Peri-cellular oxygen was measured using emission ratiometry. An integrated system comprised of an LED light source and an RGB camera (VisiSensTD, PreSens) imaged changes in the luminescence of optical oxygen sensors placed at the bottom of each well of a 96-well glass-bottom plate (Figure 1A). The plate was placed on top of the VisiSensTD system for continuous monitoring in a cell culture incubator. The oxygen sensor membrane (pink, Figure 1B) incorporated an oxygen-responsive ruthenium dye and an oxygen-insensitive reference dye. Blue light from the LEDs positioned around the camera lens excited the two dyes. The camera imaged the entire plate to acquire oxygen-dependent changes in the luminescence ratio of the two dyes within the sensor located in each well of the plate.

The oxygen sensors were semicircles that covered half the glass bottom of each well. They were laser-cut from a larger sheet (PreSens SF-RPSu4) that consisted of an oxygen sensitive layer, a polyester support layer, and a white optical isolation layer, which was placed onto a sacrificial acrylic sheet with the adhesive facing up (Figure 1C). Semicircular sensors were then laser-cut using a 30 W CO2 laser (Universal Laser Systems VLS 2.3) by placing the acrylic layer on the laser cutter bed and focusing the laser on the top of the acrylic layer. The laser cutting path was drawn in AutoCAD 2022 to cut semicircles with a radius of 2.5 mm using 15% maximum laser power and 10% maximum speed.

Oxygen sensors were attached to the bottom of the wells of 96-well plates using sterile procedures inside a laminar flow hood. Sensors were lifted with tweezers to expose the white optical blocking layer while placing a suction tube (ID < 2 mm) against this layer to hold the sensor while lowering it into a well (Figure 1D). The suction in the tube was released once the adhesive layer attached to the glass. The process was repeated to place sensors in each well (Figure 1E). Before using the plate for a cell culture experiment, the wells were sterilized with 70% ethanol inside the sterile laminar flow fume hood. After the ethanol evaporated, each well was washed three times with 1x PBS, before coating with fibronectin. After an experiment, raw images acquired by the VisiSensTD system were processed using a two-point calibration to convert the RGB values for pixels that imaged each sensor to a percentage corresponding to the peri-cellular oxygen level. An example raw luminescence image showing the optical sensors in a 96-well plate and the processed oxygen reading is illustrated in Figure 1E.

Spectra for the full range of aqueous oxygen concentration were first measured by bubbling a beaker of water with 100% O₂ gas followed by bubbling with 100% N₂ gas (Figure 2A). A sensor attached to the bottom of the beaker was illuminated with excitation light from the VisiSenseTD system and the spectrum of emitted light was acquired once every second using a spectrometer (Ocean Optics QE-Pro, Figure 2B).

Spectra for oxygen concentrations that are typical for cell cultures were acquired using a similar approach and our high throughput microfluidics-based uninterrupted perfusion system (HT-µUPS) cover for a 96-well plate (Wei et al., 2020) (Figure 2C). Nitrogen-bubbled cell culture media (0% O₂) and media equilibrated in room air (21% O₂) were loaded into two separate syringes that were each placed in one of two syringe pumps (New Era 1600 × 2). Media from each syringe flowed through a sensor (Sensirion SLI-2000) that measured the flow rate as media moved through our HT-µUPS cover to perfuse the wells of the plate (Figure 2C). Flow-through Clark electrodes (Microelectrodes Inc. MI-730) incorporated into the tubing before and after the plate measured the media oxygen concentration at those positions. The system flow rate was maintained at 200 ml/min while the flow rates of the two syringe pumps were varied to achieve mixtures of 0, 8, 12, 16, and 20 percent O₂, which were confirmed by the Clark electrodes. The same spectrometer and fiber optic cable setup as in Figure 2A was used to acquire the luminescence spectrum of a sensor within one well once every second.

Spectra collected during each characterization were plotted together to visualize changes in spectral bands corresponding to the oxygen-insensitive reference dye (centered at 510 nm, “green”) and the oxygen-responsive ruthenium dye (centered at 653nm, “red”, Figure 2D). Values at the peak wavelengths for red and green luminescence (Figure 2D, dashed lines) were used to construct the inverse Stern-Volmer relation in Figure 2E. For this relationship, the ratio of red to green emission was plotted against each oxygen concentration measured by the Clark electrode placed before the plate. A linear fit of ratio versus oxygen concentration was used due to the small oxygen concentration range of 0%–20% that corresponds to typical cell culture conditions.

