Biosensors and Bioelectronics 162 (2020) 112261

Available online 6 May 20200956-5663/© 2020 Elsevier B.V. All rights reserved.

In-situ monitoring of glucose metabolism in cancer cell microenvironments based on hollow fiber structure

Zhen Ma a, Ying Luo a,d, Qin Zhu a, Min Jiang a, Min Pan c, Tian Xie a,**, Xiaojun Huang b, Dajing Chen a,*

a Medical School, Hangzhou Normal University, China b MOE Key Laboratory of Macromolecular Synthesis and Functionalization, Department of Polymer Science and Engineering, Zhejiang University, China c College of Biomedical Engineering & Instrument Science, Zhejiang University, China d School of Traditional Chinese Medicine, Guangdong Pharmaceutical University, China

A R T I C L E I N F O

Keywords: Hollow fiber Cancer cell Metabolism microenvironment Glucose sensor

A B S T R A C T

Cancer cells alter their metabolism to promote rapid proliferation, resulting in significant amounts of glucose to be used for aerobic glycolysis in the tumor microenvironment. Due to the spatial mismatch between electrodes and cells, existing methods for evaluating cancer cell metabolism lack kinetic and microenvironmental infor-mation. In this paper, we present a hollow fiber structure loaded with sensing elements as a long-term cellular metabolism monitoring platform. The unique gradient porous structure allowed the glucose sensor to be close to cultured cells but not directly in contact to prevent adverse effects. The liquid exchange channels in the porous fiber structure ensured continuous and real-time monitoring of glucose concentration changes in the culture media. Experimental results showed high electrochemical sensitivity and stability for continuously monitoring the glucose consumption rate. Furthermore, a continuous three-day long test quantified the change in glucose consumption in response to the anticancer drug Osimertinib. In addition to traditional endpoint cell counting and analysis, this hollow fiber sensing structure provided a real-time monitoring tool for cell metabolism. Such continuous monitoring of the cell metabolism microenvironment improves in-vitro toxicology models for personalized medicine and cancer therapy.

1. Introduction

The fundamental abnormality of cancer is the uncontrolled prolif-eration of cancer cells due to genetic mutations altering cell signaling pathways (DeBerardinis and Chandel, 2016). Rather than responding appropriately to the growth factors that control normal cell behavior, cancer cells grow and divide in an uncontrolled manner. Therefore, the energy demand of the rapidly proliferating cancer cells results in higher glucose consumption compared to normal cells (Fadaka et al., 2017; Meadows et al., 2008). Monitoring cell metabolism and cell growth could provide insights into applications including drug screening, cell therapy, and toxicity tests (Hanahan and Weinberg, 2011; Pavlova and Thompson, 2016). Direct methods to determine the cell proliferating status include manual counting, automated image analysis, electronic particle counting, in situ biomass monitoring and dielectrophoretic cytometry, most of which are accurate but only provide single data

points at the end of cell culture (Butler et al., 2014; Opel et al., 2010). Indirect methods for assessing cell growth include the chemical analysis of cellular metabolic activity or cell culture media, which can be used to continuously monitor throughout the culture progress (Bavli et al., 2016; Kieninger et al., 2018; Tharmalingam et al., 2015).

Glucose concentration change in the cell culture media is an indi-cator of cancer cell metabolic activity. Numerous sensors and method-ologies have been designed to determine glucose concentration in real- time including electrochemistry (Chen et al., 2015; Zaidi and Shin, 2016), colorimetry (Xia et al., 2013), conductometry (Mahadeva and Kim, 2011), and fluorescent spectroscopy (Heo et al., 2011). Among them, the enzyme-based electrochemical glucose sensor has attracted much attention for the analysis of cell culture media due to its high sensitivity and selectivity. In addition, the electrochemical sensor continuously measures the biochemical reaction current, providing real-time data for analysis. In the material aspect, conductive materials

* Corresponding author. D307, No. 1378, West Wenyi Road, Hangzhou, 311121 , China. ** Corresponding author.

