Janus micromotors for motion-capture-ratiometric fluorescence detection of circulating tumor cells

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Janus micromotors for motion-capture-ratiometric fluorescence detection of circulating tumor cells Long Zhaoa,b, Yuan Liua, Songzhi Xiea, Pan Rana, Jiaojun Weia, Qingjie Liua, Xiaohong Lia,

⁎

a

Key Laboratory of Advanced Technologies of Materials, Ministry of Education, School of Materials Science and Engineering, Southwest Jiaotong University, Chengdu 610031, PR China b School of Bioscience and Technology, Chengdu Medical College, Chengdu 610031, PR China

HIGHLIGHTS

GRAPHICAL ABSTRACT

micromotors are developed to • Janus achieve motion-enhanced capture of circulating tumor cells.

is grafted on one side of • Catalase Janus rods to catalyze H O decom2

• • •

2

position as power source. TLS11a aptamers are conjugated on the other side of Janus rods for specific capture of cells. The competitive binding with cells induces ratiometric fluorescence changes from blue to green. Janus micromotors demonstrate the motion-capture-ratiometric fluorescence sensing capabilities.

ARTICLE INFO

ABSTRACT

Keywords: Janus micromotor Self-propelled motion Circulating tumor cell capture Aptamer labeling Ratiometric fluorescence

The rapid and sensitive detection of circulating tumor cells (CTCs) remains technical challenges due to extremely low abundance. In the current study, Janus micromotors (JMs) are developed to achieve motion-enhanced CTC capture and capture-induced ratiometric fluorescence signaling. JMs are constructed via catalase grafting on one side of Janus rods (JRs) to catalyze the decomposition of H2O2 as power source. JMs with different aspect ratios show random, spiral, rotational or linear motion trajectories, and JMs with the aspect ratio of 2 (JM-2) display rotational motion with significantly larger mean-square-displacement (MSD) values than other JMs. TLS11a aptamers are conjugated on the other side of JRs for specific capture of tumor cells, and tetraphenylethene (TPE) derivatives and fluorescein isothiocyanate (FITC) are labeled on aptamers via base-pair interactions. The competitive binding of tumor cells with aptamers causes the release of TPE and FITC from JMs, which relieves the aggregation-induced emission (AIE) effect of TPE and aggregation-caused quenching (ACQ) profile of FITC on JMs. Thus, JMs displays apparent ratiometric fluorescence changes from blue (I450) to green (I526) after capture of tumor cells. The fluorescence intensity ratios of I526/I450 could be fitted against cell levels, and the limit of detection is around 25 cells/mL within 1 min. In addition, the HepG2 detection by JM-2 exhibits no interference in the presence of other tumor cells (4T1 and H22), and the recovery rate of HepG2 cells in blood samples was over 95%. Thus, JMs demonstrate the motion-capture-ratiometric fluorescence sensing capabilities and provide a feasible strategy for real-time, rapid and sensitive detection of cells in laboratory, hospital and remote settings.

⁎ Corresponding author at: School of Materials Science and Engineering, Southwest Jiaotong University, 111 North 1st Section, 2nd Ring Road, Chengdu 610031, PR China. E-mail address: [email protected] (X. Li).

https://doi.org/10.1016/j.cej.2019.123041 Received 17 July 2019; Received in revised form 11 September 2019; Accepted 2 October 2019 1385-8947/ © 2019 Elsevier B.V. All rights reserved.

Please cite this article as: Long Zhao, et al., Chemical Engineering Journal, https://doi.org/10.1016/j.cej.2019.123041

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1. Introduction

loaded [12]. Another strategy is derived from the large towing force and surface bio-functionalization of micromotors to realize selective isolation and efficient transporting of target analyst in complex biological media, avoiding laborious and time consuming sample preparation steps [13]. Jurado-Sanchez et al. modified graphene quantum dots with phenylboronic acid and loaded into catalytic micromotors for the detection of bacteria endotoxins. The micromotor motion induced continuous mass transport to increase the interaction rate of phenylboronic acid tags with lipopolysaccharides compared to that of static counterparts, resulting in the fluorescence quenching of graphene quantum dots in the presence of target endotoxins [14]. There are several challenges in the use of micromotors for CTC detection. Most of the investigated micromotors are microtubes, which are prepared via rolling-up and template electrodeposition, and the propulsion is along the long axis [15]. The microtube carriers are efficient in the separation and enrichment of small substances, but indicate limitations in the attachment and capture of cells. In the current study, micromotors from fibrous rods are designed to offer multiple interactions as wrapping around the cells, thus decreasing the inherent steric hindrance of microtubes and microparticles for cell attachment and capture [16]. In addition, the flexible fibrous micromotors should provide more available sites to accommodate chemical cues conferring greater sensitivity and maximize cell recovery with minimal cell loss [17]. The second challenge is the cell sensing after capture by micromotors. In the current study, a signal-on-motion strategy is developed without expensive instrumentation and tedious sample preparation. “Turn-off” or “turn-on” responses of micromotors usually cause fluctuations of fluorescence intensities in the presence of analysts. The complex biological media tend to have large autofluorescence background [18], and the fluorescence measurement be affected by instrumental and environmental factors. Thus, the ratio of emission intensities at two wavelengths is utilized for ratiometric fluorescence sensing, alleviating the measurement variations. In addition, the change of fluorescent colors rather than fluorescence intensities leads to a sensitive detection by naked eyes. It should be noted that Janus fibrous micromotors and ratiometric fluorescence sensing have not been investigated up to date. In the current study, Janus rods (JRs) were prepared by cryocutting of Janus fibers, which were obtained by side-by-side electrospinning. Electrospinning has the advantages of simple equipment operation, low cost and mass production. The high surface area of electrospun fibers can provide abundant graft sites and increase the interaction rate as biosensor substrates [19]. Scheme 1a illustrates the preparation process of Janus micromotors (JMs). Catalase (CAT) was decorated on one side of JRs by covalent immobilization, while the other side was covalently bound with TLS11a aptamer. CAT-catalytic reaction could provide the propulsion force for JRs in the presence of H2O2 fuel, and TLS11a aptamer provided a high affinity to tumor cells [20]. To achieve a ratiometric fluorescence sensing, thymine and guanine were conjugated on tetraphenylethene (TPE) and fluorescein isothiocyanate (FITC) to obtain TPE-T and FITC-G, followed by grafting onto aptamers via basepair interactions. The fluorescence of TPE was turned on due to the aggregation-induced emission (AIE) characteristics, while the fluorescence of FITC was turned off due to the aggregation-caused quenching (ACQ) effect. As shown in Scheme 1b, after specific capture of CTCs, the releases of TPE and FITC from JMs restored the fluorescence of FITC and diminished the emission of TPE, and the amount of captured cells induced a ratiometric fluorescence change of FITC and TPE in JM suspensions. Thus, JMs demonstrate the capabilities of motion-captureratiometric fluorescence detection of CTCs and have desirable features such as high selectivity, rapid recognition, and low detection limit.

