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Alterations in bioenergetics due to changes in mitochondrial DNA copy number
Methods 51 (2010) 452–457
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Methods journal homepage: www.elsevier.com/locate/ymeth
Review Article
Alterations in bioenergetics due to changes in mitochondrial DNA copy number Wei Qian, Bennett Van Houten * Department of Pharmacology and Chemical Biology, University of Pittsburgh School of Medicine and The University of Pittsburgh Cancer Institute, Hillman Cancer Center, Pittsburgh, PA 15213, United States
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Article history: Accepted 22 March 2010 Available online 25 March 2010 Keywords: Mitochondria Mitochondrial DNA Oxygen consumption rate Oxidative phosphorylation Extracellular acidification rate Glycolysis ATP
a b s t r a c t All mitochondrial DNA (mtDNA)-encoded genes are involved in mitochondrial electron transport and ATP production. Alterations of mtDNA due to dysfunctional mitochondrial DNA polymerase gamma (POLG) induce loss of mitochondrial oxidative phosphorylation (OXPHOS) and mitochondrial ATP generation. Total intracellular ATP is generated by two energetic pathways, glycolysis and mitochondrial OXPHOS. Decreased ATP generation from mitochondria due to mitochondrial dysfunction induces compensatory upregulation of cytoplasmic glycolysis process, thus increasing the contribution of glycolysis to the total cellular ATP generation. Decreased mitochondrial respiration and ATP generation with concomitant enhanced glycolysis is associated with mitochondrial disease and cancer. This chapter introduces a novel assay using a pharmacological profiling strategy in combination with a Seahorse XF24 instrument, which quantifies mitochondrial oxygen consumption rate and extracellular acidification rate for the measurement of OXPHOS and glycolysis, respectively. This assay combined with an analysis of steady-state ATP levels was used to study the bioenergetics of cells depleted of mtDNA (rho0 cells). Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction Oxidative phosphorylation (OXPHOS) is the culmination of a series of bioenergetic transformations that are called cellular respiration. Mitochondrial OXPHOS and cytoplasmic glycolysis are the two main metabolic pathways to generate cellular ATP. Through glycolysis one molecule of glucose is metabolized to two molecules of pyruvate with the concomitant net production of two molecules of ATP. In the absence of oxygen or due to mitochondrial dysfunction, pyruvate is converted to lactic acid, thus completing the glycolysis cycle. Under aerobic conditions, pyruvate is transported into mitochondria. In mitochondrial matrix, pyruvate is oxidatively decarboxylated to acetyl CoA. Complete oxidation of the acetyl unit of acetyl CoA through citric acid cycle and OXPHOS generates 26 of the total 30 molecules of ATP from one molecule of glucose. During OXPHOS, oxygen is reduced to water in a four-electron reduction at cytochrome c oxidase in mitochondrial electron transport chain (ETC). The amount of intracellular ATP that is generated from OXPHOS is variable in cells throughout the body [1], and more than 90% of the body’s oxygen is consumed by the ETC. The remaining 10% is comprised of non-mitochondrial respiration, including substrate oxidation and cell surface oxygen consumption [2].
* Corresponding author. Address: Hillman Cancer Center, 5117 Centre Avenue, Research Pavilion, Suite 2.6, Pittsburgh, PA 15213-1863, United States. Fax: +1 412 6237761. E-mail address: [email protected] (B. Van Houten). 1046-2023/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2010.03.006
Increased conversion of glucose to lactic acid even in the presence of oxygen associated with decreased mitochondrial respiration is a distinct characteristic of tumors, first described by Otto Warburg (Warburg effect) in the 1920s [3,4]. Although the mechanism of Warburg effect and its role in malignant transformation remains elusive [5,6], this phenomenon demonstrates the cellular flexibility in switching the main cellular energy production from OXPHOS to glycolysis during the process of tumorigenesis [7]. OXPHOS is executed by four mitochondrial protein complexes for electron transfer (NADH-Q oxidoreductase, succinate-Q reductase, Q-cytochrome c oxidoreductase, and cytochrome c oxidase, which are also called complex I, II, III, and IV, respectively) and one complex for ATP synthesis (ATP synthase, which is also called complex V or F1–F0 ATPase). Impairment in these complexes could cause dysfunction of OXPHOS. Mitochondrial DNA (mtDNA)-encoded genes are involved in the electron transport and production of ATP to carry out OXPHOS. Mammalian mtDNA encodes genes for 2 rRNAs, 22 tRNAs, and 13 protein subunits, which are the critical components of mitochondrial ETC complexes I, III, IV, and V. The subunits include seven subunits of NADH-Q reductase, one subunit of cytochrome c reductase, three subunits of cytochrome c oxidase, and two subunits of F1–F0 ATP synthase [8]. Defects in the replication and transcription of mtDNA induce mutation, deletion, or depletion of mtDNA, cause defects in assembly of OXPHOS complexes, which can eventually cause loss of OXPHOS. Mitochondrial DNA polymerase gamma (POLG), which is the only known DNA polymerase found in mammalian mitochondria, is responsible for replication and repair of mtDNA. Mutations in
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POLG are associated with mitochondrial depletion syndrome (MDS) such as progressive external ophthalmoplegia (PEO) [9]. mtDNA depletion is also associated with cardiomyopathy [10], myopathy [11], and cancer [12]. To study the contribution of mtDNA to OXPHOS and hence cellular function, rho0 cells (cells depleted of mtDNA) have been widely used as a successful model system [13]. Rho0 cell line can be established by treating cells with low concentration of ethidium bromide (3,8-diamino-5-ethyl-6-phenylphenanthridinium bromide). Ethidium bromide inhibits mtDNA POLG more strongly than DNA polymerase alpha and beta [14], thus inhibits the replication and transcription of mtDNA without substantially affecting nuclear DNA [15–17]. In order to monitor the alterations in cellular energy metabolism as a response to mtDNA impairment and gain a better understanding of the interplay between the two main bioenergetic pathways (OXPHOS and glycolysis), in this chapter we introduce the application of Seahorse XF24 Extracellular Flux analyzer (Seahorse Bioscience, Billerica, MA) in measuring the metabolic profiles (oxygen consumption rate, OCR and extracellular acidification rate, ECAR) of cells, and the use of rho0 cells as a model system for mtDNA depletion syndromes. 2. Analysis of OXPHOS and glycolysis using the Seahorse XF24 extracellular flux analyzer 2.1. Seeding cells in XF24 cell culture plate The XF24 cell culture plate (Seahorse Bioscience, Part #100777004) is formatted like a typical 24-well plate, with rows designated A–D and columns 1–6. The seeding surface of each well is the same size as a typical 96-well plate. Even seeding and around 90% confluence of cells are extremely important for XF assay. Uneven seeding and/or too less or overgrowth of cells can cause significant well to well variation, and may not truly reflects the cellular metabolic profiles and also cause large SD value. Resuspend cells to obtain a desired final concentration to seed in 100 ll of growth media. Typical seeding densities vary from 20,000 to 60,000 cells per well depending on cell size and basal metabolic rates. Seed 100 ll of cell suspension per well gently, leaving Temperature Correction wells blank (A1, B4, C3, D6). During seeding, rest the pipet tip just below the circular rim at the top of the well for homogenous cell layer. Then place plate in an incubator and allow cells to adhere. This generally takes approximately 1 h for strongly adherent cells, but may take up to 5–6 h for less adherent cells. After cells have adhered, add 150 ll of growth media to each well, bringing the total volume of media in the well to 250 ll. When adding media to the wells, add it slowly to the sides as to not disturb the newly attached cells. Then allow the cells to grow overnight in an incubator (37 °C, 5% CO2). For the assay of non-adherent cells, the cells can be attached to the bottom of wells by centrifugation of cell suspension in a XF24 plate coated with BD Cell-Tak Cell and Tissue Adhesive (BD Biosciences, CN 354240). A swinging bucket rotor centrifuge should be equipped with plate carriers, such as the Eppendorf Centrifuge 5810R. The assay for non-adherent cells does not require overnight culture of cells after centrifugation, thus cells can be suspended directly in unbuffered media for centrifugation and proceed the step of incubation in a 37 °C incubator without CO2 (Section 2.3).
