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Although the calcium/calmodulin-activated phosphatase calcineurin may dephosphorylate many endocytic proteins, it is not considered a key molecule in mediating the major forms of endocytosis at synapses - slow, clathrin-dependent and the rapid, clathrin-independent endocytosis. Here we studied the role of calcineurin in endocytosis by reducing calcium influx, inhibiting calmodulin with pharmacological blockers and knockdown of calmodulin, and by inhibiting calcineurin with pharmacological blockers and knockout of calcineurin. These manipulations significantly inhibited both rapid and slow endocytosis at the large calyx-type synapse in 7 - 10 days old rats and mice, and slow, clathrin-dependent endocytosis at the conventional cultured hippocampal synapse of rats and mice. These results suggest that calcium influx during nerve firing activates calcium/calmodulin-dependent calcineurin, which controls the speed of both rapid and slow endocytosis at synapses by dephosphorylating endocytic proteins. The calcium/calmodulin/calcineurin signalling pathway may underlie regulation of endocytosis by nerve activity and calcium as reported at many synapses over the last several decades.
The calcium/calmodulin-dependent phosphatase calcineurin, found widely in the nervous system (Rusnak and Mertz, 2000), may dephosphorylate many endocytosis proteins, such as dynamin, synaptojanin, the adaptor protein AP180, and phosphatidylinositol phosphate kinase type Iγ (Clayton et al., 2007). This raises the possibility that calcineurin may mediate the calcium-dependent regulation of endocytosis (Cousin and Robinson, 2001), as observed at many synapses (Wu, 2004; Royle and Lagnado, 2003). Based on measurements of the FM dye release in the synaptosome preparation, an early study implicated the involvement of calcineurin in endocytosis during extremely intense stimulation, depolarization for hundreds of seconds (Marks and McMahon, 1998). Consistent with this implication, calcineurin is considered to be involved only in bulk endocytosis during very intense stimuli, but not in slow, clathrin-dependent endocytosis during milder stimuli at cerebellar synapses (Clayton and Cousin, 2009; Clayton et al., 2009; Evans and Cousin, 2007). Slow endocytosis at a calyx-type nerve terminal is triggered by more than 10 μM of calcium (Hosoi et al., 2009; Wu et al., 2009), which is much higher than the affinity of calcineurin to calcium (~1 – 1.5 μM). This result also argues against the involvement of calcineurin in slow endocytosis during milder stimuli (Hosoi et al., 2009). Rapid endocytosis, which is presumably clathrin-independent (Artalejo et al., 1995; Jockusch et al., 2005), is another form of endocytosis often observed at synapses (Wu et al., 2007). Likely owing to its fast speed, calcineurin-mediated dephosphorylation is not considered to be involved in this process. In summary, while calcineurin may dephosphorylate endocytic proteins, there has been no molecular and biophysical evidence showing the involvement of calcineurin in rapid and slow endocyotsis, two major forms of endocytosis observed in near physiological stimuli at synapses (Royle and Lagnado, 2003; Wu et al., 2007).
Recent studies at giant calyx-type synapses suggest that calcium influx triggers and regulates rapid and slow endocytosis (Wu et al., 2009; Hosoi et al., 2009). The calcium binding protein calmodulin was implied as the calcium receptor, because its blockers significantly inhibited rapid and slow endocytosis (Wu et al., 2009). However, three main issues had remained unresolved. First, pharmacological blockers may not be specific to calmodulin. Direct molecular biological evidence supporting calmodulin as the calcium sensor for endocytosis is missing, likely because calmodulin is encoded by three dispersed genes in vertebrates, making it difficult to use genetic approaches. Second, if calcium/calmodulin initiates rapid and slow endocytosis, its downstream target is unclear. Although calcineurin has been discussed as a downstream target for a long time, evidence supporting its role in rapid and slow endocytosis is missing. Third, it is unclear whether the findings obtained at giant synapses apply to the majority of synapses, the small conventional synapses. We addressed these three issues by combining quantitative measurements of endocytosis, pharmacological tools, and genetic approaches at both giant calyx-type and small cultured hippocampal synapses. We found that block of the calcium/calmodulin/calcineurin signalling pathway significantly inhibited both rapid and slow endocytosis, which calls for modification of the current endocytosis model to include calcineurin as a key player.
Parasagittal brainstem slices (200 μm thick) containing the medial nucleus of the trapezoid body were prepared from 7 - 10 days old male or female Wistar rats or mice using a vibratome (Wu et al., 2009). Whole-cell capacitance measurements were made with the EPC-9 amplifier together with the software lock-in amplifier (PULSE, HEKA, Lambrecht, Germany) that implements Lindau-Neher's technique (Sun and Wu, 2001; Sun et al., 2004). The frequency of the sinusoidal stimulus was 1000 Hz and the peak-to-peak voltage of the sine wave was ≤60 mV. We pharmacologically isolated presynaptic Ca2+ currents with a bath solution (~22 - 24 °C) containing (in mM): 105 NaCl, 20 TEA-Cl, 2.5 KCl, 1 MgCl2, 2 CaCl2, 25 NaHCO3, 1.25 NaH2PO4, 25 glucose, 0.4 ascorbic acid, 3 myo-inositol, 2 sodium pyruvate, 0.001 tetrodotoxin (TTX), 0.1 3,4-diaminopyridine, pH 7.4 when bubbled with 95% O2 and 5% CO2. The presynaptic pipette contained (in mM): 125 Cs-gluconate, 20 CsCl, 4 MgATP, 10 Na2-phosphocreatine, 0.3 GTP, 10 HEPES, 0.05 BAPTA, pH 7.2, adjusted with CsOH. Since DMSO (0.1%) was used to dissolve CsA (Sigma, St. Louis, MO) in the pipette solution, the control solution for this drug also contained 0.1% DMSO (Fig. 1A, D). CaN457-482 and scrambled CaN457-482 were purchased from Calbiochem, La Jolla, CA, and GenScript USA Inc, Piscataway, NJ, respectively.
