cytosolic Ca2+ concentration
mitochondrial Ca2+ concentration
dorsal root ganglia
mitochondrial membrane potential
excitatory postsynaptic current
carbonyl cyanide p-trifluoromethoxyphenylhydrazone
mitochondrial Ca2+ uniporter
mitochondrial calcium uptake 1
leucine zipper EF hand-containing transmembrane protein
K+-dependent Na+/Ca2+ exchanger
protein kinase A
protein kinase C
plasma membrane Ca2+-ATPase
tetramethylrhodamine 10K dextran
sarco-endoplasmic reticulum Ca2+-ATPase
transient receptor potential vanilloid 1
- • The first sensory synapse formed between the central processes of primary afferents and dorsal horn neurons plays an important role in controlling the flow of nociceptive information from the periphery to the CNS, and plasticity at this synapse contributes to centrally mediated pain hypersensitivity.
- • Although exocytosis and synaptic plasticity are regulated by presynaptic Ca2+, the mechanisms underlying presynaptic Ca2+ signalling at the first sensory synapse are not well understood.
- • In this study we show that the plasma membrane Ca2+-ATPase and mitochondria are the major regulators of presynaptic Ca2+ signalling in capsaicin-sensitive dorsal root ganglion neurons accounting for ∼47 and ∼40% of presynaptic Ca2+ clearance, respectively.
- • Quantitative analysis of changes in cytosolic and mitochondrial Ca2+ concentrations demonstrates that the mitochondrial Ca2+ uniporter is highly sensitive to cytosolic Ca2+ at this synapse.
- • These results help us understand presynaptic mechanisms at the first sensory synapse.
Abstract The central processes of primary nociceptors form synaptic connections with the second-order nociceptive neurons located in the dorsal horn of the spinal cord. These synapses gate the flow of nociceptive information from the periphery to the CNS, and plasticity at these synapses contributes to centrally mediated hyperalgesia and allodynia. Although exocytosis and synaptic plasticity are controlled by Ca2+ at the release sites, the mechanisms underlying presynaptic Ca2+ signalling at the nociceptive synapses are not well characterized. We examined the presynaptic mechanisms regulating Ca2+ clearance following electrical stimulation in capsaicin-sensitive nociceptors using a dorsal root ganglion (DRG)/spinal cord neuron co-culture system. Cytosolic Ca2+ concentration ([Ca2+]i) recovery following electrical stimulation was well approximated by a monoexponential function with a τ∼2 s. Inhibition of sarco-endoplasmic reticulum Ca2+-ATPase did not affect presynaptic [Ca2+]i recovery, and blocking plasmalemmal Na+/Ca2+ exchange produced only a small reduction in the rate of [Ca2+]i recovery (∼12%) that was independent of intracellular K+. However, [Ca2+]i recovery in presynaptic boutons strongly depended on the plasma membrane Ca2+-ATPase (PMCA) and mitochondria that accounted for ∼47 and 40%, respectively, of presynaptic Ca2+ clearance. Measurements using a mitochondria-targeted Ca2+ indicator, mtPericam, demonstrated that presynaptic mitochondria accumulated Ca2+ in response to electrical stimulation. Quantitative analysis revealed that the mitochondrial Ca2+ uptake is highly sensitive to presynaptic [Ca2+]i elevations, and occurs at [Ca2+]i levels as low as ∼200–300 nm. Using RT-PCR, we detected expression of several putative mitochondrial Ca2+ transporters in DRG, such as MCU, Letm1 and NCLX. Collectively, this work identifies PMCA and mitochondria as the major regulators of presynaptic Ca2+ signalling at the first sensory synapse, and underlines the high sensitivity of the mitochondrial Ca2+ uniporter in neurons to cytosolic Ca2+.
Primary nociceptive neurons send their central processes to the dorsal horn of the spinal cord where they form glutamatergic synaptic connections with the second-order nociceptive neurons (Woolf & Salter, 2006; Kuner, 2010). This synaptic connection, often referred to as the first sensory synapse, plays a critical role in controlling the flow of nociceptive information from the periphery into the CNS. Inhibiting transmission at this synapse with blockers of voltage-gated Ca2+ channels or opioids produces analgesic effects (Cao, 2006; McGivern, 2006; Heinke et al. 2011), whereas facilitation of sensory transmission through various forms of synaptic plasticity (including wind-up, central sensitization and long-term potentiation) contributes to the pain hypersensitivity associated with inflammation or nerve injury (Ji et al. 2003; Woolf & Salter, 2006; Kuner, 2010; Ruscheweyh et al. 2011). The mechanisms responsible for various forms of synaptic plasticity at the first sensory synapse are complex and not fully understood.
Presynaptic Ca2+ is the principal regulator of neurotransmitter release and synaptic plasticity that acts via multiple Ca2+-sensing proteins to control almost every aspect of the synaptic vesicle life cycle. For example, Ca2+ triggers synchronous transmitter release via the low-affinity Ca2+ sensors synaptotagmins 1, 2 and 9 (Sugita et al. 2002; Xu et al. 2007), whereas asynchronous and spontaneous transmitter release are mediated by the high-affinity Ca2+ sensor Doc2 (Groffen et al. 2010; Yao et al. 2011). Endocytotic retrieval of synaptic vesicles is stimulated by Ca2+/calcineurin-dependent dephosphorylation of dynamin-1 and other endocytotic proteins (Cousin & Robinson, 2001; Clayton & Cousin, 2009). Residual presynaptic [Ca2+]i accumulated during repeated electrical stimulation contributes to short-term synaptic plasticity by controlling the size of the release-ready pool of synaptic vesicles (Zucker & Regehr, 2002; Neher & Sakaba, 2008) through Ca2+-dependent activation of Munc13 and protein kinases A and C (PKA and PKC; Turner et al. 1999; Junge et al. 2004; Sørensen, 2004). To carry out these versatile synaptic functions, presynaptic Ca2+ signals must be precisely organized in time and space, which is achieved through the coordinated actions of Ca2+ channels, buffers and transporters. Therefore, identifying the specific components of presynaptic Ca2+ machinery is essential for understanding mechanisms of synaptic transmission and plasticity at any given synapse.
In spite of the key role of the first sensory synapse in transmitting sensory information and processing pain, the mechanisms that control presynaptic cytosolic Ca2+ concentration ([Ca2+]i) at this synapse are not known. Here we used cytosolic and mitochondrial Ca2+ imaging to examine mechanisms of presynaptic [Ca2+]i clearance at the synapses formed between dorsal root ganglion (DRG) and spinal cord (SC) neurons in culture. We focused our analysis on a subset of nociceptive DRG neurons that express TRPV1 (transient receptor potential vanilloid 1) receptors because of their well-established role in acute and chronic pain (Caterina & Julius, 2001; Szallasi et al. 2007). Our data demonstrate that plasma membrane Ca2+-ATPase (PMCA) and mitochondria are the major regulators of presynaptic [Ca2+]i at this synapse.