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waveguides. The waveguide delivers light at various excitation wavelengths and transmits a fluorescent signal through optical filters to a set of highly sensitive photodetectors. Each sensor cartridge is also equipped with four reagent delivery ports per well for injecting testing compounds into wells during the assay. Prior to the start of the XF assay, the array of biosensors should be each independently calibrated using an automated routine that determines a unique sensor gain based on the sensor output measured in a calibration reagent of known pH and oxygen concentration. To prepare a sensor cartridge for calibration, add 1.0 ml of Seahorse Bioscience XF24 Calibrant (pH7.4) (Seahorse Bioscience, Part #100840-000) to each well of a Seahorse Bioscience 24-well plate, then place sensor cartridge on top of the plate and incubate at 37 °C without CO2 overnight or for up to 72 h. If the cartridge is being hydrated for more than 24 h, wrap parafilm around the edges to prevent evaporation. Finally, turn on the instrument and start XF software to allow instrument to stabilize at 37 °C. 2.3. Preparing cell plate for XF24 assay Cells must be equilibrated with running media (unbuffered media) prior to the XF24 measurements. Depending on the assay types, appropriate running medium such as unbuffered DMEM, RPMI or KHB should be used. Here we describe the preparation of a commonly used unbuffered DMEM as a sample. Dissolve DMEM base (Sigma D5030-1L) 8.3 g in 500 ml dH2O. Separately, dissolve 1.85 g NaCl (Sigma S3014) in 500 ml dH2O. Combine NaCl solution with DMEM Base solution. Remove 20 ml from the combined solution, add 10 ml 100 GlutaMax-1 (Gibco 35050-061), 10 ml 100 mM Sodium Pyruvate (Sigma S8636), and 15 mg Phenol Red (Sigma P5530). Add glucose (Sigma G8270) powder for desired concentration (4.5 g = 25 mM glucose). Warm media to 37 °C, adjust pH to 7.4 using 5 M NaOH, and then filter sterilize the media. Media can then be stored at 4 °C for later usage. Depending on the purpose of the XF assay, to study the effect of metabolic substrate of OXPHOS and glycolysis on OCR and ECAR, unbuffered DMEM could also be prepared with or without containing glucose, sodium pyruvate, or L-glutamine. The next day after overnight incubation of the cells plated in the XF24 cell culture plate, inspect the cells under a microscope to assure 80% confluence and even seeding. When cell condition is confirmed ready for assay, warm unbuffered media to 37 °C. Using an aspirator remove growth media from each well and leave approximately 50 ll of media behind in each well, taking care not to touch the bottom of the wells. It is also important not to remove the media completely to prevent exposure of the cells to air and potentially drying out. Then rinse wells by adding 1.0 ml of warm unbuffered media to each well. Remove unbuffered media, again leaving behind approximately 50 ll of media. Add 625 ll (final volume in each well will be 675 ll) of unbuffered media to each well (typically 1.0 ml for baseline runs, or a minimum of 0.5 ml for runs with compound injections). Finally, incubate the XF24 cell culture plate in a 37 °C incubator without CO2 until ready for use or at least 60 min prior to loading the cell plate in XF24 instrument. After changing the growth media to running media, load the appropriate XF template file onto the instrument software so it is ready to start just after loading the sensor cartridge with compounds.
2.2. Preparing sensor cartridge for the XF24 assay and turning on instrument
2.4. Loading the sensor cartridge with compounds
Each sensor cartridge is embedded with 24 pairs of fluorescent biosensors (oxygen and pH), which are coupled to fiber-optic
There are four injection ports located on the sensor cartridge for each well, designated as Port A, Port B, Port C, and Port D.
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Testing compounds should be loaded into the ports prior to beginning calibration of the sensor cartridge. Depending on the assay types, different compounds and different combinations of selected compounds can be used. We describe here the application of a combination of four metabolic inhibitors, oligomycin (Sigma O4876), Carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone (FCCP) (Sigma C2920), 2-deoxyglucose (2-DG) (Sigma D6134), and rotenone (Sigma R8875). The final concentration for each inhibitor is oligomycin (1 lg/ml), FCCP (300 nM), 2-DG (100 mM), and rotenone (1 lM). For new cell line analysis, the concentration for each inhibitor should be optimized base on cell types, especially FCCP concentration. Inappropriate dose of FCCP will not maximally activate oxygen consumption after oligomycin administration, thus will not be able to measure maximum total reserved mitochondrial respiration capacity, which will be described in Section 3.2. Because the compounds will be injected sequentially from Port A, B, C to D, and the recommended injection volume is 75 ll (although the injection volume can range from 25 to 100 ll), the injected compound will be diluted 10 for compound A, 11 for compound B, 12 for compound C, and 13 for compound D. Prepare a 10, 11, 12, and 13 fold higher concentration than the final concentration of compound A, B, C, and D with unbuffered media. All wells including Temperature Correction or blank wells need to have a control, therefore media or compounds should be loaded in all the ports to ensure proper injection in all wells. The notch on the bottom left of the cartridge denotes the front of the cartridge. To load the compounds, angling the pipet tips into the ports and gently loading the solution, making sure not to cause any leakage into the well below. Loading may be done with either a single channel pipet (one port at a time) or with an 8 or 12-channel pipet (all ports in one column or row at once, respectively).
Protocol Start 1. Calibrate probes 2. Loop 3 3. (1–3) Mix for 3 Min. 0 s 4. Time Delay of 2 Min. 0 s 5. (1–3) Measure for 3 Min. 0 s 6. Loop End 7. Inject Port A 8. Loop 3 9. (4–6) Mix for 2 Min. 0 s 10. Time Delay of 2 Min. 0 s 11. (4–6) Measure for 4 Min. 0 s 12. Loop End 13. Inject Port B 14. Loop 3 15. (7–9) Mix for 3 Min. 0 s 16. Time Delay of 2 Min. 0 s 18. (7–9) Measure for 3 Min. 0 s 19. Loop End 20. Inject Port C 21. Loop 3 22. (10–12) Mix for 4 Min. 0 s 23. Time Delay of 2 Min. 0 s 24. (10–12) Measure for 2 Min. 0 s 25. Loop End 26. Inject Port D 27. Loop 3 28. (13–15) Mix for 2 Min. 0 s 29. Time Delay of 2 Min. 0 s 30. (13–15) Measure for 4 Min. 0 s 31. Loop End Program End
2.5. Calibrating the sensors and running the experiment
2.6. Normalization of XF assay results to cell number
Login to the XF24 software and allow the XF24 analyzer to warm to 37 °C. The XF24 analyzer should be left on overnight so the instrument is at the proper temperature. Set up an experimental template by using the Assay Wizard. This template can be prepared in advance so it is ready to be loaded into the XF24 software. Open the appropriate assay (.xls) template by clicking the ‘‘Open” icon on the bottom of the screen and choosing the desired template. If using the Assay Wizard to design an XF experiment just before running, then this template will already be loaded. Then click ‘‘start” icon to start the program and load the sensor cartridge for calibration. After the calibration process, replace the sensor cartridge with cell plate that has been placed in a 37 °C incubator without CO2. At last, click ‘‘continue” icon to start measuring process. Prior to each rate measurement, the running medium in each well should be mixed for 2–5 min followed by a time delay for 2–5 min to allow the oxygen partial pressure and pH in the microenvironment surrounding the cells to reach equilibrium. Following mixing and time delay processes, OCR and ECAR are measured simultaneously for 2–5 min to establish a baseline rate. The running medium is then gently mixed again for 2–5 min between each rate measurement to restore normal oxygen tension and pH in the medium. After the baseline measurement, 25–100 ll of a testing compound solution will be injected into each well. To expedite the compound exposure to cells, the medium is then mixed for another 2–5 min. After mixing and time delay processes, OCR and ECAR measurements after compound injection are then performed. Measurements of OCR and ECAR at baseline and after each compound injection, may be repeated three-four times. The following is a typical measuring protocol that we use in our laboratory.
