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GABAB receptor auxiliary subunits modulate Cav2.3-mediated release from medial habenula terminals

  1. Pradeep Bhandari
  2. David Vandael
  3. Diego Fernández-Fernández
  4. Thorsten Fritzius
  5. David Kleindienst
  6. Cihan Önal
  7. Jacqueline Montanaro
  8. Martin Gassmann
  9. Peter Jonas
  10. Akos Kulik
  11. Bernhard Bettler
  12. Ryuichi Shigemoto  Is a corresponding author
  13. Peter Koppensteiner  Is a corresponding author
  1. Institute of Science and Technology Austria (IST Austria), Austria
  2. Department of Biomedicine, University of Basel, Switzerland
  3. Institute of Physiology II, Faculty of Medicine, Germany
  4. BIOSS Centre for Biological Signalling Studies, University of Freiburg, Germany
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Cite this article as: eLife 2021;10:e68274 doi: 10.7554/eLife.68274

Abstract

The synaptic connection from medial habenula (MHb) to interpeduncular nucleus (IPN) is critical for emotion-related behaviors and uniquely expresses R-type Ca2+ channels (Cav2.3) and auxiliary GABAB receptor (GBR) subunits, the K+-channel tetramerization domain-containing proteins (KCTDs). Activation of GBRs facilitates or inhibits transmitter release from MHb terminals depending on the IPN subnucleus, but the role of KCTDs is unknown. We therefore examined the localization and function of Cav2.3, GBRs, and KCTDs in this pathway in mice. We show in heterologous cells that KCTD8 and KCTD12b directly bind to Cav2.3 and that KCTD8 potentiates Cav2.3 currents in the absence of GBRs. In the rostral IPN, KCTD8, KCTD12b, and Cav2.3 co-localize at the presynaptic active zone. Genetic deletion indicated a bidirectional modulation of Cav2.3-mediated release by these KCTDs with a compensatory increase of KCTD8 in the active zone in KCTD12b-deficient mice. The interaction of Cav2.3 with KCTDs therefore scales synaptic strength independent of GBR activation.

Introduction

The medial habenula (MHb) is an epithalamic structure that exclusively projects to the interpeduncular nucleus (IPN), with the dorsal MHb projecting to the lateral IPN and the ventral MHb projecting to the rostral and central subnuclei of the IPN (Figure 1A). This pathway is involved in various behaviors, including nicotine addiction and aversion (Agetsuma et al., 2010; Koppensteiner et al., 2016; Koppensteiner et al., 2017; Melani et al., 2019; Zhang et al., 2016; Zhao-Shea et al., 2013). A striking property of the MHb-IPN pathway is the prominent presynaptic localization of the R-type voltage-gated Ca2+ channel 2.3 (Cav2.3), a channel mainly located in postsynaptic elements in other brain areas (Parajuli et al., 2012). Furthermore, activation of presynaptic GABAB receptors (GBRs) on MHb terminals exerts an unusual facilitatory effect by increasing neurotransmitter release up to 10-fold (Zhang et al., 2016), and this effect appears to be involved in synaptic plasticity (Koppensteiner et al., 2017). Interestingly, this potentiation via presynaptic GBR activation only occurs in the glutamatergic/cholinergic pathway from the ventral MHb to the rostral/central IPN, whereas in the glutamatergic/substance-P-ergic pathway from the dorsal MHb to the lateral IPN, the release is attenuated by GBR activation (Melani et al., 2019). Although GBRs are ubiquitously expressed in these MHb-IPN pathways (Margeta-Mitrovic et al., 1999), the presynaptic localization and function of their auxiliary KCTD subunits is unknown.

Figure 1 with 1 supplement see all
Expression and function of GABAB receptors and Cav2.3 at two parallel MHb-IPN pathways.

(A) Schematic drawing of the two MHb-IPN pathways. In red: the dorsal part of the MHb projects to the lateral subnuclei of the IPN. In blue: the ventral part of the MHb projects to the rostral/central subnuclei of the IPN. (B) Confocal image of GABAB1 immunofluorescence signal indicates the presence of GABAB receptors (GBRs) in all IPN subnuclei. (C) In whole-cell recordings of rostral IPN neurons, activation of GBRs by baclofen (1 µM) produced a potentiation of electrically evoked EPSC amplitudes. Left: example EPSC traces before (black) and during the application of baclofen (red) and after washout of baclofen (blue); middle: example time course of EPSC amplitudes in one cell; right: averaged time course of relative EPSC amplitude change after baclofen (n = 13 cells/9 mice). (D) Baclofen reduced the amplitude of light-evoked glutamatergic EPSCs in lateral IPN neurons (n = 10 cells/5 mice). (E) Confocal image of Cav2.3 immunofluorescence signal indicates Cav2.3 presence in MHb axonal projections of both MHb-IPN pathways. (F) Pharmacological inhibition of Cav2.3 with SNX-482 in whole-cell recordings of rostral IPN neurons. Left: example traces before and after the application of SNX-482; middle: example time course of EPSC amplitude reduction by SNX-482; right: averaged time course of relative EPSC amplitude reduction by SNX-482. EPSC amplitudes were reduced by 83% on average (n = 9 cells/9 mice). (G) In Tac1-ChR2-EYFP mice, SNX-482 reduced light-evoked glutamatergic EPSC amplitudes on average by 52% (n = 8 cells/4 mice). (H, I) Left: Positions of recording and stimulating electrodes in acute thick-slice preparations from Cav2.3 KO mice. The stimulating electrode was placed on the fasciculus retroflexus just below the MHb, 2–3 mm from the recording sites. The position of the stimulating electrode and the stimulation intensity remained unchanged between rostral and lateral IPN recordings. Right: 10 Hz EPSC traces of all recorded neurons (9 cells/3 mice). (J) Cumulative EPSC amplitude plot shows significantly higher EPSC amplitudes in lateral compared to rostral IPN neurons. ***p<0.0001 two-way ANOVA; (K) FR stimulation and lateral IPN recording in 1 mm thick slice of wild-type mice. Sequential application of SNX-482, ω-conotoxin GVIA, and CdCl2. (L) Time course overlay of (F), (G), and (K) to compare SNX-482 time courses between minutes 0 and 25. For simplicity, application of ω-conotoxin GVIA starting from minute 15 in (K) not indicated in the graph. ***p<0.0001 two-way ANOVA with Tukey’s post hoc test. Scale bars in(B), (E), (H), and (I) are 100 µm. Averaged data is presented as mean ± SEM. See also Figure 1—figure supplement 1.

Figure 1—source data 1

Expression and function of GABAB receptors and Cav2.3 at two parallel MHb-IPN pathways.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig1-data1-v2.xlsx

Four KCTD subunits, KCTD8, 12, 12b, and 16, serve as GBR auxiliary subunits in the brain (Schwenk et al., 2010). KCTD subunits bind as hetero- and homo-pentamers to GBRs and modulate their signaling kinetics (Fritzius et al., 2017; Zheng et al., 2019). Structurally, KCTD subunits are composed of an N-terminal T1 domain and an H1 domain. The T1 domain binds to the GABAB2 subunit, while the H1 domain binds to the Gβ subunit of the guanine nucleotide-binding protein (G-protein) (Schwenk et al., 2010; Turecek et al., 2014; Zheng et al., 2019). X-ray crystallography of the T1 and H1 domains of KCTD12 and KCTD16 (Lepore et al., 2019; Zheng et al., 2019; Zuo et al., 2019) revealed that the secondary and tertiary protein structures of those domains are highly conserved in the KCTD proteins. However, the T1 domain possesses a highly divergent loop between helix α2 and strand β3 (following the nomenclature from Zuo et al., 2019), while the H1 domain features a divergent loop between the strands β4 and β5 (following the nomenclature from Zheng et al., 2019). These loops are hydrophilic, exposed at the protein surface, and potentially allow the different KCTD subunits to bind proteins other than GABAB2 or Gβ. KCTD8 and 16, but not KCTD12 and 12b, additionally encode a C-terminal H2 domain. The absence of the H2 domain enables KCTD12 and 12b to rapidly desensitize GBR responses by uncoupling the Gβγ subunits of the G-protein from effector channels (Turecek et al., 2014). KCTD12 and 12b can form hetero-pentamers with each other but also with KCTD8 and 16 (Fritzius et al., 2017). Compared to the rest of the brain, the expression patterns of KCTD subunits in the MHb are unique: With the exception of a weak expression in the cerebellum and superior colliculus, KCTD8 is exclusively and strongly expressed in MHb and, to a lesser extent, IPN neurons. Furthermore, KCTD12b is exclusively expressed in the ventral part of the MHb (Metz et al., 2011). In contrast, KCTD12 is weakly expressed in the ventral part of the MHb, while KCTD16 is expressed in most brain areas but not the MHb. Based on proteomics studies, voltage-gated Ca2+ channels co-precipitate GBRs and their auxiliary KCTD subunits together with release machinery proteins of the presynaptic active zone, including SNAP-25, synaptotagmins, synaptobrevin-2, Munc13-1, syntaxins, RIM1, and synapsins, among others (Müller et al., 2010). Furthermore, KCTD8 and KCTD16 were consistently found to co-purify with presynaptic Cav2.2 Ca2+ channels (Müller et al., 2010; Schwenk et al., 2016). However, the functional consequences of these interactions and whether other voltage-gated Ca2+ channels interact with KCTDs remain unknown.

Here, we studied the nano-anatomy of Cav2.3, GBRs, and KCTDs and their roles in the modulation of neurotransmission from the MHb to the IPN. Our results demonstrate that Cav2.3 is located in the presynaptic active zone of habenular terminals and required for fast neurotransmitter release. In heterologous cells, we found that KCTD8 and KCTD12b, but not KCTD12, bind to Cav2.3 at the plasma membrane. Furthermore, Cav2.3 currents were enhanced by co-expression of KCTD8, but not KCTD12b in the absence of GBRs. Strikingly, genetic deletion of KCTD8 or KCTD12b reduced or increased, respectively, the probability of neurotransmitter release in the ventral MHb to the rostral IPN pathway without affecting GBR-mediated potentiation. The increase of release in KCTD12b knock-out (KO) mice was accompanied by a compensatory recruitment of KCTD8 into the active zone, where it closely associates with Cav2.3. Furthermore, viral expression of KCTD12b in MHb neurons reduced the increased release probability in KCTD12b KO mice, whereas overexpression of KCTD8 in MHb neurons of wild-type mice increased release probability. These results support that synaptic strength at the MHb-IPN pathway is scaled via GBR-independent Cav2.3–KCTD interactions in the presynaptic active zone.

