Real-time local oxygen measurements for high resolution cellular imaging

Abstract

Single-cell metabolic investigations are hampered by the absence of flexible tools to measure local partial pressure of O₂ (pO₂) at high spatial-temporal resolution. To this end, we developed an optical sensor capable of measuring local pericellular pO₂ for subcellular resolution measurements with confocal imaging while simultaneously carrying out electrophysiological and/or chemo-mechanical single cell experiments. Here we present the OxySplot optrode, a ratiometric fluorescent O₂-micro-sensor created by adsorbing O₂-sensitive and O₂-insensitive fluorophores onto micro-particles of silica. To protect the OxySplot optrode from the components and reactants of liquid environment without compromising access to O₂, the microparticles are coated with an optically clear silicone polymer (PDMS, polydimethylsiloxane). The PDMS coated OxySplot micro-particles are used alone or in a thin (~50 micron) PDMS layer of arbitrary shape referred to as the OxyMat. Additional top coatings on the OxyMat (e.g., fibronectin, laminin, polylysine, special photoactivatable surfaces etc.) facilitate adherence of cells. The OxySplots report the cellular pO₂ and micro-gradients of pO₂ without disrupting the flow of extracellular solutions or interfering with patch-clamp pipettes, mechanical attachments, and micro-superfusion. Since OxySplots and a cell can be imaged and spatially resolved, calibrated changes of pO₂ and intracellular events can be imaged simultaneously. In addition, the response-time (t_0.5 = 0.7 s, 0 - 160 mm Hg) of OxySplots is ~100 times faster than amperometric Clark-type polarization microelectrodes. Two usage example of OxySplots with cardiomyocytes show (1) OxySplots measuring pericellular pO₂ while tetramethylrhodamine methyl-ester (TMRM) was used to measure mitochondrial membrane potential (ΔΨ_m); and (2) OxySplots measuring pO₂ during ischemia and reperfusion while rhod-2 was used to measure cytosolic [Ca²⁺]_i levels simultaneously. The OxySplot/OxyMat optrode system provides an affordable and highly adaptable optical sensor system for monitoring pO₂ with a diverse array of imaging systems, including high-speed, high-resolution confocal microscopes while physiological features are measured simultaneously.

1. Introduction

Under normal physiological conditions, the mitochondrial electron transport chain consumes cellular oxygen (O₂) to generate a proton motive force, which powers ATP synthesis by complex V, the ATP synthase. Thus, many cellular processes that depend on [ATP] are either directly or indirectly regulated by the availability of O₂ to the cells. Restriction of tissue blood flow reduces the O₂ supply and produces tissue ischemia which, when prolonged, is detrimental to virtually all cell types. The time and spatial details of the O₂ deprivation associated with ischemia matter. Thus, to investigate how ischemia causes cellular damage it is important to obtain rapid, accurate, and reliable measurements as the O₂ level declines, remains low, and rises again. For example, although restoration of blood flow can fix an ischemic event, it can also have adverse effects. Cell damage or death may occur as a result of the reperfusion due to “Ischemia-Reperfusion” (IR) injury [1–5]. The extent of IR damage may depend on the time and degree of O₂ deprivation and the circumstances surrounding the reperfusion. In the intact heart, IR injury that follows acute myocardial infarction, for example, can cause massive loss of cardiomyocytes. Since cardiomyocytes as a rule do not regenerate following cell death, IR cell damage or loss may underlie contractile dysfunction and/or lethal arrhythmia[1, 6]. Yet, despite decades of study and the severity of IR damage, it is not clear what the primary processes that develop during the ischemic phase are, and it is also unclear how this damage predisposes the myocytes to abrupt cell death on reperfusion. The tools described here enable one to control and measure the partial pressure of O₂ (pO₂) in experiments carried out in single cells or small clusters of myocytes -- while other cellular features are measured simultaneously. This method thus enables one to accurately investigate the details of the cellular changes associated with ischemia and also the details of cellular pathology produced by IR injury. These same tools could be used by others to study nearly any cell type and how O₂ affects them quantitatively.