DRG/SC co-cultures were prepared as previously described (Medvedeva et al. 2008, 2009). In brief, newborn (postnatal days 0–2) Sprague–Dawley rats were killed by decapitation with sharp scissors, according to a protocol approved by the University of Iowa Institutional Animal Care and Use Committee and in accordance with the guidelines of the National Institutes of Health. Every effort was made to minimize the number of animals used. DRG were dissected from the thoracic and lumbar segments and incubated in pronase E dissolved in DMEM (1 mg ml−1) containing 20 mm Hepes (pH 7.4) and penicillin–streptomycin (100 U ml−1 and 100 μg ml−1, respectively) for 7 min at 37°C in a 10% CO2 incubator. Ganglia were washed twice in cold DMEM containing Hepes (20 mm; pH 7.4) and then mechanically dissociated by trituration with flame-constricted Pasteur pipettes of decreasing diameter. The spinal cord was dissected into small segments (∼0.5 mm) and then digested in DMEM containing 0.025% trypsin, Hepes (20 mm, pH 7.4) and penicillin–streptomycin (100 U ml−1 and 100 μg ml−1, respectively) for 8 min at 37°C in a 10% CO2 incubator. SC segments were then washed with cold complete DMEM medium supplemented with 5% heat-inactivated horse serum, 5% fetal bovine serum and penicillin–streptomycin (100 U ml−1 and 100 μg ml−1, respectively) and dissociated using the procedure described above for DRG dissociation. Suspensions of SC and DRG cells were plated onto 25 mm glass coverslips coated with poly-l-ornithine and laminin. Cells were grown in DMEM supplemented with 5% heat-inactivated horse serum, 5% fetal bovine serum, insulin (6 μg ml−1) and penicillin–streptomycin (100 U ml−1 and 100 μg ml−1, respectively) in a 10% CO2 incubator at 37°C. After 48 h, 5 μm cytosine β-d-arabinofuranoside (AraC) was added to cultures for 24 h. Two days later, cells were again treated with 5 μm AraC for 24 h. Cultures were grown for 10–16 days before use; 50% of the culture medium was replaced every 5–6 days.
The methods for monitoring whole-cell Ca2+ currents and excitatory postsynaptic currents (EPSCs) were similar to those previously described by our group (Medvedeva et al. 2008, 2009). Whole-cell patch-clamp recordings were obtained using a Axopatch 200B patch-clamp amplifier and a Digidata 1322A analog-to-digital converter (Molecular Devices, Union City, CA, USA). Data were collected (filtered at 2 kHz and sampled at 5 kHz) and analysed using the pClamp 9 software (Molecular Devices). Patch pipettes were pulled from borosilicate glass (Narishige, New York, USA; 3–5 mΩ) on a Sutter Instruments (Novato, CA, USA) P-87 micropipette puller. For EPSC recordings, spinal cord neurons were voltage-clamped at –60 mV using patch pipettes that were filled with the following solution (mm): 125 potassium gluconate, 10 KCl, 3 Mg-ATP, 1 MgCl2, 5 EGTA and 10 Hepes (pH 7.25 adjusted with KOH; 290 mosmol kg−1 with sucrose). The standard extracellular recording solution contained (mm): 140 NaCl, 5 KCl, 1.3 CaCl2, 0.4 MgSO4, 0.5 MgCl2, 0.4 KH2PO4, 0.6 Na2HPO4, 3 NaHCO3, 10 glucose and 10 Hepes (pH 7.4 with NaOH; 310 mosmol kg−1 with sucrose). The extracellular solution also contained 10 μm bicuculline and 2 μm strychnine to block inhibitory postsynaptic currents mediated by GABAA and glycine receptors, respectively. Evoked EPSCs in SC neurons were elicited every 4 s using a glass extracellular stimulating electrode (0.2–0.4 ms pulse) positioned near the cell body of a nearby DRG neuron. For Ca2+ current recordings, extracellular solution contained (mm): 115 choline chloride, 30 TEACl, 1.3 CaCl2, 1 MgCl2, 10 glucose, 10 Hepes and 1 μm tetrodotoxin (pH 7.4 with TEAOH; 310 mosmol kg−1 with sucrose); and the patch-pipette solution had the following composition (mm): 135 caesium gluconate, 3 Mg-ATP, 1 MgCl2, 10 EGTA and 10 Hepes (pH 7.25 adjusted with CsOH; 290 mosmol kg−1 with sucrose). Voltage-gated Ca2+ currents were evoked by step depolarizations from –60 to +10 mV (100 ms, every 20 s).
Measurements of [Ca2+]i in axonal boutons
The protocols for monitoring presynaptic [Ca2+]i were similar to those we have previously described (Medvedeva et al. 2008, 2009). Cells were placed in a flow-through chamber mounted on the stage of an inverted IX-71 microscope (Olympus, Tokyo, Japan). For [Ca2+]i measurements, DRG neurons were loaded with the low affinity (Kd∼ 5.5 μm) Ca2+ indicator Fura-FF (200 μm) or, in some experiments involving small-amplitude [Ca2+]i measurements (see Figs 4D, E and 8C, D) the high-affinity (Kd = 225 nm) Ca2+ indicator Fura-2 (200 μm), via patch pipettes by holding neurons in the whole-cell configuration for 3–5 min; recordings began within 15–20 min of the pipette withdrawing. Only neurons that had membrane potential more negative than –50 mV immediately after patch membrane rupture were used for further experimentation. Fluorescence of Fura-FF or Fura-2 was alternately excited at 340 nm (12 nm bandpass) and 380 nm (12 nm) using a Polychrome IV monochromator (TILL Photonics, Gräfelfing, Germany) via a 40× oil-immersion objective (NA = 1.35, Olympus). Emitted fluorescence was collected at 510 (80) nm using an IMAGO CCD camera (TILL Photonics). Pairs of 340/380 nm images were sampled at 10 Hz. The fluorescence ratio (R = F340/F380) was converted to [Ca2+]i according to the formula: [Ca2+]i = Kdβ(R–Rmin)/(Rmax–R) (Grynkiewicz et al. 1985). The dissociation constants (Kd) used for Fura-2 and Fura-FF were 225 and 5500 nm, respectively (Molecular Probes Handbook). Rmin, Rmax and β were determined by applying 10 μm ionomycin in Ca2+-free buffer (1 mm EGTA) and saturating Ca2+ (1.3 mm Ca2+). For Fura-2, calibration constants were: Rmin = 0.23, Rmax = 2.70 and β = 4.75. For Fura-FF, calibration constants were: Rmin = 0.21, Rmax = 2.25 and β = 5.30. Fluorescence was corrected for background, as determined prior to loading of cells with the indicators.
After dye loading and patch pipette withdrawal, intact DRG neurons were stimulated with trains of action potentials using extracellular field stimulation as previously described (Usachev & Thayer, 1999). In brief, field potentials were generated by passing current between two platinum electrodes via a Grass SS stimulator and a stimulus isolation unit (Quincy, MA, USA). Trains of 1 ms pulses were delivered at 10 Hz. The stimulus voltage threshold sufficient to elicit a detectable increase in [Ca2+]i from neuronal processes and axonal boutons was determined before beginning an experiment, and the stimulus voltage for further experimentation was set at 20 V higher than the threshold voltage. An increase of the stimulus voltage above the threshold did not lead to a change in the amplitude of the resulting [Ca2+]i transients. For the experiments involving substitution of intracellular K+ with Cs+ to examine the role of K+-dependent Na+/Ca2+ exchange (see Fig. 2E), DRG neurons were stimulated under the whole-cell voltage-clamp mode by applying 20 5-ms voltage pulses from a holding potential of –60 mV to +10 mV at a rate of 10 Hz.
