Cell Microenvironment pH Sensing in 3D Microgels Using Fluorescent

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Cell microenvironment pH sensing in 3D microgels using fluorescent carbon dots Anil Chandra, and Neetu Singh ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.7b00740 • Publication Date (Web): 01 Nov 2017 Downloaded from http://pubs.acs.org on November 2, 2017

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ACS Biomaterials Science & Engineering

Cell microenvironment pH sensing in 3D microgels using fluorescent carbon dots Anil Chandra a and Neetu Singh a,b*

a

Centre for Biomedical Engineering, Indian Institute of Technology Delhi, Hauz Khas, New Delhi-110016, India

b

Department of Biomedical Engineering , All India Institute of Medical Sciences, Ansari Nagar, New Delhi-110029, India *

E-mail: [email protected] Fax: + 91-11-26582037 Tel: + 91-11-2659-1422

KEYWORDS: Carbon dots, Microgel, 3D cell culture, Microenvironment, pH-sensing,

ABSTRACT We report here a 3D cell culture microgel based system containing carbon dots capable of sensing the pH changes in the cellular microenvironment. We have utilized a simple droplet based microfluidics methodology for encapsulating cells and fluorescent pH sensitive carbon dots in polyethyleneglycol microgels. Since, the microfluidics assembly is developed from simple components that can be modified easily to yield microgels of different size, composition and architecture; it can be utilized to develop complex 3D cell culture scaffolds of desired

ACS Paragon Plus Environment

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composition along with spatial control on the polymer composition.

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The synthesized pH

sensitive carbon dots possess green fluorescence emission, which increases as the pH is lowered from neutral to acidic. Since the probe sensitivity to pH change is well within the physiologically relevant range (pH 5.8- pH 7.7) the probe can be used for detecting lowering of pH as the cells proliferate or undergo various biological processes. We demonstrate that the nanoprobes as well as the process of forming the microgel beads with nanoprobes and mammalian cells is biocompatible and the cells easily proliferate inside the microgels. The changes in pH as the mammalian cells grow in the microgels is easily monitored via fluorescence microscopy suggesting that the platform can be used to study time dependent changes in cellular microenvironment pH and can be easily utilized to monitor cellular growth, disease progression etc.

INTRODUCTION Cellular microenvironment pH plays an important role in controlling various biological processes and in deciding the fate of cell to a large extent. In a stressed microenvironment of a tumor, cells are always under lack of oxygen and nutrients due to poor vascular perfusion. In these conditions, cancer cells are inclined to rely on lactic acid pathway to generate energy, resulting in production of lactate in excess (Warburg effect), which eventually gets released by the cell into the extracellular environment to maintain the intracellular pH1. The continuous production and release of lactic acid due to high glycolytic flux in tumor decreases the extracellular pH (pHe) in tumor core to pH 6.5-6.9.2 This low pH microenvironment of cancer cells not only affects the efficacy of some chemotherapeutics but also helps cancer cell in protecting themselves from immune response.3,4 In addition, acidic microenvironment is also known to enhance activity of different proteases involved in tissue remodeling and hence tumor


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invasion.2,5 It has been found that acidic pH around a tumor can remodel extracellular matrix of the neighboring normal tissue and make them more prone to invasion by cancer cells. It was found that the most acidic regions around the tumor were the first to get invaded by the cancer cells from the tumor.2,6 Thus, monitoring of the microenvironment pH can also help in forecasting invasiveness and the metastasis potential of cancer cells. Additionally, perturbations in pH can also be used as an indication of progression of various diseases.7 Therefore, monitoring the pH changes can be a useful strategy for quick detection of changes happening in the microenvironment (either tissue or cellular) and can be used to correlate it with cellular growth and disease progression. For investigating or monitoring the extracellular pH,

fluorescent dyes8,9, pH

microelectrodes10, inorganic quantum dots11,12 and many advanced imaging techniques such as positron emission tomography (PET)13, magnetic resonance spectroscopy (1H MRS, 31P MRS, 19F MRS)14–16 have been utilized. These techniques have their own advantages and limitations.17 For instance, a pH electrode can be useful only for localized pH measurements by movement of probe tip to different depths, which can damage the cells.18 Also, most of PET and MRI based techniques require contrasting agents, which can be toxic19,20 for the cells as well as can consume sampling time from 1 to more than 30 minutes thus decreasing the possibility of high throughput screening.21 Fluorescence microscopy is yet another alternative that can be used for sensing pH of the cell microenvironment with higher spatial resolution in short time frame. There are fluorescent dye molecules available for pH sensing of cellular microenvironment, but most of the dyes suffer from problem of cytotoxicity, photobleaching and narrow to not biologically relevant pH detection range. For example, fluorescein, a commonly used dye for pH sensing can be easily photo-bleached and is useful only for sensing pH in the range 6 to 7.2.22,23 Its derivatives,