The responsiveness of the oxygen sensing system to changes in media oxygen concentration and flow rate was characterized using 96-well plates and a new oxygen-impermeant version of the HT-µUPS cover described in our previous work (Wei et al., 2020). This cover was assembled in two components, one soft PDMS sealing gasket and one acrylic perfusion base that was CNC-milled from cast acrylic (McMaster-Carr 8560K354) and thermally bonded using a heat press (Rosineer Grip Twist) at 130°C for 3 h. The sealing gasket was fabricated by pouring a mixture of Sylgard 184 (Dow) and Dragonskin 10 (Smooth-On) in a 1:2 ratio into an acrylic mold. Finally, an inlet and outlet were tapped (10–32 thread) for each well and fitted with stainless steel 10–32 barb-to-thread connectors (Pneumadyne).

The sensor response within the perfusion system HT-µUPS was validated using the setup in Figure 3A. Clark electrodes were placed before and after the cover. An optical oxygen sensor was placed at the bottom of the leftmost well of the cover. The Clark electrodes and VisiSensTD system were calibrated with water at 100% O₂ and 0% O₂. Water bubbled with 100% O₂ was loaded into a syringe pump and perfused through the cover. Flow rate was measured using a flow meter (Sensirion SLI-2000) and varied between 500 ml/min and 200 ml/min to determine the system’s response to changes in flow.

A two-point calibration converted the ratio of red to green luminescence intensity imaged from the sensors into percentage of air saturation similar to (Peniche Silva et al., 2020). The terms ‘Cal0’ and ‘Cal100’ were used in system characterization experiments for 0% and 100% air saturation, and oxygen saturation levels ranging from 0% to 18.6% were displayed in cell experiments (the equilibrium oxygen partial pressure in a humidified 37°C, 5% CO₂ incubator is 18.6%).

Two types of culture media, CDI iCell Cardiomyocytes² maintenance medium (Fujifilm CDI) and cardiac fibroblasts growth medium (Cell Applications, Inc.), were used. The air-saturated medium (Cal100) was medium straight from the bottle, warmed to around 37°C. And the 0% air-saturated medium (Cal0) was medium with 5% completely dissolved Na₂SO₃ and placed in the 37°C water bath for 30 min. In testing pH’s influence on oxygen readout, the media were adjusted with NaOH or HCL to pH 10 or pH 4.

Human iPSC-derived cardiomyocytes (iCell Cardiomyocytes² CMC-100-012-001 from a female Caucasian donor) from Fujifilm Cellular Dynamics International (CDI), and human cardiac fibroblasts, CF (Cell Applications, Inc.) were thawed according to manufacturer’s instructions. Cells were plated (50,000 cells per well) in the wells of a 96-well glass-bottom plate containing half-moon shaped oxygen sensors, that have been sterilized and fibronectin-coated (at 50 μg/ml). Culture medium exchange was done every 48 h. In some experiments, hypoxia was induced on day five after plating by filling the wells to the top with culture medium and sealing them with oxygen-impermeable tape before readout in the VisiSense system.

The VisiSenseTD system was temperature-equilibrated in the cell culture incubator at least an hour before the start of measurements. For the whole plate oxygen monitoring in human iPSC-CMs, the peri-cellular oxygen measurements started 5 h after the cell plating, after the switch from the cell plating medium to the cell maintenance medium. And for the hiPSC-CMs and cardiac fibroblast hypoxia comparison experiment, oxygen monitoring started right after the hypoxic condition was established. Oxygen recordings were set to continue for 24–48 h, with a 10 min sampling interval and 0.8 s exposure time.

Cells grown on the non-transparent oxygen sensors were visualized using an upright Leica TCS SP8 microscope with a ×25 water immersion objective (for this imaging, sensors were attached to the glass bottom 35 mm dishes). Cell imaging of the glass-bottom portion of the 96 wells was performed using an inverted confocal microscope (Zeiss LSM 710) or inverted Nikon Eclipse Ti2 microscope. Samples were formaldehyde-fixed, permeabilized and labeled as described previously (Li et al., 2020; Wei et al., 2020). Nuclei were labeled with Hoechst (H3570, Thermo-Fisher Scientific), the cytoskeleton was labeled either for F-actin using Alexa-488 phalloidin (A12379, Thermo-Fisher Scientific) or using α-actinin antibody (A7811, Millipore-Sigma), and some samples were genetically modified to express the optogenetic actuator Channelrhodopsin-2 with a fluorescent reporter eYFP.