Analysis of the rate of [Ca2+]i clearance
The recovery phase of each [Ca2+]i transient was fitted with a monoexponential function defined by the formula [Ca2+]i = [Ca2+]i0+Aexp(–(t–t0)/τ) using a non-linear, least-squares curve fitting algorithm (Origin 7 software), where [Ca2+]i0 is the basal [Ca2+]i level, A is the amplitude of the [Ca2+]i response, t0 is the time at the peak [Ca2+]i and τ is the time constant describing [Ca2+]i recovery. For a monoexponential [Ca2+]i recovery process the rate of [Ca2+]i clearance (–d[Ca2+]i/dt) is a linear function of [Ca2+]i that can be determined by the formula: –d[Ca2+]i/dt = k[Ca2+]i, where the rate constant k = 1/τ. To calculate the contribution of each specific Ca2+ transporting system, τ and k were calculated for each presynaptic [Ca2+]i transient under control conditions (τcontrol, kcontrol) and after inhibition of the Ca2+ transporter (τinhibit, kinhibit) of interest. The rate constants were averaged, and the partial contribution of a specific Ca2+ transport was calculated as:
All the data are presented as mean ± SEM.
Measurements of mitochondrial Ca2+ concentrations ([Ca2+]mt) in axonal boutons
[Ca2+]mt was measured using the mitochondria-targeted Ca2+ indicator mtPericam (Nagai et al. 2001) transferred to DRG neurons using lentivirus as previously described (Medvedeva et al. 2008). To visualize axonal boutons, DRG neurons overexpressing mtPericam were loaded with tetramethylrhodamine 10K dextran (Rhod10K; Invitrogen, Carlsbad, CA, USA) via patch pipettes (200 μm). mtPericam fluorescence was excited at 410 (12) nm via a 40× oil-immersion objective (NA = 1.35, Olympus) and measured at 530 (44) nm (sampling rate = 5 Hz). At 410 nm, a decrease in mtPericam fluorescence corresponds to the [Ca2+]mt increase. [Ca2+]mt changes were presented as
where F is the current fluorescence intensity and F0 is the fluorescence intensity in the resting cell. The initial rate of [Ca2+]mt increase was determined by dividing half of the [Ca2+]mt amplitude by the time that was required to reach one half of [Ca2+]mt peak.
Fura-2, Fura-FF and Rhod10K were obtained from Invitrogen; capsaicin was from Tocris (Ellisville, MO, USA); ionomycin was from Calbiochem (San Diego, CA, USA); agatoxin IVA was from Bachem (Torrance, CA, USA); and ω-conotoxin GVIA was from Alomone Labs (Jerusalem, Israel). All other reagents were purchased from Sigma (St Louis, MO, USA).
Enriched neuronal DRG cultures were prepared as previously described (Usachev et al. 2002). In brief, DRG cultures were grown in the presence of AraC (5 μm) for 5–6 days to eliminate non-neuronal cells. To provide trophic support for this glial cell-deficient culture, fresh culture medium was mixed with an equal volume of medium conditioned by DRG cultures grown in the absence of AraC for 4–6 days. These growth conditions yield cultures containing ∼75% of neurons and do not alter [Ca2+]i homeostasis in DRG neurons (Usachev et al. 2002). Total RNA was isolated using the RNase easy mini kit (Qiagen, Valencia, CA, USA) and reverse transcribed and amplified using the one-step RT-PCR kit (Qiagen) according to the manufacturer's protocols. RT-PCR reactions were performed (Hybaid PCR Sprint Thermal Cycler, MA, USA) under the following conditions: 50°C for 30 min for reverse transcription, followed by 95°C for 15 min for initial PCR activation, then 30 cycles of 95°C for 1 min, 55°C for 1 min and 72°C for 1 min, and 72°C for 10 min. Amplified transcripts were separated by electrophoresis on 1% agarose gels and detected using ethidium bromide. The sets of PCR primers are listed in Table 1.
|Target gene (accession no.)||Primer sequence|
DRG neurons were co-cultured with spinal cord neurons for 10–16 days as previously described, to enable formation of sensory synapses in vitro (Medvedeva et al. 2008, 2009). Synapses formed by DRG and SC neurons grown in co-culture possess many important characteristics of the first sensory synapse, such as: (1) EPSCs are inhibited by the antagonists of 2-amino-3-(3-hydroxy-5-methyl-isoxazol-4-yl)propanoic acid (AMPA)-type glutamate receptors (Gruner & Silva, 1994; Gu & Macdermott, 1997; Bao et al. 1998; Medvedeva et al. 2008; Heinke et al. 2011); (2) N-type voltage-gated Ca2+ channels (CaV2.2) play the predominant role in controlling evoked glutamate release (Fig. 1A and B; Gruner & Silva, 1994; Bao et al. 1998; Heinke et al. 2004; Cao, 2006); (3) a large proportion of these synapses are sensitive to TRPV1 agonists such as capsaicin and NADA that produce dramatic increases in the frequency of spontaneous EPSCs (Fig. 1C–E; Yang et al. 1998; Huang et al. 2002; Sikand & Premkumar, 2007; Medvedeva et al. 2008); and (4) these synapses demonstrate wind-up and long-term potentiation (LTP)-like forms of synaptic plasticity (Vikman et al. 2001; Ji et al. 2003; Woolf & Salter, 2006). Consequently, the DRG/SC co-culture system has been widely used as a reliable tool for studying the first sensory synapse under highly defined recording conditions (Gu & Macdermott, 1997; Vikman et al. 2001; Tsuzuki et al. 2004; Sikand & Premkumar, 2007; Medvedeva et al. 2008, 2009).
[Ca2+]i transients in presynaptic boutons of DRG neurons were measured using the low-affinity fluorescent Ca2+ indicator Fura-FF (Kd∼ 5.5 μm). This dye is ideally suited for monitoring [Ca2+]i changes predicted to be in the micromolar range, while the low Ca2+ affinity of the indicator minimizes its influence on Ca2+ buffering processes (Neher & Augustine, 1992). Fura-FF was loaded into small to medium-sized DRG neurons (18–32 μm) via a patch pipette, which enabled visualization of multiple axonal boutons along the processes (Figs 1–6). We have previously shown that these structures are associated with the presynaptic proteins synaptophysin and bassoon as well as the postsynaptic marker PSD95 (Medvedeva et al. 2008, 2009). [Ca2+]i transients were evoked by a train of 20 action potentials applied at 10 Hz, consistent with firing patterns of primary sensory neurons in vivo (Meyer & Campbell, 1981; Slugg et al. 2000). Such stimulation produced [Ca2+]i elevation from the basal level of 0.12 ± 0.05 μm to a peak of 2.14 ± 0.12 μm (n = 79 boutons/24 cells; Fig. 1F). After reaching its maximum, [Ca2+]i rapidly recovered to the prestimulus level. For the majority of axonal boutons (79 of 84 tested), [Ca2+]i recovery was well approximated using a monoexponential function with a time constant τ = 2.02 ± 0.11 s (n = 79 boutons/24 cells; Fig. 1G). Each cell was stimulated four times as described in Fig. 1F, and τ and the rate constant (k = 1/τ) for each [Ca2+]i transient were calculated. The first two stimuli were used as controls, and treatments directed at inhibiting various Ca2+ transporters were applied during the third and fourth stimuli. The contribution of a specific Ca2+ clearance mechanism was calculated as (kcontrol–kinhibit)/kcontrol100%, where kcontrol is the average rate constant in the control, and kinhibit is the average rate constant under conditions when a specific Ca2+ transporter is inhibited. In the absence of treatment, τ and k did not change significantly throughout the course of the experiment (Fig. 1I). Functional expression of TRPV1 was determined at the end of each experiment by applying 1 μm capsaicin (30 s; Fig. 1H), and only capsaicin-responding cells (mean presynaptic [Ca2+]i elevation ≥ 100% above baseline) were used for further analysis. We used the described approach to determine the roles of plasma membrane Na+/Ca2+ exchangers, PMCA, sarco-endoplasmic reticulum Ca2+-ATPase (SERCA) and mitochondria in clearing Ca2+ from the presynaptic boutons of DRG neurons.
