Mitochondria play a multitude of roles in cell life and death [1
]. In cell life, mitochondria produce the majority of cellular energy Adenosine triphosphate (ATP) and sequester harmful reactive oxygen species; in cell death, mitochondria play an active role in the so-called “intrinsic” pathway of apoptosis or programmed cell death [3
]. It logically follows that mitochondrial dysfunctions resulting in defective energy metabolism and resistance to apoptosis have been linked to various disorders such as Alzheimer’s disease, cardiovascular diseases, diabetes, Parkinson’s disease, and cancer [5
]. To assess mitochondrial dysfunction, current state-of-the-art experiments often report the mitochondrial respiration rate because it directly reflects the functional status and the health of the mitochondrion [6
Convincing evidence exists that mitochondria have genetic heteroplasmy at multiple levels: as an organism ages, genetic (especially mitochondrial DNA based) defects in mitochondrial function become more prevalent, resulting in a heterogeneous distribution of mitochondrial DNA within the organism and within a given tissue [7
]. During germline maturation, defective mitochondria are screened through unknown mechanisms to prevent offsprings from carrying serious mutations [8
]. While technologies for assaying genetic variation among mitochondria are relatively mature [9
], technologies to measure functional
heteroplasmy are still lacking. One of the most significant functional assays, the quantification of the oxygen consumption rate, is usually reported as an average of thousands of cells or isolated mitochondria [6
]; hence, the dynamics and variability of respiration rate from single cells or single mitochondria are lost. Furthermore, rare cell behavior such as stem cells or circulating tumor cells [7
] is masked in such an ensemble approach.
To this end, we have developed a technology to assay the respiration of a single mitochondrion. Our “micro-respirometer” consists of micron sized chambers etched out of borofloat glass substrates and coated with an oxygen sensitive phosphorescent dye Pt(II)-meso-tetra(pentafluorophenyl)porphine (PtTFPP) mixed with polystyrene. The chambers are sealed with a polydimethylsiloxane (PDMS) layer coated with oxygen impermeable Viton rubber to prevent diffusion of oxygen from the environment. As the mitochondria consume oxygen in the chamber, the phosphorescence signal increases, allowing direct determination of the respiration rate. Experiments with coupled vs. uncoupled mitochondria showed a significant difference in respiration, confirming the validity of the microchambers as single mitochondrial respirometers. This demonstration could enable future high-throughput assays of mitochondrial respiration and benefit the study of mitochondrial functional heterogeneity, and its role in health and disease.
2. Materials and Methods
2.1. Mitochondrial Isolation
The mammalian cell line HeLa (American Type Cell Culture) was maintained in the log growth phase using minimum essential medium (MEM) supplemented with 10% fetal bovine serum (FBS). All cell culture related supplies were obtained from Thermofisher Scientific (Waltham, MA, USA). 107 cells were typically harvested for mitochondria isolation.
Before isolation, the confluent cells were stained with 100 nM MitoTracker Green FM for one hour. The cells were then washed with fresh media.
Our complete isolation buffer contains 225 mM mannitol, 75 mM sucrose, 0.5 mM EGTA, 20 mM HEPES, 0.5% (w
) Bovine Serum Albumin (BSA), 1X protease inhibitor, pH 7.2 with 1 M KOH. All buffer chemicals were purchased from Sigma Aldrich (Saint Louis, MO, USA). The stock isolation buffer was prepared without BSA and protease inhibitor and stored at 4 °C. Mitochondria from the cultured cells were isolated using differential centrifugation. After collection, the cells were transferred to a glass homogenization tube in 3 mL complete isolation buffer and homogenized with 30 strokes on ice. The cells were then transferred into 2-mL Eppendorf tubes and centrifuged at a low speed of 2000 × g
for 4 min at 4 °C. The resulting supernatant was collected and centrifuged at a high speed of 12,000 × g
× 10 min at 4 °C. The supernatant as well as the light-colored fluffy sediment containing damaged mitochondria were aspirated and the resulting pellet was resuspended in respiration buffer (140 mM KCl, 2 mM MgCl2
, 10 mM NaCl, 0.5 mM EGTA, 0.5 mM KH2
