was investigated (Figure 6c). The capture efficiencies of HeLa, A549, and MCF-7 cells all having CD44 receptor overexpression are over 89%, while the capture efficiency of U87MG cells (with low CD44 receptor expression) was only 35%. This suggests that our microfluidic device could specifically capture CD44 receptor-expressing cancer cells with a considerable efficiency. Our results also imply that the fiber-integrated microfluidic chip could be employed to separate CTCs of different phenotypes from the whole blood. Clinical Utility Assessment. With the excellent cancer cell capture efficiency achieved, blood samples of patients were next investigated to isolate CTCs using our device. The isolated CTCs were collected through GSH treatment, immune-stained and observed using fluorescence microscopy. From Figure 7a, we can see that the collected CTCs are CK+(red)/4’,6-diamidino-2-phenylindole (DAPI)+(blue)/FITC-(green) cells, while WBCs are CK-/DAPI+/CD45+ cells, demonstrating that the CTCs have been effectively sorted. The CTC capture results are summarized in Figure 7b. We show that our developed microfluidic chip can isolate CTCs (1-18/mL) from whole blood of 9 of 10 non-small cell lung cancer patients and 5 of 5 breast cancer patients.

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Figure 7. (a) Fluorescence microscopic images of CTCs captured from lung cancer patient blood using the fiber-integrated microfluidic device. Cells (with blue nucleus staining) stained positive for cytokeratin 7 (CK, red) and negative for the WBC marker CD45 (green) are identified as CTCs. (b) CTC enumeration results obtained from 1 mL of blood samples of breast cancer patients (red columns, Bn) and lung cancer patients (blue columns, Ln).

CONCLUSIONS To the end, we reported a facile method to fabricate functionalized CNF-embedded microfluidic chip for highly effective and specific capture of cancer cells and CTCs. CNFs are able to be functionalized with zwitterion CBAA via Michael addition with the chitosan primary amines and with targeting ligand HA via silanization of the chitosan or PEO hydroxyl groups to have redox-sensitive disulfide linkages. With the simultaneous modification of CBAA and HA, the formed CBAA-CNF-HA mats display excellent antifouling property, can specifically capture CD44 receptor-overexpressing cancer cells with an efficiency up to 81% under a static condition, and can non-destructively release the captured cancer cells by cleaving the disulfide bond via GSH treatment. Using the fiber-integrated microfluidic device, a high cancer cell capture efficiency of 91% can be 17



achieved, and CTCs from both breast and non-small cell lung cancer patients can be isolated and identified to prove the potential clinical utility. Our study suggests that the developed CBAA-CNFs-HA-integrated microfluidic chip system may be used as a promising platform for sorting and identifying CTCs for clinical diagnosis applications.

EXPERIMENTAL PROCEDURES Functionalization of CBAA onto CNF Mat. Crosslinked CNFs were formed according to the literature (see details in Supporting Information).47 CBAA was synthesized according to the literature42 and characterized by 1H NMR spectroscopy (Figure S10, Supporting Information). Then, CBAA was grafted onto CNFs via Michael addition. In brief, CNF mats (20 pieces in total) were immersed in 100 mL of methanol solution and 16 mg of CBAA dissolved in sodium chloride water solution (100 mL, 0.138 M) was added into the above methanol solution under continuous stirring. 2, 6-Bis(1,1-dimethylethyl)-4-methylpheno (0.05 mg) was then added into the above reaction mixture to inhibit CBAA polymerization. After stirring for 48 h at room temperature, the CNF mats were taken out and then cleaned with water for 3 times. The CBAA-CNFs were obtained after drying in vacuum for 48 h. Immobilization of HA-Cys-MPTMS onto CBAA-CNFs. Redox-sensitive targeting ligand (HA-Cys-MPTMS) segment was synthesized according to the literature.46 HA-Cys-MPTMS was immobilized onto the surface of CBAA-CNFs through the silanization reaction between MPTMS and the chitosan/PEO hydroxyl groups. In brief, HA-Cys-MPTMS segment (58.6 mg) was dissolved in ethanol (200 mL), and then CBAA-CNFs were immersed in the ethanol solution under stirring overnight at room temperature.46 Finally, the nanofibrous mats were washed with water for 4 times and dried in vacuum for 48 h. The formed CBAA-CNFs-HA were kept in vacuum before use. 18