Presynaptic Ca2+ clearance in DRG neurons depends to a small extent on plasma membrane Na+/Ca2+ exchange
Plasma membrane Na+/Ca2+ exchangers are low-affinity Ca2+ transporters that are expressed in excitable and non-excitable cells (Blaustein & Lederer, 1999; Lytton, 2007). Two large families of the exchangers have been identified: the K+-independent Na+/Ca2+ exchangers (NCX proteins), and the K+ -dependent Na+/Ca2+ exchangers (NCKX proteins). The NCX transporters utilize the transmembrane Na+ gradient to extrude 1 Ca2+ ion from the cell in exchange for 3 or 4 Na+ ions moved into the cell, whereas the NCKX transporters extrude 1 Ca2+ and 1 K+ ion in exchange for 4 Na+ ions (Blaustein & Lederer, 1999; Lytton, 2007). We blocked plasma membrane Na+/Ca2+ exchange by replacing extracellular Na+ with Li+. This treatment produced a small but significant slowing of presynaptic [Ca2+]i recovery (Fig. 2). In these experiments, the time constant of [Ca2+]i recovery increased from 1.99 ± 0.24 s in control to 2.43 ± 0.31 s in Na+-free external solution (n = 15 boutons/5 cells; P < 0.05, paired Student's t test). The corresponding rate constants were 0.59 ± 0.06 and 0.52 ± 0.07 s−1, respectively (n = 15 boutons/5 cells; P < 0.05, paired Student's t test). The overall contribution of plasma membrane Na+/Ca2+ exchange to presynaptic Ca2+ clearance in DRG neurons was ∼12%. Using the high-affinity Ca2+ indicator Fura-2 we found that the Na+/Ca2+ exchanger also controls presynaptic [Ca2+]i at rest, as replacement of extracellular Na+ with Li+ resulted in a small but significant [Ca2+]i elevation from 107 ± 4 to 134 ± 8 nm (n = 20 boutons/6 cells; P < 0.001, paired Student's t test; data not shown).
The NCX protein family comprises three isoforms (NCX1–3) and the NCKX family five (NCKX1–5). With the exception of NCKX1 (rod-specific isoform), all the NCX and NCKX isoforms are found in the brain (Annunziato et al. 2004; Lytton, 2007). DRG neurons express NCX2 and NCX3, whereas NCX1 is found primarily in satellite cells in the DRG (Persson et al. 2010). Which NCKX isoforms are expressed in DRG neurons is unknown. Using RT-PCR, we detected transcripts of NCX1, NCX2, NCX3 and NCKX2 in neuron-enriched DRG cultures (Fig. 2D). A very weak band corresponding to NCKX3 was also detected. Given the presence of some NCKX isoforms in DRG neurons, we examined whether presynaptic Ca2+ extrusion mediated by Na+/Ca2+ exchange depends on K+. Substitution of intracellular K+ with Cs+ inhibits NCKX2 and NCKX3 by 80 and 95%, respectively (Lee et al. 2007b). Therefore, we replaced K+ with an equimolar amount of Cs+ in the patch pipette solution (Fig. 2E). Since action potentials could not be evoked under these conditions, we stimulated the neurons by applying 20 short (5 ms) depolarization pulses (from –60 to +10 mV) at 10 Hz using the patch clamp technique. In the absence of intracellular K+, the slowing of [Ca2+]i recovery induced by Na+ removal was similar to that observed in the presence of intracellular K+ (compare graphs in Fig. 2C and E), suggesting that NCKX exchangers do not significantly contribute to Ca2+ clearance.
Presynaptic Ca2+ clearance in DRG neurons is highly dependent on PMCA
PMCAs are key components of Ca2+ extrusion in central and peripheral neurons (Benham et al. 1992; Usachev et al. 2002; Wanaverbecq et al. 2003; Empson et al. 2007; Gover et al. 2007b). Four major PMCA isoforms (1–4) have been identified (Carafoli, 1991; Strehler & Zacharias, 2001), and we previously showed that PMCA2 and PMCA4 are the major isoforms expressed in DRG neurons (Usachev et al. 2002). In addition, strong PMCA4 immunoreactivity was detected in the superficial layers of the dorsal horn (Tachibana et al. 2004), the area where the majority of TRPV1-positive central processes terminate (Guo et al. 1999; Cavanaugh et al. 2011). Here we examined the functional significance of PMCAs in presynaptic boutons of DRG neurons. Ca2+ extrusion by PMCA requires the countertransport of protons, and thus can be blocked by raising extracellular pH (Carafoli, 1991; Schwiening et al. 1993; Usachev et al. 2002; Wanaverbecq et al. 2003). Using Fura-2, we found that blocking PMCA by raising extracellular pH to 8.8 led to baseline [Ca2+]i elevation from 98 ± 5 to 178 ± 12 nm (n = 29 boutons/6 cells; P < 0.001, Student's t test; data not shown). Fura-FF-based measurements determined that PMCA inhibition markedly slowed [Ca2+]i recovery in the axonal boutons of DRG neurons (Fig. 3). The time constant increased from 2.01 ± 0.19 s in control to 4.29 ± 0.53 s at pH 8.8 (n = 27 boutons/7 cells; P < 0.001, paired Student's t test). The corresponding rate constants were 0.60 ± 0.06 and 0.32 ± 0.03 s−1, respectively (n = 27 boutons/7 cells; P < 0.001, paired Student's t test). The contribution of PMCA to presynaptic [Ca2+]i clearance was ∼47%.
We also found that PMCA blockade resulted in a mild but significant increase of the amplitude of electrically evoked presynaptic [Ca2+]i responses from 1.83 ± 0.14 μm in control to 2.25 ± 0.19 μm at pH 8.8 (n = 27 boutons/7 cells; P < 0.01, paired Student's t test). Raising extracellular pH to 8.8 did not have any significant effect on voltage-gated Ca2+ currents in DRG neurons, as determined by whole-cell patch-clamp recordings (Fig. 3D). Thus, PMCA strongly contributes to multiple aspects of presynaptic [Ca2+]i signalling in DRG neurons, including maintenance of low [Ca2+]i at rest, regulation of the [Ca2+]i clearance rate and setting the amplitude of [Ca2+]i elevation in response to action potentials.
Presynaptic Ca2+ clearance does not depend on SERCA
SERCA is a high-affinity Ca2+ pump that serves to lower cytosolic Ca2+ and replenish intracellular Ca2+ stores for subsequent Ca2+ release via the ryanodine or inositol 1,4,5-trisphosphate (IP3) receptors (Henzi & MacDermott, 1992; Pozzan et al. 1994; Thomas & Hanley, 1994). Three SERCA isoforms (1–3) have been identified, two of which (SERCA2 and SERCA3) are present in the brain (Mata & Sepulveda, 2005; Vangheluwe et al. 2009). SERCA plays an important role in regulating [Ca2+]i in the somata of DRG and trigeminal neurons, and its contribution to [Ca2+]i clearance can be 50% and higher, depending on the stimulation protocol and sensory neuron population (Shmigol et al. 1994; Usachev & Thayer, 1999; Lu et al. 2006; Gover et al. 2007a). We tested the effects of a selective SERCA inhibitor, cyclopiazonic acid (CPA, 5 μm), on [Ca2+]i in the presynaptic boutons of DRG neurons. Application of 5 μm CPA did not affect the recovery kinetics of presynaptic [Ca2+]i transients (Fig. 4A–C), and the time constants were 1.97 ± 0.27 and 2.21 ± 0.31 s in control and in the presence of CPA, respectively (n = 18 boutons/5 cells; P = 0.28, paired Student's t test). The corresponding rate constants were 0.66 ± 0.08 and 0.62 ± 0.09 s−1, respectively (P = 0.39, paired Student's t test). Similar results were obtained using another SERCA inhibitor, thapsigargin (Tg) (Thomas & Hanley, 1994; Thayer et al. 2002). Treatment with 1 μm Tg did not significantly change the time constant, which was 2.18 ± 0.17 s in the absence and 2.13 ± 0.19 s in the presence of 1 μm Tg (n = 16 boutons/4 cells; P = 0.72, paired Student's t test; data not shown). Thus, Ca2+ clearance in the presynaptic boutons of DRG neurons is independent of SERCA.