, 2 mM HEPES, 5 mM succinate, 2 μM rotenone, pH 7.2 adjusted with KOH). However, for mitochondria protein analysis, the mitochondria were resuspended in KCl buffer without EGTA. The protein analysis was done with BCA assay kit supplied by Thermo Fisher Scientific, and the result indicated a typical yield of 140 µg mitochondrial protein. As a quality control, we used a Hansatech Oxytherm to measure the respiratory control ratio of a typical mitochondrial preparation with this protocol and found the ratio to be around 3.1, which is good for cultured cells [14
2.2. Microchambers Fabrication
Borofloat glass wafers (3″ × 3″ × 0.7 mm) with a pre-deposited 120 nm Cr layer were supplied by TELIC Company (Valencia, CA, USA). On top of the Cr layer, there was also a 530 nm preprocessed layer of AZ1500 positive photoresist. Using photolithography, Cr etching, and 48% HF wet etching, we created microchambers in the borofloat glass substrate with a depth of 7.5 µm and a surface diameter of 15 µm. The chamber’s depth was measured by a Dektak 3 surface profilometer. A scanning electron micrograph of one microchamber is shown in Figure A3
The oxygen-sensitive dye was obtained from Frontier Scientific (Logan, UT, USA) (Catalog # PtT975) and was incorporated with polystyrene (PS), average Mw
280,000 Sigma Aldrich (Saint Louis, MO, USA) (Catalog# 182427). The PtTFPP/PS oxygen sensing mixture was prepared in toluene with 150 µM PtTFPP and 64 µM PS final concentrations. A few drops of this mixture were quickly transferred onto the substrate containing the microchambers using a cotton swab and the wafer was heated at 120 °C for 1 min. Immediately after the wafer was sufficiently cooled, we used scotch tape to extract the dye from the wafer surface, leaving PtTFPP only at the bottoms of the microchambers. Due to the capillary effect [15
], the deposited dye formed a ring around the bottom of each chamber.
2.3. PDMS and Viton-Coated Top Insulating Layer
PDMS was prepared according to the manufacturer’s (Dow Corning, Midland, MI, USA) recommendation in 10:1 ratio with the curing agent. The PDMS were prepared into 3 cm × 2 cm × 3 mm slabs. Viton rubber was obtained from Pelseal Technology (Bensalem, PA, USA) (PLV 2000) company and mixed with its accompanying accelerator #4 in 44:1 ratio. Viton was paint brushed onto the top and the sides of each PDMS slab and allowed to cure at room temperature (25 °C) overnight.
2.4. Oxygen Calibration and Sealing Test
An inverted microscope Olympus IX71 was used to capture the phosphorescent intensity from PtTFPP/PS when exposed to 0% O2 (from 100% N2 standard laboratory line), 21% O2 (composition of O2 was assumed from the standard laboratory house air line), and 100% O2 (from an industrial cylinder). The air was flown approximately 5 cm from the top of the devices. Each gas was connected to a separate tubing and could be switched quickly. The microscope was equipped with a 395 nm excitation LED and the appropriate filter cube for 650 nm emission capture by QImaging QIClick camera.
To test the effectiveness of the insulating layer, oxygen sealing test was carried out by periodically exposing the microchambers with 100% O2 and measuring the changes in the phosphorescent intensity. Three tests were performed: (1) Microchambers; (2) Microchambers covered with a PDMS slab; (3) Microchambers covered with a Viton coated PDMS slab. Measurements were performed in the dark at room temperature (25 °C).
2.5. Experimental Procedure
Experiments were done at the UCI optical biology core with a Zeiss LSM 780 confocal scanning microscope. Excitation wavelengths were 405 nm and 488 nm by lasers for PtTFFP/PS and Mitotracker Green, respectively. The LSM 780 can simultaneously monitor both red and green channels. Exposure time was set at 200 ms and images were taken every one second.
Microscope focus to the bottom of the chambers were first achieved before mitochondria deposition. Following the addition of 50 µM ADP to the mitochondrial suspension to initiate respiration, 2 µL of 0.2 µg/mL of mitochondrial protein was dropped and spread to the wafer’s surface. After the mitochondria were deposited, a Viton coated PDMS slab was quickly placed on the wafer, effectively sealing and separating the microchambers. We then performed image acquisition for the durations specified in the results. To decouple the mitochondria or to induce maximal respiration rate, carbonyl cyanide 3-chlorophenylhydrazon (CCCP) was added along with ADP. Measurements were performed in the dark at room temperature (25 °C).