Due to different cell growth rate among comparing cell types and possible different cell number between well to well during cell seeding process, the cell number in each well should be counted after XF assay for normalization of OCR and ECAR results. To count the cell number of each well, remove media from each well to a 1.5 ml tube, then wash each well with 100 ll of 1 PBS. And then transfer PBS back to the same 1.5 ml tube. To detach cells, add 50 ll of trypsin–EDTA solution to each well and incubate in a 37 °C incubator. Inspect cells under a microscope and when cells appear rounded and are in suspension, remove the solution from each well to the corresponding 1.5 ml tube. Rinse each well using the media from the corresponding 1.5 ml tube. Remove the media back to the 1.5 ml tube, and monitor wells under a microscope to make sure all the cells are removed from the wells. Count cell number for each well using a haemocytometer and then normalize the XF assay results with cell number using XF assay software. 3. Application of XF24 assay in evaluating the metabolic profiles of cells depleted of mtDNA To study the effect of mtDNA alterations on OXPHOS and glycolysis, we used Seahorse XF24 analyzer together with ATP assay to monitor the metabolic profiles of cells depleted of mtDNA (rho0 cells). 3.1. Establishment of rho0 cells and confirmation of their mtDNA depletion MCF7 rho0 cell line was established by culturing human breast carcinoma MCF7 (ATCC HTB-22) cells in DMEM supplemented
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with 10% fetal bovine serum, 1% penicillin–streptomycin, 1 mM sodium pyruvate, 4.5 g/l glucose, 50 lg/ml uridine (Sigma U3003), and 50 ng/ml ethidium bromide (Sigma E1510) for four weeks [18,19]. The absence of mtDNA in rho0 cells was confirmed by quantitative polymerase chain reaction (QPCR) developed in our lab [20]. The primers used for the amplification of specific mitochondrial and nuclear DNA sequences were: Human Small mito primers HSMITO14620SENSE–CCCCACAAACCCCATTACTAAACCCA HMITO14841ANTI–TTTCATCATGCGGAGATGTTGGATGG Human Large mito primers HLMITO5999SENSE–TCTAAGCCTCCTTATTCGAGCCGA HMITO14841ANTI–TTTCATCATGCGGAGATGTTGGATGG Human polymerase beta primers HBPOL2372SENSE–CATGTCACCACTGGACTCTGCAC HBPOL3927ANTI–CCTGGAGTAGGAACAAAAATTGCTG As shown in Fig. 1, after culturing MCF7 cells in the presence of ethidium bromide for four weeks, neither small (0.22 kb) mtDNA nor large (8.9 kb) mtDNA sequences were amplified in MCF7 rho0 cells when compared with the strong amplification of both sequences in their parental control MCF7 cells. No difference was observed in the amplification of nuclear DNA sequence (polymerase beta gene, 12.2 kb). This result indicated the complete depletion of mtDNA in MCF7 rho0 cells. Mitochondrial morphology change due to the lack of mtDNA was analyzed by staining cells with MitoTracker Red (Invitrogen, MitoTrackerÒ Red CMXRos *special packaging*, Cat. No. M-7512). As shown in Fig. 2, depletion of mtDNA induced mitochondria undergoing dramatic fragmentation. 3.2. Profiles of cellular energy metabolism elucidated by the Seahorse XF24 extracellular flux analyzer To gain more insight into the effect of depletion of mtDNA on cellular energy metabolism, we performed an analysis of oxygen consumption rate (OCR, a measure of OXPHOS) and extracellular acidification rate (ECAR, a measure of lactate production and glycolysis) using the Seahorse XF24 extracellular flux analyzer.
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Fig. 1. PCR confirmation of mtDNA depletion in MCF7 rho0 cells. DNA was extracted from 1 106 cells using QIAGEN Genomic Tip and Genomic Buffer Set (QIAGEN, 10,323 and 19,060, respectively). Equal amount of 15 ng of DNA from each cell line was used to perform QPCR assay using GeneAmp XL PCR Kit (Applied Biosystems, N8080193). The QPCR cycle number for 0.22 kb small mito sequence was 27 cycles, for 8.9 kb large mito sequence was 20 cycles, and for 12.2 kb DNA polymerase beta was 27 cycles. The reaction without DNA was used as a negative control.
Fig. 2. Mitochondrial fragmentation due to depletion of mtDNA. MCF7 parental and rho0 cells were incubated with 100 nM MitoTracker Red solution for 20 min in a 37 °C incubator. Then the cells were fixed with 4% paraformaldehyde and permeabilized with 0.2% Triton X-100 in PBS. The cells were also counterstained with DAPI for nuclear. The cells were photographed with an OLYMPUS fluorescence microscope. (A) MCF7 parental cell. (B) MCF7 rho0 cell. Mitochondria (red), nuclear (blue).
We have developed a pharmacological profiling approach using the Seahorse instrument that uses four metabolic inhibitors to measure responses of OCR and ECAR to bioenergetic stress and applied this assay to parental and rho0 cells. OCR is reported in the unit of pico-moles (pMoles) per minute and ECAR in milli-pH (mpH) units per minute. The value of OCR and ECAR without any inhibitor administration is called basal OCR and basal ECAR (indicated in Fig. 3C). Compared with parental MCF7 cells, mtDNA depleted rho0 cells have significantly lower basal OCR (Fig. 3A), and higher basal ECAR (Fig. 3B), indicating greatly impaired mitochondrial respiration and compensatory increase in glycolysis for ATP generation in rho0 cells. After getting a basal level of OCR and ECAR, four compounds were injected sequentially at different time points into each well from Port A, B, C, and D, respectively. The first injection from Port A was oligomycin (1.0 lg/ml) at 20.05 min after starting the measuring protocol. Oligomycin inhibits ATP synthase causing a backup of protons in the intermembrane space, which causes subsequent loss of electron flow through the electron transport chain and loss of oxygen consumption. Administration of oligomycin invoked a concomitant decrease in OCR (Fig. 3A) and also a simultaneous increase in ECAR (Fig. 3B) in parental cells while no change was observed in rho0 cells. Analysis of steady-state levels of ATP (Fig. 4) showed that oligomycin treatment actually caused a slight, but significant increase in ATP, suggesting that glycolysis is capable of completely keeping up with the energy demands of
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Fig. 4. ATP generation in response to metabolic inhibitors. MCF7 parental and rho0 cells were seeded in a 96-well black culture plate and treated with 1 lg/ml oligomycin, 100 mM 2-DG and a combination of 1 lg/ml oligomycin with 100 mM 2-DG for 45 min. Then the ATP levels were determined with an ATPlite™ Luminescence Assay System. (A) Absolute ATP levels per cell. (B) Relative percentage of ATP levels compared with untreated cells (control). (*p < 0.05, **p < 0.01).
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Fig. 3. Pharmacological profile of oxygen consumption and lactate production. Oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were determined with a Seahorse XF24 analyzer. The metabolic inhibitors were injected sequentially from Port A (oligomycin), B (FCCP), C (2-DG), and D (rotenone), at different time point (20.05 min for Port A, 45.53 min for Port B, 71 min for Port C, and 96.48 min for Port D). The final concentration of each inhibitor is oligomycin (1 lg/ml), FCCP (300 nM), 2-DG (100 mM) and rotenone (1 lM). (A) OCR. (B) ECAR. (C) Detailed description of OCR profile elucidated through the use of metabolic inhibitors.
these MCF7 cells in the absence of ATP generation by oxidative phosphorylation. Although the functional significance and the mechanism of the increased ATP level as a response to mitochondrial dysfunction is not known. The quick response of increase in ECAR after oligomycin administration in parental cells indicated the high potential of cells in upregulating glycolysis to meet their energy need in the case of mitochondrial dysfunction. The OCR value after oligomycin treatment shows the amount of oxygen consumption that is not coupled to ATP production (proton leak). The basal OCR minus the oligomycin OCR level is equivalent to the amount of ATP-coupled oxygen consumption (indicated in Fig. 3C). The next injection from Port B was FCCP (300 nM) at 45.53 min, which elicited a rapid increase in OCR (Fig. 3A) in parental cells, while no change was observed in rho0 cells. FCCP uncouples mitochondrial respiration by carrying protons across the inner membrane, thus re-stimulating mitochondrial respiration by
