A Cell-Surface-Specific Ratiometric Fluorescent Probe for Extracellular

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A cell-surface-specific ratiometric fluorescent probe for extracellular pH sensing with solid-state fluorophore Yan Yang, Mengchan Xia, Hansen Zhao, Sichun Zhang, and Xinrong Zhang ACS Sens., Just Accepted Manuscript • DOI: 10.1021/acssensors.8b00514 • Publication Date (Web): 16 Oct 2018 Downloaded from http://pubs.acs.org on October 17, 2018

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A cell‐surface‐specific ratiometric fluorescent probe for extracellular pH sensing with solid‐state fluorophore Yan Yang, Mengchan Xia, Hansen Zhao, Sichun Zhang*, Xinrong Zhang Department of Chemistry, Beijing Key Laboratory of Microanalytical Methods and Instrumentation, Tsinghua University, Beijing, 100084, P.R. China. KEYWORDS： Extracellular pH, Ratiometric fluorescent probe, Cationic peptide, Cell‐surface‐specific, Solid‐state fluorophore ABSTRACT: Extracellular acidity is correlated with the development of various pathological states and bulk pH measurements couldn’t report surface acidity. In this study, we have developed a ratiometric fluorescent probe that aggregates upon interaction with cells, allowing persistent labelling of cells and in situ measurement of cell surface pH. The ternary nanoplatform is constructed by a convenient noncovalent combination of bovine serum albumin protected gold nanoclusters (BSA‐AuNCs), fluorescein isothiocyanate (FITC) labeled cationic peptides (CPs) and FITC‐free CPs. The red fluorescent AuNCs serve as reference fluorophore, while FITC labeled peptides act as specific recognition element for H+ and FITC unlabeled peptides are used for delivery. The probe displays a sensitive fluorescence ratiometric response for pH in the range of 5.0‐9.5 with calculated pKa of 7.2. Further studies have demonstrated that this nanosensor also has properties of high selectivity, reversibility to pH fluctuations as well as low cytotoxicity. The new surface pH‐ measurement tool was validated in mapping extracellular pH and monitoring acidification regarding cell metabolism, demonstrating its potential for bioimaging and biosensing.

The pH near the cell surface is known as extracellular pH (pHe), which is a critical parameter associated with various physiological and pathological processes1‐2. For example, acidic pH of the cellular surface is proposed to be one of the significant indications of malignant tumors related to tumor metastasis3 and chemotherapy resistance4‐5. Extracellular pH has long been considered to affect the rate and magnitude of ion transit processes through regulating ion channel activity6. Virus infection can acidify the extracellular surroundings7‐8 because the higher rate of ATP consumption would enhance glycolysis efficiency and lactate in virus‐infected cells9. Moreover, as a representative biological event, alterations in homeostasis of the extracellular environment occur very early in a disease cascade and can be used for the early diagnosis of many diseases, including cancer10. These findings described above suggest that imaging and monitoring the pH fluctuation of extracellular microenvironment could provide important clues to understand and investigate the pH‐dependent cell behaviors which may be beneficial for guiding therapeutic choices and aiding therapeutic agent research and development. Optical approaches are considered to be a noninvasive

diagnostic techniques that are competent to provide the accurate analysis at cellular level11‐14. Fluorescent imaging have been widely used for biological applications and the ratiometric strategy is especially powerful because it makes measurements independent of environmental interferences, probe concentration and instrumental factors, allowing more accurate measurement15‐17. So far, several pH‐sensitive fluorescent indicators have been reported for ratiometric detection in living cells18‐20, however, the challenge of using these probes for pHe determination is to retain them on cell membrane and record pH changes near the cell surface. In most cases, the pH‐responsive agents are small molecule dyes distributed in an entire cell and are washed out very quickly21‐22. Some ratiometric pH sensors based on nano‐ scaffolds are often easily internalized by cells via endocytosis pathways, thus are not suitable for determination of extracellular pH23‐24. The imaging strategies based on antibody or receptor over‐expressed on the cell surface would also result in the internalization of probe and taking probe away from the surface25. Attempts have been made to establish pHe ratiometric sensing system with amphiphilic polymers or pH low insertion peptide (PHLIP) and pH‐sensitive indicator,