Hippocampal cultures, stimulation and fluorescence imaging were similar to those described previously (Sankaranarayanan and Ryan, 2000). Hippocampal CA1-CA3 regions from p0 - 2 Sprague Dawley rats (if not mentioned) or p0 mice were dissected, dissociated, and plated on Matrigel-coated glass coverslips (BD Biosciences Clontech, Palo Alto, CA). Cells were maintained at 37°C in a 5% CO2 humidified incubator with a culture media consisting of MEM (Invitrogen, Carlsbad, CA), 0.5% glucose, 0.1 g/l bovine transferrin (Calbiochem, La Jolla, CA), 0.3 g/l glutamine, 10% fetal bovine serum (Invitrogen), 2% B-27 (Invitrogen), and 3 μM cytosine β-D-arabinofuranoside (Sigma, St. Louis, MO). On 6 - 8 days after plating, calcium-phosphate-mediated gene transfer was used to transfect cultures with synaptopHluorin (SpH, kindly provided by Dr. Gero Miesenböck), calmodulin shRNA plasmid, or calmodulin shRNA-resistant plasmid. After transfection, cultures were maintained at 37°C in a 5% CO2 humidified incubator for another 6 - 8 days before use. Unless otherwise indicated, all chemicals were obtained from Sigma.
Coverslips were mounted in a stimulation chamber (RC-21BRFS chamber, Warner Instruments, Hamden, CT) 6 – 8 days after transfection. The action potential was evoked by passing a 1 ms current pulse of 20 mA via platinum electrodes in the chamber. The bath solution (~22 - 24 °C) contained (in mM): 119 NaCl, 2.5 KCl, 2 CaCl2, 2 MgCl2, 25 HEPES (buffered to pH 7.4), 30 glucose, 0.01 6-cyano-7-nitroquinoxaline-2, 3-dione (CNQX) and 0.05 d, l-2-amino-5-phosphonovaleric acid (AP-5). When lowering the CaCl2 concentration, MgCl2 was increased to keep the divalent ion concentration constant.
SpH images were acquired at 1 Hz using the Zeiss LSM 510 META confocal microscope with a 40×, 1.3 NA oil immersion objective. Images were analyzed using Zeiss LSM510 software. All functionally visible varicosities were selected for analysis by testing their responsiveness to stimulation. The fluorescence intensity within a region of at least 1.5 μm × 1.5 μm were averaged together for each bouton, which avoided fluorescence decay caused by faster diffusive processes (Granseth et al., 2006). Each group of data was obtained from at least 3 different batches of cultures.
Calmodulin shRNA and calmodulin shRNA-resistant plasmids are described recently (Pang et al., 2010). We made only one modification, i.e., the GFP was cut off from these plasmids to avoid the fluorescence conflict with co-transfected SpH. Both plasmids include two RNA-polymerase III promoters (human H1 and human U6) in tandem and the Ubiquitin C promoter downstream of U6 promoter. For the calmodulin shRNA plasmid, a short-hairpin sequence targeting a common sequence found in the calmodulin 1 and calmodulin 2 mRNAs (CTGACTGAAGAGCAGATTGC; full shRNA sequence: TCGACCCCTGACTGAAGAGCAGATTGCTTCAAGAGAGCAATCTGCTCTTCAGTCAGTTTTTGGAAAT) was inserted into the downstream of the H1 promoter. A second short-hairpin sequence targeting the calmodulin 3 mRNA (sequence: CGCGCCCACGGAGCTGCAGGACATGATTATTCAAGAGATAATCATGTCCTGCAGCTCCGTTTTTTGGAAA) was inserted into the downstream of the U6 promoter.
The calmodulin shRNA-resistant plasmid includes not only the two short-hairpin sequences described above to knockdown calmodulin, but also a mutant calmodulin sequence to rescue calmodulin expression. The BamHI-EcoRI sites downstream of ubiquitin C promoter are for the insertion of rescue calmodulin cDNA. The targeted sequences in the rescue calmodulin cDNA were mutated to TTAACGGAAGAACAAATCGC and CAGAACTTCAAGATATGATCA to create a maximum difference between the shRNAs and rescue cDNA without changing the calmodulin protein sequence.
For immunostaining, neurons were fixed with 4% paraformaldehyde, permeabilized with 0.2% Triton X-100, and subsequently incubated with the primary and secondary antibodies. The antisera were diluted in PBS with 2% bovine serum albumin and incubated with cells overnight at 4 °C. After several rinses in PBS, cells were incubated with fluorescence-conjugated donkey anti-rabbit IgG (1:100) and rhodamine-conjugated donkey anti-mouse or donkey anti-goat IgG (1:100, Jackson ImmunoResearch Lab, West Grove, PA) for 30 min at 37 °C. The following antibodies were used for immunocytochemistry: polyclonal rabbit anti-GFP (1:1000, Invitrogen, Carlsbad, CA), and monoclonal mouse anti-calmodulin (1:500, Santa Cruz Biotechnology Inc., Santa Cruz, CA). Calmodulin expression level was measured at cell bodies and compared with the fluorescence intensity in un-transfected (SpH-negative) neurons. Although calmodulin was also found in neuronal branches, the immunostaining signal was weak and difficult to quantify. Thus, we did not quantify calmodulin level in neuronal branches.
For Western blot of PC12 cells, cells were washed three times with ice-cold PBS. Cell lysates were prepared in the modified RIPA buffer including protease inhibitors. Equal protein amounts were analyzed by SDS-PAGE and immunoblotting using antibodies against calmodulin (1: 1000, Santa Cruz Biotechnology Inc.) and actin (used as an internal control, 1: 10,000, Chemicon, Temecula, CA).