E-mail addresses: xbs@hznu.edu.cn (T. Xie), djchen@hznu.edu.cn (D. Chen).

Contents lists available at ScienceDirect

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journal homepage: http://www.elsevier.com/locate/bios

https://doi.org/10.1016/j.bios.2020.112261 Received 8 December 2019; Received in revised form 18 April 2020; Accepted 28 April 2020

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including metal nanoparticles (Fang et al., 2016), polymers (Li et al., 2015), carbon materials (Mehmeti et al., 2017; Zhu et al., 2012), and composite nanostructures (Zhai et al., 2013) have been explored as enzyme immobilization interfaces exhibiting enhanced sensitivity, selectivity and stability. In the system structure aspect, electrochemical analysis has been implemented in microfluidic cell culture systems for on-chip analysis (Hiramoto et al., 2019; Ino et al., 2017). A microfluidic tumor-on-a-chip can mimic the key features of physiological environ-ments and be used to better predict drug response (Skardal et al., 2016). However, electrochemical glucose sensing in the cell culture system has two fundamental limitations:

1 In order to avoid glucose concentration gradient caused by cell metabolism, sensors need to be placed close to the cells to obtain a precise measurement in the cell microenvironment (Rodrigues et al., 2008; Weltin et al., 2014). Sensors located at the downstream outlet or far from cells can only detect general glucose changes in the bulk culture media, not accurately reflecting the local concentration changes in the cell microenvironment.

2 The enzymatic reaction catalyzed by Glucose Oxidase (Gox) will oxidize glucose to hydrogen peroxide (H2O2) and D-glucono- δ-lactone. Therefore, in enzyme-based continuous electrochemical monitoring, H2O2 will accumulate and cause adverse effects on the cells (Chang et al., 2003). One solution is increasing the distance between the sensors that produce H2O2 and the cultured cells, but this is in contradiction with the above requirement.

Therefore, developing a sensing platform that overcomes the above issues and enables continuous non-destructive monitoring of the cell metabolism microenvironment is still needed. In this paper, we devel-oped an enzyme-based Polysulfone hollow fiber (PHF) composite which served both as a sensing structure and a cell culture substrate. The cells grow on the outer surface of the PHF, while sensing materials including Multi-walled carbon nanotubes (MWNT), Prussian Blue (PB), and Glucose Oxidase (Gox) were immobilized inside the hollow fiber. The sensing layer and the cells, separated by porous structure, are not in direct contact. The porous structure also provides fluid exchange channels, so that the cell microenvironment changes can be detected by the sensor in real-time. The PB layer was used as the electron transport layer to reduce the working voltage and avoid interferences. Addition-ally, PB decomposed the hydrogen peroxide generated during the enzymatic reaction to lower the adverse effects on the cells. Experi-mental results showed a high electrochemical stability that enables long- term monitoring of the effect of anticancer drugs on cell metabolism activities. This work developed a unique platform for cell metabolism analysis and can be further applied in drug testing and compound screening.

2. Materials and methods

2.1. Reagents and materials

Glucose oxides (Gox), Polysulfone (PSF), poly-l-lysine solution (PLL), bovine serum albumin (BSA), glucose, Dimethylacetamide (DMAC), K3Fe(CN)6, KCl and FeCl3 were purchased from Sinopharm Chemical Reagent (Shanghai, China). Multi-walled carbon nanotubes (MWNT) dispersion were provided by Xfnano Materials Tech Co., Ltd (Nanjing, China). Dulbecco’s modified eagle’s media (high glucose), Trypsin- EDTA (0.25%) and phosphate buffer solution (PBS) were purchased from HyClone (Waltham, USA). Fetal bovine serum (FBS) was obtained from Gibco (USA). All aqueous solutions were prepared with Milli-Q water (18.2 MΩ cm, Millipore).