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where nature and synthetic polymers such as PEG‐ phospholipid or peptides of the PHLIP family are used as the cell membrane‐targeting ligand, while a pair of pH‐ sensitive and insensitive fluorophores or fluorescence resonance energy transfer (FRET) molecular lead to ratiometric readout, thereby ensuring accurate measurement of extracellular pH26‐27. The use of recombinant DNA technology has made it possible to target fluorescence proteins to specific subcellular compartments, especially cell membrane. In the case of the genetically encoded ratiometric sensor28, the artificial gene encoding membrane probe consisting of a pH‐ sensitive fluorescent protein and a pH‐insensitive cyan fluorescent protein was expressed near the plasma membrane to determine the extracellular pH fluctuation. Although the genetically engineered probes could realize cell‐surface identification and ratiometric measurement, it suffers from the drawbacks of complex manipulation. Wang29 et al. have described a FRET‐based ratiometric i‐ motif sensor for extracellular pH detection, which anchored on the cell surface through streptavidin‐biotin interactions and responded to pH via pH‐dependent single stranded DNA (ssDNA) structural‐switch. Considering the rapid degradation of DNA via the serum nucleases30, stability of the probe need to be carefully assessed in vivo. As a consequence, there is still an urgent need for simpler strategies to in situ analyze cell surface pH of living cells with accuracy, high‐sensitivity and resolution. Recent research in the use of solid‐state fluorochrome for the preparation of molecular probe capable of providing in situ and high‐resolution fluorescence imaging have demonstrated the tremendous potential in overcoming the problem of diffusive signal dilution31. Cationic peptides are a class of small peptides with the high number of lysine and arginine amino acids present in their sequence. Owing to their high biocompatibility and diverse nature, such molecules, in particular cell penetrating peptides (CPPs), gained considerable attention as one of the promising carriers for transporting bioactive compounds32. Normally, cargoes can be associated with peptides through the covalent link or electrostatic interaction and the translocation process involves three steps: concentrating CPP/cargo complexes on membranes, penetrating into cells and finally releasing into cytoplasm or specific organelles33. FITC dye is previously tagged on peptide sequence to visualize the localization of peptide in cells34. Meanwhile, the pH‐ indicative property of FITC fluorophore endows FITC‐ peptides with the potential of pH detection35. We therefore sought to determine if FITC labeled highly charged cationic peptides could form stable electrostatic complexes with the negatively charged protein BSA templated AuNCs and realize ratiometric pH sensing at cellular level.

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Benefiting from the affinity of CPs for plasma membrane and the outstanding photoluminescence properties of FITC and gold nanoclusters (AuNCs), herein, we proposed a fluorescent nanosensor (termed as FITC‐ FPen/FPen@AuNC) for ratiometric monitoring of extracellular pH. As shown in scheme 1, the nanocomposites are well designed to fulfil ratiometric detection in which the pH‐insensitive AuNCs are used as an internal reference, while FITC labeled cationic peptide sequence serve as specific recognition element for H+. In order to concentrate the nanocomposites from the extravesicular medium to the cell surface and keep them on membrane, we introduce large amount of unlabeled cationic FPen peptide to increase the affinity of nanosensor for cell membrane. The nanocomposites are stable and well dispersed in culture media, but accumulated in the juxtamembrane region with large aggregations upon interacting with cells, thus making persistent stain of plasma membrane and possible diffusion‐resistant in situ detection of extracellular pH of live cells. To the best of our knowledge, this is the first reported fluorescent probe with solid‐state fluorophore for ratiometric detecting pHe of living cells. Compared to previous reported extracellular pH indicators, the ternary nanosensor has the following properties: (i) The capability of retaining and precipitating on cell membrane specifically produce a local and consistent fluorescent signal close to cell membrane. (ii) The ratiometric mode can minimize the interference of the complex biological environment, resulting in a more accurate analysis. (iii) The excellent biocompatibility of BSA‐AuNCs can reduce the cytotoxicity of peptides, resulting in a more favorable system for bioimaging and biosensing. Furthermore, the convenient noncovalent synthesis avoids the complicated chemical coupling and modification employed in conventional fluorescent probe. This new surface pH‐measurement tool has been applied to ratiometric map and monitor the pH changes in the extracellular microenvironment of HeLa cells. Besides, the probe was also validated in real‐time tracking the cancer cell surface acidification correlating with normal and high glucose metabolism.