Nowadays cancer is becoming one of the most devastating diseases, and over 90% of cancer-related death is related with cancer metastasis. Metastasis is resulted from the escape of tumor cells from the primary site to arrest and grow at another site into metastatic tumor colonies. The tumor metastasis involves several steps and one of the essential steps is travelling in the circulation of blood or lymph system, adhering to local vascular endothelium and invasion into different tissues of the body [1]. In addition, the break-away of tumor cells from primary tumor mass usually indicates malignant progressions of cancers. Thus, the detection of circulating tumor cells (CTCs) in peripheral blood is of great value in monitoring the tumor progression, therapeutic efficacy of treatment options, cancer recurrence, and patient survival [2]. It should be noted that there are only several CTCs in 1 mL of blood, which contains huge number of haematological cells, such as around 5 × 109 erythrocytes, 1.0 × 107 leukocytes and 3 × 108 platelets. The extremely low level in blood causes great challenges to develop ultrasensitive methods for CTC detection [3]. Up to now, various methods for CTC detection have been reported, and most of the strategies include CTC capture, enrichment, release and detection. Due to the low abundance and fragile properties of CTCs in blood, a simple and reliable capture method is highly desirable to promote the recovery rate and detection accuracy. First, various affinity ligands, such as folic acid and antibodies against epithelial cell adhesion molecules (anti-EpCAM) have been conjugated on a solid support (e.g. magnetic beads) for specific enrichment of CTCs from peripheral blood [4]. Alternatively, aptamers have been used as functional ligands to specifically recognize not only small molecules and proteins but also entire cells [5]. Aptamers are single-stranded oligonucleotides and are employed as “chemical antibodies”, indicating advantages over antibodies in reproducible production, high stability and low immunogenicity [6]. Second, the enriched CTCs are further processed for detection from the whole cells or nucleic acids after cell lysis. The whole cells are detected via various methods such as microcantilever measurement, inductively coupled plasma mass spectrometry (ICP-MS), and flow cytometry, while the nucleic acids are analyzed via electrochemical analysis and real-time quantitative reverse-transcriptase polymerase chain reaction (RQ-PCR) [7]. The only commercial product for CTC detection is Cellsearch® Systems, where anti-EpCAM-coated magnetic nanoparticles capture CTCs followed by immunostainning to differentiate CTCs from normal cells. The detection method has not been widely accepted due to the time-consuming and complicated enrichment with low selectivity, and the immunostainning suffers from cumbersome processing, strict conditions and high cost [7]. It should be noted that fluorescent methods have recently attracted many attentions due to their simplicity and effectiveness. Cui et al. developed a turn-on fluorescent detection system composed of graphene quantum dots and molybdenum disulfide nanosheets. The binding of quantum dots with EpCAM of CTCs could separate the probe from molybdenum disulfide, leading to the fluorescence recovery [8]. Self-propelled micromotors could provide efficient fluid mixing and have been employed for the capture, pickup and transport of target substances. The propulsion mechanisms of micromotors are mainly resulted from the conversion of chemical or external energy into mechanical motion [9]. External energy sources such as high magnetic fields, strong electric fields and ultrasound have been utilized to control the movement of nanomotors, but these techniques should be localized to the hospital environment and require specialists and equipment [10]. The self-propulsion of micromotors based on a catalytic mechanism of bubble propulsion has received a lot of attention [11]. One of the sensing strategies is based on the tracking of moving speed in the presence of target substances. Yu et al. designed micromotors with capture antibodies to selectively recognize the target protein, which could bind microspheres with the secondary antibodies. The loading of microspheres should slow down the micromotor motion, and the motion velocity reflected the protein levels and the amount of microsphere

2. Materials and methods 2.1. Materials Polystyrene-co-maleic anhydride (PSMA, Mw: 170 kDa; Maleic 2

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Scheme 1. (a) Preparation of self-propelled JMs. JRs were constructed by cryocutting of aligned Janus fibers prepared by side-by-side electrospinning (1). The PSMA side of JRs was grafted with aptamer (2), and the PSMA-HDA-Boc side was treated with trifluoroacetic acid to remove Boc groups (3), followed by PEI conjugation (4) and CAT immobilization (5). The resulting JRs were grafted with TPE-T and FITC-G fluorophores to obtain JMs (6). (b) Ratiometric fluorescence response of JMs after capture of CTCs. The binding between fluorophores with aptamers on JMs resulted in the ACQ effect of FITC-G and AIE of TPE-T, leading to blue fluorescence emission. In the presence of CTCs, the fluorescence of TPE-T was weakened rapidly and that of FITC-G was restored to emit green fluorescence.

anhydride content: 14.8 wt%) and polystyrene (PS, Mw: 400 kDa) were purchased from Shanghai Zhaocheng Scientific Development Corp. (Shanghai, China). Deuterated dimethyl sulfoxide (DMSO‑d6), toluidine blue O (TBO), TiCl4, azobis(isobutyronitrile) and N-bromobutanimide were procured from Heowns Biochemical Technology Co., Ltd. (Tianjin, China). Poly(ethylene imine) (PEI, Mw: 25 kDa), N-tert-butoxycarbonyl1,6-hexanediamine, 4-methylbenzophenone and di-tert-butyldicarbonate (Boc2O) CAT from bovine liver (EC 1.11.16) were supplied from Sigma (St Louis, MO). TLS11a aptamer (5′-ACAGCATCCCCATGTGAA CAATCGCATTGTGATTGTTACGGTTTCCGCCTCATGGACGTGCTG-3′) with terminated 5′ amino groups was obtained from Shanghai Sheng Bioengineering Co., Ltd. (Shanghai, China), and sulfo-cyanine5 NHS ester and BCA reagent for protein assays were from Invitrogen (Shanghai, China). 4-Dimethylaminopyridine, benzophenone, guanine, thymine, rhodamine B, coumarin 6 and FITC were purchased from Shanghai Aladdin Reagent Co., Ltd. (Shanghai, China). DMEM and fetal bovine serum (FBS) were obtained from Gibco (Waltham, MA). All other chemicals were received from Changzheng Regents Company (Chengdu, China), unless otherwise indicated.