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carboxyfluorescein24 and

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seminaphtharhodafluor (SNARF) can detect changes from pH 6 to

7.4 and from pH 7 to 8 respectively. Another class of probe, which resists photobleaching23, the semiconducting quantum dots, can also be used for pH sensing.12,26–28 However, they suffer from production of toxic heavy metal degradation products, which can harm the cell and ultimately the biological system.29,30 Carbon based fluorescent nanoprobes are another class of fluorophores that have been reported by various groups for pH sensing.31–38 Most of them involve use of a pH sensitive

small

molecules

like,

(2′,7′–bis-(Carboxyethyl)-5-(and-6)-carboxyfluorescein)

(BCECF), SNARF and fluorescein to be directly linked with the carbon dot for imparting pH sensitivity. However, not many probes exist for sensing in-situ pH over prolonged periods in an in-vivo like microenvironment. Thus, a biocompatible non-photobleachable probe that can sense pH changes in physiologically relevant range, will be indispensable for developing various biomedically relevant detection & monitoring tools. Till date there are not many techniques/probes that can effectively monitor the pH changes occurring in the cellular microenvironment, which can further aid in understanding how the cellular growth, proliferation, disease progression and other processes depend on pH changes. To be able to monitor or detect the pH of cellular microenvironment in real time, the challenges are two fold, firstly, there is a need to develop a biocompatible pH sensitive probe, preferably a nanoparticle based, that can also offer its surface for adding multiple functionalities and secondly the real time monitoring should happen, in an in-vivo like environment, such as a 3D culture which is closest in-vitro counterpart to simulate in-vivo like conditions. For culturing cells in 3D, hydrogels have shown advantages like transparency, tailorable stiffness and composition in addition to high water retention capacity compared to other cell


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culture matrices. However, there are some limitations with use of hydrogels, like poor diffusion of media and oxygen in bulk hydrogel matrix. As the cells need continuous supply of nutrients, exchange of gases and removal of waste metabolites for proper growth and division, improper diffusion can cause accumulation of waste metabolites and scarcity of media around cells. Another disadvantage of bulk hydrogels is their transparency, which decreases with thickness, and they lose their applicability for microscopic studies of cells encapsulated in them. These problems can be easily eliminated if we decrease the hydrogel size from bulk to micron for culturing cells. Among different types of hydrogels, polyethylene glycol (PEG) based hydrogels have emerged as potential material for 3D cell culture as the PEG based hydrogel system is immune to batch to batch variation observed with hydrogels made from biological substrates like collagen, gelatin, fibrin, that can significantly change the outcome of the experiment.39,40 PEG based hydrogels can be easily incorporated with biomolecules by using co-monomer linked with desired molecule during polymerization. Thus desired functionality similar to biological substrates can be easily incorporated in PEG based hydrogels in a controlled and highly reproducible manner.41,42 Here, we report development of a 3D microgel system with carbon dots based pH nanoprobes. We demonstrate that mammalian cells can be easily entrapped into the PEG microgel beads along with the nanoprobes that can offer a platform to detect cell microenvironment pH changes. The microgels with small diameter have large surface area to volume ratio compared to bulk gels, thus, are ideal for microscopic studies of encapsulated cells with less diffusion limitations. Further, the size, architecture and composition of the microgels can be easily modified. The pH sensitive carbon dots are prepared from economical source such as Agaricus bisporus (common mushroom) using hydrothermal method, which is simple,


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inexpensive and requires minimal processing. Several groups have reported the synthesis of carbon dots using natural substrate for sensing multitude of analytes43,44. They are proven to be cytocompatible due to extremely low toxicity45. In addition their resistance towards photobleaching makes them more useful compared to organic dyes.46

MATERIALS AND METHODS 2.1 Materials Poly(ethyleneglycol)