For brain tissue Western blot, dissociated hippocampal CA1-CA3 regions of 9 days old mice were homogenized in the ice-cold, modified RIPA buffer, which included protease inhibitors. The homogenates were centrifuged at 13,000 rpm at 4°C for 20 min. The supernatants were loaded to SDS-PAGE for immunoblotting using antibodies against calcineurin Aα subunit (1:200), calcineurin Aβ subunit (1:1000, Santa Cruz Biotechnology Inc.), and actin (1:10,000).
Calcineurin Aα+/- and Aβ+/- mice were provided by Dr. Jennifer L. Gooch (Zhang et al., 1996) and Jeffery D. Molkentin (Bueno et al., 2002), respectively. Calcineurin Aα-/- and Aβ-/- mice were obtained by heterozygous breeding using standard mouse husbandry procedures. Mouse genotypes were determined by PCR reaction with primers described previously (Gooch et al., 2004).
The statistical test was t-test. Means are presented as ± SE. For capacitance measurements, the Ratedecay was measured as the rate of decay in the first 2 - 10 s after stimulation. When endocytosis was inhibited, the Ratedecay was measured as the mean decay rate within 10 - 30 s after stimulation, because the capacitance decay was approximately linear within this time window. For SpH signal, the Ratedecay was measured as the decay rate in the first 4 - 10 s after stimulation. When endocytosis was inhibited, the Ratedecay was measured from the first 10 – 30 s after stimulation.
The whole-cell capacitance was measured at the calyx in 7 – 10 days old rats. We induced slow and rapid endocytosis with 1 and 10 pulses of 20 ms depolarization (from -80 to +10 mV, if not mentioned) at 10 Hz, respectively (Wu et al., 2005; Wu et al., 2009). In control, at 4 – 10 min after whole-cell break in (0.1% DMSO in pipette), a 20 ms depolarization induced a capacitance jump (ΔCm) of 462 ± 31 fF (n = 12), followed by a mono-exponential decay with a time constant (τ) of 18.6 ± 1.0 s (n = 12) and an initial endocytosis rate (Ratedecay) of 28 ± 3 fF/s (n = 12, Fig. 1A). Ten depolarizing pulses of 20 ms at 10 Hz induced a ΔCm of 1669 ± 109 fF (n = 12), followed by a bi-exponential decay with τ of 2.8 ± 0.3 s (44 ± 4%) and 23.0 ± 2.1 s (n = 12, Fig. 1D), respectively. The Ratedecay after 10 depolarizing pulses was 270 ± 20 fF/s (n = 12, Fig. 1D), which reflected mostly (>80%) the rapid component of endocytosis as demonstrated previously (Wu et al., 2005; Wu et al., 2009). This was confirmed in the present study, because the mean Ratedecay of the rapid component of endocytosis was approximately 262 fF/s, as calculated from the ratio between its mean amplitude and mean time constant (1669 fF * 0.44/2.8 s = 262 fF/s), whereas the mean Ratedecay of the slow component of endocytosis was only ~41 fF/s (= 1669 fF * 0.56/23 s). In brief, these control experimental results were similar to previous reports (Wu et al., 2005; Wu et al., 2009).
We have previously shown that calcium influx triggers endocytosis and calmodulin blockers inhibited endocytosis (Wu et al., 2009). Consistent with this finding, replacing the extracellular calcium with barium, which barely activates calmodulin, also significantly inhibited endocytosis after 10 pulses of 20 ms depolarization at 10 Hz (n = 5, data not shown). To determine whether the calcium/calmodulin-activated calcineurin is involved in endocytosis, we measured endocytosis at 4 – 10 min after whole-cell break in with a pipette containing the calcineurin inhibitor cyclosporine A (CsA, 20 μM) or calcineurin auto-inhibitory peptide (CaN457-482, 150 μM) (Oliveria et al., 2007). We found that CsA and CaN457-482, reduced the Ratedecay after 1 or 10 depolarizing pulses to only ~24 - 32% of control (e.g., Fig. 1A-B, D-E, summarized in Fig. 1C, F). We did not quantify the time constant, because we often did not observe any fast component of endocytosis, and slow endocytosis was often nearly blocked completely, which made quantification of the time constant impossible. Thus, throughout the study, we did not measure the time constant when endocytosis was inhibited.
Since the Ratedecay after a 20 ms depolarization reflected slow endocytosis, whereas > 80% of the Ratedecay after the 10 pulse train was due to the rapid component of endocytosis, both calcineurin blockers significantly inhibited both slow and rapid endocytosis. The inhibition was not due to changes in calcium currents or exocytosis, because calcium currents did not change significantly, and ΔCm changed by < 20% (Supplementary Information 1). These results suggest the involvement of calcineurin in both rapid and slow endocytosis.