chloroform at 18% (w/w). The two polymer solutions were loaded into a side-by-side spinneret containing two parallel metal capillaries [21]. The electrospinning was performed at a flow rate of 2 mL/h under a high voltage difference of 18 kV/15 cm and deposited onto a rotating frame cylinder. The collected aligned fibers were embedded in a cryomold at −20 °C, and frozenly sectioned at thickness of 2, 5, 10 and 20 μm, followed by ultrasonication and centrifugation to obtain JRs [22]. 2.3. Synthesis of TPE-T Fig. 1a shows the synthesis process of TPE-T by conjugation of thymine with 4-methyl-tetraphenyl ethylene (TPE-CH3). Briefly, zinc dust (3.6 g, 55 mmol), 4-methylbenzophenone (2.94 g, 15 mmol) and benzophenone (2.73 g, 15 mmol) were mixed in 100 mL of dry tetrahydrofuran (THF) under argon atmosphere, followed by injection of TiCl4 (3.3 mL, 30 mmol). The mixture was refluxed overnight and the reaction was terminated by adding 10% K2CO3 solution, followed by extraction with ethyl acetate. The organic layer was dried over anhydrous sodium sulfate and purified by silica gel column chromatography using dichloromethane/petroleum ether (1/80, v/v) as eluent to obtain a white solid of TPE-CH3 [23]. Yield: 63%. The 1H NMR spectrum was recorded on a Bruker AM 400 apparatus, using tetramethylsilane as the internal reference. 1H NMR (CDCl3, ppm, δ): 7.27–7.14 (m, 6H), 7.14–6.91 (m, 8H), 6.88–6.81 (m, 5H), 2.23–2.17 (m, 3H). TPE-CH3 (0.7 g, 2 mmol), N-bromobutanimide (0.712 g, 4 mmol) and a catalytic amount of azobis(isobutyronitrile) were mixed in 20 mL of CCl4 and heated to 80 °C under argon atmosphere. After reaction for 10 h, the mixture was filtrated and the filtrate was concentrated by rotary evaporation. The crude product was purified by silica gel column chromatography using dichloromethane/hexane (1/4, v/v) as eluent to obtain a faint yellow solid of TPE-Br [23]. Yield: 50%. 1H NMR (CDCl3, ppm, δ): 7.37–7.23 (m, 6H), 7.22–7.03 (m, 8H), 7.02–6.94 (m, 5H), 4.70–4.62 (m, 2H). TPE-Br (213 mg, 0.5 mmol), thymine (194 mg, 1.5 mmol) and K2CO3 (207 mg, 1.5 mmol) were mixed in dry dimethylformamide

2.2. Preparation of JRs Janus fibers were constructed by side-by-side electrospinning [21], followed by cryocutting into JRs with different lengths (Scheme 1). One side of JRs consisted of mixtures of PSMA and PS to introduce different amount of anhydride groups, and N-tert-butoxycarbonyl-protected PSMA-HDA polymers (PSMA-HDA-Boc) was used as the matrix polymer of the other side. Briefly, N-tert-Butoxycarbonyl-1,6-hexanediamine (0.22 g, 1 mmol) in dichloromethane (5 mL) was added dropwise to a dichloromethane solution (20 mL) of PSMA (3.4 g, 0.2 mmol) under argon atmosphere. After reaction for 48 h, the mixture was concentrated under reduced pressure, precipitated with methanol and vacuum dried to give PSMA-HDA-Boc. PSMA/PS mixtures with different PSMA fractions of 60%, 70%, 80%, 90%, 95% and 100% were dissolved in chloroform at a concentration of 14% (w/w), while PSMA-HDA-Boc was dissolved in 3

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Fig. 1. (a) Synthetic route of TPE-T. TPE-T was prepared by conjugation of thymine with bromo-substituted 4-methyl-tetraphenyl ethylene (TPE-CH3), which was synthesized from 4-methylbenzophenoneand benzophenone via McMurry reaction. (b) Synthetic route of FITC-G. FITC-G was prepared from FITC and Boc-guanine, which was synthesized from Boc-protected glyoxal-guanine. (c) 1H NMR spectra of TPE-T and (d) FITC-G. (e) Fluorescence spectra of TPE-T and (f) FITC-G after incubation with aptamer solutions from 0 to 1.0 μM.

(15 mL) and refluxed under argon for 24 h. After solvent removal under reduced pressure, the residue was purified by silica gel using dichloromethane/ethyl acetate (3/1, v/v) as eluent to give a white powder of TPE-T. Yield: 38%. 1H NMR (CDCl3, ppm, δ): 9.85 (s, 1H), 7.68 (s, 1H), 7.18–6.82 (m, 19H), 4.48 (m, 2H), 1.57 (m, 3H) (Fig. 1c).

Boc-protected glyoxal-guanine (0.4 g, 0.67 mmol) was dissolved in 25 mL of THF/water (1/1, v/v), followed by adding 6 mL of NH4OH and stirring at room temperature for 4 h. The excess ammonia and THF was removed by rotary evaporation, and the white precipitate was washed with water to give a gray powder of Boc-guanine. Yield: 97%. 1H NMR (DMSO‑d6, ppm, δ): 7.94 (s, 1H), 1.38 (s, 9H). Boc-guanine (126 mg, 0.5 mmol), FITC (100 mg, 0.8 mmol) and K2CO3 (207 mg, 1.5 mmol) were mixed in dry dimethylformamide (20 mL) and refluxed for 8 h under argon, followed by solvent evaporation under reduced pressure. To remove Boc groups, the resulting precipitate was dispersed in deionized water (5 mL), followed by adding dropwise trifluoroacetic acid (1 mL). After stirring for 8 h at room temperature, the mixture was concentrated by rotary evaporation and the residue was purified by silica gel column chromatography using methanol/ethyl acetate (5/1, v/v) as eluent to give a yellow powder of FITC-G. Yield: 21%. 1H NMR (DMSO‑d6, ppm, δ): 8.87 (s, 1H), 8.55 (s, 1H), 8.06 (m, 2H), 7.69 (s, 1H), 7.51 (s, 1H), 7.18(m, 2H), 6.73–6.53 (m, 4H), 4.82 (t, 2H) (Fig. 1d).