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Antifouling Assay. The antifouling property of nanofibrous mats was investigated by protein absorption test. First, the standard fibrinogen concentration-absorbance (280 nm) calibration curve was obtained. Afterwards, circular coverslips (diameter = 14 mm) covered with CNFs, CBAA-CNFs and CBAA-CNFs-HA were placed in a 24-well tissue culture plate and fixed using stainless steel rings. All samples were prepared in quadruplicate and equilibrated with 500 µL of PBS for 1 h at room tempreture. Subsequently, the PBS solution was sucked out and then 500 µL of fibrinogen solution (1 mg/mL, in 1 mL PBS) was added into each well and incubated for 1 h at room temperature. Then, the supernant in each well was collected and measured by UV-vis spectrophotometer to record the absorbance at 280 nm. The adsorption rate of fibrinogen was calculated according to Equation (1):

Adsorption % = − ⁄ × 100%

(Eq. 1)

where CI is the initial concentration of fibrinogen solution, CF the final concentration of fibrinogen solution after incubation for 1 h. The fibrinogen concentration was determined according to the concentration-absorbance calibration curve of fibrinogen. Blood-cell adhesion assay was next carried out. Typically, each circular coverslip covered with CNFs, CBAA-CNFs or CBAA-CNFs-HA (each sample in quadruplicate) was placed in cell cuture plate and treated under the same conditions as described above. Calcein AM (green) was used to stained the freshly isolated WBCs prior to blood-cell attachment experiment. WBC suspension (1 mL, 1 × 106 cells/mL) was seeded into each well, followed by incubation at 37 oC for 1 h. Afterwards, the fibrous mats were gently rinsed with PBS for 3 times. The PBS solution was collected and concentrated, and then the cells in the recovery solution were counted by a Scepter 2.0 Handheld Automated Cell Counter (Merck Millipore, Darmstadt, Germany). Fluorescence microscopy (Carl Zeiss, Axio Vert. A1, Germany) was adopted to observe the captured cells on the 19



Page 20 of 32

three different nanofibrous mats using a 10 × objective lens. The blood-cell attachment percentage (β) was calculated according to Equation (2):

β = − ⁄ × 100%

(Eq. 2)

where n0 is the total number of WBCs seeded in each well and n1 the number of WBCs in the recovery solution. Static Cancer Cell Capture and Release Assays. CD44-overexpressing A549 cells were selected as model cancer cells to conduct the cancer cell capture and release assays under static conditions. Firstly, each circular coverslip covered with CNFs, CBAA-CNFs or CBAA-CNFs-HA (each sample in quadruplicate) was placed in wells of cell culture plate and treated under the same conditions as described above. Then the Calcein red-labeled A549 cells (red) were spiked into Calcein AM-prestained WBCs (green) with a WBC density of 106/mL and a A549 density of 300/mL, respectively. Subsequently, the mixed cell suspension (1 mL) was added into each well, followed by incubation at 37 oC and 5% CO2 for a certain time period (10, 20, 40, or 60 min, respectively). At each time point, the nanofibrous mats were rinsed with PBS for 3 times. The A549 cells captured on the nanofibrous mats were imaged and counted under a fluorescence microscope. The capture efficiency (η) of cells was calculated according to Equation (3):

η = ⁄ × 100%

(Eq. 3)

where n0 is the amount of A549 cells added into each well and n2 the amount of A549 cells captured on the nanofibrous mats. For static release experiments, each nanofibrous mat was placed into cell culture plat wells and treated under the same conditions as described above, then Calcein AM-prestained A549 cells (1 × 105 cells per well) were added into the culture plate, followed by incubation at 37 oC and 5% CO2 for 1 h. Afterwards, the nanofibrous mats were rinsed with PBS for 3 times to retrieve unbound A549 20


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cells. The collected A549 cells in the recovery soluiton were concentrated and counted using a Automated Cell Counter. After capture of A549 cells, GSH solution was adopted to release the captured A549 cells. Firstly, in order to optimize the GSH concentration, different concentrations of GSH solution (10 mM, 30 mM, 50 mM, 70 mM and 90 mM, respectively) were added into each well of the 24-well culture plate, followed by incubation at 37 oC for 1 h, and then the nanofibrous mats were rinsed with PBS for 3 times. Afterwards, the cell suspension was transferred to a centrifuge tube, and cells were counted using a Automated Cell Counter. The cancer cell release efficiency (ξ) was calculated according to Equation (4):