Results

Cav2.3 mediates neurotransmission in two distinct MHb–IPN pathways

The MHb to IPN pathway comprises two major projections (Figure 1A). The dorsal MHb projects to the lateral subnuclei of the IPN and releases glutamate and substance P, whereas the ventral MHb projects to the rostral and central IPN subnuclei and co-releases glutamate and acetylcholine (Aizawa et al., 2012; Melani et al., 2019; Molas et al., 2017; Ren et al., 2011). In confocal light microscopy, immunofluorescence signal for GBRs was detected in all IPN subnuclei (Figure 1B). GBR activation has been reported to facilitate both electrically and optogenetically evoked neurotransmitter release in the ventral MHb-rostral/central IPN pathway (Koppensteiner et al., 2017; Zhang et al., 2016), whereas it inhibits release in the dorsal MHb-lateral IPN (Melani et al., 2019). We confirmed that rostral IPN neurons exhibit a strong increase in excitatory postsynaptic current (EPSC) amplitudes following GBR activation with 1 µM baclofen (Figure 1C), whereas EPSC amplitudes were reduced by baclofen in lateral IPN neurons (Figure 1D). Both the rostral/central and lateral subnuclei showed prominent Cav2.3 immunofluorescence signals (Figure 1E), in accordance with a previous study showing strong and exclusive presynaptic Cav2.3 localization in the IPN (Parajuli et al., 2012). A previous report tested the functional involvement of Cav2.3 in neurotransmission from the ventral MHb to the IPN by applying Ni2+ (Zhang et al., 2016), an ion known to also inhibit other Ca2+ channels (Lee et al., 1999). To test Cav2.3 dependency of neurotransmission in both pathways, we performed whole-cell recordings from IPN neurons in acute brain slices and applied the Cav2.3 blocker SNX-482 (Newcomb et al., 1998). In rostral and lateral IPN neurons, 1 µM SNX-482 strongly reduced the amplitude of EPSCs by 83% and 52%, respectively (Figure 1F,G). The SNX-482-mediated reduction in EPSC amplitudes was significantly stronger in the ventral MHb to rostral IPN pathway compared to the dorsal MHb to lateral IPN pathway (main effect of IPN subnucleus F1, 395 = 213.7, p<0.0001; two-way ANOVA). In addition, SNX-482 did not significantly affect action potential peak times or full-widths at half maximum in ventral MHb neurons (Figure 1—figure supplement 1A; peak times: control: 0.86 ± 0.03 ms, n = 3; SNX-482: 0.93 ± 0.05 ms, n = 4; t5 = 1.083, p=0.3282; full-widths: control 1.08 ± 0.07 ms, n = 3; SNX-482: 1.06 ± 0.03 ms, n = 4, t5 = 0.3852, p=0.7159; unpaired t-test), suggesting that the reduction of neurotransmitter release by SNX-482 was not caused by a blockade of voltage-gated Na+ or K+ channels. To corroborate the involvement of Cav2.3 in release from MHb terminals, we next performed electrical stimulation of the MHb-derived fiber tract, the fasciculus retroflexus (FR), in Cav2.3 KO mice and recorded EPSCs in rostral and lateral IPN neurons. To obtain maximally intact FR, slices were cut at 1 mm thickness, which allowed for the stimulation of the fiber tract at a distance of 2–3 mm from the IPN (Figure 1H,I). The MHb is a ventricular structure and therefore well supplied with freshly oxygenated ACSF, even in thick slices, assuring long-lasting viability of intact axons and their terminals within the IPN. In wild-type (WT) animals, electrical stimulation of the FR in slices prepared this way produced reliable responses in all recorded rostral and lateral IPN neurons (Figure 1—figure supplement 1B,D). Pathway specificity was confirmed by baclofen, which enhanced responses in the rostral but depressed responses in the lateral IPN (Figure 1—figure supplement 1C–D). In order to compare responses between rostral and lateral IPN recordings in Cav2.3 KO mice, the position of the stimulating electrode and the stimulus intensity remained unchanged between recordings. From each KO animal, three neurons each from the rostral and lateral IPN were recorded in an alternating manner (Figure 1H,I). Responses were approved when (1) evoked events were observed in response to at least six consecutive stimuli in the train, (2) the amplitude of evoked responses exceeded three times the standard deviation of the baseline noise, and (3) responses showed the characteristic shapes of a fast rising and slowly decaying synaptic response. Applying these criteria, 10 Hz electrical stimulation for 1 s failed to evoke synaptic responses in the nine of nine recorded rostral IPN neurons (Figure 1H,J). In contrast, seven of nine recorded lateral IPN neurons showed EPSC responses (Figure 1I,J; main effect of IPN subnucleus: F1, 160 = 120.4, p<0.0001; two-way ANOVA).

To investigate the identity of additional Ca2+ channels involved in release from dorsal MHb terminals, we performed FR stimulation to measure EPSCs in lateral IPN neurons (seven recordings/five mice) in WT mice and sequentially applied SNX-482, the Cav2.2 blocker ω-conotoxin GVIA (1 µM) and CdCl2 (200 µM) (Figure 1K). While SNX-482 produced a similar decrease in EPSC amplitude as in the optogenetic stimulation experiment (Figure 1K,L, effect of electrical vs. optogenetic stimulation: F1, 305 = 1.830, p=0.1772; main effect of SNX: F23, 305 = 17.41, p<0.0001; Interaction F23, 305 = 1.842, p>0.9999, two-way ANOVA of time courses between minutes 0–25), ω-conotoxin did not reduce EPSC amplitudes further. The remaining EPSC amplitude was completely blocked by CdCl2, suggesting an involvement of either P/Q- or L-type Ca2+ channels. Comparison of the SNX-482 effects on electrically and optogenetically evoked EPSCs in the lateral IPN with that on electrically evoked EPSCs in the rostral IPN confirmed a significantly weaker effect in dorsal MHb terminals in the lateral IPN than in ventral MHb terminals in the rostral IPN (Figure 1L; electrical stim. rostral vs. electrical stim. lateral: p<0.0001; electrical stim. rostral vs. optogenetic stim. lateral: p<0.0001; electrical stim. lateral vs. optogenetic stim. lateral p=0.3174; two-way ANOVA with Tukey’s post hoc test). Overall, these results suggest that ventral MHb terminals in the rostral IPN rely exclusively on Cav2.3 for release, whereas dorsal MHb terminals in the lateral IPN rely on Cav2.3 and additional Ca2+ channels.

KCTD8 and KCTD12b directly bind Cav2.3 and KCTD8 enhances currents through Cav2.3 in HEK cells

GBRs strongly modulate release in both pathways. Therefore, we next investigated the expression of GBR auxiliary subunits, KCTDs, in the IPN. Immunoreactivity for KCTD8 appeared strong in all IPN subnuclei (Figure 2A), consistent with its strong expression throughout MHb and presynaptic localization in IPN. KCTD12 immunofluorescence appeared strong in the rostral and central subnuclei but faint in the lateral subnucleus (Figure 2B). In contrast, immunoreactivity for KCTD12b was only present in the rostral and central but not the lateral IPN subnuclei, indicating its absence in the dorsal MHb-IPN pathway (Figure 2C). While KCTD12 immunofluorescence patterns in the rostral region suggested a mostly postsynaptic expression, KCTD12b signal showed the characteristic pattern of cholinergic MHb axons inside the rostral/central IPN, suggesting presynaptic expression. Antibody specificity for all KCTD antibodies was confirmed using the corresponding KO animals (Figure 2A–C), while the specificity of the anti-Cav2.3 and anti-GABAB1 antibodies has been confirmed previously (Kulik et al., 2002; Parajuli et al., 2012).

Figure 2 with 1 supplement see all
KCTD subtype expression in the IPN and interaction of Cav2.3 with KCTD8 and KCTD12b in vitro.

(A–C) Confocal images of immunofluorescence signals of KCTD8, KCTD12, and KCTD12b in the IPN of WT (upper panels) and respective KO mice (lower panels). KCTD8 immunofluorescence was present in all IPN subnuclei, whereas KCTD12 and KCTD12b signals were observed only in the rostral/central but not the lateral IPN subnuclei. Scale bars: 100 µm. (D) Co-immunoprecipitation from total cell lysates of HEK293T cells transfected with Flag-tagged KCTDs and Cav2.3. Immunoprecipitation of Cav2.3 co-precipitated KCTD8 and KCTD12b, but not KCTD12. Input lanes (bottom) indicate expression of the tagged proteins in the cell lysates. (E) KCTDs are co-localized and interact with Cav2.3 at the cell surface of HEK293T cells. The three input lanes to the right show expression of Flag-tagged KCTD8 (top) and Flag-tagged KCTD12b (bottom) in the cytosol, the total membrane fraction (‘total membrane’) and the plasma membrane fraction, from left to right. The two IP lanes to the left show that immunoprecipitation of Cav2.3 in the total membrane fraction (‘total membranes’) and the plasma membrane fraction co-precipitated KCTD8 (top) and KCTD12b (bottom), from left to right. Membrane-bound Cav2.3 (bottom lanes) is expressed in the total membrane fraction and the plasma membrane fraction, but absent from the cytosol fraction. (F) Whole-cell recordings from HEK293 cells stably expressing Cav2.3. Ba2+ current densities measured in response to a single depolarizing voltage step from −80 to 10 mV were significantly increased in KCTD8 co-transfected cells. *p<0.05, **p<0.01 one-way ANOVA with Tukey post hoc test. (G) Current density-to-voltage relationship demonstrating higher current densities in KCTD8-transfected cells compared with Control- and KCTD12b-transfected cells. ****p<0.0001 two-way ANOVA with Tukey post hoc test. (H) Activation and inactivation curves in Control-, KCTD8-, and KCTD12b-transfected cells. **p<0.01, two-way ANOVA with Tukey post hoc test. See also Figure 2—figure supplement 1.

Figure 2—source data 1

KCTD subtype expression in the IPN and interaction of Cav2.3 with KCTD8 and KCTD12b in vitro.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig2-data1-v2.xlsx

A previous study positioned presynaptic Ca2+ channels at the center of large, macromolecular complexes that also contained GBRs and KCTDs (Müller et al., 2010), and some KCTDs were found to co-purify with Cav2.2 subunit of N-type Ca2+ channels in the absence of GBRs (Schwenk et al., 2016). To test whether presynaptic KCTDs in MHb terminals may directly interact with Cav2.3, we performed co-immunoprecipitation (co-IP) experiments in HEK293T cells transiently expressing Cav2.3 and either KCTD8, KCTD12, or KCTD12b. Interestingly, we found selective binding of Cav2.3 to KCTD8 and KCTD12b, but not to KCTD12 (Figure 2D). Plasma membrane preparations (see Materials and methods) revealed that in HEK293T cells co-expressing Cav2.3 and KCTDs, both KCTD8 and KCTD12b are highly and similarly enriched in plasma membrane extracts, when compared to total membrane extracts (Figure 2—figure supplement 1; KCTD8: 10.7-fold [±1.9], KCTD12b: 12.7-fold [±2.6], no significant difference between KCTD8 and KCTD12b, p=0.34, unpaired t-test). We next performed co-IP experiments using total and plasma membrane extracts of transfected HEK293T cells and found that both KCTD8 and KCTD12b associate with Cav2.3 at the plasma membrane (Figure 2E). These results suggest that KCTD8 and KCTD12b in MHb-derived axon terminals may directly interact with presynaptic Cav2.3, even in the absence of GBRs.

Using a cell line stably expressing human Cav2.3 (Dai et al., 2008), we next co-expressed either an empty vector (Control), KCTD8, or KCTD12b, and measured Ba2+ currents through Cav2.3. Current densities resulting from a single voltage step (–80 mV to 10 mV) were significantly increased in cells co-expressing KCTD8 compared with control- and KCTD12b-transfected cells (Figure 2F, F2, 37 = 5.614; p=0.0074; Control vs. KCTD8: p=0.0086; Control vs. KCTD12b: p=0.8959; KCTD8 vs. KCTD12b: p=0.0382; one-way ANOVA with Tukey post hoc test). The current density-to-voltage relationship showed a significant difference between KCTD8 and both Control- and KCTD12b-transfected cells (Figure 2G; main effect of transfection: F2, 406 = 29.23, p<0.0001; Control vs. KCTD8: p<0.0001; Control vs. KCTD12b: p=0.5793; KCTD8 vs. KCTD12b: p<0.0001; two-way ANOVA with Tukey post hoc test; see also Supplementary file 1). Furthermore, there was a slight hyperpolarization of the steady-state inactivation in the KCTD8-expressing cells compared with Control (Figure 2H; main effect of transfection: F2, 252 = 4.831, p=0.0087; Control vs. KCTD8: p=0.0059; Control vs. KCTD12b: p=0.2667; two-way ANOVA with Tukey post hoc test), whereas the voltage dependence of channel activation was not different between groups (Figure 2H; F2, 201 = 2.303; p=0.1025, two-way ANOVA). These results suggest that the interaction of Cav2.3 with KCTD8, but not KCTD12b, increases the current through Cav2.3 in heterologous cells.

Pre-embedding and freeze-fracture replica immunolabeling of presynaptic Cav2.3, GABAB1, and KCTDs in habenular terminals in rostral and lateral IPN subnuclei

To obtain insight into the nanoscale distribution of presynaptic molecules, we investigated the electron microscopic localization of Cav2.3, GABAB1, and KCTDs in ventral and dorsal MHb terminals. We performed pre-embedding immunogold labeling and quantified the number of silver-intensified particles in the active zone, peri-synaptic region (0–50 nm distance from the edge of the active zone), and extrasynaptic regions (50–100, 100–150, and 150–200 nm distance from the edge of the active zone; Figure 3, Figure 3—figure supplement 1). Immunogold particles for Cav2.3 were concentrated in the active zone and, to a lesser extent, the immediate peri-synaptic region (Figure 3A). In accordance with our immunohistochemical data (Figure 2B,C), KCTD12 and KCTD12b labeling was detected only in MHb terminals in the rostral but not lateral IPN subnuclei (Figure 3D,E). In addition, particles for KCTD12 were found mostly on the postsynaptic side of asymmetrical synapses and only few synapses exhibited presynaptic KCTD12 labeling along synaptic and extrasynaptic membrane (Figure 3D, Figure 3—figure supplement 2). In MHb terminals in the rostral IPN, Cav2.3 and KCTD12b showed similar localization patterns, with peak particle densities in the active zone and a gradual decrease in density with increased distance from the active zone (Figure 3A,E). In contrast, GABAB1, KCTD8, and KCTD12 showed peak localization in the peri-synaptic region with lower particle densities inside the active zone (Figure 3B–D). These results suggest that KCTD12b prominently localized to the active zone of ventral MHb terminals in the rostral IPN, whereas KCTD8 dominates the active zone of dorsal MHb terminals in the lateral IPN (Figure 3F).