The core method widely used in biology to measure O₂ is the Clark-type electrode, a polarization electrode, the output of which is proportional to pO₂ [7–9]. The technology is robust and widely used. There are two primary problems for single cell experiments: (1) the large size and (2) the slow response of the Clark-type electrodes. More recently custom fabrication of miniaturized Clark-type electrode system was described [10] and additional industrial and scientific sensors have been developed. This includes the solid-state zirconium dioxide O₂ sensor [11], or a proprietary fiber optic method by Ocean Optics that has improved response time and diverse uses [12]. However, such sensors require dedicated instrumentation, custom fabrication, and an associated electronics package. Other optical sensors, such as MitoXpress, do not require such specialized instrumentation. These are phosphorescent O₂ sensors that can be simply added to the extracellular superfusion solution for liposomal cellular uptake or coupled to a biopolymer carrier that can be endocytosed [13]. To photo-stabilize the phosphorescent molecules in the intracellular environment further improvement of these sensors included imbedding the sensors into nano-beads[14]. However, the cytotoxicity [15, 16] of the nano-beads via unclear mechanisms and other practical drawbacks such as the need to load the beads by permeabilization and the partial intracellular dispersion have limited the extent to which this approach is applied. Here, we describe a non-invasive method for measurements of O₂ in experiments with single cells that is ideally suited for virtually any adherent cell type. We make use of Ruthenium based fluorescent optical sensors, which were originally developed for large scale process control [17–20]. With miniaturization, these sensors form the basis of the system presented here and those marketed by others in proprietary formulations. They are robust fluorescent sensors for O₂ and have been used widely in other applications[20, 21]. Agilent/Seahorse uses a proprietary micro well plate technology with this indicator for metabolic investigations of populations of cells or isolated mitochondria in a micro-well configuration[22]. Here, we describe a ruthenium/nile blue based ratiometric method that has been miniaturized for use with isolated single cells being imaged on a microscope. While readily used with glass coverslips and other microscopy tools, it is also adaptable by the imaginative scientist. Briefly, OxySplots is the name given to the new optical tool described here that permits rapid oxygen measurements of pO₂. We provide information needed to make and use OxySplots. It consists of silicone coated micron sized silica gel powder to which two fluorescent chemicals have been adsorbed: (1) the O₂-sensitive Ru(Ph₂phen₃)Cl₂ and (2) the O₂-insensitive Nile blue chloride. Thin (~50 microns) clear PDMS coating containing OxySplots polymerizes on a #1 glass coverslip. The cells placed above the PDMS are imaged as are the OxySplots. This approach allows for the measurement of the partial pressure of O₂ (pO₂) over the range of 0 – 160 mmHg in the cell’s local microenvironment (i.e., pericellular region). The measurements of pO₂ are carried out along with critical intracellular processes (e.g., [Ca²⁺]_i transients, mitochondrial membrane potential (ΔΨ_m), etc.) at fast rates and high spatial resolution.

2. Methods

A detailed description of OxySplots preparation and other materials and methods used here can be found in the expanded Online Data Supplement. Briefly, imaging was done with Nikon A1R inverted confocal microscope (Nikon, Japan) and with a Zeiss LSM 510 inverted confocal microscope (Carl Zeiss, Jena, Germany). Ventricular myocytes were isolated using standard enzymatic approaches [23] from rats or rabbits. ΔΨ_m was measured by supplementing the extracellular bathing solution with 30 nM of TMRM (Tetramethylrhodamine Methyl Ester Perchlorate). [Ca²⁺]_i transients were measured in cardiomyocytes using the acetoxymethyl (AM) ester form of the Ca²⁺ indicator Rhod-2 (Rhod-2 AM) to load the cells with Rhod-2, and were acquired using the confocal line-scan mode. All experiments were conducted at room temperature.