dissipating mitochondrial proton gradient allowing electrons to flow to complex IV and allowing the resumption of oxygen consumption. Some cell types show a large increase in oxygen consumption after FCCP treatment [21]. No significant change was observed in ECAR in both parental and rho0 cells after FCCP administration. The third compound injected from Port C was 2-DG (100 mM) at 71 min, which inhibits hexokinase and the utilization of glucose in glycolysis or the TCA cycle. As expected this treatment decreased ECAR in both parental and rho0 cells (Fig. 3B). Surprisingly, 2-DG also induced a further increase of OCR in parental cells, but not in rho0 cells (Fig. 3A). These data suggest that in these cells, 2-DG administration after the other two inhibitors uncovers a total reserve capacity of mitochondria to increase respiration either in response to loss of ATP generation or through altered source of carbons for the TCA cycle. This increase in respiration potential, which is stimulated by administration with FCCP and 2-DG after oligomycin administration is the maximum total mitochondrial reserved respiration capacity (indicated in Fig. 3C). The rho0 cells show little response against 2-DG indicating that mitochondrial respiration capacity of rho0 cells is severely impaired, and these cells are therefore, not be able to upregulate mitochondrial respiration to compensate for the ATP depletion caused by 2-DG inhibition of glycolysis. The fourth and final compound that was injected through Port D was rotenone (1.0 lM) at 96.48 min. Administration of rotenone after exposure of the same cells to oligomycin, FCCP and 2-DG completely diminished OCR (Fig. 3A) and somewhat surprisingly further reduced ECAR (Fig. 3B). Rotenone is a complex I inhibitor, thus shuts down all total electron flow, diminishing all oxygen
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consumption through the mitochondria. Any residual oxygen consumption is through other cellular process such as P450 cytochromes or NADPH oxidases. 3.3. Profiles of cellular energy metabolism elucidated by ATP assay Depletion of mtDNA induces dysfunction of mitochondrial OXPHOS, decreases mitochondrial ATP generation, therefore, induces a compensatory increase in glycolysis contribution to total intracellular ATP generation. Oligomycin completely inhibits ATP synthase [22], thus oligomycin-inhibitable ATP value is usually considered as mitochondrial contribution to total intracellular ATP generation [23]. 2-DG blocks the first step of glycolysis pathway, thus 2-DG-inhibitable ATP value indicates the contribution of glycolysis to total intracellular ATP generation. And the combination of both oligomycin and 2-DG inhibits the total intracellular ATP generation. To study the effect of mtDNA depletion on ATP generation and the interplay between mitochondrial and glycolytic ATP generation, the steady-state intracellular ATP levels in MCF7 parental and rho0 cells and their responses to oligomycin and 2-DG administration were determined by using an ATPlite™ Luminescence Assay System (PerkinElmer, product number: 6016941). This system is based on the production of light caused by the reaction of ATP with added firefly (Photinus pyralis) luciferase and D-luciferin. The emitted light is proportional to the ATP concentration. The luminescence signal is measured using a Synergy 2 microplate reader (BioTek instruments, Winooski, VT). As shown in Fig. 4A, the absolute value of ATP level per cell indicated that depletion of mtDNA in rho0 cells increased the total ATP level when compared with the parental cells. The percentage value of ATP level shown in Fig. 4B indicated the difference between the contribution of mitochondria and glycolysis to total ATP generation. After the exposure of cells to oligomycin for 45 min, which inhibits the mitochondrial ATP generation, parental cells showed an increase in ATP generation. The increased ATP level observed after oligomycin administration in parental cells was similar to the ATP levels observed in rho0 cells without any inhibitor treatment (Fig. 4A), suggested that MCF7 cells have ability to increase the compensatory ATP generation through glycolysis in response to mitochondrial dysfunction, either as a chronic (mtDNA depletion by ethidium bromide treatment) or acute (oligomycin treatment) response. The change in the amount of ATP is consistent with the difference in ECAR value (Fig. 3B). While in rho0 cells, because of the mitochondrial dysfunction due to mtDNA depletion, cells were no longer able to respond to oligomycin. As shown in Fig. 4B, no significant change in ATP level in rho0 cells after treatment with oligomycin was observed. 2-DG treatment caused a decline in steady-state ATP levels to a greater degree in rho0 cells when compared with parental MCF7 cells (Fig. 4B), thus indicating that rho0 cells are more dependent on glycolysis for ATP generation. In parental cells, almost 60% of ATP was inhibited after 45 min treatment with 2-DG indicated that parental MCF7 cells also have a high reliance on glycolysis for their ATP generation, which is consistent with Warburg effect. The combination of oligomycin and 2-DG inhibits all ATP generation. Thus the value after administration with both inhibitors indicated the cellular ATP usage within 45 min (Fig. 4B). The further decrease of ATP level after the treatment with the combination of oligomycin and 2-DG when compared with 2-DG only treatment in parental cells, while no further decrease in rho0 cells, indicated that 2-DG inhibited all the ATP generation in rho0 cells,
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thus indicating that rho0 cells were solely dependent on glycolysis for ATP generation. In conclusion, mtDNA depletion eliminated mitochondrial respiration capacity and caused the inability of mitochondria to modulate OXPHOS in response to metabolic stresses. Simultaneously, mtDNA-depleted cells exhibited a large compensatory increase in glycolysis for the generation of ATP. 4. Closing remarks Prior to the Seahorse extracellular flux analyzer, cellular metabolic analysis was performed through monitoring separately the rate of oxygen consumption using Clark electrodes [24], or the rate of extracellular acidification using microphysiometer [25]. The using of Clark electrode is not amenable to multiple samples and requires large amounts of material. More recently, plate reader assays in conjunction with oxygen sensitive fluorophores have been developed [26,27]. The Seahorse XF analyzer is a novel instrument that enables the simultaneous and realtime measurements of the flux of oxygen and proton from low numbers of living cells (20,000–40,000) in a microplate over time. Furthermore, the multiwell plate based assay increases the throughput for each assay, and broadens the range of specimens from floating and attached cell cultures [7,28], tissue slices [29] to isolated mitochondria [30]. References [1] D.F. Rolfe, G.C. Brown, Physiol. Rev. 77 (1997) 731–758. [2] P.M. Herst, A.S. Tan, D.J. Scarlett, M.V. Berridge, Biochim. Biophys. Acta 1656 (2004) 79–87. [3] O. Warburg, F. Wind, E. Negelein, J. Gen. Physiol. 8 (1927) 519–530. [4] O. Warburg, Science 123 (1956) 309–314. [5] M.G. Vander Heiden, L.C. Cantley, C.B. Thompson, Science 324 (2009) 1029– 1033. [6] P.L. Pedersen, J. Bioenerg. Biomembr. 39 (2007) 211–222. [7] M. Wu, A. Neilson, A.L. Swift, R. Moran, J. Tamagnine, D. Parslow, S. Armistead, K. Lemire, J. Orrell, J. Teich, S. Chomicz, D.A. Ferrick, Am. J. Physiol. Cell Physiol. 292 (2007) C125–136. [8] G. Attardi, G. Schatz, Annu. Rev. Cell Biol. 4 (1988) 289–333. [9] W.C. Copeland, Annu. Rev. Med. 59 (2008) 131–146. [10] J. Lasota, Lab. Invest. 87 (2007) 316. [11] S.E. Durham, D.T. Brown, D.M. Turnbull, P.F. Chinnery, Neurology 67 (2006) 502–504. [12] H.C. Lee, P.H. Yin, J.C. Lin, C.C. Wu, C.Y. Chen, C.W. Wu, C.W. Chi, T.N. Tam, Y.H. Wei, Ann. NY Acad. Sci. 1042 (2005) 109–122. [13] N.S. Chandel, P.T. Schumacker, FEBS Lett. 454 (1999) 173–176. [14] L. Tarrago-Litvak, O. Viratelle, D. Darriet, R. Dalibart, P.V. Graves, S. Litvak, Nucleic Acids Res. 5 (1978) 2197–2210. [15] M.M. Nass, Exp. Cell Res. 72 (1972) 211–222. [16] R.D. Leibowitz, J. Cell Biol. 51 (1971) 116–122. [17] E. Zylber, C. Vesco, S. Penman, J. Mol. Biol. 44 (1969) 195–204. [18] M.P. King, G. Attardi, Science 246 (1989) 500–503. [19] M.P. King, G. Attardi, Methods Enzymol. 264 (1996) 304–313. [20] J.H. Santos, J.N. Meyer, B.S. Mandavilli, B. Van Houten, Methods Mol. Biol. 314 (2006) 183–199. [21] D.G. Nicholls, L. Johnson-Cadwell, S. Vesce, M. Jekabsons, N. Yadava, J. Neurosci. Res. 85 (2007) 3206–3212. [22] H.A. Lardy, D. Johnson, M.W. Mc, Arch. Biochem. Biophys. 78 (1958) 587–597. [23] A.L. Nieminen, A.K. Saylor, B. Herman, J.J. Lemasters, Am. J. Physiol. 267 (1994) C67–74. [24] L.C. Clark, Trans. Am. Soc. Artif. Int. Organs 2 (1956) 41–48. [25] J.C. Owicki, J.W. Parce, Biosens. Bioelectron. 7 (1992) 255–272. [26] F. Li, Y. Wang, K.I. Zeller, J.J. Potter, D.R. Wonsey, K.A. O’Donnell, J.W. Kim, J.T. Yustein, L.A. Lee, C.V. Dang, Mol. Cell Biol. 25 (2005) 6225–6234. [27] S.M. Schieke, D. Phillips, J.P. McCoy Jr., A.M. Aponte, R.F. Shen, R.S. Balaban, T. Finkel, J. Biol. Chem. 281 (2006) 27643–27652. [28] M. Watanabe, S.M. Houten, C. Mataki, M.A. Christoffolete, B.W. Kim, H. Sato, N. Messaddeq, J.W. Harney, O. Ezaki, T. Kodama, K. Schoonjans, A.C. Bianco, J. Auwerx, Nature 439 (2006) 484–489. [29] S.W. Choi, A.A. Gerencser, D.G. Nicholls, J. Neurochem. 109 (2009) 1179–1191. [30] J.A. Dykens, J.D. Jamieson, L.D. Marroquin, S. Nadanaciva, J.J. Xu, M.C. Dunn, A.R. Smith, Y. Will, Toxicol. Sci. 103 (2008) 335–345.