Figure 3 with 2 supplements see all
Quantification of the presynaptic localization of Cav2.3, GBRs, and KCTDs along the ventral and dorsal MHb-IPN pathways.

Transmission electron microscopy images of 70 nm thick sections following pre-embedding immunolabeled IPN slices for Cav2.3 (A), GABAB1 (B), KCTD8 (C), KCTD12 (D), and KCTD12b (E) from synapses in the rostral (left images) and lateral (right image) IPN subnuclei. Scale bars: 200 nm. Graph on the right displays quantification of relative and absolute silver-enhanced gold particle densities in the active zone and at distances of 50–200 nm from the edge of the active zone (50 nm bins). (F) Absolute labeling densities are summarized for synapses in the rostral (left) and lateral IPN (right). Note the absence of KCTD12 and KCTD12b particles in presynaptic terminals inside the lateral IPN subnuclei. KCTD12 was not included in panel F because of predominantly postsynaptic localization inside the rostral IPN. Data was pooled from two animals, showing no significant difference in gold particle distribution patterns with Kolmogorov-Smirnov test (see Figure 3—figure supplement 1 and Figure 3—figure supplement 2).

Figure 3—source data 1

Quantification of the presynaptic localization of Cav2.3, GBRs, and KCTDs along the ventral and dorsal MHb-IPN pathways.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig3-data1-v2.xlsx

To circumvent potential antigen-masking effects due to the protein-dense active zone region in conventional pre-embedding immunolabeling, we performed SDS-digested freeze-fracture replica labeling (SDS-FRL) (Fujimoto, 1995). This method enables unhindered access of antibodies to proteins inside/close to the pre- or postsynaptic membrane specialization and allows for multiple labeling with gold particles of distinct sizes (Hagiwara et al., 2005; Indriati et al., 2013; Miki et al., 2017; Tanaka et al., 2005). In order to distinguish IPN subnuclei, we used an improved version of the grid-glued SDS-FRL method (Harada and Shigemoto, 2016), which facilitates the preservation of complete replicas during the handling procedures, a critical requirement for the identification of IPN subnuclei in the electron microscope (Figure 4A). We detected gold particles for Cav2.3 on the P-face of the presynaptic active zone and confirmed antibody specificity using Cav2.3 KO mice (Figure 4B,C,F; average density WT: 147.8 ± 23.9 particles/µm², n = 4 replicas; KO: 3.4 ± 0.99 particles/µm², n = 4 replicas; WT vs. KO: t3 = 5.87, p=0.0099, paired t-test). In addition, we confirmed the concentrated localization of Cav2.3 within the active zone using co-immunolabelings with a mixture of antibodies against marker proteins for the active zone, RIM1/2, neurexin, and CAST (Figure 4D; Miki et al., 2017). In the absence of marker protein labeling, demarcation of the presynaptic active zone was based on multiple criteria, including P-face curvature and intramembrane particle (IMP) size and density. The area of active zones demarcated without marker protein labeling (AZunmarked) was not significantly different from that with marker labeling (AZunmarked: 0.080 ± 0.0064 µm², n = 46 active zones, AZmarked: 0.077 ± 0.0040 µm², n = 80 active zones; p=0.8692, Kolmogorov–Smirnov test) or from the area of postsynaptic IMP clusters on the E-face, the replica equivalent of the postsynaptic density seen in conventional ultrathin sections (IMP cluster: 0.077 ± 0.0059 µm², n = 68 active zones; p>0.9999, Kolmogorov–Smirnov test), verifying our criteria for active zone demarcation.

SDS-digested freeze-fracture replica labeling confirms Cav2.3 in the active zone of medial habenula terminals in the IPN.

(A) Example image of a grid-glued replica containing the whole IPN. White dashed line indicates demarcation of rostral/central and lateral subnuclei. Scale bar: 20 µm. (B) Example image of a presynaptic P-face and a postsynaptic E-face of a habenular synapse in the rostral IPN that was double labeled with antibodies against AMPA receptors (10 nm gold) and Cav2.3 (5 nm gold). Scale bar: 100 nm. (C) Example image of a similar synaptic profile double labeled with antibodies against AMPA receptors (10 nm gold) and Cav2.3 (5 nm gold) in the rostral IPN of a Cav2.3 KO mouse. Scale bar: 100 nm. (D) Left: double labeling of a WT carbon-only replica with antibodies against Cav2.3 (5 nm gold) and a mixture of active zone proteins (2 nm gold), including RIM1/2, CAST, and neurexin. Right: the same image with additional coloring of 2 nm (red) and 5 nm (blue) particles and demarcation of the active zone area based on active zone-marker labeling. Scale bars: 100 nm. (F) Left: quantification of Cav2.3 labeling densities in the presynaptic P-face in WT and Cav2.3 KO mice. ***p<0.001, unpaired t-test. Right: areas of demarcated active zones, including incomplete profiles, were not significantly different between replicas from WT and Cav2.3 KO mice. Data were obtained from four replicas from four mice of each genotype. (G) Comparison of active zone area demarcated with or without active zone markers and the size of the glutamatergic postsynaptic IMP clusters. p-value indicates result of Kolmogorov–Smirnov test.

Figure 4—source data 1

SDS-digested freeze-fracture replica labeling confirms Cav2.3 in the active zone of medial habenula terminals in the IPN.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig4-data1-v2.xlsx

The result of our pre-embedding immunolabeling suggested that Cav2.3, GBR, and KCTDs are localized in and around the active zone of MHb terminals (Figure 3). In order to confirm that these molecules are actually co-localized inside the same terminals, we performed double immunolabelings for Cav2.3 and either GABAB1, KCTD8, or KCTD12b in SDS-FRL (Figure 5). We focused on these main presynaptic KCTDs because immunogold labeling for KCTD12 was observed in few presynaptic terminals (Figure 3D) and was located mostly postsynaptically in the rostral IPN (Figure 3—figure supplement 2). We found that in ventral MHb terminals in the rostral IPN, Cav2.3 is co-localized with GABAB1, KCTD8, and KCTD12b in over 97% of all Cav2.3-positive active zones (Figure 5A,B; GABAB1: 98 ± 0.70%, n = 3 replicas; KCTD8: 98 ± 1.10%, n = 3 replicas; KCTD12b: 97 ± 1.8%, n = 4 replicas). Similar co-localization patterns were seen in dorsal MHb terminals located inside the lateral IPN (Figure 5A,B; GABAB1: 99 ± 0.69%, n = 3 replicas; KCTD8: 97 ± 0.57%, n = 3 replicas), with the exception of an absence of KCTD12b. Particle numbers and densities of all tested molecules, except KCTD12b, were comparable between MHb terminals in the rostral and lateral IPN (Figure 5B). In addition, the nearest-neighbor distances (NND) of all tested molecules were significantly smaller than those obtained from simulations of randomly distributed particles (Figure 5C), suggesting that Cav2.3, GABAB1, and KCTDs are clustered inside the active zone.

Co-localization of Cav2.3 with GBR and KCTDs in the active zone of medial habenula terminals.

(A) Active zones double labeled for Cav2.3 and either GABAB1 (left), KCTD8 (middle), or KCTD12b (right) in IPN replicas. Top row images are from presynaptic terminals in the rostral IPN; bottom row images are from presynaptic terminals in the lateral IPN. Scale bar: 100 nm. (B) Quantification of active zone immunolabeling in the rostral and lateral IPN. With the exception of the absence of KCTD12b in lateral IPN terminals, absolute particle numbers per active zone (left graph) and particle densities (middle graph) are comparable between MHb terminals in the rostral and lateral IPN. Right graph: Over 97% of active zones positive for Cav2.3 labeling also show labeling for one of the other molecules (GABAB1, KCTD8, or KCTD12b), suggesting co-localization of all presynaptic molecules inside the same active zone. Numbers inside the bars indicate the number of replicas used for each quantification. (C) Nearest-neighbor distance (NND) for all presynaptic molecules in MHb terminals inside the rostral and lateral IPN based on the real (black line) and simulated random distribution (blue line). Smaller NND values in real distributions compared to simulation suggest clustering of all presynaptic molecules. p-values calculated via Kolmogorov–Smirnov test.

Figure 5—source data 1

Co-localization of Cav2.3 with GBR and KCTDs in the active zone of medial habenula terminals.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig5-data1-v2.xlsx

KCTDs modulate release probability at ventral MHb terminals

Based on the exclusive requirement of Cav2.3 for release and the selective expression of KCTD12b co-localized with Cav2.3 in the rostral/central but not lateral IPN, we tested the impact of different KCTDs on neurotransmitter release from ventral MHb terminals in the rostral IPN. We measured the paired-pulse ratio (PPR) of consecutive electrically evoked EPSCs at 20 Hz in WT and KCTD KO mice (Figure 6A). PPR was significantly lower in KCTD12b KO mice (1.22 ± 0.11, n = 24 cells) compared with WT (2.40 ± 0.31, n = 27 cells; p=0.0051, Kruskal–Wallis test), KCTD8 KO (2.26 ± 0.27, n = 17 cells; p=0.0080), and KCTD8/12b KO mice (2.76 ± 0.37, n = 11 cells; p=0.0010). In contrast to these changes in basal release, application of 1 µM baclofen still potentiated EPSC amplitudes in all KCTD KO lines with similar intensities (Figure 6—figure supplement 1; WT: 615.1 ± 148.9% of baseline; n = 13; KCTD8 KO: 470.8 ± 73.5%; n = 9; KCTD12b KO: 485.4 ± 146.6%; n = 12; KCTD8/12b double KO: 445.8 ± 76.2%; n = 10; F4, 51 = 0.4723, p=0.7558, one-way ANOVA), suggesting that KCTDs are not involved in the GBR-mediated enhancement of EPSC amplitudes.

Figure 6 with 1 supplement see all
KCTDs modulate release probability of ventral MHb terminals.

(A) In whole-cell recordings from rostral IPN neurons, paired-pulse ratios (PPR) of electrically evoked glutamatergic excitatory postsynaptic currents (EPSCs) were significantly lower in KCTD12b KO mice compared with WT, KCTD8 KO, and KCTD8/12b double KO mice; **p<0.01, ***p<0.001, Kruskal–Wallis with Dunn’s post hoc test. (B–D) Example variance-mean measurements of electrically evoked EPSC amplitudes at varying external Ca2+ concentrations in recorded in rostral IPN neurons of WT (B), KCTD8 KO (C), and KCTD12b KO mice (D). (E) Quantification of release probability (left graph) and quantal size (right graph) indicates significantly reduced release probability in KCTD8 KO mice and increased release probability in KCTD12b KO mice. Quantal size remained unaffected. *p<0.05, ***p<0.001, ****p<0.0001 one-way ANOVA with Tukey’s post hoc test. (F) WT animals were injected with lentivirus expressing either EGFP (WT LV-EGFP) or KCTD8/EGFP (WT LV-KCTD8) into the MHb. Left: Example images of acute IPN slices after electrophysiological recordings showing EGFP signal in fibers inside the IPN. Right: Overexpression of KCTD8 resulted in significantly lower PPR values compared with EGFP controls **p<0.01, unpaired t-test. (G) Left: Lentiviral expression of EGFP or KCTD12b/EGFP in acute slices of KCTD12b KO mice. Right: Lentiviral expression of KCTD12b significantly increased PPR values compared with EGFP controls **p<0.01, unpaired t-test. See also Figure 6—figure supplement 1.

Figure 6—source data 1

KCTDs modulate release probability of ventral MHb terminals.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig6-data1-v2.xlsx

PPR values are generally thought to be inversely correlated with release probability (Dobrunz and Stevens, 1997). Therefore, our findings suggest that basal release in KCTD12b KO mice is higher than in WT and KCTD8 KO mice and that this effect is KCTD8 dependent. To confirm the increase in release probability in KCTD12b KO mice, we performed variance-mean analysis (Figure 6B–E) to estimate the values of release probability (at 2.5 mM external Ca2+) and quantal size (Figure 6EClements and Silver, 2000). Compared with WT mice (WT release probability: 0.26 ± 0.04, n = 16 cells), the release probability from ventral MHb terminals of KCTD12b KO mice (0.49 ± 0.05, n = 9 cells) was significantly increased (Figure 6E; F2, 34 = 21.23, p<0.0001; WT vs. KCTD12b KO: p=0.0005, one-way ANOVA with Tukey post hoc test). In contrast, the release probability in KCTD8 KO mice (0.12 ± 0.02, n = 12 cells) was significantly lower than that of WT or KCTD12b KO mice (WT vs. KCTD8 KO: p=0.0176; KCTD8 KO vs. KCTD12b KO: p<0.0001; Tukey post hoc test). The quantal size remained unaffected by genotype (F2, 34 = 1.613, p=0.2142).

