Measurements of Threshold of Mitochondrial Permeability Transition Pore Opening in Intact and Permeabilized Cells by Flash Photolysis of Caged Calcium

Protocol
Part of the Methods in Molecular Biology book series (MIMB, volume 793)

Abstract

Changes in intracellular calcium concentration play a major role both in signal transduction and in cell death. In particular, mitochondrial Ca2+ overload is critically important as a determinant of irreversible cell injury. When accumulated above a threshold, matrix Ca2+ triggers opening of the mitochondrial permeability transition pore (mPTP), initiating ATP depletion and cell death via necrosis or by promoting cytochrome c release and initiating the apoptotic cascade. Measurement of mitochondrial Ca2+ uptake capacity (or the threshold for mPTP opening) is, therefore, important for understanding the mechanisms of pathophysiology in a variety of disease models and also for testing neuro- or cardioprotective drugs. We have, therefore, devised an approach that delivers Ca2+ directly to the matrix of mitochondria independently of uptake and therefore independently of potential (Δψm) that allows direct study both of the Ca2+ efflux pathway and of the specific sensitivity of mPTP to Ca2+. This is achieved using the photolytic release of Ca2+ by flash photolysis of caged Ca2+ using compounds, such as o-nitrophenyl EGTA, introduced into the cell as the acetoxymethyl (AM) ester (NP-EGTA, AM). This method can be used in both intact and permeabilized cells.

Key words

Caged Ca2+ Flash photolysis Mitochondria Permeability transition pore 

1 Introduction

Changes in intracellular Ca2+ concentration [Ca2+]c play a major role both in signal transduction and in cell death. In particular, mitochondrial Ca2+ overload is critically important as a determinant of irreversible cell injury. This principle has emerged as a common theme recapitulated in a number of widely different pathological states, perhaps best established as a mechanism of ischaemic reperfusion injury in the heart and in the CNS (1, 2), but also in widely different disease states, including the collagen VI deficiencies in Ullrich and Bethlem myopathies (3) and in acute pancreatitis (4). Therefore, measurement and analysis of mitochondrial responses to Ca2+ overload is a useful part of the experimental repertoire in trying to address mechanisms of cell injury in many model systems, including age-related neurodegenerative diseases, such as Alzheimer’s and Parkinson’s diseases (5).

When [Ca2+]c rises close to mitochondria, they accumulate Ca2+, which moves down its electrochemical gradient through an electrogenic uniporter that facilitates Ca2+ transport across the inner mitochondrial membrane into the matrix. The electrochemical gradient favours Ca2+ uptake as the matrix [Ca2+]m is kept low by the activity of Ca2+/nNa+ and/or Ca2+/2H+ exchangers (6, 7) and by the negative mitochondrial membrane potential (Δψm). Net Ca2+ accumulation occurs when the rate of influx exceeds the capacity of the exchangers to remove Ca2+. This seems to occur at a cytosolic [Ca2+] of around 500 nM, often referred to as the set point for Ca2+ accumulation, but it also depends on the mitochondrial membrane potential and the cytosolic Na+ concentration.

Under physiological conditions, raised intramitochondrial [Ca2+]m stimulates the enzyme activity of the tricarboxylic acid (TCA) cycle, thereby increasing mitochondrial oxidative phosphorylation and energy production (8). This is thought to represent a valuable mechanism that matches energy supply to demand. Accumulated above a certain threshold, matrix Ca2+ can trigger opening of the mitochondrial permeability transition pore (mPTP), especially if a rise in [Ca2+]c is associated with oxidative or nitrosative stress (9). The PTP opening usually triggers cell death, either by precipitating rapid ATP depletion and necrosis or – in cells with a high glycolytic capacity – by causing mitochondrial cytochrome c release and initiating an apoptotic cascade. In either case, this is a catastrophic event that signals cell death.