3. Results

A method is presented and demonstrated that enables high-resolution single cell experiments to be carried out while simultaneously measuring the partial pressure of O₂ (pO₂) locally at high speed. We are able to do this by coating the removable bath bottom (a 25 mm #1 glass coverslip) with a thin layer (~50 micron) of oxygen-permeable PDMS (polydimethylsiloxane) into which we have embedded OxySplots (See Fig. 1A). After applying an optional coating to the PDMS (e.g. fibronectin, polylysine, etc.), the freshly isolated cells (cardiac ventricular myocytes in the examples shown here) are added in a physiological solution to the bath and allowed to settle (See Fig. 1B). We can then carry out our experiments. For some experiments, rods are attached to the non-adherent cells for chemo-mechanical experiments and they are lifted several microns from the PDMS surface; for other experiments the cell are allowed to attach to the PDMS or its coating. We can thus carry out a multitude of distinct and separate experiments with the OxySplots under the cells being investigated. The reporting on local pO₂ is from only tens of microns away from the cells and thus exquisitely relevant to the cells (See Fig. 1B). In addition to a gas-tight global bath superfusion source that can be switched between tunable pO₂ sources, a gas-tight glass microperfusion system is available to delivery local solution if needed with variable pO₂ to a specific cell under study. This combination of definable pO₂ sources enables pO₂ to be changed quickly as demanded by the planned experiments and measured in real time by the OxySplots. The plume from the microperfusion can be adjusted to partially or fully cover the cell under investigation and the PDMS and the nearby OxySplots.

Figure 1. Simultaneous measurement of local pO₂ and high resolution cellular imaging.

A, Diagrammatic illustration of the method showing a single, isolated cardiomyocyte in close proximity to the OxySplot particles. B, High-resolution 3D reconstruction of confocal Z-stack images of a cardiomyocyte stained with di-8-ANEPPS to label the sarcolemmal membrane (red), Hoechst-33342 to label nuclear DNA (blue), and OxySplots (purple).

This system enables one to carry out physiological experiments on single cells with a defined level of pO₂ and to change the pO₂ level as required by the experiments. The method thus allows one to measure pO₂ within the microenvironment of isolated or adherent cells (e.g., cardiomyocytes, smooth muscle, model cells, and small tissue components) and investigate how pO₂ may affect cell function and other variables. Since the system is designed to work on confocal, multiphoton, structured illumination, widefield or super-resolution microscopes, the cell biology, physiology and pathophysiology of these cells can be studied simultaneously.

3.1. Calibration of the OxySplots.

To provide quantitative pO₂ measurements, we first carried out a calibration of the OxySplots using a traditional O₂-microelectrode as a pO₂ reference. The tip of the O₂ microelectrode is bathed in the chamber’s solution (total bath volume 250–300 μL), and positioned approximately 50 pm above the confocal imaging plane of the silica gel microparticles. To rapidly equilibrate the O₂-microelectrode and the fluorescent O₂-probe with a set level of pO₂, the flow-rate of the gassed solution into the chamber is 20 mL min”¹. The solution reservoir is bubbled with argon and with air. The flow ratio of the two gasses made it possible to vary the bath pO₂ in controlled increments.

Because the OxySplots are plentiful and randomly distributed each OxySplots is identified by binary image conversion (see Supplemental Fig. 2D) with respect to its location. The fluorescence signal of each OxySplots and how it changes over time is then obtained from the time-lapse confocal images of multiple OxySplots. Note, this was performed using the particle identifier plug-in from the ImageJ software. The fluorescence signals obtained from the OxySplots during the calibration procedure are shown in the inset of Supplemental Fig. 1. From this, a time-dependent fluorescence ratio (R) is measured from each OxySplot. The ratio consists of the signal from the O₂-sensitive fluorophore Ru(Ph₂phen₃)Cl₂ divided by the O₂-insensitive fluorophore Nile blue chloride and allows for correction of non-oxygen related changes (e.g., movement artifacts). Figure 2A shows the pO₂ measurements with the fluorescent O₂-probe (n=107 particles), and the simultaneous pO₂ reading from the O₂-microelectrode. To facilitate the comparison, the measurements are normalized to ambient room conditions (i.e., pO₂ = 160 mmHg, R₁₆₀). Note that the fluorescence emission intensity of an immobilized transition metal-complex, such as Ru(Ph₂phen₃)Cl₂, becomes quenched due to O₂ binding to Ru(Ph₂phen₃)Cl₂ (see Supplemental Fig. 1. and Supplemental Movie 2). The dependency of the fluorescence ratio for the OxySplots (R) on pO₂ can therefore be described by a 2-site Stern-Volmer model[24],