Rapid and slow endocytosis can be induced not only by depolarizing pulses of 20 ms, but also by trains of 1 ms depolarization that mimic action potential trains (Sun et al., 2002; Wu et al., 2005; Wu et al., 2009). For example, in the control condition with scrambled CaN457-482 (150 μM) in the pipette, 20 pulses of 1 ms depolarization to +7 mV at 200 Hz (AP-e), which mimicked a train of action potentials (Sun et al., 2002), induced a capacitance jump of 421 ± 16 fF (n = 8), followed by a mono-exponential decay with a time constant of 17.3 ± 1.2 s (n = 8) and a Ratedecay of 30 ± 1.6 fF/s (n = 8, Fig. 2A). After 200 AP-e at 200 Hz, the capacitance jump was 1331 ± 85 fF (n = 8), followed by a bi-exponential decay with time constants of 2.3 ± 0.3 s (46 ± 3%, n = 8) and 18.4 ± 1.6 s (n = 8), respectively (Fig. 2B). The Ratedecay after 200 AP-e was 252 ± 23 fF/s (n = 8, Fig. 2B). Thus, slow and rapid endocytosis induced by 20 and 200 AP-e at 200 Hz were similar to those induced by 1 and 10 pulses of 20 ms depolarization at 10 Hz, respectively. Compared to the Ratedecay in the presence of scrambled CaN457-482, CaN457-482 (150 μM in the pipette) significantly inhibited the Ratedecay to 28 ± 5% (n = 8, Fig. 2A, C) after 20 AP-e at 200 Hz, and to 34 ± 11% (n = 8, Fig. 2B, C) after 200 AP-e at 200 Hz (p < 0.01). These results suggest that calcineurin blockers inhibit endocytosis not only after trains of 20 ms depolarization, but also after trains of 1 ms depolarization that mimic action potential trains.
The calcineurin blocker specificity is often a concern that might discount the significance of pharmacological experiments. To address this issue, we used 7 – 10 days old mice lacking calcineurin Aα or Aβ subunit. Calcineurin is composed of a catalytic A and a regulatory B subunit. Among three isoforms of the A subunit, Aα and Aβ are expressed in the brain (Rusnak and Mertz, 2000). Aα-/- or Aβ-/- mice had been generated (Zhang et al., 1996; Bueno et al., 2002), from which we could not generate double knockout mice (Aα-/- Aβ-/-), likely because they die in the embryonic stage as observed in calcineurin B knockout (Chang et al., 2004).
In wild-type (WT) mice, the Ratedecay was 28 ± 4 fF/s (n = 10, Fig. 3A) and 157 ± 26 fF/s (n = 10, Fig. 3B) after 1 and 10 depolarizing pulses at 10 Hz, respectively. Similar to rat calyces (Wu et al., 2009), > 80% of the Ratedecay after the 10 pulse train was due to rapid endocytosis. Compared to WT mice, the Ratedecay after 1 (Fig. 3A) or 10 depolarizing pulses (Fig. 3B) was reduced by > 50% in Aα-/- mice (p < 0.01), but did not change significantly in Aβ-/- mice (p > 0.5). The Ratedecay reduction in Aα-/- mice was not due to changes in calcium currents or ΔCm (Supplementary Information 2). Thus, calcineurin Aα, but not Aβ subunit, is involved in mediating both rapid and slow endocytosis at calyces.
The calyx-type synapse is much larger than the conventional synapse. Whether our findings at calyces apply to conventional synapses is unclear. We addressed this issue at cultured hippocampal synapses by examining the roles of calcium, calmodulin, and calcineurin. SynaptopHluorin (SpH) was transfected to cultured rat hippocampal synapses (Sankaranarayanan and Ryan, 2000). Field electrical stimulation (20 mA, 1 ms) was applied to induce action potentials. In control, a 20 Hz stimulation train for 10 s (Train10s) caused exocytosis and thus a fluorescence increase (ΔFpeak) of 35 ± 5% of the baseline intensity (n = 7 experiments, each experiment contained ~10 - 30 boutons, Fig. 4A, left). The fluorescence increase was followed by a mono-exponential decay, owing to SpH endocytosis and vesicle re-acidification. The decay reflects mostly endocytosis, because endocytosis usually takes much longer than 10 s, whereas re-acidification takes only 3 - 4 s (Atluri and Ryan, 2006; Granseth et al., 2006). The rate of the initial fluorescence decay (Ratedecay) was 1.06 ± 0.18%/s (n = 7, fluorescence intensity normalized to baseline). The decay τ was 41.9 ± 2.4 s (n = 7, Fig. 4A, left). The fluorescence increase at 100 s after stimulation (ΔF100s) was -1 ± 11% (n = 7) of ΔFpeak, indicating completed endocytosis (Fig. 4A, left). Compared to Train10s, a 20 Hz train for 2 s (Train2s) induced a smaller ΔFpeak (16 ± 5% of the baseline), a smaller decay τ (20.9 ± 2.1 s), but only a slightly smaller Ratedecay (0.86 ± 0.09%/s), and a similar ΔF100s (-6 ± 7% of ΔFpeak, n = 6, Fig. 4A, right).
An early study showed that decreasing the extracellular calcium concentration ([Ca2+]o) to 0.75 mM or applying the calcium buffer EGTA-AM reduced the Ratedecay by several folds (Sankaranarayanan and Ryan, 2001). Given that the [Ca2+]o did not affect vesicle re-acidification, it was concluded that calcium influx regulates endocytosis. If calcium influx not only regulates endocytosis, but also initiates endocytosis, further reducing the [Ca2+]o should nearly abolish endocytosis as has been shown at calyces (Wu et al., 2009; Hosoi et al., 2009). Indeed, at 0.25 mM [Ca2+]o, Train10s induced a Ratedecay (0.20 ± 0.04%/s, n = 4) much smaller than that at 2 mM [Ca2+]o by Train10s or Train2s (p < 0.01), and induced a ΔF100s as large as 73 ± 7% (n = 4) of ΔFpeak (Fig. 4B). At 0.1 mM [Ca2+]o, Train10s could not induce a detectable ΔFpeak. However, a 10 s train at 100 Hz induced a ΔFpeak (22 ± 5%) between those induced by Train2s and Train10s at 2 mM [Ca2+]o, but a Ratedecay (0.07 ± 0.04%/s, Fig. 4C) 12 - 14 folds smaller than that induced by Train2s or Train10s at 2 mM [Ca2+]o, and a ΔF100s as large as 80 ± 10% of ΔFpeak (n = 5). At 2 mM [Ca2+]o, this 100 Hz train induced a much larger ΔFpeak (104 ± 5%), a Ratedecay (1.35 ± 0.06%/s) ~20 times higher than that at 0.1 mM [Ca2+]o, and a much smaller ΔF100s (14 ± 3% of ΔFpeak, n = 4, Fig. 4D). Clearly, decreasing the [Ca2+]o from 2 to 0.1 mM reduced the Ratedecay to nearly 0 (Fig. 4E), and significantly increased ΔF100s (Fig. 4F). These results suggest an essential role of calcium in controlling the rate of endocytosis, similar to results observed at the calyx of Held (Wu et al., 2009; Hosoi et al., 2009).