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Review Article
Alterations in bioenergetics due to changes in mitochondrial DNA copy number Wei Qian, Bennett Van Houten * Department of Pharmacology and Chemical Biology, University of Pittsburgh School of Medicine and The University of Pittsburgh Cancer Institute, Hillman Cancer Center, Pittsburgh, PA 15213, United States
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Article history: Accepted 22 March 2010 Available online 25 March 2010 Keywords: Mitochondria Mitochondrial DNA Oxygen consumption rate Oxidative phosphorylation Extracellular acidification rate Glycolysis ATP
a b s t r a c t All mitochondrial DNA (mtDNA)-encoded genes are involved in mitochondrial electron transport and ATP production. Alterations of mtDNA due to dysfunctional mitochondrial DNA polymerase gamma (POLG) induce loss of mitochondrial oxidative phosphorylation (OXPHOS) and mitochondrial ATP generation. Total intracellular ATP is generated by two energetic pathways, glycolysis and mitochondrial OXPHOS. Decreased ATP generation from mitochondria due to mitochondrial dysfunction induces compensatory upregulation of cytoplasmic glycolysis process, thus increasing the contribution of glycolysis to the total cellular ATP generation. Decreased mitochondrial respiration and ATP generation with concomitant enhanced glycolysis is associated with mitochondrial disease and cancer. This chapter introduces a novel assay using a pharmacological profiling strategy in combination with a Seahorse XF24 instrument, which quantifies mitochondrial oxygen consumption rate and extracellular acidification rate for the measurement of OXPHOS and glycolysis, respectively. This assay combined with an analysis of steady-state ATP levels was used to study the bioenergetics of cells depleted of mtDNA (rho0 cells). Ó 2010 Elsevier Inc. All rights reserved.
1. Introduction Oxidative phosphorylation (OXPHOS) is the culmination of a series of bioenergetic transformations that are called cellular respiration. Mitochondrial OXPHOS and cytoplasmic glycolysis are the two main metabolic pathways to generate cellular ATP. Through glycolysis one molecule of glucose is metabolized to two molecules of pyruvate with the concomitant net production of two molecules of ATP. In the absence of oxygen or due to mitochondrial dysfunction, pyruvate is converted to lactic acid, thus completing the glycolysis cycle. Under aerobic conditions, pyruvate is transported into mitochondria. In mitochondrial matrix, pyruvate is oxidatively decarboxylated to acetyl CoA. Complete oxidation of the acetyl unit of acetyl CoA through citric acid cycle and OXPHOS generates 26 of the total 30 molecules of ATP from one molecule of glucose. During OXPHOS, oxygen is reduced to water in a four-electron reduction at cytochrome c oxidase in mitochondrial electron transport chain (ETC). The amount of intracellular ATP that is generated from OXPHOS is variable in cells throughout the body [1], and more than 90% of the body’s oxygen is consumed by the ETC. The remaining 10% is comprised of non-mitochondrial respiration, including substrate oxidation and cell surface oxygen consumption [2].
* Corresponding author. Address: Hillman Cancer Center, 5117 Centre Avenue, Research Pavilion, Suite 2.6, Pittsburgh, PA 15213-1863, United States. Fax: +1 412 6237761. E-mail address: [email protected] (B. Van Houten). 1046-2023/$ - see front matter Ó 2010 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2010.03.006
Increased conversion of glucose to lactic acid even in the presence of oxygen associated with decreased mitochondrial respiration is a distinct characteristic of tumors, first described by Otto Warburg (Warburg effect) in the 1920s [3,4]. Although the mechanism of Warburg effect and its role in malignant transformation remains elusive [5,6], this phenomenon demonstrates the cellular flexibility in switching the main cellular energy production from OXPHOS to glycolysis during the process of tumorigenesis [7]. OXPHOS is executed by four mitochondrial protein complexes for electron transfer (NADH-Q oxidoreductase, succinate-Q reductase, Q-cytochrome c oxidoreductase, and cytochrome c oxidase, which are also called complex I, II, III, and IV, respectively) and one complex for ATP synthesis (ATP synthase, which is also called complex V or F1–F0 ATPase). Impairment in these complexes could cause dysfunction of OXPHOS. Mitochondrial DNA (mtDNA)-encoded genes are involved in the electron transport and production of ATP to carry out OXPHOS. Mammalian mtDNA encodes genes for 2 rRNAs, 22 tRNAs, and 13 protein subunits, which are the critical components of mitochondrial ETC complexes I, III, IV, and V. The subunits include seven subunits of NADH-Q reductase, one subunit of cytochrome c reductase, three subunits of cytochrome c oxidase, and two subunits of F1–F0 ATP synthase [8]. Defects in the replication and transcription of mtDNA induce mutation, deletion, or depletion of mtDNA, cause defects in assembly of OXPHOS complexes, which can eventually cause loss of OXPHOS. Mitochondrial DNA polymerase gamma (POLG), which is the only known DNA polymerase found in mammalian mitochondria, is responsible for replication and repair of mtDNA. Mutations in
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POLG are associated with mitochondrial depletion syndrome (MDS) such as progressive external ophthalmoplegia (PEO) [9]. mtDNA depletion is also associated with cardiomyopathy [10], myopathy [11], and cancer [12]. To study the contribution of mtDNA to OXPHOS and hence cellular function, rho0 cells (cells depleted of mtDNA) have been widely used as a successful model system [13]. Rho0 cell line can be established by treating cells with low concentration of ethidium bromide (3,8-diamino-5-ethyl-6-phenylphenanthridinium bromide). Ethidium bromide inhibits mtDNA POLG more strongly than DNA polymerase alpha and beta [14], thus inhibits the replication and transcription of mtDNA without substantially affecting nuclear DNA [15–17]. In order to monitor the alterations in cellular energy metabolism as a response to mtDNA impairment and gain a better understanding of the interplay between the two main bioenergetic pathways (OXPHOS and glycolysis), in this chapter we introduce the application of Seahorse XF24 Extracellular Flux analyzer (Seahorse Bioscience, Billerica, MA) in measuring the metabolic profiles (oxygen consumption rate, OCR and extracellular acidification rate, ECAR) of cells, and the use of rho0 cells as a model system for mtDNA depletion syndromes. 2. Analysis of OXPHOS and glycolysis using the Seahorse XF24 extracellular flux analyzer 2.1. Seeding cells in XF24 cell culture plate The XF24 cell culture plate (Seahorse Bioscience, Part #100777004) is formatted like a typical 24-well plate, with rows designated A–D and columns 1–6. The seeding surface of each well is the same size as a typical 96-well plate. Even seeding and around 90% confluence of cells are extremely important for XF assay. Uneven seeding and/or too less or overgrowth of cells can cause significant well to well variation, and may not truly reflects the cellular metabolic profiles and also cause large SD value. Resuspend cells to obtain a desired final concentration to seed in 100 ll of growth media. Typical seeding densities vary from 20,000 to 60,000 cells per well depending on cell size and basal metabolic rates. Seed 100 ll of cell suspension per well gently, leaving Temperature Correction wells blank (A1, B4, C3, D6). During seeding, rest the pipet tip just below the circular rim at the top of the well for homogenous cell layer. Then place plate in an incubator and allow cells to adhere. This generally takes approximately 1 h for strongly adherent cells, but may take up to 5–6 h for less adherent cells. After cells have adhered, add 150 ll of growth media to each well, bringing the total volume of media in the well to 250 ll. When adding media to the wells, add it slowly to the sides as to not disturb the newly attached cells. Then allow the cells to grow overnight in an incubator (37 °C, 5% CO2). For the assay of non-adherent cells, the cells can be attached to the bottom of wells by centrifugation of cell suspension in a XF24 plate coated with BD Cell-Tak Cell and Tissue Adhesive (BD Biosciences, CN 354240). A swinging bucket rotor centrifuge should be equipped with plate carriers, such as the Eppendorf Centrifuge 5810R. The assay for non-adherent cells does not require overnight culture of cells after centrifugation, thus cells can be suspended directly in unbuffered media for centrifugation and proceed the step of incubation in a 37 °C incubator without CO2 (Section 2.3).