Scheme 1. Principle of the developed FITC‐ FPen/FPen@AuNC nanosensor for ratiometric extracellular pH sensing.

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EXPERIMENTAL SECTION Materials and reagents. Gold (III) chloride trihydrate (HAuCl4∙3H2O, 99%), amino acids, glucose were purchased from Sigma‐Aldrich. Bovine serum albumin (BSA), dialysis membrane (MWCO: 3500), Cell Counting Kit (CCK) were obtained from BioDee BioTech Co., Ltd. (Beijing, China). Metal salts, valinmycin were obtained from Aladdin Chemistry Co. Ltd. (Shanghai, China). All the other reagents were of the analytical grade and used without further purification. All Peptides used in the article were custom ordered from GL Biochem Co. Ltd. (95% purity, Shanghai) and prepared for 4 mM stock solutions by dissolving powder in ultrapure water. Ultrapure water (18.2 MΩ.cm@25℃) obtained from a Milli‐Q water purification system (Millipore) and was used throughout the experiment. Synthesis of AuNCs, Peptide@AuNC nanocomposites. BSA protected AuNCs were synthesized according to a classical bio‐mineralization method with a slight change36. To prepare the FITC‐ FPen@AuNC and FITC‐FPen/FPen@AuNC, FITC‐FPen and FPen stock solution (4 mM) was proportionally dropped into the AuNCs solution (25 mg/mL). The Peptide@AuNC mixture was incubated and gently mixed overnight at 25℃ to obtain the nanocomposites. The probe was then centrifuged at 10,000 rpm for 10 minutes and filtered using 0.22 μm syringe filters. Investigation of Size, Zeta Potential and Colloidal Stability. Dynamic light scattering (DLS) and zeta potential analyses of AuNCs and Peptide@AuNC nanocomposites were obtained from Malvern Zetasizer Nano ZS90 and were measured in 20mM B‐R buffer with pH of 7.4. For the gel electrophoresis, AuNCs, FITC‐ FPen@AuNC and FITC‐FPen/FPen@AuNC were loaded on 1% (w/v) agarose gel for 1 hour at 100 V in 0.5×TAE buffer. After migration, the bands were visualized on the gel with UV illumination at 265 nm. Fluorescence Experiments in Vitro. In the fluorescence assays, samples were excited at 488 nm and the emission was collected from 500 to 750 nm. The standard fluorescence pH titration of FITC‐FPen@AuNC conjugates was performed in B‐R buffer solutions at varied pH values. Briefly, 8 μL of FITC‐FPen (0.2 mM), 50 μL of AuNCs solution (100 mg/ml) and 150 μL of B‐R buffer solutions at a specific pH value were mixed. After gently shook overnight at 25℃, the fluorescence spectrum were obtained on a JASCO FP‐8600 fluorophotometer. Confocal Fluorescence Microscopy Imaging. The nanosensor was excited at 488 nm and the fluorescence signal of FITC part (green channel) was collected from 505 nm to 575 nm and AuNCs part (red channel) was collected from 600 nm to 700 nm. To study the location of nanosensor on the cell membrane, cells were incubated in serum‐free medium containing the FITC‐

