Cytosolic calcium regulation in rat afferent vagal neurons during anoxia

Cell Calcium 54 (2013) 416–427

Contents lists available at ScienceDirect

Cell Calcium journal homepage: www.elsevier.com/locate/ceca

Cytosolic calcium regulation in rat afferent vagal neurons during anoxia Michael Henrich a,∗,1 , Keith J. Buckler b a b

Department of Anaesthesia, Intensive Care Medicine, Pain Therapy, Justus-Liebig-University Giessen, Rudolf-Buchheim-Str. 7, D-35392 Giessen, Germany Department of Physiology, Anatomy and Genetics, Sherrington Building, Parks Road, Oxford OX1 3PT, UK

a r t i c l e

i n f o

Article history: Received 4 August 2013 Received in revised form 30 September 2013 Accepted 6 October 2013 Available online 17 October 2013 Keywords: Calcium Vagal neurons Anoxia Mitochondria ER SERCA

a b s t r a c t Sensory neurons are able to detect tissue ischaemia and both transmit information to the brainstem as well as release local vasoactive mediators. Their ability to sense tissue ischaemia is assumed to be primarily mediated through proton sensing ion channels, lack of oxygen however may also affect sensory neuron function. In this study we investigated the effects of anoxia on isolated capsaicin sensitive neurons from rat nodose ganglion. Acute anoxia triggered a reversible increase in [Ca2+ ]i that was mainly due to Ca2+ -efflux from FCCP sensitive stores and from caffeine and CPA sensitive ER stores. Prolonged anoxia resulted in complete depletion of ER Ca2+ -stores. Mitochondria were partially depolarised by acute anoxia but mitochondrial Ca2+ -uptake/buffering during voltage-gated Ca2+ -influx was unaffected. The process of Ca2+ -release from mitochondria and cytosolic Ca2+ -clearance following Ca2+ influx was however significantly slowed. Anoxia was also found to inhibit SERCA activity and, to a lesser extent, PMCA activity. Hence, anoxia has multiple influences on [Ca2+ ]i homeostasis in vagal afferent neurons, including depression of ATP-driven Ca2+ -pumps, modulation of the kinetics of mitochondrial Ca2+ buffering/release and Ca2+ -release from, and depletion of, internal Ca2+ -stores. These effects are likely to influence sensory neuronal function during ischaemia. © 2013 Elsevier Ltd. All rights reserved.

1. Introduction Sensory neurons of the nodose ganglia (NG) [1], innervate multiple visceral organs of the thorax and the abdomen and transmit various forms of sensory information to the nucleus tractus solitarii (NTS) [2–5]. Each NG contains approximately 6000 neurons [6,7], the majority of which are chemo-sensing although there are also mechano-sensitive and bimodal (mechano- and chemo-sensitive) neurones as well [8–10]. During tissue ischaemia chemo-sensing neurons not only contribute to the transmission of pain [10–12] but also release NO, calcitonin-gene related peptide (CGRP) and substance P directly into their target organ (via axonal reflexes) which can modulate arterial vasomotion [2,13–15]. Excitation of chemosensitive neurons during ischaemia is thought to be mediated, in part at least, through tissue acidosis and the activation of proton sensing ion channels including TRPV1 which cause membrane depolarisation and calcium influx [11,16–21] [22–24].

∗ Corresponding author at: Department of Anaesthesia, Intensive Care Medicine, Pain Therapy, Rudolf-Buchheim-Str. 7, 35392 Gießen, Germany. Tel.: +49 641 98544401; fax: +49 641 98544409. E-mail address: [email protected] (M. Henrich). 1 First Author. 0143-4160/$ – see front matter © 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.ceca.2013.10.001

One of the principle causes of acidosis during ischaemia is the rapid decline in tissue oxygen levels [25]. Within a few minutes the core of the ischaemic zone approaches anoxia and is surrounded by a halo of hypoxic tissue [26–28]. This oxygen depletion impairs oxidative phosphorylation leading to increased anaerobic metabolism (and thus acidosis). Lack of perfusion compounds the effects of hypoxia with the loss of supply of metabolic substrates thus endangering ATP supply and the accumulation of ions in extracellular spaces can severely alter ionic distribution across cell membranes. Hypoxia may also affect ion channel activity [29,30]. In sensory neurons of the glossopharyngeal nerve hypoxia inhibits background potassium currents causing membrane depolarisation and voltage-gated calcium-influx [31,32]. Hypoxia can also promote increased generation of reactive oxygen species [33]. The effects of ischaemia upon sensory neurons are therefore likely to be multifactorial and not limited simply to the activation of proton gated channels. In previous investigations performed on DRG neurons we found evidence that anoxia causes mitochondrial depolarisation, partial ATP depletion and intracellular acidification [34] within a few minutes. However, DRG neurons were still able to regulate intracellular Ca2+ -concentration ([Ca2+ ]i ), albeit at an elevated level. Mitochondrial Ca2+ -buffering was maintained but whilst ATP-driven Ca2+ pumps of the ER and the plasma membrane remained functional their activity was significantly diminished [35].

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

The effects of oxygen depletion on electrical signalling and Ca2+ homeostasis in vagal afferent neurons are unknown. In the present study we have used small capsaicin sensitive neurons isolated from NG. We observed multiple effects of anoxia on cytosolic [Ca2+ ]i regulation involving Ca2+ -stores, Ca2+ -pumps, the recycling of Ca2+ buffered by mitochondria and an increase in the resting level of [Ca2+ ]i ; but did not observe any significant effect upon membrane potential. These data indicate that anoxia fundamentally alters Ca2+ -signalling in these neurons in a manner that is likely to influence their response to ischaemia. 2. Materials and methods 2.1. Ganglion preparation and neuron dissociation Rats (aged between 6 and 8 weeks, of either sex, 130–170 g) were sacrificed in accordance with schedule 1 of the UK Animals (Scientific Procedures) Act 1986. Both NG were gently removed under sterile conditions and were immediately transferred into cooled Ca2+ - and Mg2+ -free phosphate-buffered saline (PBS), pH 7.4. After cleaning from connective tissue and remaining nerve bundles, the ganglia were cut into 8–10 smaller pieces which were then incubated in a medium comprising of liberase 1 (Roche, Burgess Hill, UK) with an activity of 1.4 U/g (Wünsch units), diluted in DMEM medium together with 0.75 mg trypsin (9.3 U/mg, Sigma, T-4665), in PBS and with 60 ␮M CaCl2 and 33 ␮M MgCl2 for 30 min at 37 ◦ C. Following enzyme treatment NG were washed once in PBS (Ca2+ - and Mg2+ -free) and twice in DMEM (containing 10% FBS, 1.2 mM l-glutamine), before mechanical trituration in 1.5 ml of DMEM. Afterwards the dissociated neurons were washed twice by centrifugation (at 1000 g for 5 min) in fresh DMEM. Following the final wash the cell pellet was resuspended in 400 ␮l Basal TNB100 culture medium, (containing protein–lipid-complex, 100 IU/ml penicillin, 100 ␮g/ml streptomycin and 10 ␮M NGF) and given a final trituration followed by seeding onto poly-l-lysine- and laminin-coated coverslips. These coverslips where placed in sterile culture dishes and incubated at 37 ◦ C in 5% CO2 /95% air for 2 h to allow the neurons to adhere to the coverslips before a further 3 ml of TNB was added to each culture dish. NG neurons were kept in the incubator for 30 min to 24 h before use. 2.2. Single cell fluorescence measurements Fluorescence measurements were performed as described before [34], using a Nikon Diaphot 200 (Japan) microscope equipped with a monochromator (Cairn Instruments, Kent) and cooled (−20 ◦ C) photomultiplier tubes (PMT; Thorn EMI). In individual NG neurons Fura-2 was excited alternately at 340 nm and 380 nm (±8 nm) for 250 ms at each wavelength with cycle repetition of 1 Hz. Emitted Fura-2 fluorescence was filtered at 510 nm (±20 nm). Rhodamine 123 (Rh123) was excited continuously at 495 nm and its emitted fluorescence filtered at 525 ± 10 nm. The output from the PMT was integrated over each illumination period (Fura-2) or averaged over 500 ms (Rh123) and recorded on a computer using a micro 1401 and Spike 4 software (Cambridge Electronic Design). For Fura-2 the fluorescence ratio (340 nm/380 nm) was also calculated and recorded using Spike 4 software. 2.3. Selection and superfusion of neurons Isolated NG neurons were placed in a recording chamber with a volume of approximately 100 ␮l. The perfusion rate of this chamber was adjusted at nearly 2 ml/min. All solutions were kept in glass bottles in a water bath (see solutions) and were gravity fed to the