2.4. Synthesis of FITC-G Fig. 1b shows the synthesis process of FITC-G by conjugation of FITC and Boc-guanine, which was prepared from Boc-protected glyoxalguanine by Boc deprotection. Briefly, glyoxal monohydrate (0.87 g, 18 mmol), guanine (0.155 g, 1 mmol) and glacial acetic acid (0.5 mL) were added in 100 mL of water, followed by stirring overnight at 60 °C. After kept at 5 °C for 2 days, the suspension was filtered, and the residue was water washed and vacuum dried to obtain a white powder of glyoxal-guanine with a yield of 92%. Glyoxal-guanine (0.5 g, 2.4 mmol), 4-dimethylaminopyridine (1.5 mg, 0.12 mmol) and Boc2O (3 mL, 13.1 mmol) were added into dry THF (33 mL), and the reaction was proceeded at room temperature for 5 days, followed by filtration to collect the filtrate. After removal of excess THF by rotary evaporation, a light brown oil was purified by silica gel column chromatography using ethyl acetate/hexane (7/3, v/v) to afford Boc-protected glyoxal-guanine. Yield: 75%. 1H NMR (DMSO‑d6, ppm, δ): 1.72–1.33 (m, 27H), 8.09 (s, 1H), 6.69–6.58 (m, 2H), 1.71–1.32 (m, 27H).

2.5. Fluorescence intensity changes of TPE-T and FITC-G in the presence of aptamer The AIE and ACQ properties of TPE-T and FITC-G were determined from the fluorescence intensity changes after incubation with different 4

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concentrations of aptamers. Briefly, TPE-T solution (5 μM) in THF was added into a mixture of ethanol/water (36/64, v/v), and then incubated with aptamers from 0 to 1.0 μM. The emission spectra of the obtained suspensions were measured by a fluorescence spectrophotometer (Hitachi, Japan, F-7000 type) at the excitation/emission wavelengths of 365/450 nm. The ACQ property of FITC-G was also examined following the same procedures, and the fluorescence intensities were measured at the excitation/emission wavelengths of 365/ 526 nm.

concentration added. The aptamer concentration was detected by an UV–vis spectrophotometer at 260 nm [24], and the CAT concentration was determined by a commercial BCA protein assay kit. The activities of free and immobilized CAT were determined after incubation with H2O2 solution for 10 min at 25 °C. One activity unit of CAT was defined as the amount of enzyme that consumed 1 μmol of H2O2 per minute, and the H2O2 concentration was measured by an UV–vis spectrophotometer at 240 nm [28]. The effect of pH and temperature on the enzyme activities was studied after incubation in buffer from pH 5.0 to pH 10.0 at 37 °C or in pH 7.4 buffers at temperatures ranging from 25 to 45 °C. To display the conjugation of CAT and aptamer on the opposite sides of JRs, sulfo-syanine5 NHS ester-labeled CAT was employed in the grafting process and the grafted aptamer was labeled by TPE-T. Briefly, CAT (3 mg, 0.012 μmol) and sulfo-cyanine5 NHS ester (10 μg, 0.013 μmol) were mixed in HEPES buffer (250 μL) for 6 h, followed by dialysis for 2 days against water and grafted on JRs as above. In addition, JMs (0.2 mg) were incubated in 200 μL of Tris-HCl buffer containing 0.02 mg of TPE-T for 1 h at 37 °C, followed by removal of the unbound TPE-T by intensive rinsing with Tris-HCl buffer [29]. The obtained JMs were observed with CLSM under the excitation/emission wavelengths of sulfo-syanine5 NHS ester at 646/662 nm and TPE-T at 365/450 nm.

2.6. Surface functionalization of JRs Scheme 1 summarizes the surface grafting of CAT and aptamers on the opposite sides of JRs. Aptamers were conjugated on the PSMA side by reaction of maleic anhydride with amino terminals of aptamers, and CAT was immobilized on the PSMA-HDA side via PEI linking. Briefly, JRs were suspended in Tris-HCl buffer (1 μM) containing aptamers at a final concentration of 1 mg/mL, followed by stirring at 25 °C for 3 h [24]. After water washing, JRs were collected by centrifugation and then immersed in ethanolamine solution (100 mM) to deplete the unreacted maleic anhydride on the surface, followed by water washing and drying under argon stream to get aptamer-grafted JRs. To remove Boc groups of the other side (PSMA-HDA-Boc), JRs were suspended in deionized water (10 mL), followed by adding dropwise trifluoroacetic acid (2 mL). After stirring for 12 h, the resulting JRs were washed repeatedly with water and suspended in 100 mM phosphate buffer solution (PBS, pH 7.4) containing 3% (v/v) glutaraldehyde for 4 h at room temperature. After repeated washing with PBS and water, the glutaraldehyde-activated JRs were incubated in PEI solution (5.0 mg/mL) for 5 h at 25 °C. The PEI-conjugated JRs were washed with water and added into PBS containing 0.75 mg/mL CAT for 12 h at 4 °C. After incubation with sodium borohydride solution (1 mg/mL) for 1 h at 25 °C, JRs were washed with PBS and water and vacuum dried to give aptamer and CAT-grafted JMs [25]. To graft TPE-T and FITC-G, JMs (1 mg) were dispersed in an ethanol solution (40%) containing TPE-T (5 mg/mL) for 1 h, followed by flushing with ethanol solution. The resulting JMs were collected by centrifugation and incubated with FITC-G (5 mg/mL) for 1 h, followed by washing with ethanol solution and vacuum drying to obtain TPE and FITC-labeled JMs.