Bioultra,

10,000,

(2-Hydroxy-4′-(2-hydroxyethoxy)-2-

methylpropiophenone, propidium iodide, dichloromethane, potassium carbonate, rhodamine 6G, N-Isopropylacrylamide (NIPAM), magnesium sulphate and ethylenediamine were purchased from Sigma Aldrich. Acrylic acid from Loba chemie. Paraffin liquid light, triethylamine, Span® 80 from Merck. Calcein-AM, Dulbecco’s phosphate buffered saline (PBS), antibioticantimycotic (100X), Dulbecco's modified Eagle medium (DMEM; high glucose) for cell culture, 3-(4,5-Dimethylthiazol-2-yl)-2,5-Diphenyltetrazolium Bromide (MTT), fetal bovine serum (FBS) were procured from Invitrogen. Acryloyl chloride 96% stabilized with 400 ppm phenothiazine was purchased from Alfa Aesar. The HeLa cells were obtained from National Centre for Cell Sciences Pune (India). Pressure monitoring line (150 cm, 1.40 ml) was procured from Romsons, hypodermic needle 30 GX1/2” from PRICON, Syringe pump from kd scientific USA, UV LED (365 nm) with 1200 mW flux output at 2.7W power dissipation was purchased from Mouser electronics USA. 2.2 Synthesis of Poly(ethylene glycol) diacrylate (PEGDA) PEGDA was synthesized from PEG. Briefly 0.1 M dry PEG (10 kDa) was combined with 0.4 M acryloyl chloride and 0.2 M triethylamine in anhydrous dichloromethane (DCM) and stirred


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overnight under nitrogen atmosphere at room temperature. Subsequently the solution was washed with 2 M K2CO3 to remove the precipitated triethylamine hydrochloride salt and separated into DCM and aqueous phase. Small amount of water in DCM phase was removed using anhydrous MgSO4. DCM was removed by rotavap and dried PEGDA was dissolved in deionized water followed by dialysis for 2 days using 3.5 kDa cutoff membrane tubing to remove traces of salt. The purified PEGDA was lyophilized and stored at -20 °C. 2.3 Microgel formation Microgels were synthesized using droplet based microfluidics, which consists of simple arrangement of tubes and needles to generate coaxial inner stream of prepolymer solution surrounded by outer oil phase (paraffin liquid light with 4% span 80). The monomer mix consisted of 10% (w/v) polyethyleneglycol diacrylate (PEGDA) Mol weight 10 kDa and 1% (w/v) Irgacure 2959 in PBS. The inner stream of monomer breaks into micrometer sized droplets by shear force of outer fast moving oil phase (Figure S1). The droplets generated in this way were collected and polymerized by 30 second exposure of UV light (365 nm). The microgels after polymerization were separated from the oil phase by centrifugation and washed with PBS multiple times to remove oil film sticking to their surface and to remove unreacted monomer and photoinitiator. As the growth rate of cells in PEG microgels was slower,47 for obtaining spheroids, which are in-vitro counterparts of tumors, alginate gels with cell adhesive RGD peptide (Arginine-Glycine-Aspartate) was synthesized. The carboxyl group of the alginate (0.5 mM) were used to conjugate to amino terminal of the RGD peptide (0.05 mM) via carbodiimide chemistry. Since, the alginates cannot be photo-polymerized to obtain a crosslinked gel, we modified our protocol to form the droplet of sodium alginate (2 wt% sodium alginate in 0.9%