Measurement of mitochondrial uptake Ca2+ capacity (or the threshold for mPTP opening) is, therefore, important for understanding mechanisms of the pathology and also for testing neuro- or cardioprotective drugs. Many approaches use isolated mito-chondria or bulk measurements from permeabilized cells and titration of the Ca2+ capacity by the progressive application of external Ca2+ (10, 11). In intact cells, the threshold of mPTP opening can be estimated by using different concentrations of electrogenic Ca2+ uniporters or by causing mitochondrial oxidative stress through illumination of photosensitising mitochondrially localized fluorescent dyes (12, 13). However, we have devised an approach that allows these measurements to be made at the level of a single cell, where this approach is a requirement of the experimental system. Thus, the experimental system demands that these measurements are made at the level of single cells when cell numbers are small, the population is heterogeneous, or when a modest proportion of cells are transfected and express a protein of interest.

Furthermore, it is important to bear in mind that, as mitochondrial Ca2+ uptake depends on Δψm, comparative studies of mPTP threshold may be biased by differences in effective Ca2+ uptake in populations in which Δψm varies. We have, therefore, devised an approach that delivers Ca2+ directly to the matrix of mitochondria independently of uptake and therefore independently of Δψm that allows a direct study of the efflux pathway and the specific sensitivity of mPTP to Ca2+. This is achieved by using flash photolysis of caged Ca2+ compounds, such as o-nitrophenyl EGTA, which are introduced into the cell as the acetoxymethyl (AM) ester (NP-EGTA, AM). This method can be used in intact and permeabilized cells, although results in intact cells can be misinterpreted because Ca2+ is released both in the cytosol and in the mitochondrial matrix. Permeabilization of cells with digitonin in a pseudo-intracellular solution localizes the caged Ca2+ signal to the matrix of mitochondria only.

The principle of the “caged” Ca2+ is that NP-EGTA is a photolabile Ca2+ chelator, whose affinity diminishes greatly following exposure to UV light (the Kd changes following the UV exposure from 80 nM to >1 mM). Thus, NP-EGTA acts as a chelator that binds Ca2+avidly when loaded into cells, but releases Ca2+as the affinity falls following a UV flash.

Measurement of mPTP opening can be achieved using cells co-loaded with a fluorescent Ca2+ indicator and with probes for mitochondrial membrane potential (fluo-4 and tetramethylrhodamine methylester (TMRM), respectively). The loss of mitochondrial membrane potential in response to a rise in [Ca2+]c in intact or permeabilized cells can be attributed to mPTP opening, if it can be prevented by cyclosporine A (CsA), the archetypal inhibitor of the mPTP (14).

2 Materials

  1. 1.

    Fluo-4AM (Molecular Probes/Invitrogen) is supplied in aliquots. Dissolve an aliquot by addition of 50 μl dry DMSO to a tube before use, and store at −20°C for up to 1 month.

     
  2. 2.

    TMRM (Molecular Probes/Invitrogen) is water soluble. Prepare a 5 μM stock solution either in a physiological saline or in distilled water.

     
  3. 3.

    NP-EGTA, AM (Molecular Probes/Invitrogen) is also supplied in aliquots. Dissolve in DMSO to reach a final concentration of 1 mM in the tube before use, and store at −20°C.

     
  4. 4.

    Pluronic (Molecular Probes/Invitrogen) is soluble in DMSO. Prepare a 2% stock solution.

     
  5. 5.

    Use a dish that holds a glass coverslip in which cells are grown, and buffer while imaging cells or grow cells in a Petri dish with a glass coverslip base.

     
  6. 6.

    Confocal microscope equipped with HeNe (543- or 555-nm wavelength), Argon (488-nm wavelength), and UV lasers (351 and 360 nm) and ideally with inverted optics. Use a quartz UV-compatible objective lens and make sure that the UV laser is accurately focused on the sample by setting up the collimator correctly (seeNote 1).

     
  7. 7.

    HEPES-buffered salt solution (HBSS): 156 mM NaCl, 3 mM KCl, 2 mM MgSO4, 1.25 mM KH2PO4, 2 mM CaCl2, 10 mM glucose, and 10 mM HEPES; pH adjusted to 7.35 with NaOH. This buffer is used to bathe cells in a static chamber. This avoids the need for continuous CO2 equilibrated superperfusion. The chamber is ideally heated to 37°C.