417

recording chamber via stainless steel tubing (medical grade) and short sections of pharmed tubing (Norton performance plastics, UK, 34). Change of solutions was facilitated by a mechanically driven two-way tap that was placed close to the inflow of the recording chamber. Solution temperature was maintained at 37 ◦ C by a heating element surrounding a short section of stainless steel tubing between the tap and the chamber. This setup provided tight control over temperature and gas levels in the superfusate solutions together with rapid solution exchange [34]. Sensory neurons were selected initially on the basis of soma size (12–35 ␮m) and, in experiments utilising Fura-2, their response to capsaicin (100 nM for 10 s) was tested at the end of each experiment. In these studies >60% of neurons selected by the above size criteria proved to be capsaicin positive (i.e. capsaicin evoked a robust increase in [Ca2+ ]i ). Only capsaicin positive neurons were included in our studies on [Ca2+ ]i regulation. 2.4. Loading and calibration of Fura-2 NG neurons were loaded with Fura-2 by incubating them in either a HEPES buffered saline (for in vivo calibrations) or a bicarbonate buffered saline (for experiments) containing 5 ␮M Fura-2-AM (Molecular Probes, Leiden, NL) at room temperature for 25 min in a dark chamber. The HEPES buffered saline used comprised (in mM): HEPES: 20, glucose: 11, KCl: 4.5, MgCl2 : 1, CaCl2 : 2.5, NaCl: 117, pH 7.4 at room temperature. In vivo calibrations were performed by incubating Fura-2 loaded neurons in a zero-Ca2+ calibration medium containing 150 mM KCl, 5 mM NaCl, 1 mM EDTA, 1 mM EGTA and 10 ␮M ionomycin, for 10–20 min at room temperature. After this pre-incubation the neurons were placed in the perfusion chamber of the microspectrofluorometer and perfused with zero-Ca2+ medium (+1 ␮M ionomycin) at 37 ◦ C. After a 5 min perfusion Fura-2 fluorescence was recorded in 5 identified sensory neurons. The ratio of fluorescence obtained under these conditions was deemed equivalent to the calibration constant Rmin [36]. The perfusate was then changed to a high-Ca2+ calibration medium containing 150 mM KCl, 5 mM NaCl, 10 mM CaCl2 and 1 ␮M ionomycin. The change in fluorescence ratio was followed in one of the 5 identified neurons until it reached a new stable value and then the fluorescence ratio in it, and in the other 4 identified neurons, was recorded and deemed to be equivalent to the calibration constant Rmax . The ratio of fluorescence at 380 nm in zero-Ca2+ medium divided by that obtained in high-Ca2+ medium (Sf2 /Sb2 ) was also calculated for each neuron. The mean values obtained for Rmin , Rmax and (Sf2 /Sb2 ) were then used to calibrate measurements of the fluorescence ratio in subsequent experiments using the equation [Ca2+ ] = (R − Rmin )/(Rmax − R) × (Sf2 /Sb2 ) × Kd [36]. 2.5. Measurement of mitochondrial membrane potential with Rhodamine 123 Changes in mitochondrial membrane potential (m ) were detected using Rh123. This is a membrane permeant cation that is strongly sequestered in mitochondria due to their negative membrane potential. In concentrated solutions, as occur within the mitochondrion, Rh123 fluorescence is quenched. If the mitochondria become depolarised Rh123 is redistributed from the mitochondrion to the cytosol where it becomes diluted and as a consequence fluorescence increases (de quenching). Measurements of total Rh123 fluorescence from an intact cell can therefore be used to follow changes in m [37]. Neurons were loaded with Rh123 (5 ␮M) in bicarbonate buffered medium at room temperature for 12 min. Raw data (Fig. 4) is presented with Rh123 fluorescence expressed simply as a % of the starting value.

418

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

2.6. Whole cell patch clamp recordings

2.9. Statistics

For all whole cell patch clamp recordings fire polished glass pipettes were manufactured from borosilicate glass capillaries with 1.5 mm outer and 0.86 mm inner diameter (Harvard, Edenbridge, UK). The pipettes were fire polished and had a pipette resistance of 4–7 M when filled with the solutions described below. For conventional whole cell recording the pipette filling solution contained (in mM): KCl: 110, MgCl2 : 6, EGTA: 5, CaCl2 : 1.6, HEPES: 20, NaCl: 10, KOH: 40, K2 ATP: 5. For perforated patch whole cell recording the pipette filling solution contained either (a) KCl: 110, MgCl2 : 6, EGTA: 5, CaCl2 : 1.6, HEPES: 20, NaCl: 10, KOH: 40, K2 ATP: 5, pH 7.3 at room temperature or (b) K2 SO4 : 55, KCl: 30, MgCl2 : 5, EGTA: 1, HEPES: 20, pH 7.2 at room temperature. Nystatin (25 mg/ml in DMSO) was added to these solutions within 1 h of use to a final concentration of 500 ␮g/ml. Pipettes were filled by dipping in nystatin free solution first before backfilling with the nystatin containing solution. Whole cell recordings were performed using an Axopatch 200B amplifier (Axon Instruments, USA). Series resistance (Rs ) was compensated by ≥80% and membrane currents were filtered at 2 kHz. Seal resistances ranged between 3.5 and 7.5 G. Spike 4.21 software was used to generate voltage commands (see below) and to record membrane currents (digitised at 5 kHz).

Values are expressed as mean ± standard error of mean (S.E.M.). Statistical significance was tested using the paired Students t-test, or Wilcoxon ranks signed test for experiments with non-Gaussian distribution, calculated by SPSS 12.0 software for windows. Level of significance was set at p < 0.05.

2.6.1. Ramp protocols Depolarising ramps were used to record changes of membrane current at potentials close to the cells normal resting potential (see results). The ramp protocol had a total duration of 1000 ms and comprised of the following sequential steps (1) a holding potential of −70 mV for 150 ms, (2) a step to −100 mV for 100 ms. (3) a ramp from −100 mV to −40 mV over 500 ms, (4) a holding potential of −40 mV for 50 ms, (5) a step back to a holding potential of −70 mV for 200 ms. This protocol was repeated at 1 Hz. 2.7. Solutions Standard bicarbonate buffered Tyrode solutions contained (in mM): NaCl: 117, KCl: 4.5, CaCl2 : 2.5, MgCl2 : 1, HCO3 − : 23, glucose: 11. In Ca2+ -free solutions CaCl2 was omitted and 1 mM EGTA added. High-K+ Tyrode contained 50 mM KCl and 71.5 mM NaCl, all other constituents remained the same. Equilibration of these solutions with 5% CO2 and 95% air, achieved normoxic conditions with pH 7.4 at 37 ◦ C. Hypoxic solutions were generated by equilibration with 5% CO2 and 95% N2 (pO2 = 2 Torr). Anoxic solutions were obtained by the further addition of 0.5 mM Na2 S2 O4 [38] following 15–30 min prior equilibration with 5% CO2 /95% N2 (pH: 7.39 ± 0.02, n = 23). All solutions were kept in stoppered glass bottles and equilibrated with appropriate gas mixes at 37 ◦ C in a water bath for at least 30 min before use. In some electrophysiological experiments tetraethylammonium (10 mM, TEA) and 4-aminopyridine (5 mM, 4-AP) were added to external solutions to inhibit voltage-activated K+ channels (see Section 3). 2.8. Drugs Protein–lipid-complex was from Biochrom (Berlin, Germany). Ryanodine was from Tocris (Avonmouth, UK). All other chemicals were from either Sigma (Poole, UK) or VWR/BDH (Nottingham, UK). Cyclopiazonic acid (CPA) and thapsigargin containing solutions were prepared from stock solutions in DMSO. Capsaicin and FCCP were added from stock solutions in ethanol. The maximum concentration of solvent in Tyrodes were 50 ␮M DMSO and 10 ␮M ethanol.

3. Results 3.1. Changes in resting [Ca2+ ]i during anoxia in nodose ganglion neurons Neurons investigated in this study were all capsaicin-sensitive. The resting [Ca2+ ]i in these neurons was 166 ± 9 nM (n = 53). Hypoxia (pO2 2 Torr) induced a small but significant rise in [Ca2+ ]i by 34 ± 2.6 nM (n = 8) from 156 ± 2 nM, (n = 8) to 182 ± 2 nM (n = 8; p < 0.05). When neurons were exposed to anoxia the [Ca2+ ]i rose significantly from 167 ± 6 nM by 85 ± 7.7 nM to 259 ± 11 nM (n = 38, p < 0.001). This [Ca2+ ]i response was reversible and repeatable with subsequent anoxic periods. A similar [Ca2+ ]i response to anoxia was also seen in Ca2+ -free solution ([Ca2+ ]i = 73 ± 12 nM, n = 23, p < 0.001) and in normal Tyrode containing 1 mM Ni2+ ([Ca2+ ]i = 68 ± 17 nM, n = 5, p > 0.05). Thus neither removal of extracellular calcium nor inhibition of voltage-gated Ca2+ -channels with Ni2+ significantly attenuated the [Ca2+ ]i response to anoxia (see Fig. 1). This increase in [Ca2+ ]i is unlikely to be an artefact due to MgATP hydrolysis and an increase in cytosolic Mg2+ since we have previously measured the effects of anoxia on cytosolic [Mg2+ ]i and shown an increase of about 0.23–0.36 mM from a resting level of 1.6 mM which is too small to significantly interfere with the Fura-2 signal [34,35]. 3.2. Effects of anoxia on membrane potential The mean resting membrane potential of capsaicin sensitive NG neurons was −54 ± 0.7 mV (n = 12). In less than half of neurons investigated anoxia evoked a small but distinct and sustained depolarisation of 3.2 ± 0.5 mV (n = 7, p = 0.07, Fig. 2A). Similarly in some neurons voltage clamped at a holding potential at −70 mV, anoxia evoked a very small inward current but neither of these effects reached statistical significance. Similarly measurements of membrane currents active over a range of potentials from −100 to −40 mV (Fig. 2C) failed to detect any consistent effect of anoxia. Thus under resting conditions the effects of anoxia on the electrophysiological properties of these neurons are minimal or nonexistent. Equally the calcium response to anoxia was not diminished by voltage clamp ([Ca2+ ]i 76 ± 11 nM, n = 7). In contrast capsaicin (100 nM) evoked a robust depolarisation and firing of action potentials together with a large increase in [Ca2+ ]i (Fig. 2A). Similarly acidosis (pH0 5.0) triggered a biphasic inward current that was accompanied by a biphasic rise in [Ca2+ ]i . 3.3. Role of Ca2+ -stores in the anoxic-evoked rise in [Ca2+ ]i The above data all point to Ca2+ -release from internal stores or buffers as being the principal cause of the anoxia induced rise in [Ca2+ ]i . We therefore investigated the role of the endoplasmic reticulum (ER) in anoxia induced Ca2+ -release. Caffeine-sensitive stores were depleted by application of 30 mM caffeine (in Ca2+ -free media). Subsequent anoxic exposure still evoked a rise in [Ca2+ ]i of 45 ± 10 nM (n = 5), but this was significantly smaller than the rise in [Ca2+ ]i evoked by anoxia in Ca2+ -free solution alone (p < 0.05, Fig. 3A and D). To investigate the extent to which mitochondria contribute to the rise in [Ca2+ ]i during anoxia, we depolarised mitochondrial