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NaCl) by using high voltage electrostatic attraction between needle extruding the monomer and the Petri plate containing CaCl2 solution. Due to high electrostatic voltage of 10000 volts, monomer droplets were pulled towards the CaCl2 solution, where they instantly crosslinked to form microgels. 2.4 Synthesis of carbon dots Carbon dots were prepared by hydrothermal method using Agaricus bisporus as carbon source and ethylenediamine as nitrogen source in deionized water. Dried Agaricus bisporus powder (0.6 gram) and 3 mL of 1 M ethylenediamine were dispersed in 10 mL of deionized water and ultrasonicated for 30 minutes. The solution was hydrothermally heated at 160 °C in a Teflon lined stainless steel autoclave for 12 hours. The autoclave was kept for cooling by itself at room temperature for 12 hours. The dark yellow solution obtained after cooling, containing carbon dots, was centrifuged at 12000 rpm for 30 minutes to remove debris left after the reaction. The supernatant was collected and filtered using 0.2 micron syringe filter and dialyzed for 2 days using 3.5 kDa cutoff dialysis tubing against deionized water under constant stirring. The solution containing carbon dots was lyophilized and stored at 4 °C for further studies. 2.5 Carbon dot cytotoxicity Cytotoxicity of carbon dots on HeLa cells was evaluated using MTT assay. Cells were seeded in a 96 well plate in 200 µL cell culture media (DMEM supplemented with 10% FBS) at seeding density of 104 cells/well. After overnight incubation, media was replaced with carbon dots at final concentrations ranging from 0.005 to 2 mg/mL in cell culture media. The carbon dots were added in triplicates and incubated for 24 hours at 37 °C. Carbon dot containing media was then removed and cells were washed with sterile PBS. Untreated cells were used as control to calculate percent viability. MTT (0.5 mg/mL) in DMEM media was added to the wells and were


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incubated for another 1 hour (till purple formazan crystals were formed in control wells). MTT containing media was removed and crystals were solubilized in 200 µL DMSO. Absorbance was recorded at 550 nm using Biotek synergy H1 microplate reader. Experiment was performed at least three times. The data from triplicate samples were plotted with standard deviation between the data as the error bar. 2.6 Carbon dot loading in microgels Carbon dots were loaded in microgels by dispersing 0.5 mg/mL carbon dots in prepolymer solution (10% PEGDA: 10 kDa, 1% Irgacure 2959). The droplet formation and photopolymerization was done as explained earlier. After photo-polymerization the microgels containing carbon dots were separated from oil phase by centrifugation and washed three times with PBS. 2.7 Cell encapsulation in microgels Overnight grown, trypsinized HeLa cells, were dispersed in 10 % PEGDA pre-polymer solution containing 1% Irgacure 2959 in PBS, just before droplet formation. The cell density in prepolymer solution was kept at 107 cells/mL. After droplet generation using microfluidic assembly, they were photo-polymerized by UV irradiation for 30 seconds, trapping the cells suspended in them. Microgels containing cells were separated from paraffin oil by centrifugation and washing with PBS three times. Microgels were placed in DMEM containing 1% antibiotic at 37 °C in CO2 incubator. For obtaining PK67 and PKH26 stained cells, trypsinized and PBS washed HeLa cells were incubated with 2 µM final dye concentration in serum free media at 37 °C for 5 minutes, followed by washing with PBS three times. Cells stained with the respective dyes were then re-suspended in fresh medium for encapsulation into the core and shell using the previously mentioned method.


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2.8 Co-encapsulation of carbon dots and cells in microgels The steps mentioned for cell encapsulation in microgel were followed with addition of carbon dot in the pre polymer solution (10% PEGDA: 10 kDa, 1% Irgacure 2959) to have a final concentration of 0.5 mg/mL. The photo-polymerized microgels containing both cells and carbon dots were washed multiple times with PBS and placed in DMEM containing 10% FBS and 1% antibiotic at 37 °C in CO2 incubator. 2.9 Cell viability assay Microgels containing HeLa cells were incubated with PBS containing 2 µM of Calcein-AM and 1 µM of propidium iodide for 20 minutes followed by washing by PBS 3 times before fluorescence imaging. Live cells were identified by green fluorescence, whereas dead cells with compromised plasma membrane show red emission due to uptake of propidium iodide. Cell viability assay with pH detection was done following the same protocol as mentioned above, except this time microgel also contained carbon dots making them pH sensitive. RESULTS AND DISCUSSION 3.1 Characterization of microgels We began our studies by first developing a micrometer sized system for encapsulating and culturing cells in 3D. We optimized a droplet based microfluidics technique to obtain microgels (Figure S1) via UV photo-polymerization. The technique is based on creating an aqueous droplet into an oil medium, where the aqueous phase containing photopolymerizable monomer is made to flow with a concentric outer oil phase in a narrow tube. The oil phase with relatively higher flow rate, shears the monomer jet into small droplets of uniform size that can be photopolymerized to yield microgels. This microfluidics system also allowed us to create