Peri-cellular oxygen readings were acquired as PNG images and then analyzed through VisiSensVS software by selecting regions of interest in each image. Measured ratios were calibrated to percentage oxygen readings using two-point calibration with 5% Na₂SO₃ and upon saturation with ambient air. Time-dependent calibration files were applied in the first 2 h of recording, considering the temperature-induced changes in transferring the plate from room temperature operation to the 37°C humidified incubator. Whole-plate data normalization was applied by identifying the maximum and minimum ratio readouts from the whole plate throughout the whole period of recording. A scale factor was calculated by setting the maximum oxygen reading as 18.6%, the equilibrium oxygen concentration in a cell culture incubator.

To quantify the influence of exposure time on the ratio of red to green luminescence intensity, the mean and standard deviation (mean ± SD) of ratios from a 96-well plate were computed and displayed. The spatial variation of the ratios across a plate was assessed for 0.8 s of exposure time. For time-dependent calibration and for the assessment of the effect of pH on oxygen measurements, the change in the ratio of red to green luminescence intensity over time was calculated and displayed as (mean ± SD).

Optical peri-cellular oxygen measurements were based on dynamic oxygen quenching of a ruthenium dye embedded in the sensor patches on top of which cells were grown. For quantitative ratiometric readout, red and green luminescence intensity ratios were converted to percentage of oxygen saturation (pO2) using an adapted Stern-Volmer equation: R o R = A 1 + k s v * p O 2 + 1 − A − 1 (1)

R is the measured luminescence ratio, R₀ is the luminescence ratio at 0% O₂, ksv is the Stern-Volmer constant indicating the efficiency of oxygen quenching, and A is 0.82, a parameter for the non-linearity of the sensing material (Peniche Silva et al., 2020). The ksv was computed using the two-point calibration described in the Methods, and a time-dependent ksv was applied at the beginning of the recordings to account for temperature changes. Eq. 1 was linear for the limited range of oxygenation, that is, typical for cell cultures (<20%, Figure 2E). However, when measured over a full range of 0%–100% oxygen concentration, the inverse Stern-Volmer is expected to follow a decaying exponential function (Wang et al., 2010).

The semicircular oxygen sensors covered half of each well in a 96-well glass bottom plate (Figures 1C, D). This feature enabled multiparametric optical high-throughput measurements from the other half of each well. Such measurements include cellular action potentials, intracellular calcium transients, and contractility (Klimas et al., 2020; Heinson et al., 2023; Liu et al., 2023). Laser cutting of the sensors was quick and reproducible (Figure 1E), where more than 96 half-moon sensors could be cut in less than 10 min. Subsequent attachment of the sensors in each well of a 96 well plate could be completed in less than 1 hour. Laser cutting had a negligible effect on sensor performance and provided sufficient sensor area for good ratiometric measurements after selecting a region of interest from each sensor. We found that it is essential to keep the pre-cut sensors in the dark and to use them within 6 months of laser cutting for best results. Sterilization of the sensors with ethanol for cellular experiments did not affect their performance.

The RGB images of the entire plate (1280 × 1024 pixels, 24bit) acquired by the VisiSensTD system provided sufficient contrast with approximately 1000 pixels per sensor, and 300 to 700 pixels per sensor region of interest (Figure 1E, middle). After calibration, the pseudocolor images of the plate clearly denoted differences in oxygenation between wells. In the example shown in Figure 1E, right, the quadrants of the plate were conditioned with 0% air-saturated PBS (top left), 100% air-saturated PBS (top right), room air (bottom left), and 100% air-saturated PBS with pH = 4 (bottom right). We found that solution pH had negligible influence on calibrated oxygen sensor values. The oxygen sensors also reacted differently in air than when submerged in solution.