2.2. Apparatus

Morphology of the hollow fiber sensor was investigated by Hitachi

SU-8000 field emission scanning electron microscopy (SEM). Fourier transform infrared (FTIR) spectroscopic measurements were performed on Nicolet 380 Fourier transform spectrometer. The cyclic voltammetry (CV) and amperometric response (I-t) were recorded using CHI660 electrochemical workstation with a three-electrode system. Ag/AgCl (in saturated KCl solution) was used as a reference electrode, platinum wire as counter electrode and hollow fiber sensor was used as working electrode. Fluorescence images of cells on PHF were obtained by Olympus FV3000 confocal microscope.

2.3. Fabrication of PHF structure

The PHF was prepared by a non-solvent induced phase separation method according to our group’s previous work (Zhu et al., 2016). Briefly, 18 wt% PSF, 6 wt% PVP-K17 and 16 wt% PEG-200 in DMAC solution was extruded from the spinning head to form hollow fiber, which were passed through coagulation baths and transferred to the coiler for winding. After phase separation, the hollow fibers were soaked in 15% aqueous glycerol solutions for 24 h.

2.4. Fabrication of PHF-MWNT-PB-Gox biosensor

Sensor electrode fabrication was carried out by layer-by-layer deposition on the inner surface of the PHF through dead-end filtra-tion. First, 30 μL of 1 wt% MWNT dispersion in water was injected into the hollow fiber by a syringe pump. The MWNT were filtered and trapped by the porous hollow fiber wall and formed a conductive layer on the inner surface. Next, 0.02 M of K3Fe(CN)6 and 0.2 M of KCl as a PB precursor in 20 μL HCl solution (pH 1.7) were injected into the PHF and dried at 40 �C for 15 min. A 20 μL mixture of 0.02 M FeCl3 and 0.2 M KCl was injected into the PHF to start the interfacial growth of PB on MWNT. The MWNT-PB-PHF electrode was further immobilized with the enzyme by injecting 15 μL of Gox solution (1%), followed by 0.5% glutaralde-hyde. Finally, a gold wire was inserted into the PHF to establish an electrical connection between the MWNT-PB-Gox layer and the mea-surement circuit. The electrodes were stored at 4 �C while not in use.

2.5. Electrode performance test

All in-vitro tests were carried out under ambient temperature using a CHI660 electrochemical workstation. I-t curves were recorded in 10 mL PBS (0.1M, pH ¼ 6.8) under -0.05 V working potential with 4 mM glucose added every 600 s. Cyclic voltammograms of the PHF-MWNT and PHF-MWNT-PB electrodes in a 0.1M PBS solution were performed in a potential range of -0.2 V–0.4 V with a 50 mV/s scan rate. Experi-mental details about the sensor stability test method are provided in the supporting information. For the interference test, 0.1 mM of ascorbic acid (AA), acetaminophen (AP) and uric acid (UA) was added into the PBS sequentially between two steps of 0.1 mM glucose concentration increases, and the current signal was collected for reference. The H2O2 concentration in the solution was quantified by a Hydrogen Peroxide Assay Kit (ab102500).

2.6. Cell culture and real-time monitoring

In cell culture experiments, human lung cancer PC9 cells were cultured in a CO2 incubator at 37 �C with high glucose DMEM containing 10% FBS. To investigate the cell adhesion on PHF, PC9 cells on the PHF were stained with DAPI fluorescent stain after 72 h culture. PC9 cells were fixed in 4% formaldehyde for 20 min, followed by thorough washing. The hollow fibers were then incubated with DAPI staining solution (0.1% Tween-20 and 1% BSA in PBS) for 30 min and rinsed with PBS extensively.

For continuous metabolism monitoring, PC9 cells were seeded in a PDMS chamber (fabrication procedure in the supporting information) with a density of 6 � 104/ml and incubated for 24 h for cell adhesion on

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PHF. The system was kept in the incubator environment to maintain normal cell growth. Electrodes in the PDMS chamber were connected to the electrochemical workstation via wires through an incubator access port for continuous detection as shown in Fig. S1. After 24 h, Osimertini, as a model anticancer-drug, was injected into the culture chamber. The current response reflects glucose concentration change was recorded at an applied potential of -0.05 V through the whole culture procedure. Continuous data were collected after a 1-h stabilization period. After 72 h, cells were stained with Propidium iodide (PI) and detected by flow cytometry (FCM) to obtain cell viability. At the beginning and end of the experiment, the culture media was sampled and measured by YSI biochemical analyzer to obtain reference concentrations.