At 2 mM [Ca2+]o, as the ΔFpeak increased to ~16% (induced by Train2s), the Ratedecay increased to ~0.86%/s (Fig. 4G, solid square). Further increasing the ΔFpeak to ~104% (induced by the 100 Hz train), which was ~6.5 folds larger than that (16%) induced by Train2s, only increased the Ratedecay to 1.35%/s (Fig. 4G, solid triangle). Thus, the endocytosis capacity may be partially saturated at a ΔFpeak of ≥ 16% (Sankaranarayanan and Ryan, 2001; Balaji et al., 2008). The increase in the Ratedecay might be due to an increase of the ΔFpeak and/or an increase of the frequency of stimulation. However, the decrease of the Ratedecay at 0.1 - 0.25 mM [Ca2+]o was independent of either of these changes (Fig. 4G, comparing open and solid symbols). In particular, the Ratedecay at 0.1 - 0.25 mM [Ca2+]o (Fig. 4G, open symbols) was much smaller than that at 2 mM [Ca2+]o at similar ΔFpeak values (Fig. 4G, solid square and circle). These results suggest that the reduced calcium influx at low [Ca2+]o, but not the change in the amount of exocytosis, decreased the Ratedecay.
In the presence of a calmodulin blocker, calmidazolium (CMDZ, 10 μM in the bath, 5 - 10 min), the Ratedecay after Train10s (0.28 ± 0.09%/s, n = 7) was much smaller than that (0.86 - 1.06%/s) after Train10s or Train2s in control (p < 0.01), and the ΔF100s (79 ± 14% of ΔFpeak, n = 7) was much larger (Fig. 5A). The block of the SpH fluorescence decay was not due to inhibition of vesicle re-acidification (Supplementary Information 3). Thus, CMDZ inhibits endocytosis at hippocampal synapses.
The ΔFpeak induced by Train10s in the presence of CMDZ was smaller than that induced by Train10s in control, but larger than that induced by Train2s in control (Fig. 5A). The reduction of the ΔFpeak was not responsible for the decrease of the Ratedecay, because Train2s in control induced a smaller ΔFpeak, but a much larger Ratedecay than that induced by Train10s in the presence of CMDZ (Fig. 5A). The reduction of the ΔFpeak by CMDZ was consistent with the finding that calmodulin promotes vesicle mobilization from the reserve pool to the readily releasable pool (Sakaba and Neher, 2001), likely by initiating endocytosis that clears the released vesicle proteins from the release site (Wu et al., 2009).
CMDZ might not be specific to only calmodulin. To address this issue, we used a calmodulin shRNA that can knock down calmodulin expression by ~70% in cultured cortical neurons (Pang et al., 2010). Transfection of this shRNA to PC12 cells reduced calmodulin to 32 ± 6% (n = 6) of control (Supplementary Information 4). Co-transfection of calmodulin shRNA and SpH reduced calmodulin in the soma of rat hippocamal neurons to 30 ± 2% (n = 10 neurons from 3 transfections, p < 0.01) of that in neighbor un-transfected neurons (Fig. 5B, middle). In transfected neurons, Train10s induced a Ratedecay (0.38 ± 0.04%/s, n = 15) much slower than that (0.86 - 1.06%/s) induced by Train10s or Train2s in control (p < 0.01), and a much larger ΔF100s (61 ± 8% of ΔFpeak, n = 15, Fig. 5C), suggesting an inhibition of endocytosis similar to that caused by CMDZ. The ΔFpeak induced by Train10s was also slightly reduced as compared to the control (Fig. 5C), consistent with the effects of CMDZ in blocking vesicle mobilization to the readily releasable pool (Fig. 5A) (Sakaba and Neher, 2001).
The decrease of the calmodulin level in neurons co-transfected with calmodulin shRNA and SpH (Fig. 5B, middle) was not due to transfection of SpH. This was because transfection of SpH along did not affect the calmodulin level in the soma, as compared to the neighbor un-transfected neurons (103 ± 3%, n = 7 neurons, 2 transfections, p > 0.1, Fig. 5B, upper). In neurons co-transfected with SpH and a plasmid containing both calmodulin shRNA and shRNA-resistant calmodulin, calmodulin was over rescued to 163 ± 4% (n = 11 neurons from 3 transfections, p < 0.01) of that in un-transfected neurons (Fig. 5B, lower), and the Ratedecay (1.02 ± 0.06%/s), ΔF100s (-3 ± 4% of ΔFpeak) and ΔFpeak (36 ± 4%, n = 9) induced by Train10s were similar to control (p > 0.18, Fig. 5D). Transfection of this plasmid to PC12 cells also increased the calmodulin expression to 152 ± 5% of control (n = 3, Supplementary Information 4, see also Pang et al., 2010). These results suggest that inhibition of endocytosis by calmodulin shRNA was not due to off-target shRNA effects. We concluded that the physiological level of calmodulin is sufficient and critical in mediating normal endocytosis. This result, together with a recent finding that calmodulin may enhance the release probability by activation of CaMKII at hippocampal synapses (Pang et al., 2010), suggest that calmodulin is important not only for endocytosis, but also for exocytosis.