2.8. Motion analysis of JMs The movement of JMs was observed with a light microscope (Leica DMI4000, Germany) after incubation in 3.5% H2O2 in PBS. Each video of the JM motion was recorded with a CCD camera at a phase difference of about 25 fps. All videos were recorded for the first 10 s to ensure a relatively constant fuel concentration and enzymatic reaction rate. The trajectory was tracked using Image J to measure the moving speed. The trajectory was also extracted and input into Matlab to analyze the mean-square-displacement (MSD) values according to the aforementioned procedure [30]. The motion profiles were detected on JMs with different aspect ratios and after incubation with different H2O2 concentrations (0–4.5%, w/w). 2.9. Cell detection with JMs

2.7. Characterization of JRs and JMs

HepG2 (human liver tumor cells), H22 (mouse liver tumor cells), 4T1 (mouse breast tumor cells) and ECs (human umbilical vein endothelial cells) were obtained from American Type Culture Collection (Rockville, MD) and maintained in DMEM containing FBS. To investigate the specific detection of target cells (HepG2), JM suspensions (1 mg/mL) in PBS containing 3.5% (w/w) H2O2 were mixed in cell suspensions (104 and 106 cells/mL). After incubation for different time periods, the fluorescence spectra of the mixed suspension were recorded by a fluorescence spectroscope at the excitation wavelengths of 365 nm. The fluorescence intensities were obtained at the emission wavelengths of 450 and 526 nm for TPE and FITC, which were named as I450 and I526, respectively. In order to obtain the optimal detection conditions, the effect of JM lengths, aptamer grafting densities and H2O2 concentrations was investigated on the fluorescence intensity ratios (I526/I450). In addition, cells were harvested and fixed in 4% paraformaldehyde and freeze-dried, and the cell capture by JMs was observed by SEM as above. The selectivity of HepG2 assay was evaluated after incubation with JMs in the presence of 4T1, H22, ECs cells or the mixture of HepG2 and H22 cells at 106 cells/mL. The stability of JMs was determined from the fluorescence emission changes after immersion in fresh blood. Bloods were collected from healthy volunteers in compliance with the relevant laws and institutional guidelines. In addition, a known amount of HepG2 cells were added into blood, and the I526/I450 ratios were detected as above to calculate the cell recovery rate.

The morphology of JRs and JMs was detected via scanning electron microscopy (SEM; FEI Quanta 200, The Netherlands), and the average diameter and length of JRs were measured from SEM images by using Photoshop 10.0 edition [26]. For visualization of individual compartment of JRs, small amounts (0.1%) of rhodamine B and coumarin 6 were mixed with PSMA and PSMA-HDA-Boc solution and JRs were prepared as above. The obtained JRs were observed with a confocal laser scanning microscope (CLSM, Leica TCS-SP2, Germany) under excitation/emission wavelengths of 540/625 nm for rhodamine B and 467/497 nm for coumarin 6. In order to determine the anhydrides group densities, JRs were immersed in 100 mM ethanolamine for 1 h at room temperature to form carboxyl groups, which were determined via colorimetric methods based on TBO binding [27]. Briefly, aqueous TBO solution (0.5 mM) was adjusted to pH 10 with 0.1 mM NaOH and added to suspensions of carboxylated JRs. After constant agitation for 5 h at room temperature, JRs were intensively rinsed with 0.1 mM NaOH solution to remove the noncomplexed TBO until no dye was detected in the washing solution. The complexed TBO on JRs was desorbed from the surfaces by incubating with acetic acid solution (50%, v/v) for 10 min under vortexing, followed by detection at 633 nm with an ultraviolet–visible (UV–vis) spectrophotometer (UV-2550, Shimadzu, Japan). The grafting amount of aptamer and CAT on JMs was determined from the residual in the reaction media and compared with the initial 5

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2.10. Statistics analysis

procedures showed no interference each other. Aptamers were grafted on one side of JMs to capture target cells and initiate the ratiometric fluorescent response of TPE and FITC. The AIE profiles of TPE-T and ACQ effect of FITC-G were determined from the emission spectra after interaction with aptamer grafts on JMs. As shown in Fig. 2c, TPE-T was barely fluorescent in the ethanol/water mixtures, and the addition of a small amount of JM-2 showed a dramatic increase in the fluorescence intensities at 450 nm. The interaction with aptamers led to the rotation restriction of phenyl groups of TPE-T, accompanied by a remarkable fluorescence emission. To prove the aptamer-induced ACQ effect of FITC-G, the fluorescence spectra were recorded in the absence and presence of JM-2 (Fig. 2c). The addition of JMs led to efficient fluorescence quenching of FITC-G at 526 nm.

The statistical significance of the data obtained was analyzed by the Student’s t test. Data are expressed as mean ± standard deviation (S.D.). Probability values of p < 0.05 were interpreted as denoting statistical significance. 3. Results and discussion 3.1. Characterization of TPE-T and FITC-G TPE-T was prepared by conjugation of thymine with TPE-CH3, which was synthesized from 4-methylbenzophenone and benzophenone via McMurry reaction (Fig. 1a). FITC-G was synthesized by coupling FITC on Boc-guanine, which was prepared from Boc-protected glyoxalguanine (Fig. 1b). The chemical structures of TPE-T and FITC-G were confirmed by 1H NMR spectra (Fig. 1c, d). Fig. 1e shows fluorescence properties of TPE-T after incubation with different aptamer concentrations. There was almost no emission detected for TPE-T, but the addition of aptamers led to a gradual increase in the emission intensities at 450 nm, exhibiting typical AIE phenomena. There were over 23-fold increases in emission intensities when the aptamer concentrations increased from 0.1 to 1.0 μM. The formation of hydrogen bonds with adenine induced the rotation restriction of phenyl groups thus leading to the fluorescence enhancement of TPE-T. As shown in Fig. 1e, the fluorescence intensity of FITC-G at 526 nm was weakened gradually with the increase of aptamer concentrations. FITC is a typical ACQ molecule [31], and the specific binding with cytosine on aptamers through hydrogen bonds formed an aggregated state to quench the fluorescence emission.

3.3. Characterization of CAT immobilization on JRs One CAT molecule is composed of four subunits and each subunit is a polypeptide chain with a molecular weight of around 65 kDa [32]. CAT is mainly inactivated by subunit dissociation during enzyme immobilization process [28]. In the current study, CAT was grafted on the PSMA-HDA side of JRs via branched PEI linkers. The immobilization amount of CAT was 0.14 mg/mg JRs, and the multipoint binding of enzyme molecules via PEI increased the grafting content [33]. The activity of immobilized CAT were around 4.65 U/mg, indicating the activity retention of around 72% compared with free CAT. The multipoint attachment with PEI was capable to stabilize both the quaternary structure of an enzyme molecule and the structure of each individual subunit, permitting the high stability and activity retention after immobilization [34]. The activities of immobilized enzyme were detected under different pH and compared with those of free enzymes. Fig. 2d shows the CAT activities of JMs after incubation in buffers of pH 5.0–10.0, indicating an optimal pH (7.0) for both free and immobilized CAT. When the buffer pH was below and beyond the optimal pH, significantly higher activities were detected for immobilized CAT than that of free enzyme. About 78% of activity retention was detected for immobilized enzyme at pH 10.0, while the free enzyme was fully inactivated. The effect of temperature on the activity retention was investigated in pH 7.0 buffers under 25–45 °C. As shown in Fig. 2e, similar bell-shaped curves were observed at an optimal temperature of 35 °C for both free and immobilized CAT. It was also evident that the immobilized CAT had a higher activity than free one at each temperature, and free CAT lost all the activity beyond 40 °C. Thus, the immobilized CAT of JMs could remain high catalytic activities after incubation under a broader range of temperatures and pH values than free enzyme.