3. Results and discussion

3.1. Structure concept and design

Hollow fiber-based cell culture is one of the methods for large-scale cell culture due to its large surface area to volume ratio (Unger et al., 2005). A precisely regulated gradient porous hollow fiber structure has been developed by our group with applications in enzyme immobiliza-tion (Guo et al., 2018), microfiltration (Gao et al., 2016), and implant-able biosensing (Zhou et al., 2019). In this paper, it has been utilized with sensing materials to evaluate cell growth and metabolism proper-ties during cell culture. As shown in Fig. 1, the gradient porous hollow fiber provides a unique asymmetrical structure suitable for carrying cells and sensing material on the outer and inner walls of the hollow fiber, respectively. Cells grow on the outer surface of hollow fiber, constantly consuming glucose in the culture media. The concentration of glucose inside and outside the hollow fiber is balanced due to the diffusion through the porous structure. The Gox immobilized inside the hollow fiber catalyzes the oxidation of glucose producing gluconic acid and H2O2. PB catalyzes the reduction of H2O2 at low voltage and generates electron transfers through MWNT layer to electrochemical working

station. The porous structure also creates a selective-permeable barrier between the cell culture media and sensing layer to further improve the sensing selectivity and stability.

3.2. Characterization of PHF electrode

The morphology and structure of the PHF electrodes were examined by SEM. Fig. 2(A) shows the cross-section image of a hollow fiber with an outer diameter of 520 μm and an inner diameter of 400 μm. A magnified image of hollow fiber wall is shown in Fig. 2(B). The pores gradually became smaller from the inner surface (>1 μm) to the outer surface (<100 nm). The outer layer acted as a cell adhesion layer and the porous structure allowed fluid exchange through the interconnected pores. MWNT were used to form a 3D conducting framework at the inner surface by dead-end filtration. As shown in Fig. 2(C), unlike other bulky conductive materials, the MWNT retained a three-dimensional loose structure for fluid exchange. From Fig. 2(D), it can be observed that PB was uniformly deposited in the MWNT mesh layer without pore block-ing. The formation of PB on the MWNT was further demonstrated by FT- IR spectroscopy, as shown in Fig. 2(E). The original PHF showed char-acteristic peaks at around 1151 cm-1, assigned to SO2 symmetrical stretching, a strong peak at 1241 cm-1 corresponded to O–C–O stretch-ing, and the peak at 1579 cm-1 was associated with SO2 asymmetric stretching. In the FTIR spectra of PSF/MWNT/PB, besides similar peaks to those of PHF, a sharp peak at 2100 cm-1 was observed, which corre-sponded to the stretching vibrations of Fe–CN in PB.

3.3. Electrochemical studies

The electrochemical sensing performance was measured by succes-sively increasing the glucose concentration in PBS solution. Due to the peroxidase activity of PB, H2O2 generated during glucose oxidization can be catalytically decomposed. Fig. 3(A) shows the amperometric curve of PHF responded to the successive addition of 4 mM of glucose

Fig. 1. Schematic illustration of the PHF electrode structure and sensing principle of the electrochemical reaction catalyzed by Gox and PB.

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from 0 to 32 mM. The currents increased immediately after the addition of the glucose, promptly reaching a steady state with an average response time of approximately 60 s. The sensing target of PHF was high glucose DMEM with initial concentration over 30 mM. Thus, a wide sensing range was required to perform accurate and linear cell meta-bolism monitoring. According to our group’s previous study (Zhou et al., 2019), the gradient porous structure could enhance the restriction of

glucose transmission. PHF achieved a linearity sensing range between 0 mM and 32 mM (R2 ¼ 0.989) as shown in the insert image in Fig. 3(A). In order to verify the effectiveness of the PB deposition onto MWNT, cyclic voltammetry was utilized by cycling the modified electrodes from -0.2 V to 0.4 V. Fig. 3(B) showed the CVs of PHF-MWNT-PB and PHF-MWNT electrodes in absence and presence of glucose (1 mM) in PBS. Compared with the bare MWNT, the PB-modified electrode showed a