FPen/FPen@AuNC for 5 h, followed by washing three times with PBS to remove unbound probes. For extracellular pH calibration, HeLa cells modified with probes according to above process were incubated with high K+ buffer solutions at varied pH values in the incubator for 15 min. The fluorescence images were collected on an Olympus FV1000 confocal laser scanning microscope with a 60× objective lens and the pH calibration curve was constructed according to the average fluorescence intensity ratio of green and red channel in selected ROIs on cell membrane. The fluorescence images were analyzed with Olympus software (FV10‐ASW). Pseudocolored ratiometric images were generated by matlab software. All data were expressed as mean ± standard deviation. RESULTS AND DISCUSSION Design and Characterization of Peptide@AuNC. The as‐prepared BSA‐AuNCs solution was deep brown (Fig. 1A, inset a) and emitted strong fluorescence at 638 nm (Fig. 1A, curve a). A further observation revealed that the AuNCs was mono‐dispersed with an average size of ~1 nm, which was confirmed by HR‐TEM images (Fig. S1, SI). The peptide solution was then added into the AuNCs solution to form nanocomposites via gentle shake. Since FPen is a cationic peptide with isoelectric point (pI) 12.2 and BSA‐AuNCs retains negatively charged property like template, the FPen peptide would adsorb to the AuNCs by electrostatic force. The nanocomposites, denoted as FITC‐FPen@AuNC, consisting of FITC labeled FPen peptide and AuNCs shows a yellowish‐brown color and brighter than AuNCs itself under natural light (Fig. 1A, inset b). When excited at 488 nm, FITC‐FPen@AuNC exhibited typical dual‐emission signals with two well‐ resolved fluorescence peaks at 521 and 635 nm, which derived from FITC fluorophore tagged on peptide and AuNCs, respectively (Fig. 1A, curve b). Compared to spectrums of FITC‐FPen and AuNCs, the maximum peaks of nanocomposites was almost unchanged, whereas the fluorescence intensity of FITC at 521 nm markedly decreased and intensity of AuNCs at 635 nm weakened slightly as the FITC‐FPen linked with the AuNCs. We speculate that the decrease in FITC fluorescence is due to nanosurface energy transfer (NSET) mechanism and the intensity quenching induced by NSET was demonstrated to depend on distance from dye to the surface of the nanoparticle37‐38. It is worth noting that the FPen peptide we used is a relatively long sequence with 27 amino acid residues and the dye labeled on the peptide will be away from the surface of AuNCs when they interact, thus showing a lower quenching efficiency. Additionally, increasing the concentration of FITC‐FPen would cause the continuous quenching of AuNCs’ fluorescence (Fig. S2 A, SI). The effective translocation of cargoes requires high concentration of peptides39‐40, however, excessive addition

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increased, even though the value was up to 1 mM. These data indicate that FPen can associate with AuNCs to form stable, noncovalent Peptide@AuNC.

Figure 1. The characterization of AuNCs and Peptide@AuNC (A) Fluorescence spectra of the (a) AuNCs (b) FITC‐ FPen@AuNC (c) FITC‐FPen/FPen@AuNC and (d) FITC‐FPen (2 μM) in PBS (pH=7.4) under 488 nm excitation. The inset shows the picture of AuNCs and Peptide@AuNC solution under natural light. (B) Size distribution of (a) AuNCs (b) FITC‐FPen@AuNC and (c) FITC‐FPen/FPen@AuNC determined by DLS. (C) Zeta‐potentials of (a) AuNCs (b) FITC‐FPen@AuNC and (c) FITC‐FPen/FPen@AuNC. (D) Electrophoretic mobility of (a) AuNCs (b) FITC‐FPen@AuNC and (c) FITC‐FPen/FPen@AuNC prepared at different concentration of FPen (0.05, 0.1, 0.3, 0.5, 0.7, 1mM) in agarose gel. The concentrations of the reagents used in each experiment are as follows: 25 mg/mL for AuNCs, 10 μM for FITC‐FPen and 100 μM for FPen. The error bar represents standard deviations based on three independent measurements.

of FITC‐FPen would reduce fluorescence intensity of AuNCs. This contradiction was solved by adding unlabeled FPen to form a ternary system. As shown in Fig. 1A (curve c) and Fig. S2 B, association of unlabeled FPen to FITC‐FPen@AuNC did not affect the dual emission of the nanocomposites, but rather helped to restore the fluorescence intensity. The ternary system is denoted as FITC‐FPen/FPen@AuNC and its formation was proved by size and zeta‐potential measurements. As shown in Fig. 1B, the average size of AuNCs in solution was 6.43 ± 0.13 nm with a polydispersity index (PDI) of 0.33 determined by DLS, whereas the size of FITC‐FPen@AuNC and FITC‐ FPen/FPen@AuNC were 7.25 ± 0.11 nm (PDI = 0.32), 7.66 ± 0.10 nm (PDI = 0.31), respectively. As for zeta‐potential, the augment from ‐18.2 mV for free AuNCs to ‐16.8 or ‐ 14.7 mV for Peptide@AuNC composites indicated strong electrostatic adsorption and charge neutralization between AuNCs and peptides (Fig. 1C). Moreover, stability of composite was testified by agarose‐based gel retardation assays. FITC‐FPen@AuNC exhibited a reduced mobility when incubated with FPen and the overall mobility decreased as the concentration of FPen