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waveguides. The waveguide delivers light at various excitation wavelengths and transmits a fluorescent signal through optical filters to a set of highly sensitive photodetectors. Each sensor cartridge is also equipped with four reagent delivery ports per well for injecting testing compounds into wells during the assay. Prior to the start of the XF assay, the array of biosensors should be each independently calibrated using an automated routine that determines a unique sensor gain based on the sensor output measured in a calibration reagent of known pH and oxygen concentration. To prepare a sensor cartridge for calibration, add 1.0 ml of Seahorse Bioscience XF24 Calibrant (pH7.4) (Seahorse Bioscience, Part #100840-000) to each well of a Seahorse Bioscience 24-well plate, then place sensor cartridge on top of the plate and incubate at 37 °C without CO2 overnight or for up to 72 h. If the cartridge is being hydrated for more than 24 h, wrap parafilm around the edges to prevent evaporation. Finally, turn on the instrument and start XF software to allow instrument to stabilize at 37 °C. 2.3. Preparing cell plate for XF24 assay Cells must be equilibrated with running media (unbuffered media) prior to the XF24 measurements. Depending on the assay types, appropriate running medium such as unbuffered DMEM, RPMI or KHB should be used. Here we describe the preparation of a commonly used unbuffered DMEM as a sample. Dissolve DMEM base (Sigma D5030-1L) 8.3 g in 500 ml dH2O. Separately, dissolve 1.85 g NaCl (Sigma S3014) in 500 ml dH2O. Combine NaCl solution with DMEM Base solution. Remove 20 ml from the combined solution, add 10 ml 100 GlutaMax-1 (Gibco 35050-061), 10 ml 100 mM Sodium Pyruvate (Sigma S8636), and 15 mg Phenol Red (Sigma P5530). Add glucose (Sigma G8270) powder for desired concentration (4.5 g = 25 mM glucose). Warm media to 37 °C, adjust pH to 7.4 using 5 M NaOH, and then filter sterilize the media. Media can then be stored at 4 °C for later usage. Depending on the purpose of the XF assay, to study the effect of metabolic substrate of OXPHOS and glycolysis on OCR and ECAR, unbuffered DMEM could also be prepared with or without containing glucose, sodium pyruvate, or L-glutamine. The next day after overnight incubation of the cells plated in the XF24 cell culture plate, inspect the cells under a microscope to assure 80% confluence and even seeding. When cell condition is confirmed ready for assay, warm unbuffered media to 37 °C. Using an aspirator remove growth media from each well and leave approximately 50 ll of media behind in each well, taking care not to touch the bottom of the wells. It is also important not to remove the media completely to prevent exposure of the cells to air and potentially drying out. Then rinse wells by adding 1.0 ml of warm unbuffered media to each well. Remove unbuffered media, again leaving behind approximately 50 ll of media. Add 625 ll (final volume in each well will be 675 ll) of unbuffered media to each well (typically 1.0 ml for baseline runs, or a minimum of 0.5 ml for runs with compound injections). Finally, incubate the XF24 cell culture plate in a 37 °C incubator without CO2 until ready for use or at least 60 min prior to loading the cell plate in XF24 instrument. After changing the growth media to running media, load the appropriate XF template file onto the instrument software so it is ready to start just after loading the sensor cartridge with compounds.
2.2. Preparing sensor cartridge for the XF24 assay and turning on instrument
2.4. Loading the sensor cartridge with compounds
Each sensor cartridge is embedded with 24 pairs of fluorescent biosensors (oxygen and pH), which are coupled to fiber-optic
There are four injection ports located on the sensor cartridge for each well, designated as Port A, Port B, Port C, and Port D.
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Testing compounds should be loaded into the ports prior to beginning calibration of the sensor cartridge. Depending on the assay types, different compounds and different combinations of selected compounds can be used. We describe here the application of a combination of four metabolic inhibitors, oligomycin (Sigma O4876), Carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone (FCCP) (Sigma C2920), 2-deoxyglucose (2-DG) (Sigma D6134), and rotenone (Sigma R8875). The final concentration for each inhibitor is oligomycin (1 lg/ml), FCCP (300 nM), 2-DG (100 mM), and rotenone (1 lM). For new cell line analysis, the concentration for each inhibitor should be optimized base on cell types, especially FCCP concentration. Inappropriate dose of FCCP will not maximally activate oxygen consumption after oligomycin administration, thus will not be able to measure maximum total reserved mitochondrial respiration capacity, which will be described in Section 3.2. Because the compounds will be injected sequentially from Port A, B, C to D, and the recommended injection volume is 75 ll (although the injection volume can range from 25 to 100 ll), the injected compound will be diluted 10 for compound A, 11 for compound B, 12 for compound C, and 13 for compound D. Prepare a 10, 11, 12, and 13 fold higher concentration than the final concentration of compound A, B, C, and D with unbuffered media. All wells including Temperature Correction or blank wells need to have a control, therefore media or compounds should be loaded in all the ports to ensure proper injection in all wells. The notch on the bottom left of the cartridge denotes the front of the cartridge. To load the compounds, angling the pipet tips into the ports and gently loading the solution, making sure not to cause any leakage into the well below. Loading may be done with either a single channel pipet (one port at a time) or with an 8 or 12-channel pipet (all ports in one column or row at once, respectively).
Protocol Start 1. Calibrate probes 2. Loop 3 3. (1–3) Mix for 3 Min. 0 s 4. Time Delay of 2 Min. 0 s 5. (1–3) Measure for 3 Min. 0 s 6. Loop End 7. Inject Port A 8. Loop 3 9. (4–6) Mix for 2 Min. 0 s 10. Time Delay of 2 Min. 0 s 11. (4–6) Measure for 4 Min. 0 s 12. Loop End 13. Inject Port B 14. Loop 3 15. (7–9) Mix for 3 Min. 0 s 16. Time Delay of 2 Min. 0 s 18. (7–9) Measure for 3 Min. 0 s 19. Loop End 20. Inject Port C 21. Loop 3 22. (10–12) Mix for 4 Min. 0 s 23. Time Delay of 2 Min. 0 s 24. (10–12) Measure for 2 Min. 0 s 25. Loop End 26. Inject Port D 27. Loop 3 28. (13–15) Mix for 2 Min. 0 s 29. Time Delay of 2 Min. 0 s 30. (13–15) Measure for 4 Min. 0 s 31. Loop End Program End
2.5. Calibrating the sensors and running the experiment
2.6. Normalization of XF assay results to cell number
Login to the XF24 software and allow the XF24 analyzer to warm to 37 °C. The XF24 analyzer should be left on overnight so the instrument is at the proper temperature. Set up an experimental template by using the Assay Wizard. This template can be prepared in advance so it is ready to be loaded into the XF24 software. Open the appropriate assay (.xls) template by clicking the ‘‘Open” icon on the bottom of the screen and choosing the desired template. If using the Assay Wizard to design an XF experiment just before running, then this template will already be loaded. Then click ‘‘start” icon to start the program and load the sensor cartridge for calibration. After the calibration process, replace the sensor cartridge with cell plate that has been placed in a 37 °C incubator without CO2. At last, click ‘‘continue” icon to start measuring process. Prior to each rate measurement, the running medium in each well should be mixed for 2–5 min followed by a time delay for 2–5 min to allow the oxygen partial pressure and pH in the microenvironment surrounding the cells to reach equilibrium. Following mixing and time delay processes, OCR and ECAR are measured simultaneously for 2–5 min to establish a baseline rate. The running medium is then gently mixed again for 2–5 min between each rate measurement to restore normal oxygen tension and pH in the medium. After the baseline measurement, 25–100 ll of a testing compound solution will be injected into each well. To expedite the compound exposure to cells, the medium is then mixed for another 2–5 min. After mixing and time delay processes, OCR and ECAR measurements after compound injection are then performed. Measurements of OCR and ECAR at baseline and after each compound injection, may be repeated three-four times. The following is a typical measuring protocol that we use in our laboratory.
Due to different cell growth rate among comparing cell types and possible different cell number between well to well during cell seeding process, the cell number in each well should be counted after XF assay for normalization of OCR and ECAR results. To count the cell number of each well, remove media from each well to a 1.5 ml tube, then wash each well with 100 ll of 1 PBS. And then transfer PBS back to the same 1.5 ml tube. To detach cells, add 50 ll of trypsin–EDTA solution to each well and incubate in a 37 °C incubator. Inspect cells under a microscope and when cells appear rounded and are in suspension, remove the solution from each well to the corresponding 1.5 ml tube. Rinse each well using the media from the corresponding 1.5 ml tube. Remove the media back to the 1.5 ml tube, and monitor wells under a microscope to make sure all the cells are removed from the wells. Count cell number for each well using a haemocytometer and then normalize the XF assay results with cell number using XF assay software. 3. Application of XF24 assay in evaluating the metabolic profiles of cells depleted of mtDNA To study the effect of mtDNA alterations on OXPHOS and glycolysis, we used Seahorse XF24 analyzer together with ATP assay to monitor the metabolic profiles of cells depleted of mtDNA (rho0 cells). 3.1. Establishment of rho0 cells and confirmation of their mtDNA depletion MCF7 rho0 cell line was established by culturing human breast carcinoma MCF7 (ATCC HTB-22) cells in DMEM supplemented
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with 10% fetal bovine serum, 1% penicillin–streptomycin, 1 mM sodium pyruvate, 4.5 g/l glucose, 50 lg/ml uridine (Sigma U3003), and 50 ng/ml ethidium bromide (Sigma E1510) for four weeks [18,19]. The absence of mtDNA in rho0 cells was confirmed by quantitative polymerase chain reaction (QPCR) developed in our lab [20]. The primers used for the amplification of specific mitochondrial and nuclear DNA sequences were: Human Small mito primers HSMITO14620SENSE–CCCCACAAACCCCATTACTAAACCCA HMITO14841ANTI–TTTCATCATGCGGAGATGTTGGATGG Human Large mito primers HLMITO5999SENSE–TCTAAGCCTCCTTATTCGAGCCGA HMITO14841ANTI–TTTCATCATGCGGAGATGTTGGATGG Human polymerase beta primers HBPOL2372SENSE–CATGTCACCACTGGACTCTGCAC HBPOL3927ANTI–CCTGGAGTAGGAACAAAAATTGCTG As shown in Fig. 1, after culturing MCF7 cells in the presence of ethidium bromide for four weeks, neither small (0.22 kb) mtDNA nor large (8.9 kb) mtDNA sequences were amplified in MCF7 rho0 cells when compared with the strong amplification of both sequences in their parental control MCF7 cells. No difference was observed in the amplification of nuclear DNA sequence (polymerase beta gene, 12.2 kb). This result indicated the complete depletion of mtDNA in MCF7 rho0 cells. Mitochondrial morphology change due to the lack of mtDNA was analyzed by staining cells with MitoTracker Red (Invitrogen, MitoTrackerÒ Red CMXRos *special packaging*, Cat. No. M-7512). As shown in Fig. 2, depletion of mtDNA induced mitochondria undergoing dramatic fragmentation. 3.2. Profiles of cellular energy metabolism elucidated by the Seahorse XF24 extracellular flux analyzer To gain more insight into the effect of depletion of mtDNA on cellular energy metabolism, we performed an analysis of oxygen consumption rate (OCR, a measure of OXPHOS) and extracellular acidification rate (ECAR, a measure of lactate production and glycolysis) using the Seahorse XF24 extracellular flux analyzer.