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Figure 1. Bright field (top row) and scanning electron microscopy (middle and bottom row) images showing control over size of microgels produced by manipulating the flow rate of only continuous phase. (a), (b), (c) are images of microgels produced using oil flow rate of 2, 4 and 7 mL/min respectively. (d) Plot demonstrating the correlation between microgel diameter and continuous oil phase flow rate. (e) Lyophilized microgels showing porous structure with average pore size of 13.23 ± 4.05 µm.

microgels with different size, composition (Figure S1a) and architecture (Figure S1b). The control over the microgel property is essential as it can allow development of microgels where multiple cell co-cultures, cellular differentiation factors and multiplexing of other parameters can be achieved. The control over size of monomer droplets was achieved by manipulating the flow rate of oil and aqueous phase containing the monomer. We were able to produce PEG microgels by UV photopolymerization of PEGDA (Figure S2). The diameter of the microgels can be tuned from 200 µm to 400 µm, by varying the paraffin oil flow rate keeping the monomer phase flow rate constant (Figure 1a, 1b and 1c). The size of the polymerized microgels was assessed by light microscopy and scanning electron microscopy (Figure 1, S3 and S4). The microgels were


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transparent, which is a prerequisite for observing cells that are cultured inside them by microscopy. The plot in Figure 1d shows the relationship between microgel diameter and continuous phase (paraffin oil) flow rate for a fixed aqueous phase flow rate of 1mL/hour. The diameter of microgels formed decreases as the velocity of oil phase is increased. As seen from SEM images in Figure 1e and S4, the PEGDA microgels synthesized were highly porous with an average pore size of 13.23 ± 4 µm. The high porosity helps in encapsulation of cells and proper diffusion of media and gases through the microgel structure. We next investigated if the polymer architecture and the composition of the microgels can be altered easily. With some additional channels in the microfluidic arrangement we were able to generate core-shell and janus microgels very easily with desired composition in various regions of the microgels (Figure S1b). As can be seen from Figure S5, we were able to easily conjugate rhodamine 6G to the carboxyl groups from poly acrylic acid (pAAc) in the core. This not only allowed us to visualize the core-shell architecture by fluorescence microscopy but also demonstrated that the functional groups can be easily used for further bioconjugation of molecules of interest in any region of interest in the microgel. Further, we synthesized a temperature sensitive p(NIPAm-co-PEGDA) core and PEGDA shell microgels (Figure S6a). We were also able to show pH dependent size variation in microgels with poly(ethylene glycol)co-acrylic acid in the core and PEG as outer shell (Figure S6b). These systems can be particularly useful in replicating the cancer cell microenvironment where we can change the density of the core compared to the periphery using multiple triggers like temperature (for pNIPAM-co-PEGDA core and PEGDA shell microgels) or pH (pAAC-co-PEGDA cores and PEGDA shells). Additionally our system also allows encapsulation of two different type of cells in specific regions of the microgels. As an example, we created core shell microgel with core


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containing cells stained by a green fluorescent dye (PKH67 green fluorescent cell linker) and shell containing cells stained by a red fluorescent dye (PKH26 red fluorescent cell linker). This study clearly demonstrates the possibility of developing a spatially controlled co-culture platform (Figure S6c). This is particularly interesting, as such platforms can enable experiments where effect of one cell type on the growth of other can be studied and can shed important insight for cancer research. Our strategy can also be employed to obtain janus architecture with half hemisphere made from PEGDA, which was transparent and other half from p(NIPAm-coPEGDA), which was opaque (Figure S7). Such segmented microgels with complex architecture and composition can also provide a 3D scaffold for culturing and studying interaction between different cells simultaneously in the same microgel with possibility of providing different environment to both cell types. 3.2 Cell encapsulation and viability study After confirming that the physical properties of the microgels can be easily controlled we investigated the biocompatibility of the process and the microgel environment after encapsulating mammalian cells in them. Cell viability assay was performed at different time points via calcein-AM (live cell stain) and propidium iodide (dead cell stain) staining to visualize live and dead HeLa cells (human cervical cancer cell line). Most of the cells inside the microgel were stained with calcein-AM and showed green fluorescence, indicating good viability of the cells (Figure 2a). When the cells were monitored over 12, 24 and 48 hours, slight increase in cell number was observed indicating that the cells were able to grow inside the microgels (Figure 2b). The images clearly indicate the ability of cells to proliferate in the porous microgel matrix with negligible effect of photo-polymerization on cell viability. We also studied the viability of NIH-3T3 cells (mouse fibroblast cell line) in microgels and found them to be alive (Figure S8)


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Figure 2. (a) Cell viability assay for cells encapsulated in microgel. Fluorescence image showing live cells (green) and dead cells (red) after 12 hours, 24 hours and 48 hours. (Scale bar: 500 µm). Live cells were stained with Calcein-AM and dead cells with propidium iodide (b) Plot showing number of live cells per microgel over 12, 24 and 48 hours.