Analytical Performance of FITC‐FPen@AuNC. The fluorescence pH titration was conducted in B‐R buffer. As shown in Fig. 2A, the fluorescence intensity at 521 nm from FITC‐FPen gradually enhanced as solution pH increased from 5‐9.5, while the fluorescence intensity at 635 nm ascribed to AuNCs just had a slight change. As a result, the intensity ratio of green and red channels I521/I635 continuously increased and showed good linearity with pH values in the range of 6.1 ‐ 8.7 (R2 = 0.9956) (Fig. S2 C, SI). The pKa value of probe was calculated to be 7.2 by using the Henderson‐Hasselbalch equation. Especially, the results of statistical test on I521/I635 corresponding to two adjacent pH values indicate that the FITC‐ FPen@AuNC probe could distinguish 0.1 pH unit change around its pKa (Table S2, SI), which is superior to that of other ratiometric pH sensors, e.g., FITC‐derivatized semiconductor polymer dots(0.5)41, BSA‐stabilized Ce/Au nanoclusters(0.5)42, FITC‐modified carbon dots(0.2)43, FITC‐functionalized CdSe/CdZnS quantum dots (0.5)44. The photostability of FITC‐FPen@AuNC nanocomposites was investigated under different pH conditions. As illustrated in Fig. 2C, no significant changes of fluorescence intensity ratio I521/I635 were detected during intermittent radiation for 2 hours with an interval of 10 min. To test the pH reversibility of nanosensor, high concentrations of acid and base were added to the buffer system containing probe to regulate repeated pH changes. The determination of fluorescence intensity ratio indicated favorable reversibility of FITC‐ FPen@AuNC probe when pH was switched between 6.50 and 8.59 for three cycles (Fig. 2D). Excellent selectivity is the prerequisite for application of sensors to complicated biological systems, so the selectivity experiments were performed by measuring the dual emission intensity ratio of the nanosensor in the presence of potential interfering substance under three different pH conditions. The scope of the examination covered metal cations and active small molecules, which are widespread in living organisms. As exhibited in Fig. S3 A and B, no obvious changes in the I521/I635 ratio were observed when introduced above species to the buffer solution containing FITC‐FPen@AuNC under the same pH value. These experiments suggested applicability of our nanosensor for sensing and imaging in biological systems with high resolution, good selectivity, favorable reversibility and stability. Cell Membrane Labelling with FITC‐ FPen/FPen@AuNC. The ternary nanosensor FITC‐ FPen/FPen@AuNC was applied to cell imaging. Under the optimal experiment conditions, the fluorescence signal of nanosensor was observed in both green and red channels, interestingly, which came primarily from the cell

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Figure 2. Analytical performance of FITC‐FPen@AuNC (A) Fluorescence spectra of the ratiometric nanosensor in buffer solutions at varied pH values. (B) Plot of I521/I635 versus the pH value. (C) Photostability of FITC‐FPen@AuNC under different pH value of 6.58, 7.50 and 8.46 for 2 h. (D) Fluorescence reversibility responses of FITC‐FPen@AuNC between pH 6.50 and 8.59. I521 and I635 are the fluorescence intensity at 521 nm and 635 nm, respectively. The error bar represents standard deviations based on three independent measurements.

membrane as shown in Fig. 3B, 3C. To determine if FITC‐ FPen/FPen@AuNC was located on the cell surface, HeLa cells modified with nanosensor were imaged before and after treatment with Trypan Blue, which is membrane impermeable and capable of quenching the emission of fluorophores in the range of 500‐600 nm45. The fluorescence signal of green channel (Fig. 3F) was completely quenched, while red channel (Fig. 3G) remained intact, indicating that nanocomposites is indeed located in the extracellular space. Moreover, the results of a z‐axis depth scanning experiment also revealed that the fluorescence in all z‐axis planes derived from the cell surface zone (Fig. S7, SI). To ascertain whether peptides and AuNCs dissociate after anchoring on cell membrane, we chose a region of interest (ROI) in one HeLa cell. As illustrated in Fig 3H, the fluorescence signal of green channel co‐localized with red channel in the region and the Pearson’s correlation and overlap coefficient were 0.9019 and 0.9778, respectively. This indicates that peptides remain associated with AuNCs following cellular interaction. The cytotoxicity of nanosensor was evaluated by standard CCK assay. Cells were exposed to FITC‐ FPen/FPen@AuNC and mixture of FITC‐FPen and FPen with the same peptide concentration. About 93%, 79% and 81% cell viability could be estimated after incubating with Peptide@AuNC conjugates for 6 h, 12 h and 24 h, but only 82%, 71% and 64% viability was obtained incubating with peptides alone (Fig. S4, SI). These findings