0.22 kb 8.9 kb
N
o
D
N
A
R ho 0
Pa re nt al
12.2 kb
Fig. 1. PCR confirmation of mtDNA depletion in MCF7 rho0 cells. DNA was extracted from 1 106 cells using QIAGEN Genomic Tip and Genomic Buffer Set (QIAGEN, 10,323 and 19,060, respectively). Equal amount of 15 ng of DNA from each cell line was used to perform QPCR assay using GeneAmp XL PCR Kit (Applied Biosystems, N8080193). The QPCR cycle number for 0.22 kb small mito sequence was 27 cycles, for 8.9 kb large mito sequence was 20 cycles, and for 12.2 kb DNA polymerase beta was 27 cycles. The reaction without DNA was used as a negative control.
Fig. 2. Mitochondrial fragmentation due to depletion of mtDNA. MCF7 parental and rho0 cells were incubated with 100 nM MitoTracker Red solution for 20 min in a 37 °C incubator. Then the cells were fixed with 4% paraformaldehyde and permeabilized with 0.2% Triton X-100 in PBS. The cells were also counterstained with DAPI for nuclear. The cells were photographed with an OLYMPUS fluorescence microscope. (A) MCF7 parental cell. (B) MCF7 rho0 cell. Mitochondria (red), nuclear (blue).
We have developed a pharmacological profiling approach using the Seahorse instrument that uses four metabolic inhibitors to measure responses of OCR and ECAR to bioenergetic stress and applied this assay to parental and rho0 cells. OCR is reported in the unit of pico-moles (pMoles) per minute and ECAR in milli-pH (mpH) units per minute. The value of OCR and ECAR without any inhibitor administration is called basal OCR and basal ECAR (indicated in Fig. 3C). Compared with parental MCF7 cells, mtDNA depleted rho0 cells have significantly lower basal OCR (Fig. 3A), and higher basal ECAR (Fig. 3B), indicating greatly impaired mitochondrial respiration and compensatory increase in glycolysis for ATP generation in rho0 cells. After getting a basal level of OCR and ECAR, four compounds were injected sequentially at different time points into each well from Port A, B, C, and D, respectively. The first injection from Port A was oligomycin (1.0 lg/ml) at 20.05 min after starting the measuring protocol. Oligomycin inhibits ATP synthase causing a backup of protons in the intermembrane space, which causes subsequent loss of electron flow through the electron transport chain and loss of oxygen consumption. Administration of oligomycin invoked a concomitant decrease in OCR (Fig. 3A) and also a simultaneous increase in ECAR (Fig. 3B) in parental cells while no change was observed in rho0 cells. Analysis of steady-state levels of ATP (Fig. 4) showed that oligomycin treatment actually caused a slight, but significant increase in ATP, suggesting that glycolysis is capable of completely keeping up with the energy demands of
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A
375
A
B
C
D
rho0
A
12
275 225 175 125 75
-25 8
75
A
B
C
D
parental rho0
B 140
55
*
**
parental rho0
120 45
100
35
ATP (%)
ECAR (mpH/min)
Oligomycin 2-DG Oligomycin (1 µg/ml) (1 µg/ml) (100 mM) + 2-DG(100 mM)
control
Time ( min)
25 15
80
** **
60 40
5
20
-5 0
8
17 24 33 41 52 60 69 77 86 94 101 109 118
Time (min)
A
375
B
C
ATP 2-DG coupled activated
325
OCR (pMoles/min)
4
17 24 33 41 52 60 69 77 86 94 101 109 118
65
Basal
6
0 0
C
8
2
25
B
parental rho0
**
10
ATP (fmoles/cell)
325
OCR (pMoles/min)
parental
0
D
parental rho0
275 225
Total reserved capacity
175 125
control
Oligomycin 2-DG Oligomycin (1 µg/ml) (1 µg/ml) (100 mM) + 2-DG(100 mM)
Fig. 4. ATP generation in response to metabolic inhibitors. MCF7 parental and rho0 cells were seeded in a 96-well black culture plate and treated with 1 lg/ml oligomycin, 100 mM 2-DG and a combination of 1 lg/ml oligomycin with 100 mM 2-DG for 45 min. Then the ATP levels were determined with an ATPlite™ Luminescence Assay System. (A) Absolute ATP levels per cell. (B) Relative percentage of ATP levels compared with untreated cells (control). (*p < 0.05, **p < 0.01).
75 25 -25 0
8
17 24 33 41 52 60 69 77 86 94 101 109 118
Time ( min)
Proton leak
FCCP activated
Fig. 3. Pharmacological profile of oxygen consumption and lactate production. Oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were determined with a Seahorse XF24 analyzer. The metabolic inhibitors were injected sequentially from Port A (oligomycin), B (FCCP), C (2-DG), and D (rotenone), at different time point (20.05 min for Port A, 45.53 min for Port B, 71 min for Port C, and 96.48 min for Port D). The final concentration of each inhibitor is oligomycin (1 lg/ml), FCCP (300 nM), 2-DG (100 mM) and rotenone (1 lM). (A) OCR. (B) ECAR. (C) Detailed description of OCR profile elucidated through the use of metabolic inhibitors.