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

B)

A) K+

Ca2+-free

capsaicin

600 500

anoxia

400

200 [Ca2+]i / nM

700 [Ca2+]i / nM

419

hypoxia

300

anoxia

anoxia

150

200

100

100 200 s

200 s C)

D)

***

100 Ni2+ [Ca2+]i / nM

anoxia 250 200

anoxia

***

80

Δ[Ca2+]i / nM

300

***

60 40

*

20 150

0 200 s

hypoxia (n = 8)

anoxia (n = 38)

anoxia / anoxia Ni2+ Ca2+-free (n = 5) (n = 23)

Fig. 1. In nodose ganglion neurons anoxia evoked a rise in [Ca2+ ]i independent of extracellular Ca2+ . (A) Effects of hypoxia (pO2 = 5 mmHg) and anoxia on [Ca2+ ]i. Exposure periods are indicated by horizontal bars. (B) The effects of extracellular calcium removal on the [Ca2+ ]i -response to anoxia. (C) Effects of 1 mM Ni2+ on the [Ca2+ ]i -response to anoxia and (D) Summary data showing mean (± SEM) of [Ca2+ ]i -response to anoxia in control, Ca2+ -free and 1 mM Ni2+ conditions. Significance was assessed using Students paired t-test to compare [Ca2+ ]i under anoxia with that under normoxia; *p < 0.05, ***p < 0.001; number of experiments are given in parentheses.

A)

B)

[Ca2+]i / nM

550 500 [Ca2+]i / nM

pHo 5.0

capsaicin

450 400

400

anoxia

300 200

350 300

anoxia

0.5 nA

250

10 s capsaicin C)

0.3

60

normoxia

0.25

Em (mV)

40 20

anoxia

0.15

0

I(nA)

-20 -40

re-oxygenation

0.2

0.1 0.05

anoxia

0 -90

-80

-70

-60

-50

(mV)

-0.05

-60

-0.1

-80 100 s Fig. 2. Effects of anoxia on membrane potential and membrane currents. Panel A shows a current clamp recording of membrane potential together with a simultaneous, measurement of [Ca2+ ]i . Note that anoxia causes a rapid sustained rise in [Ca2+ ]i , but only a negligible membrane depolarisation. In contrast, exposure to capsaicin (100 nM) evoked a robust increase in [Ca2+ ]i that is accompanied by membrane depolarisation and firing of action potentials. (B) Simultaneous recording of membrane current in whole cell voltage clamp (holding potential −70 mV) and [Ca2+ ]i . Note minimal inward current in response to anoxia but significant, reversible, increase in [Ca2+ ]i . For comparison the same neuron can be seen responding to an acidic stimulus (pH 5.5) with a robust (biphasic) inward current and rise in [Ca2+ ]i and (C) Membrane currents active at, and around the cells normal resting membrane potential measured using a voltage ramp protocol. The holding potential was set at −70 mV, membrane potential was then stepped to −40 mV for 250 ms (to allow deactivation of any A-type currents) and then ramped to −100 mV over 500 ms. Anoxia had little or no effect on membrane currents measured over this voltage range in 80% of the neurons investigated.

420

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

caffeine

A)

B) K+

400 [Ca2+]i / nM

350 300

anoxia

anoxia

250 200

[Ca2+]i / nM

500 400

Ca2+-free

FCCP anoxia

anoxia

300 200

150 100

Ca2+-free

200 s

200 s 2+

Ca -free

C)

D) 100 caffeine

1500

anoxia

1000

Δ[Ca2+]i / nM

[Ca2+]i / nM

FCCP 80 60

200 s

Ca2+-free

1

40 20

500

*

***

1

*** ** *

1

2

3

0 anoxia anoxia/ anoxia/ anoxia anoxia (n = 38) Ca2+-free caffeine / FCCP / FCCP (n = 23) (n = 5) (n = 12) /caffeine (n = 4)

Fig. 3. Intracellular Ca2+ -stores contribute to anoxic rise in [Ca2+ ]i . Depletion of caffeine sensitive stores significantly attenuated the anoxia induced rise in [Ca2+ ]i to 44.8 ± 10 nM (n = 5, *p < 0.05, (A) & (D)). FCCP (1 ␮M) applied prior to anoxia, significantly reduced the anoxic evoked rise in [Ca2+ ]i to 12 ± 4 nM (n = 12, ***p < 0.001 compared to anoxia in Ca2+ -free, (B) & (D)). The simultaneous application of FCCP and caffeine, further attenuated the anoxia evoked Ca2+ -response compared to FCCP alone (4.67 ± 0.475 nM, n = 4, *p < 0.05, (C) & (D)). The statistical comparisons shown in (D) are: 1 compared to anoxia Ca2+ -free; 2 compared to anoxia/caffeine; 3 compared to anoxia/FCCP (Students paired t-test, number of experiments are given in parentheses).

membrane potential ( m ) using the uncoupler FCCP [39,40] thereby releasing any Ca2+ from these stores. Application of 1 ␮M FCCP in Ca2+ -free media significantly elevated [Ca2+ ]i to 221 ± 27 nM (n = 12, p < 0.001, Fig. 3), a level similar to that seen during exposure to anoxia in Ca2+ -free solution (205 ± 1 nM, n = 23; p = 0.503 compared to FCCP). In the presence of FCCP anoxia had only a very small effect on [Ca2+ ]i ([Ca2+ ]i = 12 ± 4 nM, n = 12), which was significantly less than that seen in Ca2+ -free medium in the absence of FCCP (57 ± 7 nM, p < 0.001) (Fig. 3B and D). The simultaneous application of FCCP and caffeine to deplete both Ca2+ stores reduced the [Ca2+ ]i response to anoxia further (4.7 ± 0.5 nM, n = 4, p < 0.001 compared to anoxia/Ca2+ -free; Fig. 3C and D). Thus the anoxic induced Ca2+ -liberation appeared to derive primarily from mitochondria and ER. In another series of experiments we used Rh123 to investigate the effects of anoxia on m . Although this probe cannot be precisely calibrated (in terms of mV) we have used 1 ␮M FCCP as an index of complete mitochondrial depolarisation (Fig. 4). Using FCCP as a reference, exposure to anoxia evoked a depolarisation and a rise in Rh123 fluorescence to 31 ± 8% of the maximum evoked by FCCP (Fig. 4, n = 4), whereas hypoxia (pO2 approx. 2 Torr) had no significant effect upon m (data not shown). The effects of anoxia were reversible on re-oxygenation and were repeatable (Fig. 4). These data show that anoxia only partially depolarises m . 3.4. Mitochondrial Ca2+ -buffering in NG neurons under anoxia The previous section suggests that anoxia provokes Ca2+ release from mitochondria and from the ER. If there is significant mitochondrial depolarisation and release of mitochondrial Ca2+ during anoxia then one would also expect severe impairment of

mitochondrial Ca2+ -buffering (amount of Ca2+ that is taken up and stored) during Ca2+ -influx. In order to test the above hypothesis voltage-gated Ca2+ influx was triggered by a brief application of high-K+ (50 mM) for 5 s in a Tyrode containing 2.5 mM CaCl2 . The amplitude of the [Ca2+ ]i response to the resultant membrane depolarisation is dependent upon the rate of Ca2+ -influx through voltage-gated Ca2+ -channels, mitochondrial Ca2+ -uptake, Ca2+ -uptake or -release from other intracellular organelles and cytoplasmic Ca2+ -buffering. Termination of the high-K+ pulse results in cessation of voltagegated Ca2+ -influx and a rapid partial reduction in [Ca2+ ]i to about 443 ± 66 nM this is followed by a secondary slow rise in [Ca2+ ]i to a plateau (or shoulder) and then a slower decline to basal levels (Fig. 5A and B). This slow secondary rise in [Ca2+ ]i following high-K+ removal has been observed in many other neurons and attributed to the release of Ca2+ from mitochondria which had previously been taken up during the voltage-gated Ca2+ -entry phase [35,40–42]. The ability of mitochondria to function as a dynamic Ca2+ -buffer/store in NG neurons, taking up Ca2+ when [Ca2+ ]i is very high and then releasing it when [Ca2+ ]i falls, was confirmed by application of FCCP immediately before and during a high-K+ pulse. This resulted in a significant amplification of the [Ca2+ ]i response to depolarisation (to 184 ± 22%, n = 8, p < 0.05) and the complete absence of secondary rise in [Ca2+ ]i following high-K+ removal (Fig. 5A). To evaluate the effects of anoxia on mitochondrial Ca2+ buffering, anoxia was applied in a similar manner to FCCP, i.e. just before and during application of a high-K+ pulse. In these experiments anoxia increased the immediate [Ca2+ ]i response to high-K+ to 118 ± 6% (n = 12, p < 0.05, Fig. 5B) and reduced the secondary rise in [Ca2+ ]i following high-K+ removal to 82 ± 6% (n = 7, p < 0.05) of control. These findings are consistent with modest impairment