3.3 Characterization of pH sensitive carbon dots With a completely tunable microgel cell culture platform in hand, we focused on development of pH sensitive nanoprobes. We developed carbon dots with fluorescence emission sensitive to pH changes via a hydrothermal synthesis involving Agaricus bisporus as carbon source and ethylenediamine as nitrogen source (Figure S9 and S10). The carbon dots were highly monodispersed with diameter of ~5 nm, estimated using transmission electron microscopy (Figure S11a). The HRTEM image of carbon dots showed lattice spacing of 0.285 nm (Figure S11b), which indicates presence of 002 plane of graphitic carbon48. The spectroscopic studies of carbon dots dispersed in PBS showed two absorption peaks at 254 nm and 330 nm, which is due to π-π* and n-π* electronic transition respectively (Figure S12a). The fluorescence spectroscopy data exhibited excitation dependent emission maxima red shift (Figure S12b), which is common


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to most of the carbon based nanoparticles49,50. The surface functional groups on the carbon dots were confirmed by FTIR to be –COOH, -OH and –NH2 (Figure S12c). These carbon dots showed pH sensitive emission intensity at 550 nm, when excited using 500 nm light. The emission spectra at 500 nm excitation showed increase in emission intensity in response to decreasing pH from neutral to acidic values (Figure S13a) confirming the pH sensitivity of the carbon dots. Interestingly, when excited at 365 nm the carbon dots had an emission maximum at 450 nm, which was found to be insensitive to pH change (Figure S13b). The differential sensitivity of the carbon dots towards pH at different excitation wavelength may arise due to presence of multiple emissive states that may respond towards different environmental conditions, which is pH change in this case51. Since the surface of the carbon dot has many ionizable functional groups that are vulnerable to pH changes, they can ultimately affect the surface states, causing change in fluorescence emission, that may give rise to pH sensitive emission.51,52. The absorbance spectrum of the carbon dots did not showed significant change against variation in pH value (Figure S13c). The pH sensitivity in carbon dots due to presence of ionizable surface functional groups is a known phenomenon and has been observed by various other groups as well.34 Study of zeta potential reaffirms the effect of ionizable functional groups for imparting pH sensitivity to these carbon dots, as zeta potential of solution containing carbon dots at pH 7.4 was -3 mV, which increased to +14.5 mV at pH 3 (Figure S14a). Carbon dots reported by various other groups, which were used for ratiometric sensing show two kinds of fluorescence emission peaks where both the emissions are sensitive to pH variation but to different extents.34,53 The sensitivity of the carbon dots towards ionic strength is a major parameter that can affect its applicability in various other domains of research, therefore we studied it by measuring


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carbon dot’s blue and green fluorescence emission using 365 nm and 500 nm as excitation wavelength respectively, in NaCl solution of different ionic strength. The blue as well as green emission from the carbon dots was found to be stable under highest ionic strength of 1 M, which is much higher compared to physiological ionic strength, which is ~0.13 M (Figure S14b). As the carbon dots are known to have resistance towards photobleaching, we compared its photoquenching with a commercially available pH sensitive dye BCECF. When irradiated with 100 watt, 365 nm ultra violet mercury lamp for 3 hours, the percent decrease in ratiometric fluorescence intensity observed for BCECF was almost double to that for carbon dots (~6.5%) (Figure S15a). The quantum yield of carbon dots for excitation in UV region was 4.2 %, where the calculations were made taking quinine sulphate (QY: 54%) as reference dye. The quantum yield for excitation at 500 nm was ~13% taking fluorescein (QY: 79%) as reference dye for calculations. The range of pH detection was observed to be from pH 10 to pH 4, which is broad and also includes the physiological pH range (5.6-7.6) of human body54. Most interestingly, unlike other probes these carbon dots are “turned-on” type probes i.e. the fluorescence intensity signal increases when pH decreases from neutral to acidic pH (Figure S13a & S15b). Thus, a biological change where the pH changes from neutral to low (usually the case) will result in lighting up of the region, which can be easily observed. Also, the carbon dots can be used directly after synthesis without any modification, which is generally a necessity for semiconducting quantum dots to prevent leaching of heavy metal ions from the core. Study of the reversibility of the fluorescence emission of carbon dots against pH change was done by varying the pH of the solution containing carbon dots in a cyclic fashion with collection of emission intensities. The robustness of the carbon dots pH sensing property were found to be