Fig. 2. (A) SEM images of the PHF cross-section. (B)Magnified image of the PHF wall showing a gradient porous structure. (C) MWNT layer deposited on the hollow fiber inner wall. (D) PB synthesized in the MWNT-PB layer. (E) FT-IR of original PHF and MWNT-PB-PHF.

Fig. 3. (A) Current-time response curve of PHF sensor as a function of glucose concentration. (B) Cyclic voltammograms of PHF-MWNT and PHF-MWNT-PB electrodes in absence and presence of glucose. (C) Working stability in different concentrations of glucose. (D) Amperometric responses of the PHF sensor to 0.1 mM of Glucose (Glu), UA, AP, and AA.

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pair of redox peaks indicating the oxidation and reduction of PB are present. Fig. S5 showed the CVs of PHF sensor at different scan rates in the range of 20–200 mV/s. Square root of scan rate showed a linear relation with peak current, indicating a diffusion-controlled behavior of the sensor.

Current fluctuation interference will affect the accuracy of the analysis, so long-term measurement stability is required for continuous glucose monitoring. The structure of the PHF sensor immobilizes the electrode and enzyme inside the hollow fiber, which is more stable than other direct contact configurations. The fluid diffuses through the porous membrane also minimizing the interference in the complex cell culture environment. Fig. 3(C) shows the operational stability of the PHF sensor immersed in DMEM at different glucose concentrations (17.6, 20.3, 24.0, 27.0, 30.5 mM) for 5 h. After 1 h of the stabilization period, the sensor current stayed at a constant value and did not show unex-pected fluctuations and drifts in the subsequent 4 h test. The influence of Osimertinib on electrochemical response were verified in PBS buffer. As shown in Figs. S4 and S6, both experiments showed no interference with the electrochemical performance by adding 4 μM Osimertinib.

For analyzing the glucose metabolism, sensor selectivity is an important requirement considering the large number of electroactive species present in the cell culture environment. Ascorbic acid (AA), acetaminophen (AP), and uric acid (UA) are by-products during cell culture, which are also the typical interferences for glucose electro-chemical sensing. As shown in Fig. 3(D), the addition of interfering molecules induced only a small current, which was less than 1% of the current change magnitude caused by glucose addition. The results showed reliable selectivity attributed to the catalytic effect in the low applied potential (0.05 V vs. Ag/AgCl) of PB covered MWNT structure.

Immobilized enzyme inactivation is a significant problem that leads to frequent calibrations and shortened sensor lifetime. We compared different configurations of enzyme electrodes to optimize the structure of enzyme, electrode, and porous membrane. Gox was immobilized on the outer surface of PHF and the surface of a flat Polysulfone porous membrane for comparison test (detailed preparation procedure in the supporting information and Fig. S2). In the storage stability test, all samples were immersed in 30 mM glucose solution at 37 �C. The ac-tivities of the enzyme electrodes were evaluated every day for one week. Fig. 4(A) shows the enzyme activity in the PHF sensor retained above 94% over one-week, while the other two enzyme immobilizing config-urations declined below 80%. The better stability of PHF sensor can be attributed to the porous structure’s interspace confinement effect against enzyme inactivation. In an enzyme-based electrochemical re-action, the generated H2O2 will continuously accumulate in solution, leading to cell damage. Due to the H2O2 decomposition by PB and encapsulation structure of the hollow fiber, the H2O2 generated in PHF can be effectively decomposed. As shown in Fig. 4(B), in the three-day continuous working experiment, the sensor without the PB layer accu-mulated 14.4 μM H2O2 in the cell culture chamber. The hydrogen

peroxide content in the PHF sensor chamber was consistently kept below 2 μM. This result proves that our proposed sensor structure is more suitable for long-term electrochemical sensing in the cell microenvironment.