To test the effect of overexpression of KCTD8 on basal release from ventral MHb terminals, we injected lentivirus expressing either EGFP or KCTD8-p2A-EGFP bilaterally into the MHb of WT mice (Figure 6F). PPR values recorded in rostral IPN neurons of EGFP control-injected animals remained unchanged (LV-EGFP: 2.17 ± 0.18, n = 17, five mice), whereas KCTD8-injected animals exhibited significantly lower PPR values (LV-KCTD8: 1.41 ± 0.14, n = 13, three mice; t28 = 3.20, p=0.0034; unpaired t-test). In contrast, lentiviral expression of KCTD12b in KCTD12b KO mice significantly increased PPR values compared with EGFP control-injected mice (Figure 6G; KCTD12b KO EGFP Control: 1.38 ± 0.09, n = 11, three mice; PPR KCTD12b KO rescue: 2.18 ± 0.29, n = 14, three mice; t23 = 2.98, p=0.0068, unpaired t-test), resulting in PPR values not significantly different from those of WT mice (WT LV-EGFP vs. KCTD12b rescue: t29 = 0.40, p=0.6925, unpaired t-test), confirming that the reduced PPR in KCTD12b KO mice is due to the lack of KCTD12b and not caused by developmental alterations. Overall, these results show that KCTDs differentially modulate basal release probability at ventral MHb terminals. The increase in Cav2.3-mediated release probability by KCTD8 is consistent with its facilitatory effects on Cav2.3 current density in HEK293 cells. The decreased PPR in the absence of KCTD12b recovered to control levels when KCTD8 was additionally absent, suggesting alterations in either Cav2.3 or KCTD8 localization in the active zone of MHb terminals in KCTD12b KO mice. Thus, we next investigated nano-anatomical changes associated with the increased release probability in KCTD12b KO mice.

Absence of KCTD12b induces a compensatory increase of KCTD8 in the active zone of ventral MHb terminals

To quantify the density of Cav2.3 and KCTD8 in ventral MHb terminals in the rostral IPN, we performed SDS-FRL and co-immunolabeled Cav2.3/GABAB1 or Cav2.3/KCTD8 in replicas of WT and KCTD12b KO IPN samples (Figure 7A). Since KCTD12b is not expressed in MHb terminals in the lateral IPN, no effects on expression of these molecules are expected in the lateral IPN in KCTD12b KO mice. Thus, we normalized the densities of presynaptic molecules in the rostral IPN to the average density of the same molecule in the corresponding lateral IPN in the same replicas. Thereby, we avoided interference by technical variabilities in labeling efficiencies between individual replicas. We found no significant difference in the densities of Cav2.3 in the presynaptic active zone in the rostral relative to the lateral IPN (Figure 7B; WT rostral: 1.01 ± 0.04 fold of lateral IPN, n = 8 replicas from eight mice; KCTD12b KO: 1.06 ± 0.13 fold of lateral IPN, n = 8 replicas from eight mice; p=0.5054, Mann–Whitney test). Interestingly, the relative density of KCTD8 in the active zone of MHb terminals was increased approximately twofold in KCTD12b KO mice compared with those of WT mice (Figure 7B; WT rostral: 0.84 ± 0.12 fold of lateral IPN, n = 5 replicas from five mice; KCTD12b KO: 2.09 ± 0.45 fold of lateral IPN, n = 5 replicas from five mice; p=0.0079, Mann–Whitney test). This result indicates that the increased release probability in KCTD12b KO mice could be ascribable to a compensatory increase of KCTD8 in the active zone resulting in enhanced association of KCTD8 with Cav2.3. On the other hand, GABAB1 expression in the presynaptic active zone of MHb terminals was not significantly different between WT and KCTD12b KO (Figure 7B; WT rostral: 1.30 ± 0.18 fold of lateral IPN, n = 3 replicas from three mice; KCTD12b KO: 0.89 ± 0.07 fold of lateral IPN, n = 3 replicas from three mice; p=0.4000, Mann–Whitney test). To confirm the compensatory increase of KCTD8 in ventral MHb terminals in the absence of KCTD12b, we next performed pre-embedding immunolabeling for KCTD8 in presynaptic terminals in rostral and lateral IPN of KCTD12b KO mice (Figure 7C). The distribution pattern of KCTD8 in the lateral IPN showed peak densities in the peri-synaptic region, as seen in WT (Figure 3C). In contrast, the density of KCTD8 particles in the active zone, but not in the peri- or extrasynaptic area of rostral IPN synapses was significantly increased compared with lateral IPN synapses (main effect of presynaptic distribution: F4, 775 = 31.65, p<0.0001; rostral active zone vs. lateral active zone: p<0.0001; two-way ANOVA with Bonferroni post hoc test), showing an altered distribution pattern (compare with Figure 3C,F) and similar to that of Cav2.3 and KCTD12b in WT animals (Figure 3A,E,F). This observation confirmed that KCTD8 compensates for the loss of KCTD12b in the active zone of ventral MHb terminals.

Absence of KCTD12b leads to a compensatory increase of KCTD8 inside the active zone of ventral MHb terminals.

(A) Example images of active zones containing Cav2.3 and either GABAB1 (left) or KCTD8 (right) in replicas of WT (upper row) and KCTD12b KO IPN tissue (lower row). Scale bars: 100 nm. (B) Quantification of relative densities for Cav2.3, KCTD8, and GABAB1 in active zones located in the rostral IPN of WT and KCTD12b KO mice. Densities were normalized to the average density in MHb terminals inside the lateral IPN of the same replica. The number inside the bars indicates the number of replicas used for quantification. **p<0.01 in a Mann–Whitney test. (C) Pre-embedding EM labeling for KCTD8 in KCTD12b KO mice (data from three mice). The density of KCTD8-labeled gold particles was significantly higher in the AZ of ventral MHb terminals in the rostral IPN compared with dorsal MHb terminals in the lateral IPN. ***p<0.0001 two-way ANOVA with Bonferroni post hoc test. (D, E) Comparison of pre-embedding EM labeling for Cav2.3 in MHb terminals in KCTD12b KO mice of control sections (standard labeling, four mice) with sections pre-incubated with anti-KCTD8 primary antibody and biotinylated secondary antibody (three mice). Pre-incubation resulted in significant reduction in labeling densities for Cav2.3 selectively in the active zone of ventral MHb terminals in the rostral IPN as well as in the peri-synaptic region of dorsal MHb terminals in the lateral IPN. ***p<0.001, *p<0.05 two-way ANOVA with Bonferroni post hoc test.

Figure 7—source data 1

Absence of KCTD12b leads to a compensatory increase of KCTD8 inside the active zone of ventral MHb terminals.

https://cdn.elifesciences.org/articles/68274/elife-68274-fig7-data1-v2.xlsx

In order to examine further the spatial proximity of KCTD8 to Cav2.3 in the active zone, we tested whether antibodies bound to KCTD8 might interfere with the labeling for Cav2.3 by steric hindrance in KCTD12b KO mice. We hypothesized that the compensatory increase in KCTD8 in the active zone might lead to increased binding of KCTD8 to Cav2.3 and, therefore, pre-incubation with anti-KCTD8 antibody might reduce Cav2.3 labeling in the active zone. In rostral IPN synapses, we found that the pre-incubation significantly reduced Cav2.3 labeling densities in the active zone, but not the peri- or extrasynaptic regions (Figure 7D; main effect of synaptic distribution: F4, 880 = 31.42, p<0.0001; control active zone vs. pre-incubation active zone: p=0.0006; two-way ANOVA with Bonferroni post hoc test). In the lateral IPN, pre-incubation slightly but significantly reduced Cav2.3 labeling density only in the peri-synaptic region (Figure 7E; main effect of synaptic distribution: F4, 890 = 39.01; control <50 nm vs. pre-incubation <50 nm: p=0.0320). The peri-synaptic region is the location exhibiting peak expression for KCTD8 in dorsal MHb terminals. The significant reduction in Cav2.3 labeling density at this position therefore validates our steric hindrance approach. These results support that KCTDs in the active zone dynamically regulate the release probability, which may be important for scaling synaptic strength independent from GBRs in the MHb-IPN pathway.

In summary, the active zone of ventral MHb terminals predominantly contains Cav2.3 and KCTD12b, whereas the peri-synaptic region exhibits high levels of KCTD8 and GBRs (Figure 8, WT terminal). In the absence of KCTD12b, KCTD8 enters the active zone to interact with Cav2.3, thereby increasing Ca2+ influx and ultimately release (Figure 8, KCTD12b KO terminal).

Figure 8 with 1 supplement see all
Summary of presynaptic localization of Cav2.3, KCTDs, and GBRs in ventral MHb neurons of WT and KCTD12b KO mice.

Schematic representation of the distribution and function of Cav2.3, GBRs, and KCTDs in ventral MHb terminals of WT and KCTD12b KO mice. Left: In WT terminals, the active zone contains Cav2.3 and hetero-pentameric rings comprising KCTD12b in excess over KCTD8, whereas KCTD8 and GBRs are located peri-synaptically. Right: In absence of KCTD12b, KCTD8 invades the active zone and compensates for the loss of KCTD12b, resulting in increased release probability, potentially via increased Ca2+ influx. See Figure 8—figure supplement 1.

Discussion

We used a combination of anatomical, biochemical, and functional methods to study the role of KCTDs at two parallel pathways from the MHb to the IPN and identified an unexpected function of KCTDs in the modulation of neurotransmitter release. We confirmed that Cav2.3 is involved in mediating transmitter release at both pathways, but only release at the ventral MHb-IPN pathway exclusively relies on Cav2.3. KCTD8 and KCTD12b show distinct expression along these pathways, with KCTD12b specifically expressed in the cholinergic ventral MHb-IPN pathway. In HEK293T cells, we found that KCTD8 and KCTD12b bind to Cav2.3 at the plasma membrane and that specifically KCTD8 increases Cav2.3 currents. Using electron microscopy, we verified that Cav2.3 and KCTDs co-localize within the presynaptic active zone of MHb terminals. Genetic deletion of the KCTDs indicated that KCTD8 increases basal release probability and that KCTD12b prevents this effect. Synapses lacking KCTD12b showed an increased density of KCTD8 in the active zone, resulting in a close association of KCTD8 with Cav2.3 and a consequent increase in release probability. Viral expression of KCTD12b in KCTD12b KO mice reduced release probability to WT levels, whereas overexpression of KCTD8 in WT mice mimicked the phenotype of KCTD12b KO mice. Our results support that the KCTDs scale synaptic strength at the MHb-IPN pathway independent of GBRs.

Dominant presynaptic localization and function of Cav2.3 in MHb-IPN pathway

In contrast to other brain areas, R-type Ca2+ channels may be the exclusive source of presynaptic Ca2+ influx to trigger release from ventral MHb terminals, given that neither P/Q- nor N-type Ca2+ channels are expressed in MHb neurons (Ludwig et al., 1997). The finding by Ludwig et al., 1997 fits the data of the Allen Brain Atlas, which shows no expression of Cav2.1 and Cav2.2 in MHb neurons (Figure 8—figure supplement 1; images from Allen Institute for Brain Science, 2004) (Lein et al., 2007). Although the predominant presynaptic localization of Cav2.3 immunoreactivity was previously reported (Parajuli et al., 2012), the functional implications of this localization were not determined. Our data confirmed that both MHb-IPN pathways heavily rely on Cav2.3 for release. Interestingly, projections from the ventral MHb to the rostral IPN were more affected by SNX-482 than the dorsal MHb to lateral IPN pathway. Using Cav2.3 KO mice, we found residual EPSC responses in lateral but not rostral IPN neurons, confirming that exclusively Cav2.3 mediates transmitter release from ventral MHb terminals. The remaining release from dorsal MHb terminals in the lateral IPN of Cav2.3 KO mice could be explained either by a compensatory upregulation of other Ca2+ channels or by the presence of additional Ca2+ channels in WT mice. In dorsal MHb terminals of WT mice, the SNX-482-insensitive EPSC component was also insensitive to ω-conotoxin GVIA, ruling out N-type Ca2+ channels, but was completely abolished by CdCl2. Therefore, dorsal MHb terminals likely express Cav2.3 together with other Ca2+ channels, potentially L-type Ca2+ channels.