200

FCCP

FCCP

421

A) K+

3000

K+

250

2500

anoxia

2000 1500

150 100

1000 50

500

100

0

100 s

3.5. ER Ca2+ -stores are affected by anoxia In the previous section we demonstrated that anoxia evoked a rise in [Ca2+ ]i that mainly derived from internal stores. Since ER Ca2+ -store refilling is dependent on energy driven processes we investigated the extent to which anoxia might empty ER stores of Ca2+ . As previously described for DRG neurons [35] caffeine exposure (30 mM, for 2 min) was used to deplete ER Ca2+ -stores and the measured resultant rise in [Ca2+ ]i was used as an index of ER Ca2+ content. Exposure to caffeine was carried out in Ca2+ -free solutions to prevent any Ca2+ -influx. To facilitate ER store filling, a 5 s pulse of high-K+ Tyrode (50 mM K+ + 2.5 mM CaCl2 ) was given (3 min) prior to caffeine application. The maximum amplitudes of the subsequent caffeine responses were measured and normalised to the amplitude of the initial (control) caffeine response (set as 100%). Exposure to anoxia for 3 min reduced the amplitude of the [Ca2+ ]i response to caffeine to 53 ± 11% (n = 6, p < 0.01) compared to control (Fig. 6A and C). Upon return to normoxia the caffeineinduced Ca2+ -response recovered to 121 ± 29% (n = 6, p = 0.494) compared with control (Fig. 6A and C). A more prolonged exposure to anoxia (18 min) abolished any response to caffeine (0 ± 0%,

FCCP

B) K+

K+

* 100

1000

(%)

[Ca2+]i / nM

control FCCP (n = 8) (n = 8)

150

1500

shoulder

500

50 duration

0

0 control anoxia (n = 12) (n = 12)

anoxia

200 s C) 1200

K+

K+

K+ 150

1000 800

(%)

of mitochondrial Ca2+ -buffering during anoxia. However, the most prominent effect of anoxia was an unexpected prolongation of the mitochondrial Ca2+ -release phase from 36 ± 5 s under normoxic conditions to 75 ± 12 s (n = 6, p < 0.05, duration measured at half height see Fig. 5B). The Ca2+ -clearance/-extrusion rate during the final phase of [Ca2+ ]i recovery (measured between 200 and 350 nM of [Ca2+ ]i ) was also significantly slowed from 5.9 ± 1 nM/s during normoxia (n = 6) to 0.9 ± 0.2 nM/s under anoxia (n = 6, p < 0.01). These data are similar to findings in DRG neurons and may indicate a larger amount of Ca2+ being buffered during anoxia, a slower release of Ca2+ from these organelles, or a slower cytosolic clearance rate [35]. We semi-quantified the amount of Ca2+ taken up by mitochondria during exposure to high K+ by using the uncoupler FCCP to dump mitochondrial Ca2+ -stores as previously reported [35]. The application of FCCP soon after the high-K+ pulse leads to a substantive rise in [Ca2+ ]i that is assumed to be due to the release of Ca2+ stored in mitochondria during the high-K+ pulse. The amplitude of this response to FCCP (relative to base line [Ca2+ ]i ) was therefore taken as an index of mitochondrial Ca2+ -load. Using this technique, anoxia was found to have no measureable effect on the ability of mitochondria to take up Ca2+ during high K+ induced Ca2+ -influx (n = 7, p = 0.312, Fig. 5C).

0 200 s

[Ca2+]i / nM

Fig. 4. Anoxia affects mitochondrial membrane potential  in NG neurons. Measurement of  performed using Rh123. Fluorescence intensity of Rh123 is reported as a percentage of its initial (baseline) fluorescence. As a reference, in each experiment a brief application of FCCP (1 ␮M) was used to determine the increase in Rh123 fluorescence corresponding to complete depolarisation of . Effects of anoxia were then investigated and compared with the FCCP evoked rise in Rh123 fluorescence. Anoxia evoked a robust and reversible depolarisation of  equivalent to 30 (±8%) of the maximum change in fluorescence evoked by FCCP, n = 4.

**

(%)

anoxia

200 amplitude

150

[Ca2+]i / nM

rhodamine 123-fluorescence (%)

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

100

600 50 400 200 200 s

FCCP

FCCP anoxia

Ca2+-free

0 FCCP anoxia/ (n = 7) FCCP (n = 7)

Fig. 5. Effects of anoxia on mitochondrial Ca2+ buffering in nodose ganglion neurons. (A) Brief exposure to 50 mM KCl (5 s) triggers a biphasic Ca2+ -transient. Under control conditions high K+ solution causes an initial spike in [Ca2+ ]i representing voltage-gated Ca2+ -influx. Upon removal of high K+ solution this is followed by an initial rapid partial recovery in [Ca2+ ]i and then a second slow phase in which [Ca2+ ]i may increase again slightly and then remain elevated for a prolonged period of time before finally recovering back to baseline. This second phase is referred to as a “shoulder” (see text for further details). In the presence of FCCP (1 ␮M) the Ca2+ transients became monophasic, the initial phase/spike was significantly amplified and there was no discernible second phase or shoulder. The right hand panel compares the amplitude of the initial [Ca2+ ]i spike in response to high K+ under control conditions and in the presence of FCCP. (B) Effects of anoxia on [Ca2+ ]i -response to high K+ . Note that anoxia significantly enhanced the amplitude of the initial spike, compared in the right hand panel, and prolonged the duration of the second phase and (C) Estimation of mitochondrial Ca2+ -uptake during high-K+ -application. Neurons were briefly (5 s) exposed to high-K+ solution containing 2.5 mM Ca2+ and then transferred to a Ca2+ -free medium. After recovery of the initial Ca2+ -spike FCCP (1 ␮M, 30 s) was applied to provoke mitochondrial depolarisation and Ca2+ -release. This FCCP-triggered Ca2+ -release was quantified as the difference between baseline immediately before the exposure to the high-K+ solution and the maximal level of [Ca2+ ]i observed in the presence of FCCP. Note that anoxia slightly enhanced the initial [Ca2+ ]i –spike upon depolarisation but did not alter the response to FCCP (compared in the right hand panel). Thus the amount of Ca2+ stored in mitochondria during high-K+ -application (mitochondrial Ca2+ -buffering) is not affected by brief periods of anoxia. Statistical significance was assessed by Students paired t-test (*p < 0.05, **p < 0.001, numbers of experiments are given in parentheses). Exposure periods are indicated by horizontal barks.

422

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

normoxic conditions (following high-K+ application) only reduced the response to caffeine to 76 ± 7% (n = 5) of control.

A) 700 K+

[Ca2+]i / nM

600

K+

anoxia

K+

caffeine

caffeine

c a ffein e

500 400 300 200 100 Ca2+-free

200 s B)

K+ 350

K+

[Ca2+]i / nM

300 250

anoxia caffeine

caffeine caffeine

200 150 100 50 Ca2+-free 200 s

maximum caffeine response [%]

C)

160 140 120 100 80

**

60 40 20

***

0 control

anoxia recovery (3 min) (n = 6) (n = 6)

anoxia recovery (18 min) (n = 6) (n = 6)

Fig. 6. Anoxia induced depletion of ER Ca2+ -stores. Effects of anoxia on ER store filling. The peak [Ca2+ ]i -response to 30 mM caffeine was used as an index of ER Ca2+ -store filling. Application of a brief KCl pulse (50 mM for 5 s) before each caffeine exposure was used to provide sufficient Ca2+ by voltage-gated Ca2+ -influx to allow ER Ca2+ -store refilling. (A) The caffeine response was significantly reduced during brief anoxia (3 min, (A) & (C)). (B) Prolonged exposure to anoxia (18 min) resulted in a sustained elevation of resting [Ca2+ ]i , and the complete loss of any [Ca2+ ]i -response to caffeine. The caffeine response subsequently recovered partially following re-oxygenation ((B) & (C)). (C) The bar charts depict means + S.E.M of [Ca2+ ]i –responses to caffeine under control, anoxic and post anoxic condition. Statistical significance was assessed by Students paired t-test (**p < 0.01, ***p < 0.0001, numbers of experiments are given in parentheses). Exposure periods are indicated by horizontal bars.

n = 6, p < 0.001 Fig. 6B and C). These data suggest that prolonged anoxia completely depletes ER Ca2+ -store content. This effect was reversible; following re-oxygenation and triggering Ca2+ -influx through the application of high-K+ Tyrode the caffeine sensitive Ca2+ -store recovered to 81 ± 11% of control (n = 6, p = 0.326, Fig. 6B and C). In comparison a 20 min incubation in Ca2+ -free media under