To test whether functional endoplasmic reticulum (ER) Ca2+ stores are present in the axonal boutons of DRG neurons, we applied an activator of Ca2+ release from ryanodine-sensitive stores, caffeine, in a Ca2+-free buffer. To improve the detection of potentially small [Ca2+]i changes in response to caffeine, we used a high-affinity Ca2+ indicator, Fura-2 (Kd = 225 nM), in these experiments. Application of 10 mm caffeine produced a small but detectable Ca2+ release in presynaptic boutons with an amplitude of 76 ± 11 nm (n = 27 boutons/5 cells; Fig. 4D and E). This value was significantly smaller than that for the caffeine-induced [Ca2+]i responses in the somata of DRG neurons (159 ± 39 nm; n = 5; P < 0.01, non-paired Student's t test). In contrast, [Ca2+]i transients induced by electrical stimulation were larger in the axonal boutons than those in the somata (Fig. 4D; arrow). Notably, after caffeine-induced depletion of the ER Ca2+ stores, switching from a Ca2+-free to normal solution containing 1.3 mm Ca2+ led to a pronounced [Ca2+]i overshoot in both presynaptic boutons and somata. This transient [Ca2+]i elevation is probably the result of activation of store-operated Ca2+ channels, as previously described for DRG neurons (Usachev & Thayer, 1999; Lu et al. 2006; Gemes et al. 2011). Moreover, depleting Ca2+ stores with 5 μm CPA or 1 μm Tg induced small but significant presynaptic [Ca2+]i elevations from 106 ± 3 to 152 ± 5 nm for CPA (n = 38 boutons/7 cells; P < 0.001 paired Student's t test) and from 116 ± 8 to 134 ± 7 nm for Tg (n = 14 boutons/4 cells; P < 0.001 paired Student's t test), consistent with the functional presence of store-operated Ca2+ channels in presynaptic boutons of DRG neurons.
Mitochondria regulate the amplitude and duration of presynaptic [Ca2+]i transients in DRG neurons
Presynaptic terminals and boutons are densely packed with mitochondria that support high energy demand at the synapses (Hollenbeck, 2005; Ly & Verstreken, 2006). Mitochondria also contribute to presynaptic Ca2+ signalling and synaptic plasticity at several central synapses, as well as at the neuromuscular junction (Billups & Forsythe, 2002; David & Barrett, 2003; Jonas, 2004; Garcia-Chacon et al. 2006; Lee et al. 2007a). We recently reported that mitochondria control TRPV1-mediated prolonged elevations of presynaptic [Ca2+]i and glutamate release at the first sensory synapse (Medvedeva et al. 2008). Here we examined whether mitochondria are also involved in shaping presynaptic [Ca2+]i transients in response to brief trains of action potentials. Ca2+ uptake by mitochondria is mediated by the mitochondrial Ca2+ uniporter (MCU), and the mitochondrial membrane potential ΔΨmt (∼–150 mV) provides the driving force for the Ca2+ uptake (Bernardi, 1999; Nicholls & Budd, 2000; Thayer et al. 2002). Mitochondrial Ca2+ uptake can be blocked by depolarizing mitochondria with a protonophore, carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP). We examined the effects of 1 μm FCCP on presynaptic [Ca2+]i transients induced by electrical stimulation (10 Hz for 2 s). FCCP treatment was combined with application of the F1,F0 ATP-synthase inhibitor oligomycin (2 μm) to prevent ATP depletion via the reverse mode of the synthase (Nicholls & Ward, 2000; Toescu & Verkhratsky, 2003). Using Fura-2, we found that FCCP+oligomycin increased baseline presynaptic [Ca2+]i from 126 ± 6 to 163 ± 7 nm (n = 17 boutons/4 cells; P < 0.001 Student's t test). Fura-FF-based measurements revealed that FCCP produced a marked slowing of [Ca2+]i recovery in the axonal boutons of DRG neurons (Fig. 5). The time constants were 2.11 ± 0.21 s in control and 4.60 ± 0.39 s after the FCCP treatment (n = 17 boutons/5 cells; P < 0.001, paired Student's t test). The corresponding rate constants were 0.54 ± 0.05 and 0.25 ± 0.02 s−1, respectively (n = 17 boutons/5 cells; P < 0.001, paired Student's t test). We also found that the amplitudes of the [Ca2+]i responses increased from 2.17 ± 0.24 μm in control to 3.45 ± 0.42 μm in the presence of FCCP (n = 17 boutons/5 cells; P < 0.01, paired Student's t test). Whole-cell patch-clamp recordings showed that FCCP+oligomycin treatment reduced voltage-gated Ca2+ currents by ∼30% in DRG neurons (Fig. 5D), probably as a result of reduced Ca2+ buffering and, consequently, increased Ca2+-dependent inhibition of voltage-gated Ca2+ channels (Hernandez-Guijo et al. 2001). Overall, our data suggest that mitochondria play an important role in buffering presynaptic Ca2+ influx and clearing [Ca2+]i elevations after periods of neuronal activity.
MCU-mediated Ca2+ transport can be blocked directly by ruthenium red (RuR; Bernardi, 1999; Thayer et al. 2002). An important advantage of this approach is that it does not disrupt either the mitochondrial electrochemical gradient or the ΔΨmt. RuR is membrane impermeable and thus was delivered to the cell through a patch pipette (10 μm) along with Fura-FF (see Methods). The amplitudes of action potential-evoked [Ca2+]i transients were significantly larger in RuR-loaded boutons (4.63 ± 0.45 μm; 33 boutons/7 cells) than in control cells (2.02 ± 0.12 μm; 79 boutons/24 cells; P < 0.001, unpaired Student's t test). As shown in Fig. 6, treatment with RuR also led to a marked slowing of presynaptic [Ca2+]i recovery. The time constants were 2.06 ± 0.10 s in control cells (n = 79 boutons/24 cells) and 3.35 ± 0.16 s in RuR-treated cells (n = 33 boutons/7 cells; P < 0.001, unpaired Student's t test). The corresponding rate constants were 0.58 ± 0.03 s−1 in control cells (n = 79 boutons/24 cells) and 0.35 ± 0.02 s−1 in RuR-loaded cells (n = 33 boutons/7 cells; P < 0.001, unpaired Student's t test). Thus, mitochondria blunt presynaptic [Ca2+]i elevations and play important role in presynaptic Ca2+ clearance in DRG neurons. The mitochondrial contribution to [Ca2+]i recovery process ranges between 40% (based on RuR experiments) and 54% (based on FCCP experiments).