3.4. Cell culture application test

The cell culture on the outer surface of the PHF was examined through SEM and confocal microscopy. To verify the cell proliferation on the surface of PHF, PC9 cells were cultured with PHF for three days and viewed with SEM. As shown in Fig. 5(A) and (B), the cells could adhere to and spread on the PHF surface in a normal morphology, suggesting that the PHF nanoporous surface was a biocompatible sub-strate for cell adhesion and proliferation. Moreover, the cells cultured directly on PHF for three days were stained by DAPI and observed by a confocal microscope. Fig. 5(C and D) show the 3D volume rendering fluorescence images of the PC9 cells’ distribution along the entire length of the PHF surface. The cell fluorescence image showed a typical curved geometry structure indicating the close cell-PHF adhesion. In Fig. S3, fluorescence microscope images of FITC stained cells showed the cell density increased throughout the three-day culture, indicating the cell viability on the PHF surface. Cell culture morphology characteristics demonstrated that PHF could be used as a proper platform for cell cul-ture in the subsequent studies.

3.4.1. Long-term monitoring in cell culture To verify the continuous sensing and drug screening functionality,

PC9 cancer cells were cultured on PHF and treated by anticancer drugs. First, the response of the sensor in the culture media without cells was measured for three days. The PHF sensor showed a stable current output during the three-day test. After 72 h of continuous work, the sensor current retained 99.7% of the initial value, shown as the red curve in Fig. 6(A). The glucose concentrations measured by biochemistry analyzer at the beginning and the end of the continuous experiment were 30.4 mM and 30.2 mM, respectively, in accordance with PHF measured values. It also indicated that the amount of glucose consumed by the PHF electrode operation was much smaller than the amount consumed by the cells, confirming that the continuous electrochemical monitoring does not act as a nutrient competitor with the cells.

The practical applications of the PHF sensor were demonstrated in a 72 h continuous anticancer drug screening experiment. Osimertinib, also known as AZD9291, was used as an anticancer drug in this exper-iment. Osimertinib potently inhibits signaling pathways and cellular growth in both EGFRmþ (Epidermal growth factor receptor) and EGFRmþ/T790M þ mutant cell lines in vitro, translating into profound and sustained tumor regression in EGFR-mutant tumors (Cross et al., 2014). PC9 cells were seeded in the PDMS chamber with a density of 6 �104/mL and incubated for cell adhesion on the PHF. Cell metabolic rates were reflected by glucose concentration change, which was quantified

Fig. 4. (A) Enzyme stability study of Gox immobilized in different configurations with polymer membranes. (B) H2O2 generated during three-day continuous work.

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by sensor current data. After the first 24 h, drugs were added to the media to trigger cellular metabolism changes. In Fig. 6(A), as a control group, only PBS was injected into the chamber so that cells normally grew over 72 h of culture. In this control group, the PHF sensor current dropped 12.5% from its initial value, which is equivalent to the con-sumption of 5.8 mM glucose concentration in the media. The glucose consumption measured by the biochemistry analysis is 5.65 mM, and the error from PHF sensor was within 3%. In the anticancer drug test groups, a high dose (4000 nM) and a low dose (60 nM) of Osimertinib were added to the respective cultured media after the first 24 h. In the low dose group the sensor current dropped 3.1% in the first 24 h, as shown in Fig. 6(B). In the subsequent 48h after drug addition, the current dropped 2.4% (equivalent to 1.11 mM glucose concentration consumption), which is only 25% of the control group consumption in the same period, showing the drug inhibition effect on cancer cell proliferation. In Fig. 6 (C), before high dose drug addition, the current dropped 3.0% in the first 24 h. In the subsequent 48 h, the current of the PHF sensor in the high dose group slightly dropped 0.9%, equivalent to 0.45 mM glucose con-centration consumption after drug addition, indicating the strong inhi-bition effect on PC9 cell growth. Fig. 6(G) compared the daily glucose consumption in different groups. In the first 24 h before drug addition, all three samples showed similar glucose concentration consumption around 1.4 mM. After drug addition, both samples exposed to Osi-mertinib showed significant metabolite changes. The glucose concen-tration consumptions of the two samples on the second and third day were 0.79 mM, 0.37 mM and 0.36 mM, 0.09 mM, respectively. Cell proliferation and growth inhibition were further analyzed by flow cytometry. After 72 h culture, cells were washed off from the culture chamber and stained with PI. Dead cells displayed a high level of PI staining compared to viable cells. The percentage of viable cells in the three samples dropped according to the higher drug exposure concentration.