In the presence of the calcineurin blocker cyclosporin A (CsA, 20 μM in the bath, 5 - 10 min), Train10s induced a ΔFpeak (66 ± 8%, n = 13) nearly two times the control, but a Ratedecay (0.72 ± 0.14%/s, n = 13) smaller than the control (1.06 ± 0.18%/s, n = 7, p < 0.05), and a much larger ΔF100s (62 ± 8%, n = 13, Fig. 6A). The initial rate of endocytosis (Ratedecay) increases as the amount of exocytosis (ΔFpeak) increases (Balaji et al., 2008) until the latter reaches the endocytic capacity (Sankaranarayanan and Ryan, 2000; Wu and Betz, 1996; Sun et al., 2002) (see also Fig. 4G, solid symbols). Thus, an increase of the ΔFpeak by CsA might cause an increase of the Ratedecay, leading to an underestimate of the inhibition of Ratedecay by CsA. To examine this possibility, we divided the CsA experiments into two groups with ΔFpeak smaller or larger than 50% of the baseline. The reason we used 50% to divide the data was that the group with a smaller ΔFpeak had a ΔFpeak (40 ± 3%, n = 5) similar to that induced by Train10s in control. This group had about 7 folds smaller Ratedecay (0.16 ± 0.04%/s, p < 0.01), and a much larger ΔF100s (84 ± 13%; Fig. 6B, left, comparing solid and dotted traces). The group with a larger ΔFpeak had a mean ΔFpeak (82 ± 8%, n = 8) close to that induced by the 100 Hz train for 10 s in control (103 ± 5%, n = 4), but had a smaller Ratedecay (0.85 ± 0.06%/s, n = 8, p < 0.01) and a larger ΔF100s (48 ± 5%, n = 8, p < 0.01) as compared to that induced by the 100 Hz train in control (Ratedecay: 1.35 ± 0.06%/s; ΔF100s: 14 ± 3%; n = 4, Fig. 6B, right).
Clearly, CsA was more effective in blocking endocytosis at smaller ΔFpeak (Fig. 6B). Consistent with this result, a 4 s stimulation train at 20 Hz in the presence of CsA induced a ΔFpeak (23 ± 4%, n = 6) between those induced by Train10s and Train2s in control, but an ~3 - 4 fold smaller Ratedecay (0.26 ± 0.08%/s), and a much larger ΔF100s (63 ± 11%, Fig. 6C) than those induced by Train10s or Train2s in control. Large ΔFpeak may force the endocytic machinery to operate at near maximal capacity (Sankaranarayanan and Ryan, 2000), at which inhibition could be more difficult. These results, and the observation that CsA did not inhibit vesicle re-acidification (Supplementary Information 5), suggest that CsA significantly inhibited endocytosis.
The increase of ΔFpeak by CsA (Fig. 6A) could be due to a block of endocytosis and/or an increase of release. To distinguish these possibilities, a dynamin inhibitor, dynasore (100 μM) was applied to the bath for 5 - 10 min, which essentially blocked endocytosis after Train10s (Fig. 6D) (Newton et al., 2006). In this condition, Train10s induced a ΔFpeak (44 ± 4%, n = 12, Fig. 6D) higher than that (35 ± 5%, n = 7, p < 0.05) in control, but smaller than that (66 ± 8%, n = 13, p < 0.05, Fig. 6A) in the presence of CsA. These results suggest that CsA may also increase release, consistent with previous reports that block of calcineurin increases transmitter release by an as yet unidentified mechanism (Sihra et al., 1995; Lin and Lin-Shiau, 1999; Chi et al., 2003).
Next, we studied endocytosis in hippocampal cultures of calcineurin Aβ-/- or Aα-/- mice where the block of calcineurin function is more specific. In WT mice, Train10s induced a ΔFpeak of 36 ± 3%, a Ratedecay of 0.95 ± 0.05%/s, and a ΔF100s of 3 ± 7% (n = 4), which were nearly the same as those obtained in control rats (comparing the dotted trace in Fig. 7A & 6A). In Aβ-/- mice, Train10s induced a ΔFpeak (79 ± 8%, n = 21) much larger than the WT (p < 0.01, Fig. 7A), which was similar to the effects of CsA (Fig. 6A). Similar to the CsA experiments (Fig. 6B), we divided the data into two groups depending on whether the ΔFpeak was smaller or larger than 50% (Fig. 7B). The group with a smaller ΔFpeak had a ΔFpeak (40 ± 4%, n = 5) similar to that induced by Train10s in WT, but a ~3 fold smaller Ratedecay (0.33 ± 0.02%/s, n = 5, p < 0.01), and a much larger ΔF100s (66 ± 9%, n = 5, p < 0.01; Fig. 7B, left). The group with a larger ΔFpeak had a mean ΔFpeak (91 ± 8%, n = 16) close to that induced by the 100 Hz train for 10 s in WT (106 ± 8%, n = 8), but a Ratedecay (0.91 ± 0.05%/s, n = 16) smaller than that induced by the 100 Hz train in WT (1.48 ± 0.02%/s, n = 8, p < 0.01), and a much larger ΔF100s (Aβ-/-: 53 ± 6%, n = 16; WT: 23 ± 5%, n = 8, p < 0.01; Fig. 7B, right).
Similar to the effect of CsA, knockout of calcineurin Aβ was more effective in blocking endocytosis at smaller ΔFpeak (Fig. 7B). Consistent with this result, a 4 s stimulation train at 20 Hz in Aβ-/- mice induced a ΔFpeak (32 ± 3%, n = 5) similar to that induced by Train10s in WT, but an ~2 - 3 fold smaller Ratedecay (0.37 ± 0.03%/s), and a much larger ΔF100s (63 ± 9%, Fig. 7C).