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similar to commercially available dye BCECF as there was negligible change in the fluorescence signal after repeated cycles of pH variation (Figure S15c and S15d). To test the applicability of the carbon dots for sensing the pH changes within the microgel environment, it was essential to assess the amount of carbon dots loaded and released from the microgel as dyes that can release faster and not retained may not be useful for such platforms. We observed an initial quick release of ~9% of the loaded carbon dots within first few hours followed by saturation within 3 hours without any further release (Figure S16). This indicated that the carbon dots present in macroporous voids of the microgel were removed due to diffusion leaving more than 90% carbon dots retained in the nanoporous voids. Interestingly, we observed almost ~85% BCECF being released compared to only ~9% release of carbon dots within 3 hours of incubation. 3.4 pH sensing by carbon dot loaded microgel Followed by convincing proof of carbon dot’s retention in microgel we incubated the microgels in PBS maintained at different pH values. The microgels were then imaged using fluorescence microscope to obtain the emission intensity at each pH value. The microgels showed increase in green fluorescence intensity as the pH of the solution decreased (Figure 3a). The average fluorescence intensity from microgels, calculated using ImageJ software, when plotted against pH values showed a linear relation between fluorescence emission and pH (Figure 3b); indicating suitability of the carbon dot loaded microgel for pH sensing. 3.5 Cell viability in microgels To monitor the in-situ pH changes the carbon dots must be encapsulated into the microgels along with cells and should not possess any cytotoxicity. The carbon dots were found to be biocompatible when cytotoxicity was assessed using a commonly used mitochondrial assay, the


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Figure 3. (a) Green emission (λex=480 nm) from the microgels due to presence of pH sensitive carbon dots in response to changing solution pH. Image is showing increased fluorescence as the pH decreases from 7.4 to 5. Scale bar is 200 µm. (b) Plot showing change in average fluorescence intensity per microgel against respective pH values.

MTT assay (Figure S17). The carbon dots were well tolerated upto 1 mg/mL by the HeLa cells and can be used as probe for sensing the pH change in the microenvironment of encapsulated cells without causing any cytotoxicity. When the carbon dot and cell loaded microgels were imaged by confocal microscopy (Figure S18a), even after 12 days the carbon dots were found to be well distributed inside the microgels and retained outside in the microenvironment without getting internalized by the cells as evident from the non-fluorescent cytosol (Figure S18b). Further, the viability of Hela cells encapsulated in the microgels with carbon dots were confirmed by cell viability assay after culturing them in the microgels for 24 hours. The cell viability assay was done at different pH buffer solution. Live HeLa cells stained by Calcein-AM were equally bright in microgels at pH 6 and pH 7.4, whereas significant emission from the surrounding microenvironment inside the microgels was observed from microgels maintained at pH 6 (Figure S19). This study confirmed that the cells were viable in the presence of carbon dots inside the microgel and the carbon dots also responds to changes in pH.


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3.6 pH sensing in microgels with cells Motivated by the sensitivity of the microgel carbon dot system towards manual pH changes, we assessed its applicability for monitoring pH changes due to cellular growth and metabolism. Therefore, we incubated microgels containing both carbon dots and cells in DMEM at 37 °C in a CO2 incubator to monitor the changes in the microenvironment pH due to cell growth over time (Figure 4a). When the fluorescence from microgel was assessed using fluorescence microscopy, it was found that compared to day 0, average fluorescence intensity per microgel at day 1 and day 2 increased by 17.5 % and 36% respectively, as shown in the plot (Figure 4b). The increase

Figure 4. (a) Fluorescence image of microgels with encapsulated pH nanoprobes and HeLa cells. The green fluorescence (λex=480 nm) from microgel increases with time indicating decrease in pH. (Scale bar: 500 µm). (b) Plot showing percent increase in average fluorescence emission from microgels with time. (c) Plot showing rate of increase of emission intensity from microgels containing cells over 12 days when incubated in paraffin oil.