$$\frac{\text{R}}{{\text{R}}_{160}}=\frac{\left(\frac{{\text{R}}_0}{{\text{R}}_{160}}\right)\left({\text{K}}_{\mathrm{sv}}\left(p{\text{O}}_2-f_{1}p{\text{O}}_2\right)+1\right)}{{\text{K}}_{\mathrm{sv}}p{\text{O}}_2+1}$$ (1)

where R₀ is the fluorescence ratio at 0 mmHg, f₁ is the fraction of the immobilized Ru(Ph₂phen₃)Cl₂ population that can become quenched, and K_sv is the Stern-Volmer quenching constant. Our tests (see Fig. 2) suggest that under these conditions the large majority of the Ru(Ph₂phen₃)Cl₂ molecules can become quenched by O₂ (f₁=0.96 ± 0.02). Other constants (R₀ / R₁₆₀=2.85 ± 0.06, K_sv= 0.012 ± 0.001 mmHg⁻¹) have comparable values to those reported by others [19, 24]. Having determined these parameters, fluorescence measurements of pO₂ can now be quantified using ambient room conditions (i.e., R₁₆₀) as a null-point (see Supplemental Table 1) by rearranging equation 1 to give,

$$p{O}_2=\frac{\left({\text{R}}_{\text{A}}-{\text{R}}_{\text{B}}\right)}{{\text{K}}_{\mathrm{sv}}\left({\text{R}}_{\text{B}}+{\text{R}}_{\text{A}}\left(f\mathit{1}-1\right)\right)}$$ (2)

where R_A = R₀/R₁₆₀ and R_B=R/R₁₆₀. Note, R_A is determined by the calibration shown in Fig. 2 and can be assumed to be a constant over the lifespan of the OxySplots. In addition, the inherent steady-state quenching constants of Ru(Ph₂phen₃)Cl₂ (i.e., f₁ and K_sv) are not expected to vary in bathing solutions of different compositions because the polydimethylsiloxane (PDMS) gel is gas-permeable but it is impermeable to ions. Note also that the diffusion rate of O₂ through PDMS is roughly the same as though H₂O (2.4×10⁻⁵ cm²/s at 25° C, see Supplemental Table 2), which makes PDMS an ideal encapsulating material for measuring local, pericellular pO₂. On the other hand, it is critical that other materials which deliver or retain the perfusion solutions be significantly less permeable to O₂ (see Supplemental Table 2) as atmospheric oxygen at ambient room conditions can rapidly contaminate hypoxic solutions. Importantly, even if anoxic solutions flow rapidly and travel relatively short distances through plastic tubing (or even metal), by the time they reach the cellular imaging chamber they are contaminated with O₂ (see Supplemental Fig. 2). This highlights the importance of continually monitoring pericellular pO₂ using a technique such as the one we present here when conducting experiments that require controlled hypoxia.

Figure 2. Formulation and Calibration of OxySplots.

A, Schematic showing OxySplot fluorophores ([Ru(Ph₂phen)₃]²⁺ and Nile blue) adsorbed to silica gel. B, Representative traces showing florescence of both OxySplot fluorophores over time for calibration shown in Fig. 2C. C, Normalized, steady-state fluorescence measurements (each color indicates repeated calibration) from OxySplot (R/R₁₆₀) versus instantaneous pO₂ measurements via a Clark-type O₂ microelectrode. Fit lines are two-site Stern–Volmer model (n=3 pO₂ calibrations). D, Simultaneous, calibrated measurements of pO₂ via OxySplot and Clark-type oxygen microelectrode (see SD for more details).