In Aα-/- mice, Train10s induced a ΔFpeak of 36 ± 3% (n = 11), a Ratedecay of 1.02 ± 0.04%/s (n = 11) and a ΔF100s of 2 ± 1% (n = 11), all of which were similar to the WT (Fig. 7D). We concluded that calcineurin Aβ, but not Aα knockout inhibits endocytosis in a similar way as CsA at hippocampal synapses (Figs. 6 - -77).
Could the lack of effect of Aα knockout on endocytosis be due to the absence of calcineurin Aα subunit in the hippocampus? To examine this possibility, mouse hippocampal CA1-CA3 regions were dissociated for Western blot using two antibodies against calcineurin Aα and Aβ, respectively (Fig. 7E). Immunoblotting results revealed that Aα and Aβ were expressed in wild-type, but not in Aα-/- and Aβ-/- mice, respectively (Fig. 7E). Consistent with early studies (Kuno et al., 1992; Hashimoto et al., 1998), these results suggest that the lack of effect of Aα knockout on endocytosis is not due to the absence of Aα subunit in the hippocampus.
The present work provided the first genetic evidence together with pharmacological evidence suggesting an important role of calmodulin and calcineurin in rapid and slow endocytosis at 7 - 10 days old calyceal synapses and cultured hippocampal synapses (Figs. 1 - -3,3, ,55 - -7).7). Consistent with results obtained at calyces, where calcium influx triggers endocytosis (Wu et al., 2009; Hosoi et al., 2009), reducing the [Ca2+]o to 0.1 mM nearly abolished endocytosis at hippocampal synapses (Fig. 4). We therefore concluded that calcium influx during nerve firings activates calmodulin/calcineurin, which initiates and up-regulates slow, clathrin-dependent and rapid, presumably clathrin-independent endocytosis.
Calmodulin/calcineurin-dependent dephosphorylation of endocytic proteins (Cousin and Robinson, 2001; Robinson et al., 1993) may be synchronously activated by calcium influx during nerve firings, which may rapidly increase the endocytosis efficiency and thus initiate endocytosis. Since calcineurin is involved in rapid endocytosis, dephosphorylation must occur within tens to hundreds of milliseconds after stimulation. Larger calcium influx may speed up endocytosis (Wu, 2004) by inducing more calmodulin/calcineurin-dependent dephosphorylation.
How dephosphorylation initiates and accelerates endocytosis is unclear. Dynamin dephosphorylation promotes its interaction with syndapin (Anggono et al., 2006). It was suggested that calcium influx accelerates endocytosis by increasing the number of endocytic sites (Balaji et al., 2008). This suggestion was obtained by reducing the [Ca2+]o to only 1 mM. The near full block of endocytosis at 0.1 mM [Ca2+]o, as shown here (Fig. 4), suggests an extremely slow endocytosis at each endocytic site, although we could not fully exclude the possibility that few endocytic sites are assembled in low calcium conditions.
Our results seem inconsistent with the observation that endocytosis is triggered by calcium at a threshold (~10 μM) higher than the affinity (dissociation constant) of calcineurin to calcium (~1 μM) (Hosoi et al., 2009; Wu et al., 2009; Rusnak and Mertz, 2000). The affinity was measured in vitro with prolonged (minutes) presence of calcium and calcineurin in the steady-state (Rusnak and Mertz, 2000), whereas in nerve terminals, the calcium increase to > 10 μM decayed in < 1 s (Hosoi et al., 2009; Bollmann and Sakmann, 2005). The binding among calcium, calmodulin and calcineurin may not reach the steady-state during transient calcium influx, explaining why > 10 μM calcium is needed to initiate endocytosis. Furthermore, calcineurin (B subunit) has four calcium binding sites, one with a high affinity (< 0.1 μM), and three with affinities at ~15 μM (Rusnak and Mertz, 2000), the later of which may help to explain the need of > 10 μM calcium.
Our finding that calcium/calmodulin/calcineurin signaling pathway controls rapid and slow endocytosis may explain regulation of endocytosis by extra- and intracellular calcium observed at many synapses and endocrine cells over the last several decades (Ceccarelli and Hurlbut, 1980; Henkel and Betz, 1995; Ramaswami et al., 1994; Marks and McMahon, 1998; Cousin and Robinson, 1998; Gad et al., 1998; Sankaranarayanan and Ryan, 2001; Balaji et al., 2008; Wu et al., 2005; Neves et al., 2001; Artalejo et al., 1996). It may also explain why endocytosis is extremely slow in resting conditions (Wu et al., 2009; Hosoi et al., 2009). Since calcium/calmodulin may initiate bulk endocytosis at calyces (Wu et al., 2009), and calcium/calcineurin may trigger bulk endocytosis at cerebellar synapses (Clayton and Cousin, 2009; Clayton et al., 2009; Evans and Cousin, 2007), it is likely that the calcium/calmodulin/calcineurin signaling pathway is a common mechanism at synapses to initiate and regulate endocytosis, including rapid, slow, and bulk endocytosis.
Our results seem inconsistent with a report of no calcineurin involvement in slow, clathrin-dependent endocytosis during relatively mild stimulation at cerebellar synapses (Clayton et al., 2009). Although synapse heterogeneity provides an explanation, this discrepancy is likely due to methodological differences. The study at cerebellar synapses was based on the ability of a stimulus to unload FM dye from nerve terminals preloaded with the dye (Clayton et al., 2009). Instead of measuring endocytosis, this method measures the vesicle cycling involving both endocytosis and vesicle reuse. Since the dye was washed out immediately after the dye loading stimulus, the analysis (the amount of dye release after dye pre-loading) could not provide the endocytosis time course, distinguish between rapid and slow endocytosis, or measure endocytosis time course after stimulation (Clayton et al., 2009). In contrast, we quantitatively measured rapid and slow endocytosis time course using SpH imaging and capacitance measurement techniques. We used not only calcineurin blockers as in previous studies, but also calcineurin knockout mice and calmodulin knockdown techniques. Furthermore, our results were verified in two types of synapses, the hippocampal and the calyx-type synapse.