3.2. Characterization of JRs Janus fibers were prepared by side-by-side electrospinning, and JRs were constructed by cryosection of aligned Janus fibers. Fig. 2a shows typical SEM images of JRs with different lengths, indicating an average diameter of 4.6 ± 0.3 μm. The cryocutting thickness was set from 2 to 20 μm to control JR lengths of 1.9 ± 0.2, 4.9 ± 0.3, 10.4 ± 0.7 and 19.8 ± 0.5 μm, and the aspect ratios were 0.4 (JR-0.4), 1 (JR-1), 2 (JR2) and 4 (JR-4), respectively. To visualize individual compartments and discern Janus morphologies, JRs were prepared by inoculation of rhodamine B and coumarin 6 in the respective electrospinning solutions. The inset of Fig. 2a shows CLSM images of JRs with different aspect ratios, displaying a common feature of red (rhodamine B) and green (coumarin 6) compartments. There was a thin yellow line between the two compartments, indicating a well-defined interface. The homogeneous color distribution indicated that the side-by-side electrospinning of two solutions was a feasible method to fabricate integral Janus structures with different phases. CAT and aptamers were grafted on the opposite sides of JRs to construct JMs, followed by loading TPE-T and FITC-G via aptamers on the JM surface (Scheme 1). Fig. 2b displays typical SEM images of JMs with different aspect ratios, showing no apparent morphological changes after grafting with functional moieties. The grafting of enzyme and aptamers on JMs led to water swelling of JRs and a slight increase in the average diameter to 5.0 ± 0.4 μm. The surface modification of JMs was confirmed by CLSM after fluorescently labeling of CAT and aptamer. CAT was labeled with sulfo-cyanine5 NHS and grafted on one side of JRs to reveal red fluorescence. The aptamer grafts on the other side of JRs were labeled with TPE-T to reveal blue fluorescence due to the inherent recognition ability. As shown in the inset of Fig. 2b, the compartmental distribution of red and blue fluorescence was clearly observed for JMs with different aspect ratios. Thus, CAT and aptamer should be grafted on the respective sides of JRs and the modification

3.4. Self-propelled motion profiles of JMs Asymmetric structure is usually created in micromotors to achieve one-way thrust to achieve the purpose of movement. In the current study, CAT was immobilized on one side of JRs to catalyze the decomposition of H2O2 for generating oxygen bubbles, which were utilized as the “engine” to provide the propulsion force [35]. The motion profiles of JMs with different aspect ratios were observed after incubation in H2O2 solutions, and the videos are included in the Supplementary Information. Fig. 3a shows typical tracking trajectories of JMs in the presence of 3.5% H2O2 during 4 s, indicating that the motion behaviors were highly dependent on the aspect ratios. For JMs with cylindrical structures, CAT was immobilized on both the side and cross-sectional areas, and the self-propelling force came from three directions. JM-0.4 had much larger areas of the upper and lower crosssections than those of the side areas, leading to the random motion (Supplementary Video 1). The spiral motion of JM-1 (Supplementary Video 2) was resulted from the deviation of the propulsion force from

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Fig. 2. (a) SEM images of JRs with different aspect ratios. Insets are CLSM images of JRs loaded with rhodamine B (red) and coumarin 6 (green) in the respective PSMA and PSMA-HDA-Boc sides. (b) SEM images of JMs with different aspect ratios. Insets are CLSM images of JMs after labeling CAT with sulfo-Cyanine5 NHS (red) and grafting aptamer with TPE-T (blue). Bars represent 20 μm. (c) Fluorescence spectra of TPE-T and FITC-G in a mixture of ethanol and water (36/64, v/v) and in the presence of JM-2. (d) The effect of pH and (e) temperature on the catalytic activities of free and immobilized CAT (n = 3). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Fig. 3. (a) Typical motion trajectories of JMs with different aspect ratios in 3.5% H2O2 during 4 s. (b) Average velocities and (c) MSD analysis from tracking trajectories of JMs with different aspect ratios in 3.5% H2O2 (n = 3). (d) Average velocities and (e) MSD analysis from tracking trajectories of JM-2 after incubation with H2O2 of different concentrations (n = 3).

the side and cross-sections [36]. Similarly, the increase in JM lengths produced a far larger driving force from the fiber side than that from the cross-sectional area. JM-2 showed both rotational and translational motion due to the combined propulsions (Supplementary Video 3). The

side area of JM-4 was much larger than that of cross sections, leading to driving force generated mainly at one side of the structure. The unidirectional propulsion force caused the motion trajectory close to a straight line (Supplementary Video 4). 7

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Supplementary Video 1.

Supplementary Video 2.

Supplementary Video 4.

Supplementary Video 3.

Fig. 3b summarizes the average moving velocities of JMs with different aspect ratios. The movement speed of JMs decreased from 69 to 19 μm/s accompanied by the increase of aspect ratios from 0.4 to 4. JMs with larger aspect ratios moved perpendicularly to the long axis and 8

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Fig. 4. (a) Anhydride group and aptamer densities on the surface of JM-2 with different PSMA fractions (n = 3). (b) I526/I450 values of JM-2 containing different amounts of PSMA after incubation with HepG2 cells (n = 3). (c) I526/I450 values of JM-2 suspensions after incubation for 70 s with HepG2 at 104 and 106 cells/mL (n = 3). Insets are photographs of JM-2 suspensions under the illumination of a hand-held UV lamp before and after incubation with HepG2 (106 cells/mL) in 3.5% H2O2. (d) I526/I450 values of JM suspensions with different aspect ratios after incubation with HepG2 cells in 3.5% H2O2. (e) I526/I450 values of JM-2 suspensions after incubation with HepG2 cells at different H2O2 concentrations (n = 3). (f) SEM image of JM-2 after capture of HepG2 cells.