3.2. Testing the spatiotemporal resolution of the OxySplots.

One significant advantage of the OxySplots over traditional approaches is the ability to carry out spatially resolved measurements of pO₂. To mimic a localized O₂ contamination we perfused solution contaminated by O₂ via a micro-perfusion manifold into a bath bulk perfused with an anoxic solution. The OxySplots enable this contamination to be visualized with high spatial resolution (see Fig. 3A). While the contaminated region is relatively small (i.e., < 0.5 mm²), this area could easily encompass multiple cells and might go unnoticed using existing pO₂ measurement techniques (e.g., micro-electrode). Furthermore, because the OxySplots are plentiful their individual measures of pO₂ can be combined via 2D interpolation into a “heatmap” of pO₂ surround the cells (see Fig. 3B) enabling the OxySplots to provide unprecedented details regarding the microenvironment of a cell.

Figure 3. Local Gradients of pO₂.

A, Calibrated pO₂ as reported by a dense layer of OxySplots B, Interpolated heat-map of pO₂ gradient. Bulk, bath perfusion using O₂ depleted solution with micro-perfusion manifold emitting O₂ rich solution positioned in the bottom left of field of view.

To estimate the response-time of the OxySplots to abrupt changes of pO₂, a local microperfusion system was devised. The OxySplots are first perfused with anoxic bathing solution then a hyperoxic solution is applied via the microperfusion system (see Fig. 3A). By adding an O₂-insensitive fluorophore, sulforhodamine, to the hyperoxic solution we can monitor the light absorption by sulforhodamine as confirmation of bathing solution exchange (see Supplemental Fig. 3). Abrupt step changes of pO₂ lead to reciprocal and rapid changes in Ru(Ph₂phen₃)Cl₂ fluorescence (see Fig. 3A). Our tests indicate that the fluorescent O₂-probe can capture an 80 mmHg pO₂ rise within 0.7 seconds, and a decay of identical magnitude within 1.63 seconds (see Fig. 4C). This confirms that our fluorescent O₂-probe is capable of accurately and rapidly reporting changes in pO₂.

Figure 4. Measuring rapid changes of pO₂ with OxySplots.

A, confocal line-scan image showing the time-dependent fluorescence of [Ru(Ph₂phen)₃]²⁺ and Nile blue. Confocal measurements are done during abrupt local (bathing) solution switch from anoxic solution to hyperoxic (160 mmHg), and back again to anoxic solution (for more details see Fig. S2). B, pO₂ measurements via OxySplots (black line, 8 silica particles indicated in A). C, Double exponential fits to the pO₂ rise (t_0.5 =0.7 sec, n=6 experiments) and pO₂ decay (t_0.5 =1.63 sec, n=6 experiments).

3.3. Parallel measurements of pericellular pO₂ and mitochondrial membrane potential (ΔΨ_m) of single cardiomyocytes.

Here we take advantage of this new approach to perform cellular time-lapse experiments with high spatial resolution. Since OxySplots are approximately 10-20 μm beneath the adherent cell, each can be resolved separately. This allows high spatial resolution confocal images to be captured from either the OxySplot (Fig. 5A) or cellular focal plane (Fig 5B). In these experiments cardiomyocytes are loaded with tetramethylrhodamine methyl ester (TMRM), which preferentially accumulate in the matrix of polarized (i.e., respiring) mitochondria. Since TMRM accumulation is directly proportional to the voltage gradient across the mitochondrial inner membrane (i.e., ΔΨ_m), its fluorescence is used for dynamic measurements ΔΨ_m. Figure 5C shows the ability of OxySplots to report pericellular pO₂ while ΔΨ_m is monitored during a brief period (400 seconds) of hypoxia (see black line in Fig. 5C). While this brief exposure to hypoxia did not produce any significant change in ΔΨ_m, TMRM is ideal for reporting rapid changes in ΔΨ_m [25]. Additionally, the ability to simultaneously monitor pericellular pO₂ and mitochondrial dynamics such as ΔΨ_m will be crucial to future investigations into the role of mitochondria in IR injury.

Figure 5. Mitochondrial inner membrane potential in single cardiac cells during changes of pericellular pO₂.