     
  8. 8.

    “Pseudo-intracellular” solution: 135 mM KCl, 10 mM NaCl, 20 mM HEPES, 5 mM pyruvate, 5 mM malate, 0.5 mM KH2PO4, 1 mM MgCl2, 5 mM EGTA, and 1.86 mM CaCl2 (to yield a free [Ca2+] of ∼100 nM), pH 7.1  +  20 μM digitonin  +  40 nM TMRM.

     

3 Methods

TMRM partitions across the cell membrane and is accumulated by polarized mitochondria due to the delocalized positive charge of this organic lipophilic cation. According to Nernstian principles, at a mitochondrial potential of −180 mV, the intramitochondrial concentration of TMRM is about 1,000-fold greater than that in the cytosol. In most cells, TMRM takes about 30 min to reach full equilibration between compartments.

Fluo-4AM (5 μM) and NP-EGTA, AM (10 μM) cross the cell membrane, where the AM ester is cleaved by cellular esterases. Both NP-EGTA and fluo-4 as AM esters distribute evenly in the cytosol and cell organelles. Once the AM ester group is cleaved, the charged moiety remains trapped within a compartment. Loading the cells at room temperature may increase AM dye loading into the mitochondria, as lower temperatures reduce the esterase activity, allowing more time for accumulation of the dye into these organelles. NP-EGTA, AM is loaded into cells without Ca2+. After intracellular cleavage of the ester group, NP-EGTA binds Ca2+ with a high affinity, forming a complex NP-EGTA: Ca2+ both in the mitochondrial matrix and in the cytosol. It seems that the Ca2+ then re-equilibrates, as the intracellular homeostatic mechanisms maintain normal levels of free [Ca2+]. Low temperatures reduce the rate of physiological processes, including mitochondrial metabolism, and limit Ca2+ “loading” of the chelator, and we have found that this reduces Ca2+ release following flash photolysis.

The optimum loading protocol depends on the cell type and has to be determined empirically. The Kd of NP-EGTA Ca2+ increases following UV illumination from 80 nM to >1 mM. UV “flashes” can be applied using a UV confocal and the intensity of the flash can be tuned to allow repeated uncaging, until mPTP opening is seen as the (CsA sensitive) release of fluo-4 and TMRM from the mitochondria. Each UV flash releases a set fraction of the caged Ca2+ (which depends on the flash intensity and efficiency of the optics), and so inevitably the absolute amount of Ca2+ released with successive flashes decreases. We have found that conditions can be established in several model systems such that wild-type cells maintain Ca2+ and membrane potential over many repeated flashes while cells expressing a mitochondrial pathology associated with neurodegeneration (e.g. in cells from the PINK1 knockout mouse; Figs. 1 and 2, and see also ref. 5) show mPTP opening after several flashes.
Fig. 1.

Uncaging Ca2+ induces [Ca2+]m overload and mitochondrial depolarization in a neuron. PINK1 knockout neurons demonstrate inhibition of the mitochondrial Na+/Ca2+ exchanger and a reduced threshold for mPTP opening (5). The images show selected frames imaging cells loaded with o-nitrophenyl EGTA, fluo-4, and TMRM. Images selected from a time series showing the responses to flash photolysis of the caged Ca2+ in the area indicated by the dashed line. The trace below shows the continuous intensity measurements as a function of time at the area marked in the images with an arrow. The arrows on the trace indicate times of UV-induced flash photolysis. The photolysis-induced rise in [Ca2+]c resulted in a dramatic increase in [Ca2+]m, as demonstrated by the fluo-4 signal in the mitochondrial area (note the co-localization of TMRM and the bright fluo-4 fluorescence in the images at 85″). This was rapidly followed by mitochondrial depolarization and the subsequent release of fluo-4 from the mitochondria. The calibration bar indicates 10 μm.

Fig. 2.