3.6. Cytosolic Ca2+ -removal is affected by anoxia The observation that [Ca2+ ]i clearance following voltage-gated Ca2+ -influx took longer under anoxic conditions suggests that anoxia may decrease Ca2+ -extrusion and/or uptake into intracellular stores. To determine the effects of anoxia on cytosolic Ca2+ -clearance we performed a series of experiments under conditions in which mitochondrial Ca2+ -buffering is assumed to be negligible. Neurons were again depolarized by a high-K+ (50 mM for 5 s) Tyrode but with reduced extracellular [Ca2+ ]o (250 ␮M). Exposure to this high-K+ -low-Ca2+ , solution triggered an immediate rise in [Ca2+ ]i to 253 ± 62 nM, (n = 11). Following withdrawal of high-K+ -low-Ca2+ solution, [Ca2+ ]i showed a monophasic decline back to baseline (Fig. 7A–D), confirming that [Ca2+ ]i did not reach the threshold for significant mitochondrial Ca2+ -buffering (which has been estimated to be around 500 nM [43]). We calculated Ca2+ clearance during this recovery phase as the rate of decline in [Ca2+ ]i (d[Ca2+ ]i /dt) in nM/s measured between 250 and 150 nM. Under normoxic conditions repeated exposure to high-K+ -low-Ca2+ solution evoked similar rise in [Ca2+ ]i and consistent Ca2+ -clearance rates (11 ± 1 nM/s, n = 11; see Fig. 7E). Under anoxic conditions the mean amplitude of the [Ca2+ ]i transients fell to 197 ± 47 nM (77% of control, n = 6, p < 0.05) and Ca2+ -clearance rates were reduced from 11 ± 1 nM/s, n = 6 in normoxia to 6.4 ± 0.4 nM/s (n = 6, p < 0.05), after 3 min of anoxia and 3.3 ± 0.4 nM/s (n = 6, p < 0.001) after 25 min of anoxia (Fig. 7). Since Ca2+ -clearance can be mediated by Ca2+ -efflux across the plasma membrane (by PMCA or Na+ /Ca2+ -exchange) and/or Ca2+ -uptake into the ER (by SERCA), we performed further experiments to determine the relative importance of these pathways to Ca2+ -clearance. Inhibition of SERCA, by CPA (10 ␮M) or thapsigargin (100 nM), reduced the Ca2+ -recovery rate under normoxic conditions from 11 ± 1 nM/s to 6 ± 0.8 nM/s (n = 6, p < 0.01). The remaining Ca2+ -clearance rate corresponds to Ca2+ -efflux across the plasma membrane (Fig. 7B, D and F). Exposure to anoxia in the presence of CPA or Thapsigargin further reduced Ca2+ clearance rate to 4.1 ± 0.6 nM/s after 3 min and 2.5± 0.4 nM/s after 25 min (n = 6, p < 0.05), indicating that Ca2+ -efflux is impaired during anoxia. To determine whether anoxia also alters SERCA function the rate of uptake into the ER was estimated by subtracting the calcium-clearance rates observed in the presence of CPA (or Thapsigargin) from those observed in their absence. This analysis was conducted under both normoxic and anoxic (3 min) conditions using the data shown in Fig. 7 and is presented in Fig. 8. Using this approach we estimated ER Ca2+ -uptake to be equivalent to 5.2 ± 0.3 nM/s (n = 6) under normoxic conditions and 2.3 ± 0.5 nM/s, (n = 6, p < 0.05) under anoxic conditions. Thus anoxia appears to inhibit both Ca2+ -efflux and ER Ca2+ -uptake/SERCA function. The effect of anoxia upon ER Ca2+ -uptake (56% inhibition) is however more pronounced than its effect upon Ca2+ -efflux (31% inhibition). Comparison of the relative importance of the two pathways suggests a 46/54% split between ER Ca2+ -uptake and Ca2+ -efflux under control conditions and a 36/64% split in anoxia (Fig. 8). Thus under anoxic conditions Ca2+ -clearance is mostly mediated by Ca2+ -efflux. To confirm the role of PMCA in mediating Ca2+ -clearance we investigated the effects of extracellular alkalosis. As PMCA normally exchanges intracellular calcium for extracellular protons it can be inhibited by lowering the extracellular proton concentration such as to make the forward operation of the pump thermodynamically unfavourable [44]. In Ca2+ -containing solutions exposure to pH0 8.8 greatly reduced the Ca2+ -clearance rate following

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

A)

B) K

+

K

+

K

+

K

+

K

+

K

+

400 300

‡

anoxia K+

K+

500

[Ca2+]i / nM

[Ca2+]i / nM

500

‡ K+

†

†

423

K+

K+

K+

K+

400 300 200

200

100 100

200 s

anoxia

CPA

Ca2+-free

Ca2+-free

200 s D)

500

500

400

400

[Ca2+]i / nM

[Ca2+]i / nM

C)

300 anoxia

200

200 100

300

100

normoxia

control 20 s

20 s E)

F) 20

16

18

14

16

12

12 10

[Ca2+] nM/s

14 [Ca2+] nM/s

CPA

control anoxia

8 6

10

anoxia / CPA

8 6 4

4

2

2 0

0 control

3

8

13

18

25

control CPA

time (min)

3

8

13

18

25

time (min)

Fig. 7. Ca2+ -extrusion rates during anoxia. Neurons were depolarised repetitively using a 5 s application of a high K+ (50 mM) solution containing low (250 ␮M) CaCl2 and then returned to a Ca2+ -free solution. This resulted in small [Ca2+ ]i –transients with an amplitude assumed to be below the threshold for mitochondrial Ca2+ -uptake and which consequently recovered monophasically. Ca2+ -clearance/recovery rates were measured between 200 and 150 nM [Ca2+ ]i during the recovery period. ((A), (C)) Monophasic [Ca2+ ]i –transients elicited using high K+ low Ca2+ solutions under normoxic and anoxic conditions. Individual transients shown in C are those identified in A with a †. ((B), (D)) Monophasic [Ca2+ ]i –transients elicited using high K+ low Ca2+ solutions under normoxic conditions in the presence of the SERCA inhibitor CPA and in the presence of both CPA and anoxia. Individual transients shown in D are those identified in B with a ‡. (E) Mean (±SEM) [Ca2+ ]i recovery rates measured under control and anoxic conditions. Solid diamonds show data from control experiments in which cells were stimulated with high K+ /low Ca2+ solutions repetitively under normoxic conditions throughout (n = 6). Note the stability of recovery rates measured under these conditions. Solid circles show data from an experiment in which cells were stimulated with high K+ /low Ca2+ solutions first in a normoxic solution and then repetitively in an anoxic solution for up to 25 min (n = 6). Note rapid decline in recovery rate during anoxia and (F) Mean (±SEM, n = 6) [Ca2+ ]i recovery rates measured under control, CPA and CPA + anoxic conditions. Note decline in recovery rate in response to both CPA and CPA + anoxia.

voltage-gated Ca2+ -entry from 5 ± 1.1 nM/s to 1.3 ± 0.5 nM/s (n = 5). This inhibition was reversed by removing extracellular calcium, thus making forward transport by PMCA energetically favourable again, as expected (Fig. 9A). In Ca2+ -containing medium addition of CPA and pHo 8.8 completely inhibited Ca2+ -clearance (Fig. 9B). We also assessed the role of Na+ /Ca2+ -exchange in mediating Ca2+ -clearance by inhibiting the forward mode of exchange (Na+ in/Ca2+ out) with Na+ -free medium. Ca2+ -clearance rates (following depolarisation in high-K+ -low-Ca2+ media) were not-significantly slower in Na+ -free Tyrodes than in control Na+ -containing Tyrode (data not shown). Thus Na+ /Ca2+ -exchange does not appear to contribute significantly to Ca2+ -clearance. These data confirm that

PMCA and SERCA are indeed the main mechanisms for Ca2+ clearance from the cytoplasm in NG neurons. 4. Discussion Sensory neurons respond to ischaemia with both the release of locally acting neurotransmitters/neuromodulators as well as sending chemosensory information back to the brain. It has been suggested that excitation and neurosecretion in response to ischaemia is largely due to the activation of proton sensitive channels in response to tissue acidosis [11,19]. There is however also growing evidence that low oxygen can independently influence

424

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

normoxia anoxia

12 10 [Ca2+] nM/s

8

SERCA + PMCA

**

SERCA

1

*

1

**

6

1

SERCA

SERCA + PMCA

4 PMCA

PMCA

2 0 CPA (n = 6)

control (n = 11)

control (n = 6)

CPA (n = 6)

Fig. 8. Exposure to anoxia reduces mainly SERCA Ca2+ -clearance. Ca2+ -clearance in NG neurons is carried out by PMCA and SERCA. Calcium clearance in the presence of CPA (black bars) is significantly reduced compared to control (white bar), demonstrating a major role for SERCA. The magnitude of the SERCA flux may thus be estimated as the difference between the control rate of clearance and that in the presence of CPA. Even in the presence of CPA however a substantial rate of Ca2+ -clearance remains which is assumed to correspond to PMCA activity (see text). Data (Mean ± SEM) are derived from experimental measurements shown in Fig. 7. Numbers of experiments are given in parentheses. Effects of CPA, anoxia and anoxia plus CPA are compared to control using Students paired t-test (*p < 0.05; **p < 0.01). The estimated magnitude of flux on the individual pathways, PMCA and SERCA, are indicated by arrows.

neuronal function. For example hypoxia depolarises sensory neurons of the glossopharyngeal nerve [31,32]. In the present study we have investigated the effects of anoxia on Ca2+ -regulation in nodose ganglion neurons. In a previous study we demonstrated that ATP in NG neurons undergoes a rapid 21%

A)