Role of KCTDs in potentiation of Cav2.3-mediated release by GBR activation

A previous report of the potentiating action of presynaptic GBRs on cholinergic MHb terminals identified Cav2.3 as a critical mediator of this effect, based on the observations that both genetic ablation of Cav2.3 and pharmacological inhibition of Cav2.3 with Ni+ prevented the potentiation (Zhang et al., 2016). Given the modulatory roles of KCTDs in shaping GBR effector responses (Fritzius et al., 2017), we initially hypothesized that the unique repertoire of KCTDs in ventral MHb neurons may be involved in the facilitatory action of GBRs. However, deletion of neither KCTD8, KCTD12b, nor both prevented the potentiation of EPSC amplitudes by baclofen, suggesting a KCTD-independent effect.

Regulation of Cav2.3-mediated release by KCTDs

In addition to the co-localization of Cav2.3 with KCTDs at the active zone in habenular terminals, we identified a hitherto unknown direct interaction of Cav2.3 with KCTD8 and KCTD12b in vitro. Previous proteomics studies revealed co-precipitations of N-type/Cav2.2 Ca2+ channels with KCTD8 and KCTD16, but no other KCTD−Cav2 interactions were reported (Müller et al., 2010; Schwenk et al., 2016). Possibly, the use of total brain extracts may have limited detection to the most prominent protein interactions. Thus, brain-wide approaches to study presynaptic Ca2+ channel-interacting proteins may have failed to detect interactions uniquely occurring inside MHb terminals. In contrast, our pre-embedding EM approach utilizing interference in Cav2.3 immunogold labeling by steric hindrance with KCTD8 antibody in the AZ of MHb terminals (Figure 7D) indicates a close association in situ of these molecules within 16 nm (two times of IgG size 8 nm), if not a direct binding.

The selective KO of KCTD8 lowered basal levels of release probability from ventral MHb terminals below that of WT synapses. On the other hand, KO of KCTD12b resulted in a significant increase of basal release probabilities. This finding of modulation of release independent of GBR activation could be explained by the current-enhancing effects of KCTD8 on Cav2.3 observed in a heterologous expression system. Since the probability of neurotransmitter release is relatively low in WT terminals (0.26), the additional reduction in release probability in KCTD8 KO mice (0.12) may not be detectable by PPR measures. An important indication that KCTD8 functionally increases synaptic strength was the observation that viral overexpression of KCTD8 in WT ventral MHb neurons significantly lowered PPR values compared with EGFP controls. This result suggests that KCTD8 and KCTD12b may be in a state of dynamic balance. Creating an imbalance appears to result in alterations in synaptic strength. This possibility would be in line with our pre-embedding EM result, with KCTD12b showing peak densities inside the active zone of ventral MHb terminals, whereas KCTD8 shows peak expression in the peri-synaptic region. This could be an indication for the preferred interaction of synaptic Cav2.3 with KCTD12b over KCTD8 due to spatial proximity or higher binding affinity. Although the increase in release probability at ventral MHb terminals is likely caused by a direct effect of KCTD8 on Cav2.3, it is unknown whether other Ca2+ channels involved in transmitter release from dorsal MHb terminals are equally affected by KCTD8.

It is unclear whether a compensatory increase of KCTD12b occurs in KCTD8 KO mice. Our initial results from WT pre-embedding EM indicate that KCTD12b displays peak densities in the active zone, whereas KCTD8 shows peak densities in the peri-synaptic region. Therefore, although it may be possible to see a compensatory increase in KCTD12b in the active zone of KCTD8 KO mice, due to the weaker expression of KCTD8 in the WT active zone, such an increase would be expected to be smaller than the compensatory increase seen in KCTD12b KO animals. Furthermore, the functional consequence of a compensatory increase in KCTD12b in the active zone in KCTD8 KO mice would not be detected in our functional experiments, as KCTD12b did not alter Cav2.3 function in vitro and KCTD12b has no effects on PPR in the absence of KCTD8 (no significant difference between KCTD8 KO and KCTD8/12b double KO in Figure 6A). Our results suggest that the relatively mild reduction in release probability seen in KCTD8 KO mice resulted from removal of the comparatively low levels of KCTD8 from the WT active zone, rather than from a compensatory increase in KCTD12b.

To induce synaptic plasticity in the ventral MHb to IPN pathway, release of glutamate, activation of postsynaptic Ca2+-permeable AMPA receptors on GABAergic IPN neurons, and subsequent retrograde release of GABA to activate presynaptic GABAB receptors on MHb terminals are required (Koppensteiner et al., 2017). Therefore, KCTD-mediated modulation of release probability could determine a ventral MHb synapse’s propensity to undergo and express activity-dependent plasticity. Future studies may provide additional insights into whether physiological learning paradigms affecting the MHb-IPN pathway, such as the formation and extinction of aversive memories (Agetsuma et al., 2010; Koppensteiner et al., 2017; Melani et al., 2019; Zhang et al., 2016), could alter the ratio of KCTD8 to 12b.

Overall, our study provided new insights into the physiological role of presynaptic Cav2.3, GBRs, and their auxiliary KCTD subunits in an evolutionary conserved neuronal circuit. It remains to be determined whether the prominent presence of presynaptic KCTDs at other synapses (Müller et al., 2010) could be an indication of similar neuromodulatory function of KCTDs in different pathways of the brain.

Materials and methods

Key resources table
Reagent type
(species) or
resource
DesignationSource or
reference
IdentifiersAdditional
information
Strain, strain background (Mus musculus, ♂)C57BL/6JThe Jackson Laboratory#000664
Strain, strain background (Mus musculus, ♂)BALB/cJThe Jackson Laboratory#000651
Strain, strain background (Mus musculus, ♂)Cav2.3 KO (gene: Cacna1e)Prof. Tsutomu Tanabe, (Saegusa et al., 2000)
Strain, strain background (Mus musculus, ♂)KCTD8 KO (gene: Kctd8)Prof. Bernhard Bettler, (Schwenk et al., 2010)
Strain, strain background (Mus musculus, ♂)KCTD12 KO (gene: Kctd12)Prof. Bernhard Bettler, (Schwenk et al., 2010)
Strain, strain background (Mus musculus, ♂)KCTD12b KO (gene: Kctd12b)Prof. Bernhard Bettler, (Schwenk et al., 2010)
Strain, strain background (Mus musculus, ♂)KCTD8/12b double KO (genes: Kctd8/Kctd12 b)This paper
Strain, strain background (Mus musculus, ♂♀)Tac1-IRES-Cre (B6;129S-Tac1tm1.1(cre)Hze/J)The Jackson Laboratory#021877
Strain, strain background (Mus musculus, ♂♀)Ai32 (B6;129S-Gt(ROSA)26Sortm32(CAG-COP4*H134R/EYFP)Hze/J)The Jackson Laboratory#012569
Strain, strain background (Mus musculus, ♂)Tac1-ChR2-EYFPOffspring of Tac1-IRES-Cre and Ai32 mice
Genetic reagent (virus)LV–hSyn–3xFlag/mKCTD8[NM_175519.5]/P2A/EGFPVectorBuilder GmbHCustom-made
Genetic reagent (virus)LV–hSyn–3xFlag/mKCTD12b[NM_175429.4]/P2A/EGFPVectorBuilder GmbHCustom-made
Genetic reagent (virus)LV–hSyn–EGFPVectorBuilder GmbHCustom-made
Cell line (human)HEK293TATCC, (Seddik et al., 2012)https://web.expasy.org/cellosaurus/CVCL_0063
Cell line (human)HEK293 cells stably expressing α1E-3Prof. David J. Adams, (Berecki et al., 2014)
AntibodyGuinea pig polyclonal anti-Cav2.3Genovac, (Parajuli et al., 2012)Custom-made antibody, 1 µg/ml for IHC, 8 µg/ml for SDS-FRL
AntibodyRabbit polyclonal anti-GABAB1Prof. Akos Kulik, (Kulik et al., 2002)Custom-made antibody, final concentration: (1 µg/ml) for IHC, (2 µg/ml) for SDS-FRL
Antibodyrabbit polyclonal anti-KCTD8Prof. Bernhard Bettler, (Schwenk et al., 2010)Custom-made antibody, final concentration: (1 µg/ml) for IHC, (4 µg/ml) for SDS-FRL
AntibodyRabbit polyclonal anti-KCTD12Prof. Bernhard Bettler, (Schwenk et al., 2010)Custom-made antibody, final concentration: (1 µg/ml) for IHC, (4 µg/ml) for SDS-FRL
AntibodyRabbit polyclonal anti-KCTD12bProf. Bernhard Bettler, (Schwenk et al., 2010)Custom-made antibody, final concentration: (1 µg/ml) for IHC, (4 µg/ml) for SDS-FRL
AntibodyRabbit polyclonal anti-RIM1/2Synaptic Systems140 203final concentration: (5 µg/ml) for SDS-FRL
AntibodyRabbit polyclonal anti-CASTProf. Watanabe, (Hagiwara et al., 2018)final concentration: (3 µg/ml) for SDS-FRL
AntibodyRabbit polyclonal anti-NeurexinProf. Watanabe, (Miki et al., 2017)final concentration: (5 µg/ml) for SDS-FRL
Peptide, recombinant proteinSNX-482hellobioHB1235
Peptide, recombinant proteinω-Conotoxin GVIAAlomone Labs106375-28-4
commercial assay or kitPlasma membrane extraction kitAbcamab 65400
Chemical compound, drugR(+)-Baclofen hydrochlorideMerckG013
Chemical compound, drug1(S),9(R)-(−)-Bicucullin-MethiodidMerck14343
Chemical compound, drugHexamethonium bromideTocris4111
Chemical compound, drugMecamylamine hydrochlorideTocris2843
Chemical compound, drugCadmiumchlorideMerck202908
Software, algorithmGraphpad Prism 8Graphpadhttps://www.graphpad.com/scientific-software/prism/
Software, algorithmMATLABMathWorkshttps://www.mathworks.com/products/matlab.html?s_tid=hp_products_matlab
Software, algorithmReconstructJohn C. Fiala, Ph.D.https://synapseweb.clm.utexas.edu/software-0

Animals

Wild-type C57BL/6J (Jax, Bar Harbor, ME; #000664) and BALB/cJ (Jax, #000651) mice were initially purchased from Jackson Laboratory. Homozygous KCTD KO lines were generated by the lab of Bernhard Bettler at the University of Basel (Schwenk et al., 2010). For KCTD8 KO line generation, exon one containing ATG and most of the open reading frame (ORF) of the Kctd8 gene (MGI:2443804) was replaced with a loxP-flanked neo. To generate KCTD12 KO mice, 5′ part of exon one containing complete ORF of the Kctd12 gene (MGI:2145823) was replaced with a loxP-flanked neo (Cathomas et al., 2015). Similarly, KCTD12b KO line was generated by replacing the 5′ part of exon three containing complete ORF of the Kctd12b gene (MGI:2444667) with a loxP-flanked neo. All KCTD KO lines had the neo removed by crossing founders with a Cre-deleter line, leaving one loxP site behind. The KCTD8/12d double KO line was generated by mating F2 hybrids of the parental lines. Background strains of KCTD KO lines were as follows: KCTD8 (C57BL/6J and 129), KCTD12 (C57BL/6J, 129, BALB/cJ), KCTD12b (BALB/cJ), and KCTD8/12b (C57BL/6J, 129, BALB/cJ). Cav2.3 KO mice were generated by the lab of Tsutomu Tanabe (Saegusa et al., 2000). To obtain Tachykinin1 (Tac1)-ChR2-EYFP mice, we crossed Tac1-Cre (Jax, #021877) with Ai32 (Jax, #012569) mice. All mice were bred at the preclinical facility of IST Austria on a 12:12 light–dark cycle with access to food and water ad libitum. All experiments were performed in accordance with the license approved by the Austrian Federal Ministry of Science and Research (Animal license number: BMWFW-66.018/0012-WF/V/3b/2016) and the Austrian and EU animal laws. Only male mice aged 2–5 months were used for all experiments.

Transcardial perfusion for brain fixation

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Mice were anaesthetized with a mixture of ketamine (90 mg/kg body weight) xylazine (4.5 mg/kg) solution intraperitoneally, and 25 mM ice-cold phosphate-buffered saline (PBS) was transcardially perfused through the left ventricle at a flow rate of 7 ml/min for 30–60 s. Subsequently, paraformaldehyde (PFA) solution was perfused for 12 min. PFA solutions of different concentrations were used for confocal imaging (4% PFA [TAAB Laboratories Equipment Ltd., Aldermaston, UK] and 15% picric acid in 0.1 M phosphate buffer, PB), pre-embedding (4% PFA and 15% picric acid in 0.1 M PB+0.05% glutaraldehyde [TAAB]), and SDS-digested freeze-fracture replica labeling (SDS-FRL, 2% PFA and 15% picric acid in 0.1 M PB). The pH of all PFA solutions was adjusted to 7.4 ± 0.05 with HCl. After perfusion, the brain was excised and placed in 0.1 M PB and sectioned within 3 days. Slices of different thickness (50 µm for confocal microscopy and pre-embedding, 70 µm for SDS-FRL) were cut with a vibratome (Linear-Pro7, Dosaka, Japan) in ice-cold 0.1 M PB.