Measurements of mitochondrial Ca2+capacity in permeabilized cells. Flash photolysis of permeabilized neurons co-loaded with fluo-4 and TMRM demonstrated flash-induced increases in [Ca2+]m followed by the concurrent release of Ca2+ and complete mitochondrial depolarization. The images show selected frames from a time series taken at the times indicated while the trace below shows the continuous changes in signal with time in a small area as indicated. In this experiment, the whole field of view shown was “flashed” with UV at time points indicated by the blue arrows. The calibration red bar indicates 20 μm.

Caveat. In intact cells, a UV flash increases Ca2+ concentration in both the cytosol and the mitochondria inducing elevation of [Ca2+]m both through direct release from the marix-localized caged compound and from uniporter-mediated uptake. Subsequent UV flashes in a short period of time (every 1 min) induced step-like increase of mitochondrial Ca2+ and mitochondrial depolarization (Fig. 1). This method allows detection of mPTP threshold, but also reveals the kinetics of mitochondrial Ca2+ efflux. Increases of [Ca2+]c by flash photolysis in cells, in which the Na+/Ca2+ exchanger is inhibited (either pharmacologically using the drug CGP37157 or, as we have found recently, in the case of PINK1-deficient neurons (5)), cause the appearance of a very bright fluo-4 signal co-localizing perfectly with the TMRM signal, reflecting high levels of [Ca2+]m (Fig. 1).

In permeabilized cells, TMRM can be used as an indicator of Δψm, but fluo-4 is generally useful only to indicate mitochondrial integrity, as fluo-4 is a relatively high-affinity Ca2+ indicator (Kd  =  345 nM) and the signal tends to saturate (seeNote 2). It is possible to use a low-affinity indicator for the same purpose (fluo-4 FF or Calcium Green 5N), although the Ca2+ threshold for mPTP opening in healthy mitochondria (it may be higher than 300 μM) is likely higher than the Kd of these indicators as well.

With mPTP opening, the disappearance of individual mitochondria can be detected by the loss of both fluo-4 and TMRM fluorescence.

3.1 Preparing the Cells

  1. 1.

    Place glass coverslips at the bottom of 6-well tissue culture dishes (seeNote 3).

     
  2. 2.

    Cells should be plated so that the density of the cells in the coverslip achieves ∼70% confluence when they are imaged. In the mixed hippocampal cultures that we frequently use, containing a mixture of neurons and glia, this refers primarily to the astrocytes, as the neurons tend to grow unevenly over a coverslip.

     
  3. 3.

    Cells should be trypsinized and plated at least 24 h before the experiment to allow them to attach to the coverslip and recover.

     

3.2 Loading Fluo-4 AM, o-Nitrophenyl EGTA, AM, and TMRM into the Cells

  1. 1.

    In a 1.5-ml Eppendorf tube, add 5 μM fluo-4 AM, 10 μM NP-EGTA, AM, 25 nM TMRM  +  0.05% pluronic, then add 1 ml HBSS buffer, and mix well.

     
  2. 2.

    Replace the buffer on the cells with HBSS buffer containing fluo-4 AM, o-nitrophenyl EGTA, AM, TMRM, and pluronic.

     
  3. 3.

    Leave the cells for 30 min at room temperature to equilibrate the dyes.

     
  4. 4.

    Place the cells in a tissue culture incubator at 37°C for 20 min for de-esterification of the dyes.

     
  5. 5.

    The buffer used for imaging should contain 25 nM TMRM for the whole duration of the experiment and in all solutions used prior to permeabilization, when the concentration is increased to compensate for the loss of the concentrating power of the plasma membrane potential.

     

3.3 Confocal Imaging of Fluo-4 and TMRM Fluorescence

  1. 1.

    Transfer the coverslip containing the cells to the imaging chamber.

     
  2. 2.

    Wipe any liquid from underneath the coverslip using a tissue and place the chamber on the objective of the microscope, adding a drop of oil to the objective, if necessary. Ensure that a UV-compatible quartz objective is used, as this is required for the UV-induced uncaging.

     
  3. 3.

    Using phase-contrast or transmitted light, adjust the focus until the cells are clearly visible.

     
  4. 4.