Our results seem inconsistent with the block of endocytosis by prolonged intracellular dialysis of ~1 μM calcium in ribbon-type synapses (Von Gersdorff and Matthews, 1994). Accordingly, our findings are likely limited to the transient calcium increase during brief depolarization. Prolonged calcium increase might perturb the cycle of phosphorylation and dephosphorylation, resulting in a block of endocytosis.
A study published after we finished the present work showed that the calcium buffer BAPTA abolished endocytosis in both the immature (P7 - 9) and more mature (P13 - 14) calyces (Yamashita et al., 2010), consistent with previous studies (Wu et al., 2009; Hosoi et al., 2009). This study also showed that calcineurin inhibitors (FK506 and CsA) inhibited rapid and slow endocytosis in P7 – 9 calyces (Yamashita et al., 2010), consistent with the present work. Surprisingly, calcineurin inhibitors did not block endocytosis in P13 - 14 calyces, suggesting that the calcium sensor for endocytosis changes developmentally from calcineurin to an unknown sensor (Yamashita et al., 2010). Accordingly, our results might be limited to immature synapses. However, this important suggestion may need further scrutiny for two reasons. First, it is based solely on pharmacological manipulation. Second, the same calcium influx triggers both exocytosis and endocytosis (Wu et al., 2009; Hosoi et al., 2009). Calcium channels are more tightly coupled to release in P13 -14 than P7 - 9 calyces, likely because calcium channels are located closer to the release site (Fedchyshyn and Wang, 2005; Wang et al., 2008; Kochubey et al., 2009; Yang et al., 2010). Tight coupling may produce a higher local calcium concentration during the same stimulus, which may accelerate endocytosis to a saturating speed (Wu et al., 2009). At such high concentration of calcium, the possibility that calcineurin blockers are not as effective in inhibiting endocytosis as in normal conditions has not been ruled out.
Rapid endocytosis is considered clathrin-independent (Artalejo et al., 1995; Jockusch et al., 2005). Its underlying mechanisms are poorly understood. The present work identified calcineurin as an important player in rapid endocytosis. Both rapid and slow endocytosis are regulated by the same calcium/calmodulin/calcineurin signaling pathway (Figs. 1 - -7)7) (Wu et al., 2009), and require dynamin in most, but not some stimulation conditions (Xu et al., 2008). Neither of them recycles vesicles to the readily releasable pool (Wu and Wu, 2009). These observations suggest that rapid and slow endocytosis share similar mechanisms of initiation, fission, and recycling. Rapid endocytosis is triggered by a higher calcium concentration (Wu et al., 2009; Beutner et al., 2001), likely because high calcium induces more calcineurin-dependent dephosphorylation.
Incomplete inhibition of endocytosis by calmodulin and calcineurin blockers, calmodulin knockdown, or calcineurin Aα or Aβ knockout (Figs. 1 - -3,3, ,55 - -7)7) is likely due to the inefficiency of blockers in vivo, the incomplete knockdown of calmodulin, or the remaining calcineurin A subunit. Although the involvement of other calcium-dependent pathway(s) could not be excluded, the calcium/calmodulin/calcineurin pathway must be a major signaling mechanism, because inhibition of calcineurin reduced the Ratedecay by up to ~4 - 7 folds (Figs. 1, ,66).
Knockout of calcineurin Aα and Aβ inhibited endocytosis at calyces and hippocampal synapses, respectively (Figs. 3, ,7).7). The reason for this difference is unclear. It is not because of the lack of Aβ at calyces and Aα in the hippocampus, because both isoforms are present in the hippocampus (Fig. 7E) (Kuno et al., 1992; Hashimoto et al., 1998). A difference in the relative abundance or sub-cellular localization of Aα and Aβ isoforms might provide an explanation.
CsA and calcineurin Aβ knockout increased ΔFpeak by enhancing transmitter release at hippocampal synapses (Figs. 6 - -7),7), whereas calcineurin inhibitors and calcineurin Aα knockout did not increase ΔCm at calyces (Figs. 1 - -3).3). The reason for this difference is unclear. Synapse heterogeneity could provide an explanation. The difference in the stimulation protocol might provide another explanation. If block of calcineurin increases the release probability, but not the readily releasable pool size, it might increase release during action potential stimulation at hippocampal synapses, but not the ΔCm induced by 20 ms depolarization that depleted the readily releasable pool at calyces (Wu and Wu, 2009).
Knockout of endocytosis genes often causes behavioral defect. Although we did not examine the behavior of Aα-/- and Aβ-/- mice, we noticed that we could not generate double knockout mice (Aα-/-, Aβ-/-) from Aα-/- and Aβ-/- mice, likely because they die in the embryonic stage. Consistent with this possibility, knockout of calcineurin B, the only calcineurin regulatory subunit, results in embryonic death (Chang et al., 2004). Furthermore, most Aα-/- mice die within a few months after birth owing to the heart failure (Molkentin et al., 1998). These results suggest the importance of calcineurin for animal survival.
This work was supported by the National Institute of Neurological Disorders and Stroke Intramural Research Program. We thank Drs. Jonathan G. Seidman (Harvard Medical School, Boston, MA 02114) and Jennifer L. Gooch (Emory University School of Medicine, Atlanta, GA 30322) for providing us with calcineurin Aα+/- mice. We thank Dr. Gero Miesenböck (University of Oxford, Oxford, UK) for providing us with the synaptopHluorin plasmid.