Immunohistochemistry

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Brain slices containing the IPN were washed with phosphate-buffered saline (PBS) and subsequently incubated in blocking buffer (10% normal goat serum, 2% BSA, 0.5% Triton-X100 in 0.1 M PBS) for 1 hr prior to incubation with primary antibodies: guinea pig anti-Cav2.3 (1 µg/ml, two overnight [O/N], Genovac), rabbit anti-GABAB1 (B17, 1 µg/µl, 1 O/N [Kulik et al., 2002]), rabbit anti-KCTD8 (1 µg/µl, Bettler lab, Univ. Basel), rabbit anti-KCTD12 (1 µg/µl, 1 O/N, Bettler lab, Univ. Basel), and rabbit anti-KCTD12b (polyclonal, raised against a synthetic peptide comprised of the N-terminal amino acids 1–16 of KCTD12b) (1 µg/µl, 1 O/N, Bettler lab, Univ. Basel) (Metz et al., 2011; Schwenk et al., 2010). Brain sections were washed in PBS and subsequently incubated for 1 hr in secondary antibody (1:500, Alexa-488 anti-guinea pig [Molecular Probes, Eugene, OR] or Alexa-488 anti-rabbit [Molecular Probes]). Sections were mounted onto glass slides, and images were taken with an LSM 800 (Zeiss, Oberkochen, Germany) confocal microscope.

Pre-embedding immunolabeling

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Brain slices were washed in 0.1 M PB (two times, 10 min each) and cryo-protected by incubation in 0.1 M PB containing 20% sucrose O/N at 4°C. The next day, slices underwent three cycles of freeze-thawing by freezing the slices on liquid nitrogen for 1 min and thawing them in 20% sucrose on a hot plate (50°C) for 2 min. Slices were washed in 0.1 M PB (10 min) followed by washing in TBS (three times, 20 min each). Free aldehydes were quenched by incubating slices in 50 mM glycine (Sigma–Aldrich, St. Louis, MO) in TBS (10 min). After washing in TBS (three times, 20 min each), slices were blocked with blocking buffer (10% NGS, 2% BSA in TBS, 1 hr). Primary antibody incubation was done with respective antibodies in 2% BSA solution for 2 O/N at 4°C. The concentration of antibodies was 1 µg/ml for Cav2.3, KCTD8, KCTD12, KCTD12b, and GABAB1. For steric hindrance experiments, slices were pre-incubated with 2 µg/ml of anti-KCTD8 antibody (2 O/N), followed by incubation with biotinylated anti-rabbit secondary antibody (1:100, 2 O/N) before incubation with anti-Cav2.3 antibody (2 O/N). Subsequently, slices were rinsed in TBS (three times, 20 min each) and incubated in respective secondary antibodies (1:100) O/N at 4°C in 2% BSA in TBS. For Cav2.3, 1.4 nm gold-conjugated anti-guinea pig antibody (Nanoprobes, Yaphank, NY) and for all other antibodies, i.e. GABAB1, KCTD8, KCTD12, and KCTD12b, 1.4 nm gold-conjugated anti-rabbit antibody (Nanoprobes) was used. Slices were washed in TBS and PBS (two times, 20 min each) followed by post-fixation in 1% glutaraldehyde in PBS (10 min), washing in PBS (three times, 10 min), and quenching of free glutaraldehyde in 50 mM glycine in PBS (10 min). Finally, slices were washed in PBS (three times, 10 min each) and Milli-Q (MQ) H2O (three times, 5 min each).

For silver intensification of immunogold particles, Nanoprobes silver intensification (Nanoprobes) component A (initiator) and B (moderator) were mixed and vortexed, followed by the addition of component C (activator). After vortexing, slices were incubated in the mixture for 9 min 15 s in the dark. Tubes were tapped every 2 min for uniform intensification. Slices were washed with MQ water (three times, 10 min each) and in 0.1 M PB (10 min), followed by post-fixation in 1% OsO4 in 0.1 M PB (20 min in the dark). After osmification, slices were washed in 0.1 M PB (10 min) and in MQ water (three times, 5 min each) and counter-stained in 1% uranyl acetate (Al-labortechnik, Zeillern, Germany) in MQ H2O (35 min in the dark). Subsequently, slices were serially dehydrated in ethanol solutions of different concentrations in ascending order up to 100% (50–95% ethanol in five steps, 5 min each; 100% ethanol two times, 10 min each) and then washed with propylene oxide (Sigma–Aldrich; two times, 10 min each). Slices were then submerged in Durcupan resin (Sigma –Aldrich; mixture of components A, B, C, and D in proportion of 10:10:0.3:0.3, respectively) for 1 O/N at room temperature (RT).

For flat embedding, each slice was isolated on a silicon-coated glass slide, covered with an ACLAR fluoropolymer film (Science Services, Munich, Germany) and incubated at 37°C (1 hr) followed by incubation at 60°C (2 O/N). For re-embedding, tissue containing the rostral or lateral IPN was excised with a surgical blade, placed into the lid of a plastic tube (TAAB), which was then filled with Durcupan resin and incubated at 60°C (2 O/N). Each resin block was trimmed using a Leica EM TRIM2 to remove the resin surrounding the sample. The resin above the sample in the trimmed block was further cut at 200 nm increments using a glass knife in the ultramicrotome Leica EM UC7 until the sample was exposed. Seventy nanometer sections were cut with a diamond knife (Diatome Ultra 45°). A small ribbon of floating sections was collected and mounted onto a copper-grid coated with formvar. Once the grid was dry, it was stored in a grid box for further observation in Tecnai10 (FEI; accelerating voltage 80 kV) or Tecnai 12 (FEI; accelerating voltage 120 kV) transmission electron microscopes.

SDS-digested freeze-fracture replica preparation and labeling

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Brain slices (70 µm) of mice transcardially perfused with 2% PFA in 0.1 M PB were prepared, and the whole IPN was manually excised. Tissue was then incubated in 30% glycerol in 0.1 M PB O/N for cryo-protection. The next day, tissue samples were transferred into gold or copper carriers and frozen under high pressure (>300 bar) using an HPM010 (Leica, Wetzlar, Germany). Frozen samples were stored in liquid nitrogen until further processing. To craft freeze-fracture replicas, two gold carriers containing frozen tissue samples were placed in a carrier holder in liquid nitrogen, which was inserted into the freeze-fracture machine (BAF060, Leica) and left to equilibrate to ‒117°C under high vacuum (2.0 × 10−7–1.0 × 10−6 mbar) for 20 min. Subsequently, tissue was fractured and a carbon layer (5 nm at a rate of 0.1–0.3 nm/s) was evaporated onto the tissue at a 90° angle, followed by a platinum/carbon layer (2 nm at a rate of 0.06–0.1 nm/s) applied at a 60° angle and another carbon layer (20 nm at a rate of 0.3–0.6 nm/s) applied at a 90° angle. For the preparation of carbon-only replicas, the second layer consisted of a 5 nm carbon layer applied at a 60° angle. After evaporation, replicas were removed from the machine and placed in tris-buffered saline (TBS, 50 mM). Subsequently, replicas were glued (tissue-side up) onto a finder grid and the glue (optical adhesive 61, Norland, Cranbury, NJ) was hardened by applying UV light for 20 s and then transferred into SDS-solution containing: 2.5% SDS, 20% sucrose in 15 mM Tris buffer (pH 8.3). Tissue was subsequently digested by incubating the replicas for 48 hr at 60°C under gentle agitation (50 rpm shaker), followed by incubation for 12–15 hr at 37°C under gentle agitation.

For immunolabeling of SDS-digested replicas, replicas were washed in washing buffer (containing 0.1% Tween-20, 0.05% bovine serum albumin [BSA], 0.05% NaN3pH = 7.4) and incubated in blocking buffer (washing buffer+ 5% BSA) for 1 hr. Replicas were transferred to blocking buffer containing primary antibody (ginea pig anti-Cav2.3, 8 µg/µl) followed by incubation at 15°C O/N. Thereafter, replicas were washed and incubated in blocking solution containing secondary antibodies (2 nm gold-conjugated anti-rabbit, 5 nm gold-conjugated anti-ginea pig, or 10 nm gold-conjugated anti-rabbit, all diluted 1:30) at 15°C O/N. The following day, the antibody labeling procedure was repeated for the next primary antibody (4 µg/ml for anti-KCTD8, anti-KCTD12, anti-KCTD12b; 2 µg/ml for anti-GABAB1) and corresponding secondary antibodies. Labeled grid-glued replicas received a final carbon layer (20 nm) onto the labeled replica side using a High-Vacuum Coater ACE600 (Leica), followed by dissolution of the glue in Dynasolve 711 (Dynaloy, Indianapolis, IN) at 60°C under gentle agitation (60 rpm) for 2 hr. Solvent was subsequently removed by washing the grid in 100% methanol for 10 min followed by ethanol (100, 95, 90, 70, 50%, 5 min each). After short air drying, replicas were stored in grid boxes until observation under the transmission electron microscope.

Acute brain slice electrophysiology

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Mice were anesthetized with a mixture of ketamine (90 mg/kg) and xylazine (4.5 mg/kg) and transcardially perfused with ice-cold, oxygenated (95% O2, 5% CO2) artificial cerebrospinal fluid (ACSF) containing (in mM) 118 NaCl, 2.5 KCl, 1.5 MgSO4, 1 CaCl2, 1.25 NaH2PO4, 10 d-glucose, 30 NaHCO3, (pH = 7.4). The brain was rapidly excised, and coronal brain slices of 250 µm thickness were prepared with a Dosaka Linear-Pro7. For SNX-482 experiments in rostral IPN, brain slices were prepared at a 54° angle to allow for electrical stimulation of FR. For angled slice recordings in lateral IPN of WT and Cav2.3 KO mice (Figure 1H–K), slices were cut at 1 mm thickness to assure that FR remained intact all the way to the lateral IPN. Slices were recovered at 35°C for 20 min and thereafter slowly cooled down to RT over the course of 1 hr. After recovery, one slice was transferred to the recording chamber (RC-26GLP, Warner Instruments, Holliston, MA) and superfused with ACSF containing 2.5 mM CaCl2, 20 µM bicuculline methiodide, 50 µM hexamethonium bromide, and 5 µM mecamylamine hydrochloride at a rate of 3–4 ml/min at 32.0 ± 2.0°C. Rostral or lateral IPN nuclei were visually identified using an infrared differential interference contrast video system in a BX51 microscope (Olympus, Tokyo, Japan). Electrical signals were acquired at 10–50 kHz and filtered at 2 kHz using an EPC 10 (HEKA, Lambrecht/Pfalz, Germany) amplifier. Glass pipettes (BF150-86-10, Sutter Instrument, Novato, CA) with resistances of 3–4 MΩ were crafted using a P97 horizontal pipette puller (Sutter Instrument) and filled with internal solution containing (in mM) 130 K-Gluconate, 10 KCl, 5 MgCl2, 5 MgATP, 0.2 NaGTP, 0.5 EGTA, 5 HEPES; pH 7.4 adjusted with KOH. Whole-cell patch-clamp recordings were performed in voltage-clamp mode at a holding potential of –60 mV, and access resistance was constantly monitored via a –10 mV voltage step at the end of each sweep. Recordings with access resistances exceeding 20 MΩ or with changes in access resistance or holding current by more than 20% were discarded. To evoke glutamatergic excitatory postsynaptic currents (EPSCs) in rostral IPN neurons, voltage pulses (0.5–3.5 V, 0.2 ms duration) were applied with an ISO-Flex stimulus isolator (AMPI, Jerusalem, Israel) to a concentric bipolar stimulating electrode (CBBPC75, FHC, Bowdoin, ME) located inside the IPN, ~200–300 µm distal to the recorded neuron. For optogenetic stimulation in Tac1-ChR2 mice, blue light (λ = 465 nm) was emitted directly onto the lateral IPN through a 5 mm long mono fiber-optic cannula (fiber diameter 200 µm, total diameter 230 µm, Doric lenses, Quebec, Canada) connected to a PlexBright LED (Plexon, Dallas, TX), with an optical patch cable (fiber diameter 200 µm, total diameter 230 µm, 0.48 NA). The LED was triggered via 200–290 mA current pulse (2 ms duration) from a LED Driver (LD-1, Plexon), which was controlled directly via the HEKA EPC10 amplifier. The sweep interval of all stimulation protocols (electrical and optogenetic) was 10 s. For the application of SNX-482 (1 µM, hellobio, Bristol, UK) and ω-Conotoxin GVIA (1 µM, Alomone Labs, Jerusalem, Israel), 0.1% bovine serum albumin was added to the ACSF. For variance-mean analysis, ACSF with four different Ca2+ concentrations (1.5–8 mM) was applied to measure variance and mean EPSC amplitude at different release probabilities. Each Ca2+ concentration was washed in for 5–10 min. Once EPSC amplitudes stabilized, 15–20 consecutive EPSC responses were used for the calculation of mean EPSC amplitude and variance, followed by the wash-in of the next Ca2+ concentration. The order of application was 2.5 mM Ca2+, followed by 1.5 mM Ca2+, followed by 6–8 mM Ca2+, followed by 4–5 mM Ca2+. Baseline noise was much smaller than synaptic noise. The values for release probability and quantal size were calculated according to the equations Var = Iq – I²/N and I = Nqp, with I being the average EPSC amplitude, N the number of release sites, and q the quantal size. To measure the paired-pulse ratio (PPR) of two consecutively evoked EPSCs at 20 Hz, the PPRs of 20–30 EPSC pairs evoked at 10 s intervals were averaged. To study the effect of GBR activation on EPSC amplitude, R(+)-Baclofen hydrochloride (1 µM) was bath applied for 3 min and washed for 10 min. For PPR and variance-mean experiments, data from wild-type C57BL/6J and BALB/c mice were found to not differ significantly and thus were pooled. To measure action potential properties in ventral MHb neurons, a 50 pA current was injected for 100 ms, which reliably evoked a train of action potentials. Action potential peak times were measured as the time from the threshold potential to the peak. Action potential full-width at half maximum was measured as the time between rising and the falling phase of the action potential at half of the maximal action potential amplitude. Only the first action potential in the train was analyzed.