K+

K+

decline within 3 minutes of anoxia followed by a further slower fall of up to 40% by 20 min [34]. The ability of these neurons to maintain ATP levels at around 60% even after 20 min of anoxia reflects the continued availability of glucose [34]. Unlimited glucose availability, combined with a maintained extracellular pH, means that

K+

600

7 6 [Ca2+] nM / s

[Ca2+]i / nM

500 400 300

5 4 3

200

2

100

1 0

pHo 8.8

200 s

control / pHo 7.4 (n = 5)

pHo 8.8 2+

Ca -free

K+

B)

K+

**

K+

pHo 8.8 (n = 5)

K+ 7

250

[Ca2+] nM/s

[Ca2+]i / nM

6 200

150

5

*

4 3 2

100

1 200 s

pHo 8.8

CPA

CPA pHo 8.8

***

0 control / pHo 7.4 (n = 5)

CPA (n = 5)

CPA / pHo 8.8 (n = 5)

Fig. 9. PMCA is the main Ca2+ -extrusion pathway in nodose ganglion neurons. Ca2+ -influx was evoked by depolarisation in a high K+ medium (50 mM) containing 250 ␮M Ca2+ -ions for 5 s. (A) Extracellular alkalisation (pH0 8.8) significantly reduced Ca2+ -clearance rates in Ca2+ -containing solutions but not in Ca2+ -free solutions and (B) CPA significantly reduced neuronal Ca2+ -extrusion in Ca2+ -containing solutions. Ca2+ -extrusion was completely blocked by the combination of CPA and pH0 8.8. After washout of CPA the Ca2+ -extrusion rate recovered partially. The rates of Ca2+ -extrusion at pH0 8.8 pre and post CPA application were identical (indicated by dashed lines, *p < 0.05, **p < 0.01, ***p < 0.001, compared to control, Students paired t-test). Horizontal bars indicate exposure periods.

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

our anoxic conditions do not recapitulate all the metabolic effects of sustained ischaemia but are likely to be representative of the early stages. 4.1. Effects of anoxia on ER Ca2+ -stores Anoxia caused a rapid increase in resting intracellular calcium by approximately 80 nM. We found no evidence for voltage-gated Ca2+ -entry, instead the rise of [Ca2+ ]i was entirely dependent upon release from internal stores as has been reported for cervicothoracic DRG neurons [35]. Ca2+ -release from internal stores in response to hypoxia or metabolic stress has also been reported in hippocampal neurons and pulmonary smooth muscle cells [45–49]. Approximately 40% of the calcium released into the cytosol in anoxia was dependent upon a caffeine sensitive internal store presumed to be the ER. Similarly caffeine evoked Ca2+ -release was either diminished by brief exposure to anoxia or completely abolished following anoxic periods in excess of 18 min. These data strongly implicate ER Ca2+ -release in playing a significant role in the anoxia evoked raise in [Ca2+ ]i . In principle there are three general mechanisms that might account for this. • Firstly, anoxia could lead to activation of ryanodine receptors themselves. For example generation of reactive oxygen species (ROS) or cyclic ADP ribose in hypoxia has been reported to activate ryanodine receptors in astrocytes, cardiomyocytes, chromaffin cells and pulmonary artery smooth muscle cells [50–53]. ROS derive mainly from mitochondria or NADPH oxidase, activating the ryanodine receptor [52,54]. Alternatively NO production has also been reported to increase in hypoxia in these neurons which could also activate ryanodine receptors [13,55,56]. However, the production of either NO or ROS in anoxia seems unlikely since both require molecular oxygen as a substrate. • Secondly, activation of other Ca2+ -release channels may cause Ca2+ -efflux from the ER. In rat hippocampal neurons activation of IP3 receptors contributes to hypoxic or anoxic increase of [Ca2+ ]i via a NADPH dependent pathway [57,58]. Similarly in mouse cortical neurons the IP3 -receptor is the main Ca2+ -release channel from the ER during ischaemia [59]. A Ca2+ -leak pathway, that is insensitive to inhibitors of either ryanodine or IP3 receptors, could also facilitate Ca2+ -loss from the ER [60]. • Thirdly, the observation that anoxia depletes ER stores could be explained by reduced activity of SERCA in combination with a Ca2+ -leak which need not necessarily be enhanced. We observed a 68% reduction of SERCA activity within three minutes of anoxia (see below). 4.2. Effects of anoxia on mitochondrial Ca2+ -release/-buffering Mitochondria play a pivotal role in neuronal Ca2+ -regulation by acting as a dynamic buffering system [40,43,61–63]. In the present study application of FCCP caused only a small increase in [Ca2+ ]i which is consistent with mitochondria containing only a small pool of calcium under resting conditions. Following application of FCCP the anoxia induced increase in [Ca2+ ]i was reduced by 60% suggesting that anoxia either causes Ca2+ -release from mitochondria or some other energy dependent store. Note that while FCCP causes an intracellular acidosis, the fall in pHi measured in NG neurons (pHi −0.3) would be insufficient to interfere with the fura-2 signal so we assume that the changes in [Ca2+ ]i reported here are reasonably accurate [33,34]. Although we did not investigate the mechanisms of this effect of anoxia, if it is due to mitochondrial Ca2+ -release there are a number of possible explanations. First depolarisation of m may lead to permeability transition pore (PTP) opening, secondly mitochondrial Ca2+ /H+ -exchange could be altered by cytosolic acidification

425

[34,64] and thirdly reduction of m might reduce Ca2+ -influx by the uniporter (which is normally balanced by Ca2+ -efflux through mitochondrial Na+ /Ca2+ -exchange [65]). It should also be noted that, as we have used photometry to measure m , we cannot determine whether the observed depolarisation is uniform, or whether some mitochondria depolarise more than others. We also investigated the effects of anoxia on mitochondrial Ca2+ -buffering. When [Ca2+ ]i exceeds a threshold of approx. 500 nM Ca2+ is taken up into the mitochondria via the Ca2+ -uniporter where it is complexed with phosphate in the inner matrix [66,67]. This prevents extremely high levels of [Ca2+ ]i developing that could damage the cell [68]. In the present study we found that anoxia did not alter mitochondrial Ca2+ -buffering significantly. The initial rise in [Ca2+ ]i during high-K+ induced depolarisation was slightly enhanced but the amount of Ca2+ that appeared to be taken up into mitochondria was not altered. The only major effect of anoxia was to slow Ca2+ -clearance during the mitochondrial Ca2+ -release phase. This we attribute to mechanisms other than those of mitochondrial Ca2+ -transport per se (see below). The observations that mitochondria appear to release Ca2+ in response to anoxia under resting conditions but to retain an uninhibited ability to take up calcium when cytosolic [Ca2+ ] is elevated by voltage-gated Ca2+ -influx appear contradictory. This might reflect the presence of different mitochondrial subpopulations. In myocytes for example mitochondrial subpopulations differ by their membrane potential, Ca2+ -buffering capacity, and response to uncouplers like FCCP [69,70]. It is also possible however that the rise in [Ca2+ ]i in response to anoxia and FCCP under resting conditions is not due to mitochondrial release per se but to Ca2+ -release from some other energy dependent store or buffer system.

4.3. Cytosolic Ca2+ -clearance during oxygen depletion Ca2+ -clearance following small Ca2+ -loads was progressively inhibited by anoxia over an 8 minute period and then remained stable for up to 25 min. Under these conditions Ca2+ -clearance involves both Ca2+ -uptake into the ER via SERCA and Ca2+ -efflux from the cell via PMCA with roughly 46% of Ca2+ following the SERCA route and 54% being extruded by PMCA. Under anoxic conditions however SERCA activity was inhibited by approximately 56% whereas the PMCA activity was reduced by 31%. Thus under anoxic conditions 64% of any Ca2+ -load is cleared by PMCA and only 36% is taken up into the ER. This contrasts with DRG neurons in which anoxia reduced PMCA activity by 53% and SERCA activity by 60% compared to control [35]. In both neuron populations anoxia reduces the SERCA activity by roughly the same amount whereas PMCA activity in NG neurons seemed less sensitive to anoxia. In pancreatic acinar cells cytosolic Ca2+ -clearance is mainly driven by PMCA and declines gradually in parallel with ATP-exhaustion during inhibition of mitochondrial and glycolytic metabolism [71]. In contrast, in cerebellar neurons cytosolic Ca2+ -clearance is little affected by inhibition of mitochondrial ATP synthesis and relies, instead, on glycolysis [72]. The extent to which different cells are dependent upon PMCA vs. SERCA for Ca2+ -clearance as well as the sensitivity of these mechanisms to loss of mitochondrial ATP production therefore seem to vary. This may reflect variation in the expression of the different isoforms of these pumps [73–77]. We have not investigated how anoxia inhibits either SERCA or PMCA but there are two obvious possibilities. One is partial depletion of cellular ATP (see above) [78,79] the other is via the cytosolic acidification that also accompanies anoxia [34]. Since both SERCA and PMCA exchange Ca2+ and H+ (with stochiometric ratios of 1/2 and 1/1 respectively) pump turnover may be slowed by a rise in cytosolic H+ [80,81].