experienced a larger drag, resulting in a lower average velocity. Reddy et al. constructed Pt/Pd-Au Janus fibers via electrospinning and sputtering technique, followed by ultrasonication to produce short nanorods. The average velocity was 1–2 μm/s in 7.5% H2O2 solution, lower than those of current JMs decorated with enzyme. It was suggested that the higher density after platinum coating led to sink onto the bottom and experience a larger frictional drag [37]. As an important motion indicator of JMs, the MSD magnitude reflected the moving range and interactions with other substances in the media [38]. The position vector of moving particles determines MSD, and the MSD magnitude depends on the trajectory and average velocity [39]. Fig. 3c shows MSD values of JMs with different aspect ratios as a function of time intervals. There were no apparent MSD increases for JM-0.4 and JM-1 during 4 s, due to the random and spiral motion (Fig. 3a). The trajectories of JM-2 and JM-4 tended to be directional motion as evidenced by their linear MSD curves. The MSD value decreased with the increase in the aspect ratios due to the higher velocity of JM-2 (41 μm/s) than that of JM-4 (19 μm/s). The MSD should reflect the active range of JMs during a period of time and affect the capture efficiency of cells. The propulsion relied on the ejection of the oxygen bubbles generated from the biocatalytic decomposition of H2O2 by CAT on one side of JMs. Thus, the propelling force was dependent on the fuel concentrations, besides the amount of CAT grafted. Fig. 3d shows the moving velocity of JM-2 after incubation with H2O2 of different concentrations. There were significant increases from 14 μm/s at 1.5% H2O2 to 42 μm/s at 3.5% H2O2. At H2O2 concentrations higher than 4.5%, JM-2 displayed a slow speed because of the inactivation of CAT by a high level of hydroxyl radicals [40]. In addition, the MSD values increased with H2O2 concentrations (Fig. 3e), indicating that the enzymatic catalysis led to a fuel concentration-dependent motion.

strand DNA to bind with aptamers, double-stranded DNA was denatured and complementary strand DNA was released [41]. As shown in Scheme 1, the binding amount of TPE-T and FITC-G on the aptamer determined the initial and final fluorescence intensities of JMs after treatment with HepG2 cells. Thus, the optimal fluorescent responses of JMs could be achieved by tuning the grafting amount of aptamers. Aptamers were grafted onto PSMA sides of JRs via reaction with maleic anhydride groups. PSMA/PS mixtures at PSMA fractions of 100%, 95%, 90%, 80%, 70% and 60% (w/w) were used to adjust the aptamer densities. Fig. 4a shows the densities of anhydride groups and aptamers on JRs, indicating a gradual increase with the increase in the PSMA fractions. It should be noted that the density of surface anhydride groups was just 8% for JRs containing 60% of PSMA in the fiber matrix. The electrospinning process usually caused the distribution of chemical groups with low binding energy on the surface of fibers [42]. The surface densities of anhydride group indicated a rapid increase from 7.8 to 105.3 mmol/mg when the PSMA fractions in the fiber matrices increased from 60% to 100%. As expected, amino-terminated aptamers were sufficiently reacted with anhydride groups, indicating the increase of aptamer densities from 1.3 to 14.8 mmol/mg. Fig. 4b shows I526/I450 values of JM-2 suspensions with different aptamer densities after treatment with HepG2 cells (106 cells/mL). The specific interaction of HepG2 cells with aptamers led to stronger fluorescence restoration of FITC-G at 526 nm and concomitant decrease of TPE-T at 472 nm, resulting in significantly higher I526/I450 values. However, when the PSMA fraction increased to over 95%, there was no significant increase in the I526/I450 values in the presence of HepG2. A higher aptamer grafting density could achieve highly binding amount of TPE-T and FITC-G, but the excess amount of aptamer/TPE-FITC complexes on the JM surface led to less effective release of fluorescent molecules. It was suggested that HepG2 and aptamer grafts reached saturated interactions and no significant change in I526/I450 values when the PSMA fraction was over 95%. Thus, taking into account the ratiometric fluorescence assay of JM-2 in the presence of HepG2, the PSMA/PS ratios of 95/5 was used to construct JMs with a grafting density of aptamers at around 12 nmol/mg. Fig. 4c summarizes the time-dependent I526/I450 changes of JM-2

3.5. Fluorescence response of JMs in the presence of cells In the current study, HepG2 cells were used as the tumor cell model, and TLS11a aptamer was applied to recognize HepG2 with a high affinity [20]. When HepG2 cells competed with the complementary 9

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Fig. 5. (a) Fluorescent spectra and (b) I526/I450 ratios for JM-2 suspensions after incubation with HepG2 cells at different concentrations (n = 3). (c) I526/I450 ratios for JM-2 suspensions after incubation with blood, HepG2, 4T1, H22 and ECs cells and the cell mixtures (n = 3). (d) I526/I450 ratios for JM suspensions after incubation with HepG2 cells at different concentrations in the absence and presence of H22 (n = 3). (e) Recovery rate of HepG2 cells after adding into blood samples and detected by JM-2 (n = 3).

after incubation with HepG2 at concentrations of 104 and 106 cells/mL. The I526/I450 values increased during the incubation time period, and the increase was less significant after incubation for 1 min, which was set as the time frame for cell incubation. In addition, the increase in I526/I450 ratios became more dramatic under higher HepG2 concentrations, suggesting the potential of JM-2 as the ratiometric fluorescence probe for HepG2 monitoring. The inset of Fig. 4c shows the dramatic fluorescence contrast of JMs after incubation with HepG2 under an ultraviolet lamp. The JM-2 suspension emitted obvious blue fluorescence before treatment with HepG2, but the emission showed a distinct change to bright green fluorescence after incubation with HepG2 at 106 cells/mL. As indicated above, JMs with different aspect ratios had different motion profiles and should affect interactions with HepG2 cells. Fig. 4d shows the I526/I450 values of JMs with different aspect ratios after incubation with HepG2 cells. Compared with that without cell treatment at 0.55, I526/I450 values indicated a rapid increase with the increase in the aspect ratio up to 2. This result was consistent with the MSD of JMs (Fig. 3c), and a large MSD increased the probability of cell recognition and capture, resulting in an increase in I526/I450 ratios. Similarly, I526/I450 values of JM-2 were detected under different H2O2 concentrations to examine the effect of JM motions. As shown in Fig. 4e, I526/I450 ratios showed significant increases in the H2O2 concentration range of 0–3.5%, which was consistency to the motion profiles of JMs (Fig. 3e). Thus, self-propelled JM-2 could promote the capture of cells to increase the fluorescence intensity, and 3.5% of H2O2 was the optimal detection concentration. Fig. 4f shows SEM image of JM-2 after incubation with HepG2 cells, displaying tight integration with JMs.