Spectral characterization of the optical oxygen sensors and validation of the oxygen measurements with Clark electrodes confirmed that small changes in peri-cellular oxygen concentration could be accurately measured in 96-well plates (Figure 2 and Figure 3). The Presens oxygen sensors (SF-RPsSU4) had a peak emission of 653 nm upon excitation with 409 nm light, and this emission decreased in response to increased oxygen concentration (Figure 2B), demonstrating dynamic ruthenium fluorescence quenching by oxygen (Wang and Wolfbeis, 2014; Wolff et al., 2019). Increased fluorescence in the green band (485–570 nm) was also measured from the sensors upon illumination with 409 nm light (Figure 2D), with a midpoint of 510 nm. The intensity of this band did not change when oxygen concentration increased, corresponding to the oxygen insensitive reference dye that Presens has added to the sensor material. These controlled spectral measurements in a 96-well plate using our HT-µUPS system confirmed adequate spectral sensitivity of the sensors for the range of peri-cellular oxygen that is expected for cell cultures (<18.6%). The relationship of the ratio of red to green emission intensity was linear (Figure 2E), as predicted by Eq. 1.

A recent study cross-validated VisiSensTD peri-cellular oxygen measurements using a fiber-optics oxygen microelectrode (Peniche Silva et al., 2020). In contrast, we used in-line Clark electrodes to validate the measurement of peri-cellular oxygen using laser-cut semicircular sensors, the VisiSensTD system, and physiological flow rates through our HT-µUPS system in a 96-well plate (Figure 3A). Our system demonstrated appropriate responsiveness to changes in media oxygen concentration and flow. During a 30 min recording, media flow began at 500 ml/min, was reduced to 200 ml/min, and then restored to 500 ml/min (Figure 3A). Oxygen values from the Clark electrode positioned at the inlet of the plate indicated that media oxygenation was near 100% and remained constant (Figure 3B). Oxygen values from the Clark electrode positioned after the plate aligned with values measured optically from the sensor at the bottom of the first well. The diameter and length of tubing and the time of the peak oxygen value (85%) measured at the bottom of the well (Figure 3C, green line) corresponded to the flow rate of 500 ml/min. The drop in oxygenation at 200 ml/min indicated a loss of oxygen as the media traveled more slowly through the connection at the entrance of the HT-µUPS cover.

Excitation light was not uniformly distributed across the bottom of 96-well plates. This increased the spatial variance (well-to-well differences) of red:green emission ratios from the sensors for short excitation light exposure times. The dependence of sensor emission ratio variability on illumination exposure times between 0.15 and 1.5 s was measured to identify exposure times where spatial variation was minimized (Figure 4A). Spatial variation was highest for exposure times less than 0.4 s. Spatial variation was lowest for exposure times between 0.4 and 0.9 s. An optimal exposure time of 0.8 s was chosen within this range as a duration for the follow up experiments. The red:green ratio for different wells of a 96-well plate for an exposure time of 0.8 s at two levels of oxygenation (0% and 100% air-saturation) and 2 cell culture media (hiPSC-CMs and cardiac fibroblast) varied within ±6% (Figures 4B, C). The standard deviation for hiPSC-CM media was 6% and 2%, for Cal0 and Cal100, respectively; the standard deviation for cardiac fibroblasts media was 3.5% and 1.7% for Cal0 and Cal100, respectively. This variability was much lower than the typical variability between experimental groups of cultured cells.

Temperature had a significant effect on the red:green emission ratio for each sensor, which was evident after placing a 96-well plate at room temperature in the incubator maintained at 37°C. The effect of temperature on emission ratio, as a plate was warmed to 37°C, was measured over 4 h after placing a plate in the incubator (Figure 5A). Wells contained media for hiPSC-CMs or media for cardiac fibroblasts, and wells had an oxygen level of either 0% or 100% air-saturation. Oxygen images were acquired every 5 min. The average red:green ratio for each combination of media and oxygenation changed over the first 40–90 min (40 min for the cFB media and 90 min for the hiPSC-CM media) (Figure 5A). Ratios for wells having media at 100% air-saturation were similar throughout the 4 h and changed approximately 10% in the first hour. Ratios for wells having media at 0% changed as much as 60% in the first 2 hours and the change was greater for wells with fibroblast media. This result was likely due to differences in the content of the two types of media. The Stern-Volmer ksv values (Eq. 1) computed from the ratios for wells having media at 0% also varied within the first 40–90 min (Figure 5A, inset). These results necessitated the development of a protocol to correct for the effect of changes in well temperature as a plate was warmed to 37°C and for the effect of cell culture media content that determines media color.