It has been proposed that the Ca2+ accumulated in mitochondria can stimulate several Ca2+-dependent dehydrogenases and thereby enhance the production of ATP (Hajnoczky et al. 1995; Denton, 2009). Thus, the inhibition of mitochondrial Ca2+ uptake could potentially slow ATP-dependent transport, such as PMCA-mediated Ca2+ efflux. We therefore tested the role of PMCA in RuR-treated neurons. Inhibiting PMCA by raising extracellular pH to 8.8 produced a more than two-fold increase in the time constant, from 3.44 ± 0.20 s in RuR only-treated cells to 7.80 ± 0.53 s after a combined RuR + pH 8.8 treatment (n = 16 boutons/5 cells; P < 0.001, paired Student's t test; Fig. 6D-F). The magnitude of the pH 8.8 effect in RuR-treated cells was similar to the pH 8.8 effect in control cells (Fig. 3), indicating that PMCA-mediated Ca2+ transport was independent of the RuR-sensitive mitochondrial Ca2+ uptake under the described experimental conditions. We were unable to perform similar experiments in FCCP-treated cells because the combined application of FCCP and pH 8.8 destabilized presynaptic [Ca2+]i in DRG neurons (n = 14 boutons/3 cells; data not shown).
Properties of Ca2+ uptake by presynaptic mitochondria in DRG neurons
Given that mitochondria significantly contribute to the amplitude and time course of changes in presynaptic [Ca2+]i, we next examined the properties of Ca2+ transport by presynaptic mitochondria in capsaicin-sensitive DRG neurons. In these experiments, we directly monitored [Ca2+]mt using a mitochondria-targeted Ca2+ indicator, mtPericam (Nagai et al. 2001), as previously described (Medvedeva et al. 2008). DRG neurons transfected with mtPericam were subsequently loaded with Rhod10K through a patch pipette to enable visualization of axonal varicosities (Fig. 7A). Stimulating DRG neurons with trains of action potentials produced a rapid [Ca2+]mt elevation. Only 3–5 action potentials were required to produce detectable Ca2+ uptake by mitochondria (Fig. 7B). Comparison of the time courses of [Ca2+]i and [Ca2+]mt showed that mitochondrial Ca2+ uptake developed with a 1–2 s delay relative to the presynaptic [Ca2+]i rise. For example, in response to a stimulus with 20 action potentials (10 Hz, 2 s), [Ca2+]i and [Ca2+]mt reached their peak values within 1.9 ± 0.1 s (79 boutons/24 cells) and 3.5 ± 0.2 s (n = 22 boutons/5 cells; P < 0.001, unpaired Student's t test compared to the [Ca2+]i rising time) after beginning of the stimulation, respectively. In addition, Ca2+ removal from mitochondria occurred at a markedly slower rate (half recovery time = 9.2 ± 0.8 s; n = 22 boutons/5 cells) than Ca2+ clearance from the presynaptic cytosol (Fig. 7D). The fact that [Ca2+]mt remained elevated long after [Ca2+]i has fully recovered suggests that mitochondria serve as an integrator of consecutive presynaptic [Ca2+]i spikes produced by repeated neuronal stimulation.
Next we addressed the important and widely debated question of how mitochondrial Ca2+ uptake depends on [Ca2+]i. The reported values for the Ca2+ affinity of MCU cover a wide range (0.5–20 μm), depending on the preparation, cell type and method of monitoring mitochondrial Ca2+ uptake (Gunter & Pfeiffer, 1990; Bernardi, 1999; Spat et al. 2008). In neuronal preparations, high Ca2+ sensitivity of MCU (K0.5∼0.4–0.5 μm) was reported for presynaptic mitochondria at nerve–muscular junctions of the fruit-fly and lizard (David, 1999; Chouhan et al. 2010). In contrast, mitochondrial Ca2+ uptake in the cell body of bullfrog sympathetic neurons was estimated to have a much lower Ca2+ affinity (K0.5∼ 10 μm; Colegrove et al. 2000). Comparable quantifications in mammalian neurons are lacking. To determine the dependence of mitochondrial Ca2+ uptake on [Ca2+]i in axonal boutons of DRG neurons, we compared [Ca2+]i and [Ca2+]mt elevations induced by depolarizing pulses of incremental amplitude (extracellular buffers containing 10, 15, 20, 30, 50 or 90 mm KCl; Fig. 8), as well as by treating cells with 1 μm capsaicin, which is known to produce large presynaptic [Ca2+]i and [Ca2+]mt elevations in DRG neurons (Medvedeva et al. 2008). To facilitate detection of small presynaptic [Ca2+]i changes in response to mild to moderate depolarization (10–50 mm KCl), we used the high-affinity Ca2+ indicator Fura-2, while the low-affinity Ca2+ indicator Fura-FF was employed for monitoring [Ca2+]i produced by the stronger stimuli such as 90 mm KCl and 1 μm capsaicin. Since the excitation spectra of Fura-2 (Fura-FF) and mtPericam overlap significantly, the measurements of [Ca2+]i and [Ca2+]mt were carried out in parallel experiments. Figure 8 shows representative presynaptic [Ca2+]i and [Ca2+]mt traces in response to incremental depolarization. [Ca2+]i changes in axonal boutons were detectable in response to 10 mm KCl, whereas triggering [Ca2+]mt response required 15 mm KCl to produce a stronger depolarization. Analysis of the initial rate of the [Ca2+]mt rise as a function of presynaptic [Ca2+]i elevations showed that Ca2+ uptake by mitochondria was induced by a [Ca2+]i elevation as little as 200–300 nm (K0.5∼ 600 nm); the rate of [Ca2+]mt rise (VMCU) progressively increased with the [Ca2+]i increase, until VMCU reached its maximum once [Ca2+]i began to exceed ∼2 μm (Fig. 8E). Thus, the mitochondrial Ca2+ uptake mechanism in axonal boutons of DRG neurons is highly sensitive to cytosolic Ca2+. Moreover, the Hill coefficient was 2.3, suggesting that MCU activation by Ca2+ is cooperative (Fig. 8E).
Finally, we examined the expression of several recently identified components of mitochondrial Ca2+ transport. MCU has been proposed to mediate mitochondrial Ca2+ uptake (Baughman et al. 2011; De Stefani et al. 2011), whereas the Na+/Ca2+/Li+ exchanger (NCLX) is probably responsible for mitochondrial Na+/Ca2+ exchange (Palty et al. 2010). Leucine zipper EF hand-containing transmembrane protein (Letm1) was identified as a mitochondrial Ca2+/H+ antiporter that can contribute to both mitochondrial Ca2+ uptake and release, depending on the mitochondrial Ca2+ concentration (Jiang et al. 2009). In addition, the mitochondrial calcium uptake 1 (MICU1) has been proposed to function as an essential regulator of mitochondrial Ca2+ uptake (Perocchi et al. 2010; Mallilankaraman et al. 2012). RT-PCR analysis of RNA extracted from neuron-enriched DRG cultures demonstrated the presence of all four transcripts (Fig. 8F).