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in fluorescence intensity indicated decreasing pH inside the microgels and clearly indicates the ability of the system to monitor growth of cells and changes in microenvironmental pH by analyzing the fluorescence intensities. A similar experiment was performed for a longer period of time where the microgels were contained in oil phase so that no exchange with external media is possible. We anticipated that this would make our system more sensitive towards pH change as the buffering effect of the external medium will not interfere with pH change due to cellular growth. When monitored over 12 days, we observed that the pH decreased increasing the fluorescence emission intensity from carbon dots inside the microgels continuously. A ~7.4% increase in emission intensity after day 1 was observed, that increased to ~33% by day 12 (Figure S20a and S20b). The data suggests an exponential decrease in the pH with a dramatic change between day 5 and day 9 (Figure 4c). By comparing the percent increase in fluorescence intensity of the microgels at day 12 compared to day 0 with the standard plot (pH vs % increase in fluorescence), we were able to estimate the pH inside the microgel. The estimated value was similar to the measured experimental pH (Figure S20c). To confirm the viability of cells encapsulated in paraffin submerged microgel after 12 days of incubation, MTT assay was performed for HeLa cells. The cells showed good viability even after 12 days (Figure S21). These studies confirmed the potential of the 3D scaffold in monitoring pH changes occurring over long period of time. As an application of this platform, we cultured MCF-7 cells (human breast cancer cell line) in a spheroid forming microgel (RGD modified alginate microgels) and mapped the pH across the spheroid. Microscopy results showed formation of pH gradient between the core and the peripheral part of the microgel. In the images while the periphery was non fluorescent, the fluorescence from the acidic core was clearly evident. These studies validate


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the applicability of our platform for measuring and visualizing pH changes in a growing tumor (Figure S22a and S22b). CONCLUSION We have developed a 3D cell culture microgel system consisting pH sensitive carbon dots that can sense the cellular microenvironment pH changes. The carbon dots and the process of cell encapsulation in microgels is biocompatible. The developed platform can be used to monitor the pH changes in real time as the cells grow in in vivo like 3D environment. Simple image processing yields fluorescence intensity vs pH plots that can be correlated with pH of the microgel. With this platform we can monitor the pH of organoids or spheroids, which are closest in-vitro systems that mimics tumors found in-vivo. We believe that this platform can enable studies related to growth of tumor cells and can help screen factors affecting it.

ASSOCIATED CONTENT The Supporting Information is available free of charge on the ACS Publications website at DOI: Figure S1 to S22 containing data for Schematic for microgel formation, chemical structure of PEG hydrogels, control over size of microgels, SEM of microgels, control over microgel architecture and composition, live dead assay of NIH-3T3 cells encapsulated in microgel, pH sensitive CD synthesis steps, chemical structure of CDs, TEM of CDs showing size and lattice spacing, plots for spectroscopic properties of CDs (Absorbance, fluorescence, FTIR), plots for variation in absorbance, fluorescence emission and zeta potential with change in pH, plot showing effect of ionic strength on fluorescence emission of CDs, comparison of CD with BCECF dye for photo-quenching; pH sensitivity; robustness and release from microgel,


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cytotoxicity of CDs on HeLa cells by MTT assay, confocal Z-sectioning of microgel containing cells and CDs, images showing effect of culture time over fluorescence emission from microgels containing CD and HeLa cells submerged in paraffin oil, live dead assay for HeLa cells encapsulated in microgel after 12 days incubation under paraffin oil, spheroid culture and generation of pH gradient in alginate microgel. AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. Fax: + 91-11-26582037. Tel.: + 91-011-26591422 (N.S.). Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Acknowledgment We acknowledge Central Research facility (CRF) and Nanoscale Research Facility (NRF) at IIT, Delhi for the instrumentation facility used in this work. A.C. acknowledges support from UGC as a Senior Research Fellow. We acknowledge financial support by Indian Institute of Technology, Delhi. The work was funded by DBT, India (IYBA program). REFERENCES (1) (2)

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Cell Microenvironment pH Sensing in 3D Microgels Using Fluorescent

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