A, Left, bright-field image (112×112 μm) showing OxySplots. Also visible is the shadow of a cardiomyocyte adherent to the top of the PDMS layer 36 μm above the shown imaging plane. Middle, confocal image showing the fluorescence of [Ru(Ph₂phen)₃]²⁺. Right, confocal image showing the fluorescence of Nile blue. B, Confocal images of a TMRM loaded cardiomyocyte captured at the indicated time points during the experiments shown in panel C. C, The time course of changes in pO₂ (black) and of TMRM fluorescent signals from the whole-cell (red), from intermyofibrillar mitochondria (IFM), from subsarcolemmal mitochondria (SSM), and from perinuclear mitochondria (PNM).

3.4. Measuring [Ca²⁺]_i from single cardiomyocytes during ischemia-reperfusion.

Now we can test the capability of this new approach to provide dynamic measurements of cellular processes at high temporal resolution from single cells, and how they change during ischemia. To monitor rapid changes in cellular [Ca²⁺] ([Ca²⁺]_i) and pericellular pO₂ the confocal microscope is programmed to quickly alternate between the focal planes of the OxySplots and the cardiomyocyte (see Fig. 6A). Here [Ca²⁺]_i is measured using the Ca²⁺-sensitive fluorescent indicator Rhod-2. Throughout the experiment shown in Fig. 6A an isolated cardiomyocyte is electrically paced at 0.5 Hz via electrical field stimulation. Initially, under normal conditions (i.e., cell bathed with normal Tyrode’s solution (NT) equilibrated with room air), a [Ca²⁺]_i transient and the corresponding cell contraction it induces can be observed following each field stimulus. Following the transition to the ischemic conditions (i.e., cell bathed with ischemic Tyrode’s solution (IT)) pericellular pO₂ declines, and pronounced alterations in [Ca²⁺]_i are observed (see Fig. 6A-also see SD Fig 5). Note that the [Ca²⁺]_i transients quickly stabilize and remain in sync with the pacing rate during the 15 minutes of ischemic conditions. However, when the ischemic, extracellular solution is replaced with NT (i.e., reperfusion), asynchronous, oscillatory [Ca²⁺]_i signals occurred. Confocal images capture the cellular alterations can be captured as they occur (see Fig. 6Bi) and the final states of hypercontracture and cell necrosis (see Fig. 6Bii). By capturing pericellular pO₂ alongside critical cellular signaling processes the O₂ monitoring technique shown here allows for the careful consideration of O₂ as a key determinant in cell injury during pathophysiology.

Figure 6. Isolated ventricular cardiomyocytes as an ischemia-reperfusion injury model.

A, the time course of changes in pO₂ (blue line) and Rhod-2 fluorescence (black line). The Ca²⁺-sensitive fluorophore Rhod-2 is used for [Ca²⁺]_i measurements. Fluorescence line-scan measurements of [Ca²⁺]_i are acquired along the longitudinal axis of a cardiomyocyte. During each imaging break the automated focal plane adjustment function of the microscope switches back and forth between two planes; 1) the imaging plane of the cardiomyocyte and 2) that of the OxySplots for pO₂ measurements. The cell is initially perfused with Normal Tyrode, and then with ischemic Tyrode buffer, followed by Normal Tyrode again, as indicated. B, (i) the line-scan image acquired during the reperfusion stage, (ii) 225 μm × 225 μm bright field and confocal image of the cells captured 10 seconds after the line-scan imaging in (i) ended. During the entire time course of the experiment 0.5 Hz electrical field stimulation is applied.

4. Discussion

A new and extremely practical method is presented that enables any investigator to develop a simple, affordable, effective and rapid method to measure pO₂ on virtually any fluorescence microscope system. We have outlined our system and demonstrated its use in heart muscle cells. It can be customized for special needs and adapted to overcome unanticipated problems for virtually all cell types. The basic design of the pO₂ optrode probe is a nano-porous inert scaffold of silica gel (porous silicon dioxide) to which two fluorophores are adhered. One is quenched by O₂ (Ru(Ph₂phen₃)Cl₂) while the other (Nile blue chloride) is unaffected. The probes and scaffold structures are protected with the O₂-permeable PDMS and as functional units have been called OxySplots. As shown here the ratios of these signals can be quickly calibrated via a null-point formula (see below) so that the OxySplots accurately report pO₂ over the range of 0–160 mm Hg. Diverse experiments can be carried out when the OxySplots are incorporated in a thin layer of PDMS but for other experiments the OxySplots can be positioned in the same plane as the cells under investigation.