The present study identifies mechanisms of presynaptic [Ca2+]i regulation in an in vitro model of the first sensory synapse. This synapse controls nociceptive transmission from the periphery to the CNS, and plasticity at this synapse contributes to bidirectional modulation of pain processing (Ji et al. 2003; Woolf & Salter, 2006; Kuner, 2010; Ruscheweyh et al. 2011). We found that presynaptic [Ca2+]i in DRG is at ∼100 nm at rest, and that this low baseline [Ca2+]i is maintained by Ca2+ extrusion via PMCA and NCX, as well by Ca2+ buffering into intracellular Ca2+ organelles. Presynaptic [Ca2+]i transients induced by brief electrical stimulation rapidly recovered to the baseline with a time constant of ∼2 s. The ER Ca2+ pumps did not significantly contribute to presynaptic [Ca2+]i clearance at this synapse under the described stimulation conditions. On the other hand, PMCA-mediated Ca2+ extrusion and Ca2+ uptake by presynaptic mitochondria had major impacts on the rate of [Ca2+]i clearance, accounting for ∼47 and 40%, respectively, of the [Ca2+]i recovery. Ca2+ imaging in presynaptic mitochondria revealed a high sensitivity of MCU to [Ca2+]i elevations, with 3–5 action potentials being sufficient to produce detectable Ca2+ uptake by presynaptic mitochondria. Quantification of the rate of mitochondrial Ca2+ uptake as a function of [Ca2+]i yielded a K0.5 of ∼600 nm, and a Hill coefficient of 2.3. These values are consistent with the high Ca2+ affinity of presynaptic MCU (K0.5∼ 0.4–0.5 μm) reported for non-mammalian synapses (David, 1999; Chouhan et al. 2010). In addition, the Hill coefficient of 2.3 is similar to the previously reported value of ∼2 for isolated mitochondria and intact cells (Gunter & Pfeiffer, 1990; Colegrove et al. 2000), and suggests that Ca2+ cooperatively binds to and activates MCU in the presynaptic mitochondria of DRG neurons.
Our finding that presynaptic [Ca2+]i recovers in DRG neurons with τ∼ 2 s is in good agreement with the [Ca2+]i recovery rates reported for presynaptic boutons in the hippocampus (1–3 s) (Scotti et al. 1999; Scott & Rusakov, 2006) and neurohypophysial axonal terminals (1–2 s) (Lee et al. 2002), although faster [Ca2+]i clearance kinetics have been observed in presynaptic terminals of Purkinje and cerebellar granule cells (τ∼ 200–300 ms) (Mintz et al. 1995; Regehr & Atluri, 1995). Our data are also in good agreement with the observations in human nociceptive C-fibres, where electrically evoked [Ca2+]i transients recovered to baseline with a half-recovery time of ∼2–3 s (Mayer et al. 1999). However, a 5–10-fold slower rate of [Ca2+]i clearance was reported for sensory terminals of the rat cornea (Gover et al. 2007b), suggesting that Ca2+ clearance mechanisms at these sensory terminals originating from the trigeminal ganglia appear to be quite different from those in the peripheral and central processes of DRG neurons. This may not be surprising given that mitochondria do not participate in [Ca2+]i clearance from corneal terminals (Gover et al. 2007b), but significantly impact presynaptic [Ca2+]i recovery in DRG neurons (Figs 5 and 6). It is also possible that use of the high-affinity Ca2+ indicator Oregon Green BAPTA-1 dextran (Kd∼170 nm) to measure [Ca2+]i in the corneal terminals (Gover et al. 2007b) might have also contributed to a slowed rate of [Ca2+]i recovery in that work (Neher & Augustine, 1992; Kirischuk et al. 1999).
Our finding that SERCA does not significantly contribute to presynaptic Ca2+ clearance in DRG neurons (Fig. 4) is somewhat surprising given that SERCA plays major roles in setting the duration and amplitude of [Ca2+]i responses in the somata of DRG and trigeminal sensory neurons (Usachev & Thayer, 1999; Lu et al. 2006; Gover et al. 2007a; Rigaud et al. 2009). Just the same, our data are consistent with a negligible role of SERCA in peripheral sensory terminals of the cornea (Gover et al. 2007b), suggesting that the impact of SERCA on [Ca2+]i regulation differs substantially between the cell body and axons in sensory neurons. However, we cannot rule out the possibility that SERCA blockade led to enhanced function of PMCA or other Ca2+ transporters, thereby masking the effects of SERCA inhibitors. It is likely that the presynaptic boutons of primary sensory neurons contain functional intracellular Ca2+ stores associated with the ER. First, the SERCA inhibitor Tg was shown to modulate glutamate release at the first sensory synapse (Tsuzuki et al. 2004). Second, we found that an activator of ryanodine receptors, caffeine, induced Ca2+ release from the intracellular stores in presynaptic boutons of DRG neurons (Fig. 4D). Although SERCA pumps do not contribute significantly to the rate of presynaptic [Ca2+]i recovery, Ca2+ stores are involved in the maintenance of low [Ca2+]i at rest, as the SERCA inhibitors CPA and Tg produced 20–50 nm elevation of the [Ca2+]i baseline. The latter could be explained either by activation of Ca2+ influx through presynaptic store-operated Ca2+ channels (Usachev & Thayer, 1999; Gemes et al. 2011) or the role of SERCA pumps in controlling presynaptic [Ca2+]i at rest.
Na+/Ca2+ exchange of the plasma membrane is another mechanism that can contribute to Ca2+ clearance. Based on our RT-PCR data (Fig. 2D), DRG neurons express three Na+/Ca2+ exchanger isoforms that are K+-independent (NCX1, NCX2 and NCX3) and one that is K+-dependent (NCKX2). The functional analysis showed that plasma membrane Na+/Ca2+ exchange regulated the resting [Ca2+]i and contributed approximately 12% to presynaptic Ca2+ clearance in DRG neurons (Fig. 2), and that this transport was probably mediated by the NCX, but not NCKX, isoforms (Fig. 2E). Only limited information is available about the specific roles of NCX and NCKX at various synapses. NCKX was shown to be the major mechanism mediating Ca2+ extrusion from axonal terminals of the rat neurohypophysis (Lee et al. 2002), whereas both NCX and NCKX contribute to presynaptic [Ca2+]i clearance at the calyx of Held synapse (Kim et al. 2005). Notably, deletion of NCX2 results in increased paired-pulsed facilitation of synaptic transmission and enhanced LTP in the CA1 region of the hippocampus (Jeon et al. 2003), whereas deletion of NCKX2 leads to a profound loss of the hippocampal LTP (Li et al. 2006). Thus, it is likely that the NCX and NCKX isoforms play distinct roles at various synapses.
Identification of PMCA as a key regulator of presynaptic [Ca2+]i at the sensory synapses is in agreement with the major role of PMCA in controlling [Ca2+]i recovery in the somata of DRG and trigeminal sensory neurons (Benham et al. 1992; Usachev et al. 2002; Gover et al. 2007a), as well as in peripheral sensory terminals innervating the cornea (Gover et al. 2007b). PMCA is a high-affinity (0.1–0.2 μm) ATP-dependent Ca2+ transporter that extrudes Ca2+ from the cell (Carafoli, 1991; Elwess et al. 1997). The PMCA family of proteins consist of four major isoforms, PMCA1–4, and alternative RNA splicing of these gives rise to nearly 30 PMCA proteins that differ in their tissue distribution, function and regulatory properties (Strehler & Zacharias, 2001; Thayer et al. 2002). PMCA2 (splice variants 2a and 2b) and PMCA4 (splice variant 4b) are the major PMCA isoforms expressed in the DRG (Usachev et al. 2002). PMCA4 is highly expressed in the superficial lamina of the dorsal horn, where the majority of TRPV1-positive sensory fibres terminate; PMCA2, on the other hand, is found in the deeper lamina of the dorsal horn (Tachibana et al. 2004). PMCA2 has very high affinity for Ca2+ (∼0.09 μm for PMCA2a and ∼0.06 μm for PMCA2b), suggesting that it plays an important role in maintaining low resting [Ca2+]i (Elwess et al. 1997). PMCA4b exhibits a somewhat lower apparent affinity for Ca2+ (∼0.16 μm) (Elwess et al. 1997), but its activity can be markedly increased by PKC-mediated phosphorylation (Enyedi et al. 1996). Notably, this mechanism is responsible for a marked PKC-mediated acceleration of Ca2+ efflux from the somata and axonal boutons of DRG neurons (Usachev et al. 2002).