Cell lines

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Human Embryonic Kidney 293T (HEK293T) were directly obtained from ATCC (https://web.expasy.org/cellosaurus/CVCL_0063)and maintained in DMEM supplemented with GlutaMAX (Invitrogen) and 10% FCS in a humidified atmosphere (5% CO2) at 37°C. HEK293 cells (RRID:CVCL_0045), stably expressing the human Cav2.3 (R-type) channel α1E-3 splice variant (also called α1E-c; GenBank L29385), the Ca2+ channel auxiliary subunits human α2bδ−1 (M76559) and human β3a (NM_000725), as well as the human potassium channel KCNJ4 (Kir2.3; U07364), were a kind gift from the laboratory of David J. Adams (University of Wollongong, Australia). Cells were maintained in culture as described before (Dai et al., 2008). All cell lines were authenticated using Short Tandem Repeat (STR) analysis by Microsynth (Switzerland) and tested negative for mycoplasma contamination.

Cell culture transfection and electrophysiology

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For experiments, the Cav2.3-expressing HEK293 cells were transiently transfected with plasmids for the expression of Myc-tagged KCTD8 (AY615967) or KCTD12b (AL831725) using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) as described earlier (Schwenk et al., 2010). Plasmids for the expression of eGFP (Clontech, Kyoto, Japan) were co-transfected in order to identify cells expressing the plasmids. A total quantity of 0.75 µg of DNA was transfected in each well, and empty pCI plasmid was used to complete to this amount of total DNA. The control group contained only eGFP and empty vector. Two to four days after transfection, cells were briefly subjected to Versene (ThermoFisher, Waltham, MA) treatment for its dissociation and plated on glass coverslips and, at least 2 hr later, electrophysiology experiments were performed.

Whole-cell recordings were performed using fire-polished borosilicate patch pipettes (2–4 MΩ of tip resistance), filled with a cesium-based intracellular solution containing (mM): 110 CsMeSO3, 15 CsCl, 10 EGTA, 10 HEPES, 10 Tri–phosphocreatine, 0.1 NaGTP, 4 MgATP, pH 7.3 adjusted with CsOH. The extracellular solution contained (mM): 110 NaCl, 30 TEA-Cl, 10 HEPES, 10 d-glucose, 5 CsCl, 1 MgCl2 and 5 mM BaCl2, pH 7.4 adjusted with 40% TEA-OH. For patch-clamp experiments, isolated cells with a clear eGFP fluorescent signal were selected. The culture was maintained under continuous perfusion (~1 ml/min) at room temperature. A MultiClamp 700B (Molecular Devices, San Jose, CA) amplifier connected to a 1440A Digidata was used to acquire signals at 10–50 kHz, filtered at 2–8 kHz. After whole-cell formation, series resistance (<10 MΩ) was compensated by 85%. Linear leak currents and capacity transients were subtracted on-line using a −P/4 protocol. Current-voltage relationships were recorded from a holding potential of −80 mV using 25 ms depolarizations from −70 to 60 mV in 10 mV increments. Peak IBa was measured at each step and normalized to the capacitance of the cell to obtain the current densities, which were plotted as a function of the voltage step. I−V curves were fit with a standard Boltzmann equation as follows: IBa = Gmax(V − Vrev)/(1 + exp((V0.5actV)/k)), where IBa is the current measured at each test potential V, Gmax is the maximal conductance, Vrev is the reversal potential, V0.5act is the half-maximal voltage of activation, and k is the slope factor. To study voltage dependency of activation, a series of 25 ms voltage steps from −80 to 60 mV (with 10 mV increments) were applied, followed by a 10 ms step to 0 mV, where peaks of the evoked current tails were measured, normalized to the maximal peak, and plotted against the corresponding voltage. To the study-steady state inactivation, a series of 2 s long voltage steps from −120 to 10 mV (10 mV increments) was applied, followed by a test step to 10 mV (150 ms) where peaks of elicited currents were measured, normalized to the maximal response, and plotted against the corresponding voltage. Both activation and inactivation curves were fitted using a Boltzmann sigmoidal as follows: I = I2 + (I1I2) / (1 + exp((V0.5 act/inactVt)/k)), where V0.5 act/inact is the half-maximal activation or inactivation and k is the slope factor that was positive for activation and negative for inactivation curves.

Co-immunoprecipitation

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Plasmids encoding N-terminally Flag-tagged KCTDs were described earlier (Seddik et al., 2012). Plasmids expressing C-terminally Flag-tagged Cav2.3 were obtained from GenScript. Cultured human embryonic kidney 293T (HEK293T) cells were transfected using 2 μg/μl polyethylenimine (Sigma-Aldrich) with indicated plasmids for the expression of either Flag-KCTDs alone or in combination with Flag-Cav2.3. Cells were washed 48 hr after transfection in ice-cold PBS and lysed in NETN buffer (100 mM NaCl, 1 mM EDTA, 0.5% Nonidet P-40, 20 mM Tris–HCl, pH 7.4, supplemented with cOmplete EDTA-free protease inhibitor mixture [Roche, Basel, Switzerland]), followed by rotation for 10 min at 4°C. Cell lysates were then cleared by centrifugation at 10,000 × g (4°C, 10 min) and directly used for immunoblot analysis (Input) or for co-immunoprecipitation assay, in which they were precleared for 1 hr using 30 μl (dry volume) of a 1:1 mixture of protein-A and protein-G-agarose beads (GE Healthcare, Chicago, IL). Thereafter, lysates were incubated by rotating for 16 hr at 4°C in the presence of 2.5 μl of 0.3 μg/μl anti-Cav2.3 (CACNA1E) antibody (ACC-006, Alomone Labs, Jerusalem, Israel). The next day, 10 μl (dry volume) of a 1:1 mixture of protein-A and protein-G-agarose beads were added and incubated by rotating for 40 min at 4°C. Afterward, they were washed with 5 × 1 ml of NETN buffer and pulled down proteins were eluted with 25 μl of 4× sample loading buffer containing 200 mM DTT. Proteins were resolved using standard one-dimensional SDS–PAGE on 10% polyacrylamide gels (for 45 min at 70 mV, followed by an additional 1.5 hr at 120 mV). For immunoblotting analysis, proteins were transferred to 0.45 μM polivinylidene fluoride membranes (Milipore, Burlington, MA) for 2 hr at 200 mA and probed with the primary antibodies rabbit anti-Flag (F7425, Sigma) and anti-Cav2.3 (ACC-006, Alomone Labs) in combination with peroxidase-coupled secondary donkey anti-rabbit antibodies (NA934, GE Healthcare, 1:10,000).

For co-IP experiments using membrane extracts, we transiently transfected HEK293T cells with the pore forming α-subunit (rat Cav2.3) together with auxiliary β3 and α2δ1 subunits in order to emulate the experimental conditions present the stable cell line (Dai et al., 2008). These cells were then subjected to hypotonic lysis to separate cytosolic proteins from membrane proteins (total membrane). A fraction of the total membrane pool was then subjected to a plasma membrane purification protocol using a commercially available plasma membrane extraction kit (ab 65400, Abcam, Cambridge, UK). The plasma membrane fraction obtained after the purification as well as the total membrane fraction were subsequently solubilized in detergent-containing buffer (0.5% Nonidet P-40) and loaded on 10% SDS–PAGE gels. The presence of the KCTD proteins in the corresponding tissues was determined by western blotting using ECL detection of HRP-coupled antibodies against Flag-tagged KCTD8 (rabbit anti-KCTD8) and Flag-tagged KCTD12b (mouse anti-Flag). For co-immunoprecipitation assay, lysates were incubated by rotating for 16 hr at 4°C in the presence of 2.5 μl of 0.3 μg/μl anti-Cav2.3 (CACNA1E) antibody (ACC-006, Alomone Labs, Jerusalem, Israel), washed, and eluted as described above with subsequent immunoblotting analysis performed as described above.

Lentiviral injections

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Lentiviral vectors (LV–hSyn–EGFP, LV–hSyn–3xFlag/mKCTD8[NM_175519.5]/P2A/EGFP, and LV–hSyn–3xFlag/mKCTD12b[NM_175429.4]/P2A/EGFP) were generated by the life science facility at IST Austria and by VectorBuilder GmbH (Neu-Isenburg, Germany). Bilateral viral injections were performed in male C57Bl/6J or KCTD12b KO mice at 2 months of age. Seven hundred nanoliter of virus was injected at a rate of 50 nl/min to the following coordinates: anterior/posterior: –1.42 mm from Bregma, lateral: ±0.22 mm from the midline, dorsal/ventral: –2.5 mm from the surface of the brain. Mice were allowed to recover for 10–14 days before carrying out electrophysiology experiments.

Data analysis and statistics

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All statistical tests and graph preparations were done using Prism (GraphPad, San Diego, CA), and figure assembly was done in Photoshop (Adobe, San Jose, CA). To determine whether to use parametric or non-parametric statistical tests, Shapiro-Wilk test for normality of residuals was applied. Unless otherwise noted, averaged data is presented as mean ± SEM, and p-values<0.05 were considered to indicate statistical significance. Power analysis (power = 0.8, α error = 0.05) was used to determine the appropriate sample size for SDS-FRL and variance-mean experiments using G*Power software (Univ. Kiel, Germany). Sample size in other experiments was based on those of similar experiments in previous studies. Masking was not used for any experiment. For the analysis of pre-embedding immunolabeled ultrathin sections, the presynaptic active zone was manually demarcated based on (1) rigid alignment of pre- and postsynaptic membranes and the presence of a postsynaptic density with the same length as the presynaptic active zone, as previously described (Rubio et al., 2017). Active zone length and silver-intensified gold particle densities and distances were measured using Reconstruct software. In SDS-FRL samples, demarcation of presynaptic active zones was performed manually based on two morphological criteria: there had to be a visible alteration in surface curvature in the P-face and/or a concentration in intramembrane particles (IMPs) of variable sizes that appeared clearly different from those of the surrounding P-face. For density measurements, incomplete active zones were analyzed whenever there was a visible transition to the postsynaptic E-face that displayed the characteristic IMP clusters of a glutamatergic postsynapse (Tanaka et al., 2005). For confirmation, the complete active zone area demarcated based on protoplasmic surface depression and concentrated IMP cluster were compared with the area of complete postsynaptic E-face IMP clusters. In addition, the area of manually demarcated complete active zones was also compared with those demarcated using a mixture of active zone-marker antibodies for RIM1/2 (5 µg/ml; Synaptic Systems, Göttingen, Germany), neurexin (5 µg/ml) (Miki et al., 2017), and CAST (3 µg/ml) (Hagiwara et al., 2018). To analyze clustering of molecules (Cav2.3, GABAB1, KCTD8, and KCTD12b), we performed 100 Monte Carlo simulations using GPDQ software (Luján et al., 2018). For each simulation, the same number of particles was redistributed randomly, with each pixel having the same probability of becoming the center of a particle, on the demarcated area of interest under the condition that two particles could not be closer to each other than 10 nm. We then compared nearest neighbor distances (NNDs) between real particles with the NNDs of simulated particles using Kolmogorov–Smirnov test.