confirmed that the nanocomposites exhibited low toxicity to cells, even better than CPs, possibly due to desirable biocompatibility of BSA stabilized AuNCs. Furthermore, the confocal imaging in Fig. S8 shows the successful modification of the nanosensor on the membrane of other cell types including cancer cells (MCF‐7, A549) and normal cells (MCF‐10A), thereby proving the generality of the strategy. The distinguished large aggregations from the differential interference contrast (DIC) image in Fig. 3A indicate the probe accumulated on membrane forming solid‐state fluorophore. To further confirm that the probe retains on cell surface with solid‐state, we treated modified HeLa cells with valinomycin which is reported to be used to regulate cell membrane potential46. The affinity of CPP/cargo complexes to cell membrane is primarily initiated by electrostatic interactions between complexes and negatively charged plasma membrane47 and changes of plasma‐membrane potential will break the electrostatic adsorption between the probe and cell membrane, causing the released soluble fluorophores to diffuse away from the membrane surface. Surprisingly, even treated with valinomycin as long as 12 hours and cells presented an apoptotic state, the intense dual fluorescence of nanocomposites could still be observed on the cell surface (Fig. S9, SI), implying that the nanosensor exists as precipitates on the cell surface. The same phenomenon observed in the process of cell apoptosis induced by hydrogen peroxide also supports this

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Figure 3. Location of FITC‐FPen/FPen@AuNC. (A ‐ D) Confocal images of HeLa cells modified with FITC‐FPen/FPen@AuNC. (H) The relationship between signals of green and red channels in ROI chosen from Fig. 3D. (E ‐ G) FITC‐FPen/FPen@AuNC modified HeLa cells treated with 20 μg/ml Trypan Blue. Scale bar: 10 μm.

conclusion (Fig. S9, SI). The formation of large aggregates could be ascribed to the interaction between nearly electro‐neutral charged FITC‐FPen/FPen@AuNC nanocomposites and anionic lipid membrane48. Before aggregation, the nanosensor will first concentrate on the cell surface from bulk solution and we speculate this anchoring should be related to the special amino acid sequence of the FPen peptide. The FPen sequence is rationally designed to modify a phenylalanine at the N‐terminal of penetratin peptide (denoted as Pen), which is a classic cationic peptide with excellent transport ability for protein. Some literatures have reported the special interaction of N‐terminal phenylalanine and cells49‐50. Additionally, there is a cysteine at the C‐terminus of FPen and this reactive amino acid could contribute to complexation of peptide with BSA and also plasma membrane affinity51. In order to determine which moiety of FPen sequence is the key factor in the probe targeting cell membrane, we selected two of FPen related peptide to investigate their delivery of AuNCs (listed in Table S1, SI). These sequences include Pen, the parent peptide of FPen and a random sequence that replaces hydrophobic amino acids including phenylalanine in FPen. As illustrated in Fig. S10, only FPen@AuNC complex could target cell membrane and present red fluorescence derived from AuNCs. Two other complexes, Pen@AuNC and random@AuNC, similar to AuNCs alone, entered cells. By comparing the amino acid sequences of three peptides, we infer that the modification of N‐terminal phenylalanine in FPen peptide plays an important role in keeping probe on the cell surface. Therefore, the synergy between CP containing distal phenylalanine and electrostatic neutralization of AuNCs together induces accumulation of composites on