PMCA2 and PMCA4 are also highly sensitive to stimulation by calmodulin (CaM). Ca2+-dependent binding of CaM to the PMCA autoinhibitory domain, located in its carboxy-terminus, disinhibits the pump, which results in a marked increase of both Ca2+ affinity and maximal velocity (Vmax) of PMCA (Carafoli, 1991; Strehler & Zacharias, 2001). The two major PMCA isoforms in DRG neurons, PMCA2b and PMCA4b (Usachev et al. 2002), are characterized by unusually slow dissociation of CaM (>20 min) (Caride et al. 1999, 2001). This implies that even a brief elevation in [Ca2+]i will produce a long-lasting CaM/Ca2+-dependent stimulation of PMCA. Indeed, a priming [Ca2+]i elevation has been reported to result in long-term enhancement of Ca2+ extrusion (∼1 h) in DRG neurons (Pottorf & Thayer, 2002). Thus, PMCA-mediated Ca2+ extrusion in sensory neurons is subject to modulation by both neuronal activity and PKC. In the context of spinal pain processing, the described mechanisms are predicted to accelerate presynaptic [Ca2+]i clearance during central sensitization, which is known to be associated with heightened neuronal activity in the spinal cord and enhanced release of neuromodulators coupled to PKC signalling (e.g. bradykinin and substance P). However, direct testing of this idea is hampered by a lack of isoform-specific inhibitors of PMCA and technical difficulties associated with presynaptic Ca2+ imaging in spinal cord slices or in vivo.
Ca2+ uptake by presynaptic mitochondria is another major mechanism responsible for presynaptic [Ca2+]i clearance in DRG neurons. Electron microscopy has demonstrated numerous presynaptic mitochondria in the central processes of primary sensory neurons, including TRPV1-expressing boutons (Maxwell & Rethelyi, 1987; Rethelyi et al. 1989; Hwang et al. 2003). We also found that axonal boutons are intensely labelled by the mitochondria-targeted Ca2+ indicator mtPericam (Medvedeva et al. 2008; Figs 7 and 8). Presynaptic clustering of mitochondria is consistent with the high energy demand at the synapses. Mitochondria also play important roles in shaping presynaptic [Ca2+]i signals at various synapses (Billups & Forsythe, 2002; David & Barrett, 2003; Jonas, 2004; Garcia-Chacon et al. 2006; Lee et al. 2007a), including the first sensory synapse (Medvedeva et al. 2008; Figs 5 and 6). Mitochondria buffer excessive Ca2+ via the Ca2+ uniporter mechanism, and subsequently release accumulated Ca2+ back into the cytosol via Na+/Ca2+ exchange or other mechanisms (Bernardi, 1999). Ca2+ buffering by mitochondria limits the amplitude of the cytosolic [Ca2+]i elevation, and accelerates the initial recovery of presynaptic [Ca2+]i (Medvedeva et al. 2008; Figs 5 and 6). Our data suggest that the impact of mitochondria on the amplitude of presynaptic [Ca2+]i responses is much greater than that of PMCA (FCCP or RuR induced ∼60–100% increase in the amplitude of [Ca2+]i response compared to ∼20% increase produced by pH 8.8). One possible explanation of this difference is that Ca2+ transport by the mitochondrial uniporter is much faster than that by PMCA (Miller, 1991; Elwess et al. 1997; Gunter & Sheu, 2009).
Our comparison of the time courses of [Ca2+]i and [Ca2+]mt changes induced by electrical stimulation demonstrated that [Ca2+]mt reaches its peak by 1–2 s later than presynaptic [Ca2+]i (Fig. 7D), indicating that mitochondrial Ca2+ uptake continues during the initial phase of [Ca2+]i recovery. This observation is consistent with the role of mitochondrial Ca2+ uptake in [Ca2+]i recovery in the axonal boutons of DRG neurons. Based on the effects of the mitochondrial Ca2+ uniporter inhibitor RuR, we estimate that mitochondria account for at least 40% of the [Ca2+]i recovery in the axonal boutons of DRG neurons. Although our analysis using the protonophore FCCP suggested that MCU has a somewhat stronger effect on the rate of presynaptic Ca2+ clearance (∼54%), this difference could be due to a partial inhibition of ATP synthesis by FCCP.
Ca2+ initially accumulated by mitochondria can be subsequently released back into the cytosol. Following intense (e.g. tetanic) stimulation, strong Ca2+ efflux from mitochondria can markedly slow recovery of presynaptic [Ca2+]i leading to a [Ca2+]i plateau lasting many minutes after stimulation is terminated (Tang & Zucker, 1997; Garcia-Chacon et al. 2006; Lee et al. 2007a). We previously reported that TRPV1 activation or intense electrical stimulation (20 Hz for 30 s) results in a long-lasting presynaptic [Ca2+]i plateau associated with enhanced glutamate release in DRG neurons (Medvedeva et al. 2008). In contrast, mild electrical stimulation such as that used in the present study (10 Hz for 2 s) was insufficient to generate a [Ca2+]i plateau in the same axonal boutons, in spite of the contribution of mitochondria to Ca2+ clearance. Thus, a specific role of presynaptic mitochondria at the sensory synapse depends on the type and intensity of the stimulus.
In addition to regulating presynaptic [Ca2+]i, mitochondrial Ca2+ uptake can stimulate the production of ATP via Ca2+-dependent dehydrogenases (Hajnoczky et al. 1995; Denton, 2009; Chouhan et al. 2012) and the generation of reactive oxygen species (ROS) (Nicholls & Budd, 2000; Hongpaisan et al. 2004). Both effects require strong neuronal activation (Hongpaisan et al. 2004; Chouhan et al. 2012), and this probably explains why mitochondrial Ca2+ uptake following the mild stimulation used in our study did not detectably impact ATP-dependent Ca2+ transport (Fig. 6D–F). In pathological pain states, neuronal activity in the spinal cord is markedly intensified, and the resulting enhancement of ROS production is thought to contribute to central sensitization (Schwartz et al. 2009). Notably, recent data suggest that mitochondrial Ca2+ uptake is essential for ROS-dependent plasticity in the spinal cord, and for the development of persistent pain (Kim et al. 2011).
In summary, this study provides the first detailed characterization of Ca2+ clearance mechanisms in presynaptic boutons of DRG neurons, and identifies PMCA and mitochondria as major regulators of Ca2+ signalling at the first sensory synapse. Development of new pharmacological and genetic tools targeting specific PMCA isoforms and mitochondrial Ca2+ transporters will be required to determine their precise contribution to the various forms of synaptic plasticity involved in spinal pain processing, such as wind-up, LTP and central sensitization.
Y.M.U. and L.P.S. designed the work. L.P.S., Y.M.U., M.S.K., P.R.H. and Y.V.M. performed the experiments. Y.M.U. wrote the paper. All authors provided critical intellectual input to the text and figures and approved the final version of the article.
This work was supported by National Institutes of Health grant NS072432 and the Fraternal Order of Eagles Diabetes Research Center (FOEDRC) pilot grant to Y.M.U. and by predoctoral fellowships from the American Heart Association, Midwest Affiliate, to M.S.K. and P.R.H. We thank Dr Miyawaki for his gift of the mtPericam plasmid.