426

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

In summary the effects of anoxia upon calcium regulation in nodose ganglia could be largely explained through the inhibition of the Ca2+ -pumps SERCA and PMCA. Decreased SERCA activity would be expected to lead to a net leak of Ca2+ from the ER helping to raise [Ca2+ ]i and deplete the ER of Ca2+ at rest and would also slow the redistribution of Ca2+ -loads buffered by the mitochondrion and slow Ca2+ -clearance from the cytosol. The coincident inhibition of PMCA could also help raise resting [Ca2+ ]i and slow calcium-clearance. These effects may be combined with further Ca2+ -release from another internal FCCP sensitive (possibly mitochondrial) store. The mechanisms by which anoxia inhibits Ca2+ -pumps remains to be elucidated. Disclosures None. Acknowledgements This work was supported by a grant from the Wellcome Trust (K.J.B.) and a DFG research fellowship (HE3678/1-1 to M.H.). References [1] A. Kuntz, The autonomic nervous system (1934). [2] T. Hayakawa, S. Kuwahara-Otani, S. Maeda, K. Tanaka, M. Seki, Projections of calcitonin gene-related peptide immunoreactive neurons in the vagal ganglia of the rat, J Chem Neuroanat 41 (2011) 55–62. [3] A. Torvik, Afferent connections to the sensory trigeminal nuclei, the nucleus of the solitary tract and adjacent structures; an experimental study in the rat, J Comp Neurol 106 (1956) 51–141. [4] G.D. Housley, R.L. Martin-Body, N.J. Dawson, J.D. Sinclair, Brain stem projections of the glossopharyngeal nerve and its carotid sinus branch in the rat, Neuroscience 22 (1987) 237–250. [5] R.J. Contreras, R.M. Beckstead, R. Norgren, The central projections of the trigeminal, facial, glossopharyngeal and vagus nerves: an autoradiographic study in the rat, J Auton Nerv Syst 6 (1982) 303–322. [6] E. Cooper, Synapse formation among developing sensory neurones from rat nodose ganglia grown in tissue culture, J Physiol 351 (1984) 263–274. [7] H. Zhuo, C.J. Helke, Presence and localization of neurotrophin receptor tyrosine kinase (TrkA, TrkB TrkC) mRNAs in visceral afferent neurons of the nodose and petrosal ganglia, Brain Res Mol Brain Res 38 (1996) 63–70. [8] P.N. Thoren, Characteristics of left ventricular receptors with nonmedullated vagal afferents in cats, Circ Res 40 (1977) 415–421. [9] P.N. Thoren, W.R. Saum, A.M. Brown, Characteristics of rat aortic baroreceptors with nonmedullated afferent nerve fibers, Circ Res 40 (1977) 231–237. [10] J.A. Armour, M.H. Huang, A. Pelleg, C. Sylven, Responsiveness of in situ canine nodose ganglion afferent neurones to epicardial mechanical or chemical stimuli, Cardiovasc Res 28 (1994) 1218–1225. [11] C.J. Benson, S.P. Eckert, E.W. McCleskey, Acid-evoked currents in cardiac sensory neurons: A possible mediator of myocardial ischemic sensation, Circ Res 84 (1999) 921–928. [12] Y. Lu, X. Ma, R. Sabharwal, V. Snitsarev, D. Morgan, K. Rahmouni, H.A. Drummond, C.A. Whiteis, V. Costa, M. Price, C. Benson, M.J. Welsh, M.W. Chapleau, F.M. Abboud, The ion channel ASIC2 is required for baroreceptor and autonomic control of the circulation, Neuron 64 (2009) 885–897. [13] Y. Yamamoto, M. Henrich, R.L. Snipes, W. Kummer, Altered production of nitric oxide and reactive oxygen species in rat nodose ganglion neurons during acute hypoxia, Brain Res 961 (2003) 1–9. [14] C. Owman, Peptidergic vasodilator nerves in the peripheral circulation and in the vascular beds of the heart and brain, Blood Vessels 27 (1990) 73–93. [15] T. Strecker, S. Koulchitsky, A. Dieterle, W.L. Neuhuber, M. Weyand, K. Messlinger, Release of calcitonin gene-related peptide from the jugular-nodose ganglion complex in rats - a new model to examine the role of cardiac peptidergic and nitrergic innervation, Neuropeptides 42 (2008) 543–550. [16] S. Bevan, P. Geppetti, Protons: small stimulants of capsaicin-sensitive sensory nerves, Trends Neurosci 17 (1994) 509–512. [17] D.C. Immke, E.W. McCleskey, Lactate enhances the acid-sensing Na+ channel on ischemia-sensing neurons, Nat Neurosci 4 (2001) 869–870. [18] D.C. Immke, E.W. McCleskey, ASIC3: a lactic acid sensor for cardiac pain, Scientific World J. 1 (2001) 510–512. [19] H.L. Pan, J.C. Longhurst, J.C. Eisenach, S.R. Chen, Role of protons in activation of cardiac sympathetic C-fibre afferents during ischaemia in cats, J Physiol 518 (Pt 3) (1999) 857–866. [20] G.P. Ahern, I.M. Brooks, R.L. Miyares, X.B. Wang, Extracellular cations sensitize and gate capsaicin receptor TRPV1 modulating pain signaling, J Neurosci 25 (2005) 5109–5116.

[21] J.C. Longhurst, A.L.S.C. Tjen, L.W. Fu, Cardiac sympathetic afferent activation provoked by myocardial ischemia and reperfusion. Mechanisms and reflexes, Ann N Y Acad Sci 940 (2001) 74–95. [22] M. Liu, N.J. Willmott, G.J. Michael, J.V. Priestley, Differential pH and capsaicin responses of Griffonia simplicifolia IB4 (IB4)-positive and IB4-negative small sensory neurons, Neuroscience 127 (2004) 659–672. [23] Q. Gu, L.Y. Lee, Characterization of acid signaling in rat vagal pulmonary sensory neurons, Am J Physiol Lung Cell Mol Physiol 291 (2006) L58–L65. [24] M. Kollarik, B.J. Undem, Mechanisms of acid-induced activation of airway afferent nerve fibres in guinea-pig, J Physiol 543 (2002) 591–600. [25] G.X. Yan, A.G. Kleber, Changes in extracellular and intracellular pH in ischemic rabbit papillary muscle, Circ Res 71 (1992) 460–470. [26] M.C. Heidt, D. Sedding, S.K. Stracke, T. Stadlbauer, A. Boening, P.R. Vogt, M. Schonburg, Measurement of myocardial oxygen tension: a valid and sensitive method in the investigation of transmyocardial laser revascularization in an acute ischemia model, Thorac Cardiovasc Surg 57 (2009) 79–84. [27] W.L. Rumsey, M. Pawlowski, N. Lejavardi, D.F. Wilson, Oxygen pressure distribution in the heart in vivo and evaluation of the ischemic “border zone”, Am J Physiol 266 (1994) H1676–H1680. [28] M. Khan, P. Kwiatkowski, B.K. Rivera, P. Kuppusamy, Oxygen and oxygenation in stem-cell therapy for myocardial infarction, Life Sci 87 (2010) 269–274. [29] Y.J. Bae, J.C. Yoo, N. Park, D. Kang, J. Han, E. Hwang, J.Y. Park, S.G. Hong, Acute hypoxia activates an ENaC-like Channel in rat pheochromocytoma (PC12) Cells, Korean J Physiol Pharmacol 17 (2013) 57–64. [30] K.J. Buckler, TASK-like potassium channels and oxygen sensing in the carotid body, Respir Physiol Neurobiol 157 (2007) 55–64. [31] V.A. Campanucci, I.M. Fearon, C.A. Nurse, A novel O2 -sensing mechanism in rat glossopharyngeal neurones mediated by a halothane-inhibitable background K+ conductance, J Physiol 548 (2003) 731–743. [32] V.A. Campanucci, C.A. Nurse, Biophysical characterization of whole-cell currents in O2 -sensitive neurons from the rat glossopharyngeal nerve, Neuroscience 132 (2005) 437–451. [33] A.Y. Abramov, A. Scorziello, M.R. Duchen, Three distinct mechanisms generate oxygen free radicals in neurons and contribute to cell death during anoxia and reoxygenation, J Neurosci 27 (2007) 1129–1138. [34] M. Henrich, K.J. Buckler, Effects of anoxia, aglycemia, and acidosis on cytosolic Mg2+ , ATP, and pH in rat sensory neurons, Am J Physiol Cell Physiol 294 (2008) C280–C294. [35] M. Henrich, K.J. Buckler, Effects of anoxia and aglycemia on cytosolic calcium regulation in rat sensory neurons, J Neurophysiol 100 (2008) 456–473. [36] G. Grynkiewicz, M. Poenie, R.Y. Tsien, A new generation of Ca2+ indicators with greatly improved fluorescence properties, J Biol Chem 260 (1985) 3440–3450. [37] E.C. Toescu, A. Verkhratsky, Assessment of mitochondrial polarization status in living cells based on analysis of the spatial heterogeneity of rhodamine 123 fluorescence staining, Pflugers Arch 440 (2000) 941–947. [38] M. Sato, K. Ikeda, K. Yoshizaki, H. Koyano, Response of cytosolic calcium to anoxia and cyanide in cultured glomus cells of newborn rabbit carotid body, Brain Res 551 (1991) 327–330. [39] V. Shishkin, E. Potapenko, E. Kostyuk, O. Girnyk, N. Voitenko, P. Kostyuk, Role of mitochondria in intracellular calcium signaling in primary and secondary sensory neurones of rats, Cell Calcium 32 (2002) 121–130. [40] S.A. Thayer, R.J. Miller, Regulation of the intracellular free calcium concentration in single rat dorsal root ganglion neurones in vitro, J Physiol 425 (1990) 85–115. [41] J.L. Werth, S.A. Thayer, Mitochondria buffer physiological calcium loads in cultured rat dorsal root ganglion neurons, J Neurosci 14 (1994) 348–356. [42] J. Herrington, Y.B. Park, D.F. Babcock, B. Hille, Dominant role of mitochondria in clearance of large Ca2+ loads from rat adrenal chromaffin cells, Neuron 16 (1996) 219–228. [43] D.G. Nicholls, S.L. Budd, Mitochondria and neuronal survival, Physiol Rev 80 (2000) 315–360. [44] C.D. Benham, M.L. Evans, C.J. McBain, Ca2+ efflux mechanisms following depolarization evoked calcium transients in cultured rat sensory neurones, J Physiol 455 (1992) 567–583. [45] R.I. Jabr, H. Toland, C.H. Gelband, X.X. Wang, J.R. Hume, Prominent role of intracellular Ca2+ release in hypoxic vasoconstriction of canine pulmonary artery, Br J Pharmacol 122 (1997) 21–30. [46] J.M. Dubinsky, S.M. Rothman, Intracellular calcium concentrations during “chemical hypoxia” and excitotoxic neuronal injury, J Neurosci 11 (1991) 2545–2551. [47] T.O. Grondahl, J.J. Hablitz, I.A. Langmoen, Depletion of intracellular Ca2+ stores or lowering extracellular calcium alters intracellular Ca2+ changes during cerebral energy deprivation, Brain Res 796 (1998) 125–131. [48] M. Sato, Different effects of removing extracellular Ca2+ on cytosolic Ca2+ response to anoxia of sensory neurons and carotid chemoreceptor cells from newborn rabbits, Jpn J Physiol 45 (1995) 279–289. [49] T.M. Kang, M.K. Park, D.Y. Uhm, Characterization of hypoxia-induced [Ca2+ ]i rise in rabbit pulmonary arterial smooth muscle cells, Life Sci 70 (2002) 2321–2333. [50] W. Du, M. Frazier, T.J. McMahon, J.P. Eu, Redox activation of intracellular calcium release channels (ryanodine receptors) in the sustained phase of hypoxia-induced pulmonary vasoconstriction, Chest 128 (2005) 556S–558S. [51] P.K. Aley, H.J. Murray, J.P. Boyle, H.A. Pearson, C. Peers, Hypoxia stimulates Ca2+ release from intracellular stores in astrocytes via cyclic ADP ribose-mediated activation of ryanodine receptors, Cell Calcium 39 (2006) 95–100. [52] D. Souvannakitti, J. Nanduri, G. Yuan, G.K. Kumar, A.P. Fox, N.R. Prabhakar, NADPH oxidase-dependent regulation of T-type Ca2+ channels and ryanodine