ξ = ⁄ − × 100%

(Eq. 4)

where n0 is the amount of A549 cells initially added into each well; n3 the number of the uncaptured A549 cells; and n4 the amount of released A549 cells. Next, to obtain the optimum GSH treatment time, we further performed the release experiments with optimum concentration of GSH treatment for different time periods (10, 20, 30, 40, 50, or 60 min). The release procedures were the same as the protocols described above. In addition, fluorescence microscopy was adopted to observe the A549 cells captured onto the substrate before and after the cell release (10 × objective lens). The release efficiency of cancer cells was calculated according to Equation (4). The survival rate of released cells was investigated using a standard Live/Dead staining protocol. The viability assay of released cells is quite similiar to the above cancer cell release assay. The only difference is that A549 cells were not stained prior to seeding into a 24-well culture plate. After GSH (50 mM) treatment, the recovered solution was centrifuged (1000 rpm for 5 min) and was stained with propodium iodide (PI) solution (8 µM) and Calcein AM solution (2 µM) for 15 min, respectively. Then, the stained cells were observed by a fluorescence microscopy (10 × objective 21



Page 22 of 32

lens), and the live and dead cells were counted respectively from at least 5 different image fields for each sample. The cell viability (φ) was calculated according to Equation (5):

φ = " ⁄" + $ × 100%

(Eq. 5)

where nl and nd is the amount of live cells and dead cells in the recovery solution, respectively. Dynamic Cell Capture Using a Fiber-Integrated Microfluidic Chip. Using the designed fiber-integrated microfluidic system designed (Figure S11, Supporting Information), we firstly optimized the capture efficiency of A549 cells under different flow rates. A549 cells were incubated under the same conditions as mentioned above. Prior to the capture experiment, the microfluidic chip was infiltrated with PBS. Then, Calcein AM-stained A549 cells (300 cells/mL) were spiked into freshly isolated WBC suspension with a WBC concentration of 106/mL. After that, the cell suspension (1 mL) was pumped into the microfluidic device at different flow rates (0.5, 1.0, 2.0, 4.0, or 6.0 mL h-1, respectively). After cells were captured, PBS (500 µL) was passed through the microfluidic chip at a flow rate of 2.0 mL h-1 to remove the non-adherent cells. Then, the cells captured on the fibrous mat were counted under a fluorecence microscope. The capture efficiency of A549 cells was calculated according to Equation (3). In order to obtain the detection range of the microfluidic chip, different numbers of pre-stained A549 cells (10, 20, 50, 100, 200, or 300 cells per mL) were spiked into WBC suspension. Analogous experimental processes were carried out to investigate the capture efficiency at an optimum flow rate as described above. Then, the captured A549 cells were viewed and counted using a fluorescence microscope and the capture performance was quantified by Equation (3). Next, we studied the capture performance of various types of cancer cells. Three kinds of CD44 receptor-expressing cancer cells (A549, HeLa, and MCF-7 cells) were selected, and U87MG cells with low CD44 receptor expression were adopted as a negative control. All types of cancer cells 22


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were stained with Calcein AM and were spiked into a WBC suspension at a concentration of 300/mL. The capture experiments were carried out according to the processes described above at an optimum flow rate. Cancer cells captured on the substrate were counted under fluorescence microscope. The cell capture efficiency was calculated using Equation (3). Potential Clinical Utility Assessments. We collected peripheral venous blood from 15 patients suffering from non-small-cell lung cancer (10 patients) and breast cancer (5 patients) without radiotherapy or chemotherapy. After lysed by RBC lysis buffer, 1 mL of cancer patient blood was introduced into our microfluidic device. The recovery solution containing CTCs and WBCs was obtained after treated with GSH solution for 40 min at a flow rate of 2.0 mL/h. A commonly used three-color immunocytochemistry method was utilized to identify and count CTCs, including FITC-labeled anti-CD45 (a marker for WBCs) and Alexa Fluor 568-labeled anti-cytokeratin-7 (CK-7, a protein marker for epithelial cells) as well as DAPI nuclear staining. The stained cells were imaged and counted under fluorescence microscope. More experimental details can be seen in Supporting Information.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: xxx. Additional experimental details, reaction scheme, fiber-integrated microfluidic chip design, materials characterization data of FTIR, TGA, contact angle, and 1H NMR, and in vitro hemolysis and anticoagulant assay results.(PDF)

23



AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. ORCID Xiangyang Shi: 0000-0001-6785-6645 Notes The authors have no competing financial interest to declare.

ACKNOWLEDGEMENTS This study was financially supported by National Natural Science Foundation of China (81761148028, 21773026 and 21405012), the Science and Technology Commission of Shanghai Municipality (17540712000), the support from the Key Laboratory of Textile Science & Technology, Ministry of Education, “111 Project” (B07024), and the Fundamental Research Funds for the Central Universities.

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