4.1. Null-point.

A new and simplified calibration procedure for OxySplots is also presented here allowing ambient room conditions (i.e., pO₂ = 160 mmHg, R₁₆₀) to serve as a null-point calibration thereby avoiding repeated calibrations of the same probe. This null-point correction eliminates the effects of possible instabilities in excitation/emission light intensities and also corrects the pO₂ measurements for possible gradual bleaching of the immobilized fluorescent indicators. See “Calibration of the OxySplots” subsection located in the Results section above.

4.2. Ischemia.

The cellular, organellar, and molecular processes underlying ischemic cell injury remain poorly understood. It is clear however that many cell types -- even cardiomyocytes, which critically depend on continuous O₂ supply -- can adapt to episodes of reduces O₂ supply and ischemia. Irreversible cell injury (i.e., IR injury) does not necessarily occur after every ischemic episode. It has long been known from the work of Murry et al., 1986 [26] that cardiomyocytes have inherent adaptation mechanism(s) that, when triggered, can confer protection from subsequent ischemic episodes. Many critical questions remain regarding this discovery of ischemic adaptation, originally termed “ischemic-preconditioning”. Unfortunately, the available tools limit the ability to investigate this adaptation processes (as well as others) with high spatiotemporal resolution.

Techniques to measure pathological intracellular processes in real-time during ischemia have so far been primarily limited to the use of the halted-flow, whole-heart system. Many critical discoveries have been made possible through the frequent use of this system; however, its usage does entail several critical disadvantages. For example, inducing ischemia by halting the flow through major coronary arteries while the heart is exposed or fully exteriorized creates varying levels of ischemia through the myocardium. In addition, the signal measured in these systems is tissue level (i.e., multi-cellular) causing signals from near epicardial layers to comingle with those arising from deeper layers near the ischemic myocardial core.

Single-cell experiments such as those presented here allow the pericellular pO₂ to be tightly controlled thereby avoiding the tissue-level limitations described above. The types of single cell experiments presented here are timely, as is the ability to measure intracellular signals alongside pericellular pO₂. These data provide a direct means for the first time to link changes in pO₂ levels (surrounding the cell) with changes of critical intracellular variables. These challenges (and others) have inspired us to develop an O₂ monitoring system compatible with high resolution confocal measurements of critical cellular components (e.g., mitochondria) allowing rapid alterations and measurements of pericellular pO₂. Furthermore, the ability to resolve dynamic mitochondrial signals at high resolution with pericellular pO₂ is valuable for additional reasons. For example, explicit evidence reveals that mitochondria are damaged during cardiac IR [27] but the time-course and molecular details of this process remain elusive. It has been argued that a large conductance channel across the IMM opens during IR [28–39]. Our ability to observe and characterize this channel (the mitochondrial permeability transition pore, mPTP) during IR, will be significantly enhanced by the use of the technique presented here.

4.3. Contamination of hypoxic solutions by atmospheric O₂.

Here we also demonstrate how the OxySplots technique can provide new means to precisely control the pericellular pO₂. This is significant as single-cell experiments can be easily contaminated by atmospheric O₂. In fact, we found that even when our buffer reservoir was heavily bubbled with 100% Ar to completely remove the dissolved O₂ as measured by a dipped Clark-type electrode, the measured pericellular perfusate pO₂ was significantly higher. This required additional modifications of the system. In addition, we confirmed that even when using materials with low O₂ permeability to carry hypoxic solutions, contamination is very likely. We quantified this behavior (see Supplemental Table 2) to aid future material selections. Furthermore, clearance of pericellular O₂ is not a rapid process, even when this is done with solutions saturated with a heavy inert gas such as argon, which yields a stagnant gaseous phase limiting reentry of O₂. Thus, the ability to measure pericellular pO₂ is critical so that induced ischemic and hypoxic conditions could be confirmed and carefully adjusted.