M. Henrich, K.J. Buckler / Cell Calcium 54 (2013) 416–427

[53]

[54]

[55]

[56] [57]

[58]

[59]

[60] [61] [62]

[63]

[64]

[65]

[66]

receptors mediate the augmented exocytosis of catecholamines from intermittent hypoxia-treated neonatal rat chromaffin cells, J Neurosci 30 (2010) 10763–10772. T.G. Favero, A.C. Zable, M.B. Bowman, A. Thompson, J.J. Abramson, Metabolic end products inhibit sarcoplasmic reticulum Ca2+ release and [3H]ryanodine binding, J Appl Physiol 78 (1995) 1665–1672. F.J. Gerich, F. Funke, B. Hildebrandt, M. Fasshauer, M. Muller, H2 O2 -mediated modulation of cytosolic signaling and organelle function in rat hippocampus, Pflugers Arch 458 (2009) 937–952. J.P. Eu, J. Sun, L. Xu, J.S. Stamler, G. Meissner, The skeletal muscle calcium release channel: coupled O2 sensor and NO signaling functions, Cell 102 (2000) 499–509. J.P. Eu, L. Xu, J.S. Stamler, G. Meissner, Regulation of ryanodine receptors by reactive nitrogen species, Biochem Pharmacol 57 (1999) 1079–1084. P.E. Bickler, C.S. Fahlman, J. Gray, W. McKleroy, Inositol 1,4,5-triphosphate receptors and NAD(P)H mediate Ca2+ signaling required for hypoxic preconditioning of hippocampal neurons, Neuroscience 160 (2009) 51–60. A.B. Belousov, J.M. Godfraind, K. Krnjevic, Internal Ca2+ stores involved in anoxic responses of rat hippocampal neurons, J Physiol 486 (Pt 3) (1995) 547–556. X. Chen, D.B. Kintner, J. Luo, A. Baba, T. Matsuda, D. Sun, Endoplasmic reticulum Ca2+ dysregulation and endoplasmic reticulum stress following in vitro neuronal ischemia: role of Na+ -K+ -Cl− cotransporter, J Neurochem 106 (2008) 1563–1576. C. Camello, R. Lomax, O.H. Petersen, A.V. Tepikin, Calcium leak from intracellular stores- the enigma of calcium signalling, Cell Calcium 32 (2002) 355–361. S.L. Budd, D.G. Nicholls, A reevaluation of the role of mitochondria in neuronal Ca2+ homeostasis, J Neurochem 66 (1996) 403–411. S. Chalmers, D.G. Nicholls, The relationship between free and total calcium concentrations in the matrix of liver and brain mitochondria, J Biol Chem 278 (2003) 19062–19070. S.A. Thayer, G.J. Wang, Glutamate-induced calcium loads: effects on energy metabolism and neuronal viability, Clin Exp Pharmacol Physiol 22 (1995) 303–304. T.E. Gunter, L. Buntinas, G. Sparagna, R. Eliseev, K. Gunter, Mitochondrial calcium transport: mechanisms and functions, Cell Calcium 28 (2000) 285–296. D.R. Pfeiffer, T.E. Gunter, R. Eliseev, K.M. Broekemeier, K.K. Gunter, Release of Ca2+ from mitochondria via the saturable mechanisms and the permeability transition, IUBMB Life 52 (2001) 205–212. D.G. Nicholls, Mitochondrial calcium function and dysfunction in the central nervous system, Biochim Biophys Acta 1787 (2009) 1416–1424.

427

[67] G. Hajnoczky, G. Csordas, S. Das, C. Garcia-Perez, M. Saotome, S. Sinha Roy, M. Yi, Mitochondrial calcium signalling and cell death: approaches for assessing the role of mitochondrial Ca2+ uptake in apoptosis, Cell Calcium 40 (2006) 553–560. [68] M.R. Duchen, Roles of mitochondria in health and disease, Diabetes 53 (Suppl. 1) (2004) S96–S102. [69] J.E. Saunders, C.C. Beeson, R.G. Schnellmann, Characterization of functionally distinct mitochondrial subpopulations, J Bioenerg Biomembr 45 (2013) 87–99. [70] J.W. Palmer, B. Tandler, C.L. Hoppel, Heterogeneous response of subsarcolemmal heart mitochondria to calcium, Am J Physiol 250 (1986) H741–H748. [71] S.L. Barrow, S.G. Voronina, G. da Silva Xavier, M.A. Chvanov, R.E. Longbottom, O.V. Gerasimenko, O.H. Petersen, G.A. Rutter, A.V. Tepikin, ATP depletion inhibits Ca2+ release, influx and extrusion in pancreatic acinar cells but not pathological Ca2+ responses induced by bile, Pflugers Arch 455 (2008) 1025–1039. [72] M.V. Ivannikov, M. Sugimori, R.R. Llinas, Calcium clearance and its energy requirements in cerebellar neurons, Cell Calcium 47 (2010) 507–513. [73] F. Baba-Aissa, L. Raeymaekers, F. Wuytack, L. Dode, R. Casteels, Distribution and isoform diversity of the organellar Ca2+ pumps in the brain, Mol Chem Neuropathol 33 (1998) 199–208. [74] F. Baba-Aissa, L. Raeymaekers, F. Wuytack, C. De Greef, L. Missiaen, R. Casteels, Distribution of the organellar Ca2+ transport ATPase SERCA2 isoforms in the cat brain, Brain Res 743 (1996) 141–153. [75] M.R. Sepulveda, M. Hidalgo-Sanchez, A.M. Mata, Localization of endoplasmic reticulum and plasma membrane Ca2+ -ATPases in subcellular fractions and sections of pig cerebellum, Eur J Neurosci 19 (2004) 542–551. [76] L. Plessers, J.A. Eggermont, F. Wuytack, R. Casteels, A study of the organellar Ca2+ -transport ATPase isozymes in pig cerebellar Purkinje neurons, J Neurosci 11 (1991) 650–656. [77] A.M. Mata, M.R. Sepulveda, Calcium pumps in the central nervous system, Brain Res Brain Res Rev 49 (2005) 398–405. [78] U. De Marchi, C. Castelbou, N. Demaurex, Uncoupling protein 3 (UCP3) modulates the activity of Sarco/endoplasmic reticulum Ca2+ -ATPase (SERCA) by decreasing mitochondrial ATP production, J Biol Chem 286 (2011) 32533–32541. [79] J. Bruce, Plasma membrane calcium pump regulation by metabolic stress, World J Biol Chem 1 (2010) 221–228. [80] X. Yu, S. Carroll, J.L. Rigaud, G. Inesi, H+ countertransport and electrogenicity of the sarcoplasmic reticulum Ca2+ pump in reconstituted proteoliposomes, Biophys J 64 (1993) 1232–1242. [81] J.M. Salvador, G. Inesi, J.L. Rigaud, A.M. Mata, Ca2+ transport by reconstituted synaptosomal ATPase is associated with H+ countertransport and net charge displacement, J Biol Chem 273 (1998) 18230–18234.