Up to 15 chambers were visible in the field of view. The chambers were separated into two groups: (1) with mitochondria and (2) without. Image processing was done with ImageJ. The average red intensity and green intensity (when applicable) were measured for each chamber. The data were processed in IGOR software. To minimize the effect of photobleaching, the red intensity was divided by the average green intensity from all applicable chambers. These data were further normalized by the initial processed intensity. The linear fit feature in IGOR was used to fit a line to the plotted data.
We have demonstrated a simple, robust technology to assay respiration of individual mitochondria. The proof of concept shows that measurement of functional heteroplasmy (that is, functional differences between mitochondria) is possible. This is significant because heteroplasmy in the mitochondrial DNA content has been shown, but techniques to assay the functional differences are poorly developed.
How does this compare to existing technology? Broadly speaking, two methods are dominantly used to measure respiration: Electrochemical (Clark type electrodes) and luminescent readouts. Oxygen sensors based on a Clark-type oxygen electrode rely on the amperometric electrochemical reaction between a coated Pt-electrode and dissolved molecular oxygen, and have challenges for miniaturization such as consumption of oxygen by the sensor itself, and the requirement for a reference electrode [14
]. Our approach in this paper has been to use oxygen-sensitive phosphorescent probes, which do not consume oxygen and offer high sensitivity, specificity, fast response times down to milliseconds [28
], and demonstrated compatibility with biological samples [29
summarizes the features of selected state of the art commercially available instruments (Oroboros [24
], Warner [25
], Hansatech [31
], Seahorse [6
], MitoXpress [32
]), as well as those demonstrated in research labs (Cantebury [34
], and UWash [22
Due to the diffusion of O2
, measuring the absolute oxygen consumption rate of any biological sample requires a completely sealed and isolated chamber. The available commercial Clark-type electrode systems offer excellent sealing chambers but require a large amount of sample and as discussed previously, they are not suitable for miniaturization. For the systems based on phosphorescent probes, commercial companies offer alternatives to sealing chambers. MitoXpress uses a layer of mineral oil to minimize to back diffusion of O2
; however, this approach is limited to assessing treated versus non-treated sample and still does not solve the sealing problem. Seahorse Biosciences, on the other hand, employs a semi-closed chamber and develops a complex mathematical model to calculate the O2
back diffusion, which is deducted from their final measurements. Oxygen sensing has also been integrated in microfluidic channels [34
]; yet, these channels were designed to monitor the oxygen level in tissue cultures and not for the purpose of detecting oxygen consumption rate. Single-cell respiration with a set of sealed micro-wells have been demonstrated by UWash [38
], however, this setup requires an external pressure by a piston. Our microchambers, in contrast, require no piston for sealing and can achieve single mitochondrial resolution, which translates to approximately 1 pg worth of mitochondrial protein [39
]. In addition, our micro-chambers are only 1.5 pL, the smallest in size to our knowledge with the next smallest being 80 pL from UWash. These attributes present considerable improvements of our microchambers over the compared technologies.
While our approach demonstrates proof of concept, additional work is required to make the technology broadly useful in studies of mitochondrial heteroplasmy. First, the noise of the system needs to be critically assessed to determine the ultimate limits of respiration. At this point, while the integration time required for measurement (~10 s) is slightly better than existing technologies (~one minute), the signal to noise is inferior. One method to improve this would be to use phosphorescence lifetime rather than intensity measurement, which typically suffers from lower drift and improved signal to noise. We have demonstrated the feasibility of this approach for measuring oxygen tension in our chambers (Supplemental Information
), so this is a clear possible next step. Second, a method needs to be developed to more practically introduce reagents into the chamber over time, rather than seal the chamber only once for the entire assay. Third, the instrument needs to be built out for turnkey operation, rather than requiring a skilled operator. This should be straightforward with appropriate off the shelf electronics, optics, and control software. Finally, the technology needs to be validated against existing biological models. Such a validation demonstration, of course, would only be possible for ensemble respiration measurements, as biological models for functional heteroplasmy of mitochondria are only now possible to assess based on the prototype devices presented here. Of course, these four additional tasks are beyond the scope of this proof of concept prototype demonstration.
Intracellular probes for measuring oxygen concentrations have recently advanced in single cell applications [40
]. However, these probes are still a matter for further research [17