The substrate, salt, and chemical content of the hiPSC-CM media and the cardiac fibroblast media were different, resulting in each culture media having a distinct color. Media pH also determines media color and changes in pH could have an independent effect on sensor luminescence. The effect of media color and pH on sensor emission ratio was studied using 96-well plates and the VisiSensTD system. Wells contained either PBS, hiPSC-CM media, or cardiac fibroblast media and the pH of each well was set to be either 4 or 10 (Figure 5B, plate layout). Differences in the color of the media in each well were clearly visible (Figure 5B, bottom-up plate view). However, color differences were not evident in images of the emission ratio (Figure 5B, raw oxygen data). Color differences were also not evident in pseudocolor images after computing the percent oxygen concentration of each well using the emission ratios (Figure 5B, analyzed oxygen data). To determine if pH had a direct effect on sensor luminescence, the emission ratio of each media having 100% air-saturation at standard pH of 7 or a pH of 4 or 10 was measured once every 5 min for 1 hour using the VisiSensTD system (Figures 5C, D). No significant difference was detected between media having a pH of 7 and 4. The emission ratio was consistently lower for media having a pH of 10, which is highly alkaline and has less biological relevance for cell culture experiments.

After validation, characterization, and the development of a robust calibration protocol, the VisiSensTD oxygen imaging system was used to continuously measure peri-cellular oxygen concentration from cardiac cells cultured in a 96-well plate. Using an upright microscope with a water-immersion objective, we confirmed that human iPSC-CMs formed a dense cellular syncytium on top of the oxygen sensors after coating with fibronectin (Figure 6A, lower image). The transparent half of the glass-bottom of each well that was not covered by a sensor enabled human iPSC-CMs and human cardiac fibroblasts to be monitored to confirm healthy growth. This corroborated no adverse effects of the sensors as cells thrived as a syncytium equally well on top of each sensor and on the glass-bottom portion of the well that was not covered by a sensor. In monitoring cell function over several days, we confirmed that sensor readouts were stable and did not alter cell growth. Typical images of human iPSC-CMs and human cFBs grown on fibronectin-coated glass-bottom portion are shown in Figure 6A, upper image and Figure 6C, respectively. Studies were conducted for time intervals ranging from 24 to 48 h (Figure 6). In one experiment, peri-cellular oxygen was measured every 10 min over 48 h from hiPSC-CMs plated in every other row of a 96-well plate (Figure 6A). Two control wells had oxygen sensors and media but no cells. Peri-cellular oxygen dropped dramatically to <5% within 10 h in all wells that had cells. Oxygen concentration in control wells was constant at 18.6%, corresponding to the equilibration of the media with the incubator oxygen concentration. In wells having cells, two patterns of peri-cellular oxygen dynamics emerged after 10 h (Figure 6B): one of monotonic oxygen depletion and another of intermittent depletion. In cases of monotonic depletion, after the initial oxygen drop to <5%, peri-cellular oxygen monotonically and slowly decreased to a steady-state level. In intermittent depletion, peri-cellular oxygen oscillated intermittently after the initial oxygen drop. For the results shown in Figure 6B, monotonic depletion was observed in 11 out of 38 wells (29%) and intermittent depletion occurred in 27 out of 38 wells (71%). Oscillatory intermittent depletion occurred in different wells within the rows of a plate, confirming that oxygen dynamics were independent of the position of a well within a plate.

Dynamic patterns of peri-cellular oxygen were studied in cardiac fibroblast and hiPSC-CM cultures over 24 h to determine if depletion occurs more rapidly in cells having higher metabolic rate (Figures 6C, D). Depletion was hypothesized to occur more rapidly in hiPSC-CM cultures due to the higher oxygen consumption of contracting cells. The two cell types were cultured in separate sets of six wells of the same plate. Two of those wells were filled with the typical 200 ml of media to provide normoxia while the other four wells were filled with 300 ml of media and sealed with oxygen-impermeable tape to generate hypoxia (Figure 6C). Peri-cellular oxygen was depleted within 6 h for normoxic and hypoxic cultures of hiPSC-CMs. Oxygen depletion was much slower for fibroblasts, where over 24 h oxygen did not drop below 10% for normoxic cultures and most hypoxic cultures maintained an oxygen level above 5% (Figure 6D). These results confirm the hypothesis and also demonstrate the utility of optical oxygen sensing in providing long-term measurements that reveal cell-type differences in the baseline and fluctuations of peri-cellular oxygen concentration.