significant changes when HepG2 concentrations were higher than 106 cells/mL, resulted from saturated interactions between cells and JMs. To further quantify HepG2 concentrations, I526/I450 ratios were plotted as a function of HepG2 concentrations. As shown in Fig. 5b, I526/I450 values rapidly increased from 0.55 to 2.46 when HepG2 concentrations gradually increased and reached a plateau at 106 cells/mL. I526/I450 values were drafted against HepG2 concentrations and fit into the following equation:

Y = 2.83 + ( 2.43/(1 + (X/10496)0.3)) (R2 = 0.991) where Y represents I526/I450 values and X HepG2 concentrations (0–106 cells/mL). Thus, HepG2 concentrations can be obtained from I526/I450 values using the above equation, and the limit of detection (LOD) was around 25 cells/mL calculated from the signal-to-noise of 3 [43]. Chen et al. used TPE-labeled aptamers for specific binding with human Burkitt’s lymphoma cells, and the AIE-induced fluorescence light-up indicated an LOD of around 100 cells/mL [44]. Chang et al. developed aptamer-functionalized 1,3-phenylenediamine resin nanoparticles, and the fluorescence analysis after incubation with human acute lymphoblastic leukemia cells and human Burkitt’s lymphoma cells for 10 min indicated LOD at 44 and 79 cells/mL, respectively [45]. Thus, the LOD of the current JMs was lower than those from fluorescence “turn-on” assays toward cancer cells. Compared to static probes, the mobile JMs led to more interactions with cells, thereby not only significantly enhancing the selective capture of cells but also shortening the response time. Furthermore, the ratiometric assay was derived from the intensity ratios of dual channels, and the fluorescence background could be minimized, allowing accurate and reliable detection. In addition, the interaction area and strength between aptamers and cells should affect the fluorescence changes [46]. It should be noted that the membrane receptor density in CTCs usually correlates with the tumor stage of development [47], which provides biomarkers for predicting tumor progression but may affect the detection sensitivity in the initial stage. Thus, it is essential to understand the characteristics and evolution profiles of a specific type of tumor cells, in order to design specific aptamers and establish a well-fitted equation.

3.6. Quantitative detection of HepG2 cells by JMs HepG2 cells were determined from I526/I450 values after incubation with JM-2 in 3.5% of H2O2 for 1 min. Fig. 5a summarizes dual-channel fluorescence responses toward different HepG2 concentrations. As the HepG2 concentration increased, the emission of FITC-G at 526 nm increased with a simultaneous decrease in the emission of TPE-T at 450 nm. The fluorescence intensities from the dual channels showed no 10

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3.7. Selectivity and anti-interference of HepG2 cell detection by JMs

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The important technical indicators for CTC detections include the selective and recovery rate [7]. The selective detection of HepG2 by JMs was evaluated using 4T1, H22 and ECs as interfering cells. As shown in Fig. 5c, JM-2 suspensions showed no significant changes in I526/I450 values after incubation with 4T1, H22 and ECs cells. However, there was an almost 5-fold increase in I526/I450 after incubation with HepG2 cells. In addition, the presence of 4T1, H22 and ECs did not interfere the specific binding between HepG2 and JM-2. Fig. 5d summarizes the variation of I526/I450 ratios in the presence of HepG2 and HepG2/H22 mixtures at different concentrations. The addition of H22 cells resulted in no significant change in I526/I450 values, demonstrating the high selectivity of JMs toward HepG2 over interfering cells. The stability of JM-2 was detected after immersion in blood, and the I526/ I450 ratio remained no significant change compared with the initial value (p > 0.05). It was indicated that the base-pair interactions for labeling TPE-T and FITC-G on aptamers were stable in the presence of blood cells and proteins. In addition, known amount of HepG2 cells were added into blood samples, and then detected by JMs as above to examine the recovery rate. As shown in Fig. 5e, the detection values were close to the number of cells added, and the recovery rates were all over 95%. Thus, HepG2 cells in each blood sample were almost identified. In addition, this cell detection procedure was completed within 1 min, showing the great potential for rapid clinical screening of cancer cells. 4. Conclusion JMs were constructed via CAT grafting on one side of JRs to catalyze the decomposition of H2O2 and provide sufficient propulsive force. Aptamers were conjugated onto another side, followed by labeling of TPE-T and FITC-G to achieve ratiometric fluorescence detection of target cells. JMs with different aspect ratios shows random, spiral, rotational or linear motion trajectories, and JM-2 indicated rotational motion with significantly larger MSD than other JMs. The detection efficiency of JMs is closely related to the aspect ratios of JMs, aptamer densities and H2O2 concentrations. The capture of CTCs by JMs displays apparent changes from blue to green fluorescence. The JM-2 detection demonstrates low detection limit, rapid recognition, high selectivity and large recovery rate in blood samples. It is suggested that the continuous mixing induced by the autonomous movement of JMs enhances the CTC capture and the ratiometric fluorescence signals promote the detection sensitivity. Thus, the motion-capture-ratiometric fluorescence sensing features of JMs provide a feasible strategy for rapid, real-time and sensitive monitoring of cells. Acknowledgements This work was supported by National Natural Science Foundation of China (31771034 and 51803015), the Key Research and Development Program of Sichuan Province (2018SZ0348), and the Analytical and Testing Center of Southwest Jiaotong University for SEM and CLSM analysis. References [1] S. Wang, C. Zhang, G. Wang, B. Cheng, Y. Wang, F. Chen, Y. Chen, M. Feng, B. Xiong, Aptamer-mediated transparent-biocompatible nanostructured surfaces for hepotocellular circulating tumor cells enrichment, Theranostics 6 (2016) 1877–1886. [2] Y. Ming, Y. Li, H. Xing, M. Luo, Z. Li, J. Chen, J. Mo, S. Shi, Circulating tumor cells: from theory to nanotechnology-based detection, Front. Pharmacol. 8 (2017) 35. [3] D.H. Moon, D.P. Lindsay, S. Hong, A.Z. Wang, Clinical indications for, and the future of, circulating tumor cells, Adv. Drug Deliv. Rev. 125 (2018) 143–150. [4] L. Rao, Q.F. Meng, Q. Huang, Z. Wang, G.T. Yu, A. Li, W. Ma, N. Zhang, S.S. Guo, X.Z. Zhao, Platelet–leukocyte hybrid membrane-coated immunomagnetic beads for highly efficient and highly specific isolation of circulating tumor cells, Adv. Funct.

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