4.4. Other existing technologies for measurements of pO₂ in single cells experiments.

The acute need to understand better how the availability of O₂ is involved in physiological and pathophysiological mechanisms within cells has prompted the development of pO₂ measurement technologies, including the one presented here. The goal is the simultaneous measurement of pO₂ and other important signals from the single cells under study. However, the technologies available before OxySplots have generally been inconvenient and in some cases cumbersome. Polarization-electrodes like the classic Clark electrode [10] or fiber-optic electrodes [12] require specialized or customized fabricated instrumentation and may interfere with patch clamp hardware or micro-tools used in mechanotransduction experiments. Other intriguing technologies include “nanobead” pO₂ sensors. However, the use of nanobead phosphorescent O₂ sensors [14] has serious drawbacks. The phosphorescent molecules are embedded in the nanobeads composed of one of several plastics that must be microinjected, endocytosed or require prolonged (hours) liposomal loading or aggressive electroporation [14]. These phosphorescent nanobead sensors are not readily compatible with many isolated primary cells such as cardiomyocytes. In addition, the nanobeads may alter physiology or be cytotoxic [15, 16, 40].

4.5. Diverse potential applications.

While we present two applications for the use of OxySplots in single cell experiments, the potential uses are nearly unlimited. Beyond investigating cardiomyocytes, the layered application of OxySplots could be used to investigate nearly any adherent cell type (e.g., HeLa cells, neurons, HEK-293, etc.) using a confocal microscope. Even small organisms (e.g., C. elegans, yeast, bacteria, etc.) could be placed above the OxySplots layer. In addition to confocal measurements, the OxySplots could be arranged in small blobs (see Supplementary Data, Fig. 4, illustration) making the approach compatible with a diverse array of wide-field imaging techniques. In SD-Fig. 4, we show a schematic drawing where the cells and the OxySplots are in the same focal plane but they are laterally displaced. This arrangement enables the two signals to be spatially resolved within the same image and acquired at the same time. Additionally, if OxySplots were placed within a closed system that included an oxygen-consuming cell or organism, O₂ consumption could be measured. Such a enclosure would constitute a microscope-based respirometer. Clearly, there are many additional potential applications.

Oxysplots.

While OxySplots are unusually robust and adaptable sensors for the measurement of pO₂, they have limitations. While the adsorbed dyes on the micron-sized silica gel powder last a very long time (years) when kept cool and dry, the PDMS embedded sensors (OxySplots) has a shelf life of about 6 months. See Supplementary Data for preparation of sensors and OxySplot details. Additionally, while outstanding in reporting pO₂ over the range of 0 mm Hg to 160 mm Hg as shown in Fig. 2, the OxySplots are less sensitive at higher O₂ levels.

4.6. Overview.

A versatile and inexpensive oxygen optrode has been developed for use with single cells, small tissue samples as well as cells in culture when examined with state-of-the-art fluorescence microscopes. The nickname for the optrode is OxySplot to reflect its rapid full-range sensitivity to oxygen, its amorphous and variable shape, and small size. Simple demonstration of its calibration and utility have been provided. With this technology, there is now no longer any need to simply or blindly “set” the pO₂ at its source level and assume that it will be delivered unchanged. Instead, now, the pO₂ can be readily measured at the cellular target while simultaneously measuring electrical, mechanical and physiological properties.

Abbreviations

[Ca²⁺]_i

cytosolic free calcium

[Ca²⁺]_m

mitochondrial matrix free calcium

IMM

inner mitochondrial membrane

ΔΨ_m

the voltage gradient across the inner mitochondrial membrane

MCU

the mitochondrial Ca²⁺ uniporter

NCLX

the mitochondrial sodium calcium exchanger

jSR

the junctional sarcoplasmic reticulum

pO₂

partial pressure of O₂