    In the confocal microscope software, choose appropriate imaging optics for imaging fluo-4, i.e. excitation using the 488-nm line of the laser, and use a band-pass filter between 505 and 550 nm to collect emitted light. This is most important – if a long-pass filter is used, light from TMRM (which is also excited at 488 nm) will be included in the signal measured. For excitation of TMRM, use the 543-nm line of the laser while emitted light collected between 560 and 630 nm (or using a long-pass filter of >560 nm).

     
  5. 5.

    Keep the laser power to a minimum and the gain high as far as it is compatible with reasonable signal-to-noise ratios (we usually operate at ∼1% or less) to avoid damaging the cells.

     
  6. 6.

    Start the continuous scan, and adjust the focus until mitochondria are clearly visible.

     
  7. 7.

    Open the pinhole setting in the software to give an optical section of approximately 3 μm and decrease the laser power and/or gain until a signal of ∼50% intensity saturation is obtained. Use the lowest laser power compatible with a reasonable signal/noise to avoid phototoxicity.

     
  8. 8.

    Stop continuous scanning.

     
  9. 9.

    Increase image averaging (e.g. to average four frames) and/or decrease the scan speed to obtain a high-quality image to obtain clear resolution of mitochondria.

     
  10. 10.

    Most microscope control software have settings called “bleach” protocols. Using these settings, you are able to set up the requirements for the flash photolysis – we usually set the UV laser flash power at ∼50%, but precise conditions have to be tuned according to your optical arrangement.

     
  11. 11.

    The software also allows you to select a region to be “bleached” – use this to choose the sector of the cell(s) to be exposed to UV.

     
  12. 12.

    Start the scan to obtain the images of fluo-4 and TMRM fluorescence in the cells and flash chosen regions of interest with UV.

     

3.4 Permeabilization of the Cells

  1. 1.

    Obtain the image of the cells in the selected areas of the coverslip.

     
  2. 2.

    Start a time series scanning images repeatedly.

     
  3. 3.

    During the scanning, replace HBSS in the chamber with “pseudo-intracellular” solution as described above (Subheading 2, Item 8). We do this simply using an Eppendorf pipette. Perfusion systems with these experiments are complicated by the continuous presence of the dye in the saline – perfusion requires large volumes of dye which become expensive and also stain all the tubing used.

     
  4. 4.

    When visualizing plasma membrane permeabilization – signalled by egress of fluo-4 from the cytosol, for example, and co-localization of the remaining fluo-4 signal with TMRM in the mitochondria – wash out the digitonin and replace the buffer with the “pseudo-intracellular” solution  +  40 nM TMRM, but without digitonin.

     
  5. 5.

    Choose the sector of the cell(s) to be exposed to UV.

     
  6. 6.

    Start the scan to obtain the images of fluo-4 and TMRM fluorescence in the cells and flash chosen sectors with UV.

     
  7. 7.

    Analysis of the data is straightforward, as most software packages – ImageJ, Metamorph, The Zeiss, or Leica proprietary software – allow plotting of “Regions of Interest” intensities with time. As the dyes used are of single wavelength only, we would normally quantify the changes simply in terms of a “fold” change in signal.

     

4 Notes

  1. 1.

    It is useful to bear in mind that once mitochondria are loaded with fluo-4, the dye is trapped within the mitochondria and is released from mitochondria only in case of membrane destruction or mPTP opening.

     
  2. 2.

    In order to obtain images of sufficient resolution to distinguish mitochondria and in order to effectively release Ca2+ from the complex with NP-EGTA, it is essential to image the cells grown on a glass coverslip rather than imaging directly in the plastic tissue culture dish. An alternative is to grow cells on glass-bottomed tissue culture dishes or to use an upright microscope and image on plastic.

     
  3. 3.

    We do this by imaging NADH autofluorescence, which is excited at 351 nm and emits between 430 and 480 nm. If this is set up optimally, then the laser will be correctly aligned and will be optimal for flash photolysis.

     

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Copyright information

© Springer Science+Business Media, LLC 2011

Authors and Affiliations

  1. 1.Department of Cell and Developmental Biology and Consortium for Mitochondrial ResearchUniversity College LondonLondonUK
  2. 2.Department of Molecular NeuroscienceUCL Institute of NeurologyLondonUK

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