these MCF7 cells in the absence of ATP generation by oxidative phosphorylation. Although the functional significance and the mechanism of the increased ATP level as a response to mitochondrial dysfunction is not known. The quick response of increase in ECAR after oligomycin administration in parental cells indicated the high potential of cells in upregulating glycolysis to meet their energy need in the case of mitochondrial dysfunction. The OCR value after oligomycin treatment shows the amount of oxygen consumption that is not coupled to ATP production (proton leak). The basal OCR minus the oligomycin OCR level is equivalent to the amount of ATP-coupled oxygen consumption (indicated in Fig. 3C). The next injection from Port B was FCCP (300 nM) at 45.53 min, which elicited a rapid increase in OCR (Fig. 3A) in parental cells, while no change was observed in rho0 cells. FCCP uncouples mitochondrial respiration by carrying protons across the inner membrane, thus re-stimulating mitochondrial respiration by
dissipating mitochondrial proton gradient allowing electrons to flow to complex IV and allowing the resumption of oxygen consumption. Some cell types show a large increase in oxygen consumption after FCCP treatment [21]. No significant change was observed in ECAR in both parental and rho0 cells after FCCP administration. The third compound injected from Port C was 2-DG (100 mM) at 71 min, which inhibits hexokinase and the utilization of glucose in glycolysis or the TCA cycle. As expected this treatment decreased ECAR in both parental and rho0 cells (Fig. 3B). Surprisingly, 2-DG also induced a further increase of OCR in parental cells, but not in rho0 cells (Fig. 3A). These data suggest that in these cells, 2-DG administration after the other two inhibitors uncovers a total reserve capacity of mitochondria to increase respiration either in response to loss of ATP generation or through altered source of carbons for the TCA cycle. This increase in respiration potential, which is stimulated by administration with FCCP and 2-DG after oligomycin administration is the maximum total mitochondrial reserved respiration capacity (indicated in Fig. 3C). The rho0 cells show little response against 2-DG indicating that mitochondrial respiration capacity of rho0 cells is severely impaired, and these cells are therefore, not be able to upregulate mitochondrial respiration to compensate for the ATP depletion caused by 2-DG inhibition of glycolysis. The fourth and final compound that was injected through Port D was rotenone (1.0 lM) at 96.48 min. Administration of rotenone after exposure of the same cells to oligomycin, FCCP and 2-DG completely diminished OCR (Fig. 3A) and somewhat surprisingly further reduced ECAR (Fig. 3B). Rotenone is a complex I inhibitor, thus shuts down all total electron flow, diminishing all oxygen
W. Qian, B. Van Houten / Methods 51 (2010) 452–457
consumption through the mitochondria. Any residual oxygen consumption is through other cellular process such as P450 cytochromes or NADPH oxidases. 3.3. Profiles of cellular energy metabolism elucidated by ATP assay Depletion of mtDNA induces dysfunction of mitochondrial OXPHOS, decreases mitochondrial ATP generation, therefore, induces a compensatory increase in glycolysis contribution to total intracellular ATP generation. Oligomycin completely inhibits ATP synthase [22], thus oligomycin-inhibitable ATP value is usually considered as mitochondrial contribution to total intracellular ATP generation [23]. 2-DG blocks the first step of glycolysis pathway, thus 2-DG-inhibitable ATP value indicates the contribution of glycolysis to total intracellular ATP generation. And the combination of both oligomycin and 2-DG inhibits the total intracellular ATP generation. To study the effect of mtDNA depletion on ATP generation and the interplay between mitochondrial and glycolytic ATP generation, the steady-state intracellular ATP levels in MCF7 parental and rho0 cells and their responses to oligomycin and 2-DG administration were determined by using an ATPlite™ Luminescence Assay System (PerkinElmer, product number: 6016941). This system is based on the production of light caused by the reaction of ATP with added firefly (Photinus pyralis) luciferase and D-luciferin. The emitted light is proportional to the ATP concentration. The luminescence signal is measured using a Synergy 2 microplate reader (BioTek instruments, Winooski, VT). As shown in Fig. 4A, the absolute value of ATP level per cell indicated that depletion of mtDNA in rho0 cells increased the total ATP level when compared with the parental cells. The percentage value of ATP level shown in Fig. 4B indicated the difference between the contribution of mitochondria and glycolysis to total ATP generation. After the exposure of cells to oligomycin for 45 min, which inhibits the mitochondrial ATP generation, parental cells showed an increase in ATP generation. The increased ATP level observed after oligomycin administration in parental cells was similar to the ATP levels observed in rho0 cells without any inhibitor treatment (Fig. 4A), suggested that MCF7 cells have ability to increase the compensatory ATP generation through glycolysis in response to mitochondrial dysfunction, either as a chronic (mtDNA depletion by ethidium bromide treatment) or acute (oligomycin treatment) response. The change in the amount of ATP is consistent with the difference in ECAR value (Fig. 3B). While in rho0 cells, because of the mitochondrial dysfunction due to mtDNA depletion, cells were no longer able to respond to oligomycin. As shown in Fig. 4B, no significant change in ATP level in rho0 cells after treatment with oligomycin was observed. 2-DG treatment caused a decline in steady-state ATP levels to a greater degree in rho0 cells when compared with parental MCF7 cells (Fig. 4B), thus indicating that rho0 cells are more dependent on glycolysis for ATP generation. In parental cells, almost 60% of ATP was inhibited after 45 min treatment with 2-DG indicated that parental MCF7 cells also have a high reliance on glycolysis for their ATP generation, which is consistent with Warburg effect. The combination of oligomycin and 2-DG inhibits all ATP generation. Thus the value after administration with both inhibitors indicated the cellular ATP usage within 45 min (Fig. 4B). The further decrease of ATP level after the treatment with the combination of oligomycin and 2-DG when compared with 2-DG only treatment in parental cells, while no further decrease in rho0 cells, indicated that 2-DG inhibited all the ATP generation in rho0 cells,
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thus indicating that rho0 cells were solely dependent on glycolysis for ATP generation. In conclusion, mtDNA depletion eliminated mitochondrial respiration capacity and caused the inability of mitochondria to modulate OXPHOS in response to metabolic stresses. Simultaneously, mtDNA-depleted cells exhibited a large compensatory increase in glycolysis for the generation of ATP. 4. Closing remarks Prior to the Seahorse extracellular flux analyzer, cellular metabolic analysis was performed through monitoring separately the rate of oxygen consumption using Clark electrodes [24], or the rate of extracellular acidification using microphysiometer [25]. The using of Clark electrode is not amenable to multiple samples and requires large amounts of material. More recently, plate reader assays in conjunction with oxygen sensitive fluorophores have been developed [26,27]. The Seahorse XF analyzer is a novel instrument that enables the simultaneous and realtime measurements of the flux of oxygen and proton from low numbers of living cells (20,000–40,000) in a microplate over time. Furthermore, the multiwell plate based assay increases the throughput for each assay, and broadens the range of specimens from floating and attached cell cultures [7,28], tissue slices [29] to isolated mitochondria [30]. References [1] D.F. Rolfe, G.C. Brown, Physiol. Rev. 77 (1997) 731–758. [2] P.M. Herst, A.S. Tan, D.J. Scarlett, M.V. Berridge, Biochim. Biophys. Acta 1656 (2004) 79–87. [3] O. Warburg, F. Wind, E. Negelein, J. Gen. Physiol. 8 (1927) 519–530. [4] O. Warburg, Science 123 (1956) 309–314. [5] M.G. Vander Heiden, L.C. Cantley, C.B. Thompson, Science 324 (2009) 1029– 1033. [6] P.L. Pedersen, J. Bioenerg. Biomembr. 39 (2007) 211–222. [7] M. Wu, A. Neilson, A.L. Swift, R. Moran, J. Tamagnine, D. Parslow, S. Armistead, K. Lemire, J. Orrell, J. Teich, S. Chomicz, D.A. Ferrick, Am. J. Physiol. Cell Physiol. 292 (2007) C125–136. [8] G. Attardi, G. Schatz, Annu. Rev. Cell Biol. 4 (1988) 289–333. [9] W.C. Copeland, Annu. Rev. Med. 59 (2008) 131–146. [10] J. Lasota, Lab. Invest. 87 (2007) 316. [11] S.E. Durham, D.T. Brown, D.M. Turnbull, P.F. Chinnery, Neurology 67 (2006) 502–504. [12] H.C. Lee, P.H. Yin, J.C. Lin, C.C. Wu, C.Y. Chen, C.W. Wu, C.W. Chi, T.N. Tam, Y.H. Wei, Ann. NY Acad. Sci. 1042 (2005) 109–122. [13] N.S. Chandel, P.T. Schumacker, FEBS Lett. 454 (1999) 173–176. [14] L. Tarrago-Litvak, O. Viratelle, D. Darriet, R. Dalibart, P.V. Graves, S. Litvak, Nucleic Acids Res. 5 (1978) 2197–2210. [15] M.M. Nass, Exp. Cell Res. 72 (1972) 211–222. [16] R.D. Leibowitz, J. Cell Biol. 51 (1971) 116–122. [17] E. Zylber, C. Vesco, S. Penman, J. Mol. Biol. 44 (1969) 195–204. [18] M.P. King, G. Attardi, Science 246 (1989) 500–503. [19] M.P. King, G. Attardi, Methods Enzymol. 264 (1996) 304–313. [20] J.H. Santos, J.N. Meyer, B.S. Mandavilli, B. Van Houten, Methods Mol. Biol. 314 (2006) 183–199. [21] D.G. Nicholls, L. Johnson-Cadwell, S. Vesce, M. Jekabsons, N. Yadava, J. Neurosci. Res. 85 (2007) 3206–3212. [22] H.A. Lardy, D. Johnson, M.W. Mc, Arch. Biochem. Biophys. 78 (1958) 587–597. [23] A.L. Nieminen, A.K. Saylor, B. Herman, J.J. Lemasters, Am. J. Physiol. 267 (1994) C67–74. [24] L.C. Clark, Trans. Am. Soc. Artif. Int. Organs 2 (1956) 41–48. [25] J.C. Owicki, J.W. Parce, Biosens. Bioelectron. 7 (1992) 255–272. [26] F. Li, Y. Wang, K.I. Zeller, J.J. Potter, D.R. Wonsey, K.A. O’Donnell, J.W. Kim, J.T. Yustein, L.A. Lee, C.V. Dang, Mol. Cell Biol. 25 (2005) 6225–6234. [27] S.M. Schieke, D. Phillips, J.P. McCoy Jr., A.M. Aponte, R.F. Shen, R.S. Balaban, T. Finkel, J. Biol. Chem. 281 (2006) 27643–27652. [28] M. Watanabe, S.M. Houten, C. Mataki, M.A. Christoffolete, B.W. Kim, H. Sato, N. Messaddeq, J.W. Harney, O. Ezaki, T. Kodama, K. Schoonjans, A.C. Bianco, J. Auwerx, Nature 439 (2006) 484–489. [29] S.W. Choi, A.A. Gerencser, D.G. Nicholls, J. Neurochem. 109 (2009) 1179–1191. [30] J.A. Dykens, J.D. Jamieson, L.D. Marroquin, S. Nadanaciva, J.J. Xu, M.C. Dunn, A.R. Smith, Y. Will, Toxicol. Sci. 103 (2008) 335–345.