cell surface and eventually form larger precipitation, which hampers further transmembrane transport of the probe. Extracellular pH Calibration. Inspired by specific location of FITC‐FPen/FPen@AuNC at the plasma membrane and its outstanding performance in vitro, we tried to employ this nanosensor for ratiometric mapping and monitoring the pH changes in the extracellular microenvironment of tumor cells. FITC‐ FPen/FPen@AuNC labeled HeLa cells were subjected to confocal microscopy with changing the external buffered solutions stepwisely from pH 5.94 to 8.78. As expected, a significant reduction of the emission intensity of green channel was observed with the decrease of the extracellular pH, while the red channel barely changed, thereby presenting a distinctive ratio images (Fig. 4). For the quantitative analysis, regions of interest (ROIs) were set on the plasma membrane, in which the fluorescence intensity ratio was calculated. The fluorescence ratio of two channels showed good linearity with external pH in the range of 5.96 ‐ 8.78 as plotted in Fig. 5A. Moreover, the dual fluorescence signals respond reversibly to the repetitive external pH alteration as observed for the response in the buffer (Fig. S11, SI), thus making our nanosensor reusable during long‐term tracking the continuous pH fluctuation. Application for Tracking Cells Metabolism. To further validate the applicability in living cells, the nanosensor was exploited for tracking pH variation of extracellular environment in the process of cells growth and metabolism. HeLa cells modified with FITC‐ FPen/FPen@AuNC were washed with PBS buffer to remove free probes and then observed by confocal laser

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Figure 4. Confocal and ratio images of FITC‐FPen/FPen@AuNC modified HeLa cells exposed to external media at pH 5.94, 6.24, 6.86, 7.36, 7.90, 8.42 and 8.78. The color strip represents the pseudocolor change with pH. Scale bar: 20 μm.

scanning microscope at different times. As shown in Fig. 5B, first group, an apparent decrease in extracellular pH reflected via intensity ratio of green channel and red one was observed with the time of cells proliferation and metabolism, revealing gradual acidification of the microenvironment on the cancer cell surface. Additionally, the calibrated FITC‐FPen/FPen@AuNC was also utilized to reflect the degree of acidification produced by high glucose metabolism in cancer cells. The nanosensor modified HeLa cells were cultured in the medium supplemented with additional D‐glucose, which enhances and promotes cellular metabolism27 and then the fluorescence ratio of green and red channel at the cell surface were measured at different intervals. The second group in Fig. 5B shows the extracellular pH obviously decreased with the incubation time growing. More importantly, the change of pH values in the group added glucose (Fig. 5B second group) was more significant than that of the first group in normal culture medium, indicating that acidification of the extracellular space of cells occurred at a much greater rate after administration of glucose. These results clearly demonstrated the feasibility of our nanosensor for extracellular pH determine, making the probe a promising tool in real‐ time bioimaging. CONCLUSIONS In summary, we report a fluorescence nanosensor that allows for sustained tagging and ratiometric pH measurement at the surface of living cells. The ternary nanosensor is composed of FITC‐labeled or unlabeled FPen peptide sequence and BSA‐stabilized AuNCs through a facile noncovalent approach. It is capable of anchoring on cell surface with aggregated state to achieve

in situ extracellular pH imaging. By using the inherent excellent properties of pH‐sensitive FITC dye and pH stable AuNCs, a ratiometric fluorescence detection of extracellular pH with high sensitivity, excellent reversibility as well as low cytotoxicity was successfully achieved. As a proof‐of‐concept, the extracellular acidity mapping of HeLa cells in different pH conditions were realized. Moreover, the acidification of cell surface in normal and high glucose metabolism was successfully monitored. We expect that the probe will be widely used in subcellular detection after the replacement of appropriate indicator molecules and specific CPPs sequences.

Figure 5. (A) Plot of emission intensity ratio (Green channel / Red channel) of HeLa cells modified with FITC‐ FPen/FPen@AuNC probes vs extracellular pHs (5.94, 6.24, 6.86, 7.36, 7.90, 8.42, 8.78). (B) The variety of extracellular pH of FITC‐FPen/FPen@AuNC modified HeLa cells in normal and high glucose metabolism. The amount of glucose added is 100 mM. The error bar represents standard deviations based on fluorescence intensity ratio extracted from at least 15 individual ROIs.

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ASSOCIATED CONTENT Supporting Information. The Supporting Information is available free of charge on the ACS Publications website. Some experimental details, including cell culture, CCK assay, interference study and cell imaging condition optimization. Additional fluorescence spectra and images.

AUTHOR INFORMATION Corresponding Author Email: [email protected]

ORCID: Sichun Zhang: 0000-0001-8927-2376

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This research was supported by the Ministry of Science and Technology of China (2016YFF0100301) and the National Natural Science Foundation of China (No. 21390410, 21727813 and 21621003)

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