3.4.2. Dynamic monitoring in cell culture Besides long-term monitoring, dynamic response was evaluated by

adding 1% of TritonX-100 (TX100) to lyse cells in a short time. TX100 is one of the most widely used nonionic surfactants for permeabilizing the living cell membrane. A large amount of TX100 will result instant cell death and release of protein and other cellular organelles. As shown in Fig. 6(H), before TX100 addition, cells consumed 0.95 mM glucose during first 24 h culture, which showed a similar metabolism rate as other groups. After the addition of 1% TX100, the cells were immedi-ately lysed, so the glucose metabolism process was aborted and the subsequent sensor currents no longer decreased. However, it can be observed in Fig. 6(I) that there was a small current increase after the addition of TX100, which presumably caused by intracellular glucose release after the cells are lysed. This minor current increase also indi-cated that the sensor structure could capture subtle changes in the cellular metabolic status. As a control group, the cells were suspended in the culture media instead of adhering on the PHF surface. This PHF sensor did not record any glucose increase upon the addition of TX100, indicating that only the sensor located close to cells can detect micro-environment concentration changes. The measurement curves in Fig. 6 showed that the system can continuously monitor glucose changes in the cell metabolism environment and obtain specific time points of meta-bolic changes. Unlike endpoint analysis, the PHF sensor provided real- time dynamic information about the glucose metabolism in the cell culture microenvironment.

4. Conclusion

In this paper, a long-term cell metabolism monitoring system was built to accurately assess the cell consumption rate of glucose. PHF sensor was fabricated based on the hollow fiber structure with sensing materials immobilized in the lumen. The combination of porous struc-ture and in-situ deposited sensing layer showed remarkable

Fig. 5. (A) SEM of cells on PHF; (B) magnified view of cell adhesion on the PHF outer layer; (C) Volume view of fluorescence stained cells on the fiber surface; (D) Magnified view of cells on a single fiber surface.

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electrochemical sensitivity and stability. The gradient porous structure of the hollow fiber allowed the glucose sensor to be close to the cultured cells but not in direct contact to prevent adverse effects to cells. Comparative study of different electrode structures showed that the structure of PHF can provide a more stable enzymatic environment and greater efficiency in the decomposition of H2O2. This PHF sensor addressed some of the challenges of implementing electrochemical sensor for long-term monitoring in high glucose concentration media. Compared with traditional endpoint analysis, continuous real-time monitoring offered insight into biological information on drug regu-lated metabolic rates and cellular metabolism alterations. With further structural development and functionalization, PHF sensor can be used as a platform technology for real-time monitoring of cellular metabolism and behaviors, achieving broader applications in drug screening and cancer research.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

CRediT authorship contribution statement

Zhen Ma: Methodology, Investigation, Data curation, Writing - original draft. Ying Luo: Investigation. Qin Zhu: Investigation. Min Jiang: Investigation. Min Pan: Data curation. Tian Xie: Project administration, Resources. Xiaojun Huang: Validation, Writing - re-view & editing. Dajing Chen: Conceptualization, Project

administration, Writing - review & editing, Resources.

Acknowledgements

This work was financially funded by Natural Science Foundation of Zhejiang Province (Grant No.LY18H180009), National Natural Science Foundation of China (Grant No. 61801160) and Zhejiang Province Key R&D Project (Grant No. 2017C03029).

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi. org/10.1016/j.bios.2020.112261.

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