Data availability

For all figures, numerical data that are represented in graphs are provided as source data excel files.

References

    1. Fujimoto K
    (1995)
    Freeze-fracture replica electron microscopy combined with SDS digestion for cytochemical labeling of integral membrane proteins. Application to the immunogold labeling of intercellular junctional complexes
    Journal of Cell Science 108:3443–3449.
    1. Ludwig A
    2. Flockerzi V
    3. Hofmann F
    (1997)
    Regional expression and cellular localization of the alpha1 and beta subunit of high voltage-activated calcium channels in rat brain
    The Journal of Neuroscience 17:1339–1349.

Decision letter

  1. Yukiko Goda
    Reviewing Editor; RIKEN, Japan
  2. John R Huguenard
    Senior Editor; Stanford University School of Medicine, United States
  3. Marco Capogna
    Reviewer; University of Aarhus, Denmark

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

This study examines presynaptic regulation of the medial habenula to interpeduncular nucleus connection, which plays a crucial role in aversion and addiction related behaviors. The authors focus on the special features of these synapses, namely the presynaptic expression of R-type voltage gated calcium channel, Cav2.3, the potent modulation by the presynaptic GABAB receptors, and their auxiliary KCTD subunits whose functions have been elusive. Through electrophysiological, biochemical and ultrastructural characterizations, the paper shows that KCTD8 and KCTD12b directly interact with Cav2.3 and regulate neurotransmitter release from ventral medial habenula terminals independently of GABAB receptors, where effectively KCTD12b serves to restrain release while the recruitment of KCTD8 to the active zone potentiates release by promoting Cav2.3 currents. These findings are important as they identify novel, GABAB receptor-independent role for KCTD proteins in regulating neurotransmission and provide insights into the underlying mechanisms.

Decision letter after peer review:

[Editors’ note: the authors submitted for reconsideration following the decision after peer review. What follows is the decision letter after the first round of review.]

Thank you for submitting your work entitled "GABAB receptor auxiliary subunits modulate Cav2.3-mediated release from medial habenula terminals" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by a Senior Editor. The reviewers have opted to remain anonymous.

Our decision has been reached after consultation between the reviewers. Based on the discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife, at least in its present form.

All three reviewers recognized the data are of very high quality and the findings to be potentially interesting. However, concerns were raised with respect to uncertainties about the underlying mechanisms of GABAB receptor-mediated presynaptic modulation and the roles of KCTDs, such that the study remained premature at this stage.

Reviewer #1:

This study examines presynaptic modulation of the medial habenula (MHb) to interpeduncular nucleus (IPN) connection, which plays a crucial role in aversion and addiction related behaviors. The authors focus on the special features of these synapses, namely the expression of R-type VGCC, Cav2.3 in presynaptic active zones and the robust presynaptic GABAB receptor (GBR)-mediated potentiation. In particular, they seek to determine the mechanism of GBR-dependent potentiation of release by considering the actions of the KCTD subfamily members that serve as auxiliary subunits of GBR.

The study reveals in presynaptic terminals of ventral MHb to rostral IPN connection in KCTD12b KO mice, a gain of function activity of KCTD8 in potentiating neurotransmitter release, likely via interacting with Cav2.3 to modulate its channel activity. It remains unclear what the normal functions of KCTD8 and KCT12b might be. For example, the lateral IPN has no detectable KCTD12b but expresses high levels of KCTD8 (Figure 2), yet its response to baclofen is opposite to the response detected in the rostral IPN in KCTD12b KO mice. Given that KO mice have been analyzed in the adult stage, whether the developmental compensatory effects could have contributed to the observed effects, requires attention also.

Altogether, the study provides an informative characterization of the ultrastructural localization of KCTD proteins, GBR and Cav2.3 at the synapse, and neurotransmitter release properties of synapses lacking KCTD8 and KCTD12b either singly or in combination. The paper is written clearly and the experiments have been carefully executed. However, further experiments and analysis are required to compellingly support the main conclusions and establish further insights into the mechanism by which KCTD8 and KCTD12 regulate basal release and clarify their contributions to GBR-dependent release.

Specific points

If there is a bi-directional regulation of neurotransmitter release by KCTDs in which KCTD8 potentiates whereas KCTD12b depresses, then one should test the effects of overexpression of KCTD8 and KCTD12b in WT animals to evaluate the competitive nature of the actions of KCTD8 and KCTD12b.

In order to address whether there may be developmental contribution of KCTD8 and KCTD12b, one should test whether acute knock-down produces the same effects and/or demonstrate the sufficiency by rescuing the mutant phenotype with exogenous overexpression in the adult.

p. 4, middle paragraph: "KCTD subunits co-precipitate with release machinery proteins of the presynaptic active zone…" Please specify which active zone proteins?

In KCTD8 KO neurons, is there a compensatory recruitment of KCTD12b to the active zone, that is, the opposite of what takes place in KCTD12b KO neurons?

Figure 2 – it would be useful to test for cross-labelling to evaluate any compensatory expression for the loss of particular KCDT subtype with another KCDT subtype at the level of light microscopy.

Figure 6A. Regarding the representative plot for the double KO mice, it is worrisome that there is a drastic increase in the baseline prior to baclofen addition, such that it is difficult to interpret the rest of the data points. Does baclofen still potently increase EPSC amplitudes in KCTD8, KCTD12b KO or KCTD8/12b DKO mice if recordings are carried out in 4 mM extracellular Ca instead of 2.5 mM?

Figure 7. To determine the nature of the increase in the active zone expression of KCTD8 in KCTD12b KO terminals, one should perform (1) the distance-dependent analysis of immunogold labelling for KCTD8 in KCTD12b KO preparation as shown for Figure 3, especially with respect to whether the profile of KCTD8 changes to what is shown for KCTD12b in rostral IPN, and (2) analysis at the level of light microscopy to assess whether there is an increase in the level of expression or KCTD8 in KCTD12b KO rostral IPN (i.e. the same comment as for Figure 2 above).

Figure 8. The relative levels of surface expression of KCTD8 and KCTD12b in HEK cells needs to be quantified in order to fully interpret the preferential increase in Cav2.3-mediated currents in KCTD8-transfected cells. The direct interaction between KCTD8 and KCTD12b with Cav2.3 should be confirmed in the brain.

Reviewer #2:

This manuscript aims to examine the mechanism of the unusual potentiating effect of baclofen acting on presynaptic GABAB receptors in the MHb-IPN pathway. I was expecting an examination of underlying second messenger pathways, but this was not the route taken, as the authors investigating whether there is a role for the KCTD proteins which interact with GABAB receptors.

My impression of this manuscript is that by trying to understand the potentiating effect of baclofen on the synapses in question, the authors have uncovered a very interesting effect of KCTD8 potentiating CaV2.3, and an interplay between KCTD8 and KCTD12b, shown with several techniques including very nice EM images and analysis, which however is completely separate from the effect of baclofen acting on GABAB receptors.

In trying nevertheless to combine the two stories, the authors have overcomplicated the message of their study, in my view. I would prefer to see the results separated, with the KCTD study stripped out and potentially published here, as it is novel and well performed. Otherwise it seems to me the manuscript is over-complex and the very interesting message about the KCTDs is difficult to extract.

1. Line 99 SNX482 is not "selective". The authors are using it at 1 µM. It may be relatively selective between calcium channels, but it also blocks certain K channels in the same concentration range, in fact 500 nM was used in slices to block A-type K currents (Kimm and Bean, 2014). It also affects other Ca channels and Na channels at concentrations below 1µM (Newcomb et al., 1998). How sure are the authors this blockade of other channels is not affecting their study ?

2. Line 104. You cannot say the two MHb-IPN pathways have different sensitivities to SNX482, as no dose response curve has been performed, and the difference between the pathways at the single dose studied may indicate the involvement of other Ca channels.

3. Similarly, in the Discussion, line 310 "We confirmed that both dorsal MHb-IPN and ventral MHb-IPN pathways require Cav2.3 for transmitter release", is not completely correct, as how is release occurring for the ~50% SNX482-insensitive lateral epscs (Figure 1) ?

4. Figure 8A needs correct labelling of the panels, with MW markers. I assume the top panel is KCTDs, but this is unclear as both CaV2.3 and KCTDs have Flag tags, which is an unusual way of performing the experiment. The entire blot should be shown for the top panel, as blotting with Flag would show both KCTDs and CaV2.3.

5. If there is co-ip between CaV2.3 and KCTD8/12b, should there be actual co-localization observed in the EM images? Can this be measured?.

6. In Figure 8c, how selective is the augmentation with KCTD8 and CaV2.3, ie does it also occur for any other Ca channels? It is possible that a low level of other presynaptic Ca channels might be also augmented by KCTD8 in these terminals, increasing Ca2+ entry.

7. If this large augmentation of CaV2.3 currents by KCTD8 plays a physiological role should the epsc's in the KCTD8 KO mice be reduced? I cannot see this basic result, although I note the release probability is indeed reduced.

8. Following on from point 6, in Discussion line 329, has the 1997 study by Ludwig et al. showing no α1B, A, C mRNA in the habenula been replicated anywhere? The technique used in this early study is not very sensitive, are there other more recent studies using single cell RNAseq for example?

9. Discussion line 330 "Although this exclusivity has been described anatomically (Parajuli et al., 2012)" is not correct as it gives the impression other Ca channels were examined in this study, whereas this is not the case.

10. Discussion line 331: SNX482 is here described as specific, which is incorrect.

References

Kimm, T., and Bean, B.P. (2014). Inhibition of A-type potassium current by the peptide toxin SNX-482. J Neurosci 34, 9182-9189.

Newcomb, R., Szoke, B., Palma, A., Wang, G., Chen, X.H., Hopkins, W., Cong, R., Miller, J., Urge, L., Tarczy-Hornoch, K., et al. (1998). Selective peptide antagonist of the class E calcium channel from the venom of the tarantula Hysterocrates gigas. Biochemistry 37, 15353-15362.

Reviewer #3:

The manuscript attempts to elucidate the mechanisms underlying the potentiation of synaptic transmission between the ventral medial habenula (MH) and the rostral interpeduncular nucleus mediated by presynaptic GABA-B receptors. The topic is interesting especially because it is very unusual that GABA-B receptors gate an enhancement of neurotransmission, since usually when they are activated, they depress the release of neurotransmitter.

The manuscript contains some high quality data, for example, freeze-fracture replica immunolabelling of presynaptic proteins in habenular terminals. However, I also found some technical problems, (e.g., see comment #1 below). Overall, the manuscript is potentially interesting and covers an original topic. However, the data currently submitted fail to answer the two major questions, namely the underlying mechanisms and the physiological relevance of GABA-B receptor-mediated modulation. A lot of efforts are devoted in characterizing key protein expression in various K+ channel tetramerization domain-containing (KCTD) proteins. However, I doubt that this strategy paid off and really shed light on the mechanisms underlying GABA-B receptor mediated modulation of the synapses studied.

1. Data on the action of SNX-482 on evoked EPSCs need refinement. The presynaptic fibers of the fasciculus retroflexus were electrically stimulated and recordings were made in the rostral IPN. In contrast, stimulation of dorsal fibers were obtained via optogenetic stimulation and responses were recorded in the lateral IPN. The effect of SNX-482 was stronger at the former versus the latter synapses. However, this result is at odds with similar Cav2.3 densities in the active zones of dorsal and ventral MHb terminals. The Authors discussed this issue, and I acknowledged this mis-match in their data. Nevertheless, the problem remains: a quantitative comparison between drug modulation of synaptic responses evoked by electrical versus optogenetic stimulation is a poor choice. Amongst various problems, several recent papers report the non-physiological high release probability often caused by optogenetic stimulation of presynaptic fibers that contrasts with the electrical stimulation method. This discrepancy would affect the results obtained with SNX-482. This set of experiments should be repeated by using more uniform experimental conditions in the two synaptic pathways. Furthermore, a serious comparison of the efficacy of SNX-482 should rely on testing at least three different drug concentrations for each synapse. Furthermore, normalization assuring that approximatively an equal number of presynaptic fibers are stimulated at each synaptic connection should be employed. What ever methods of presynaptic stimulation will be used, optical or electrical, test of selective stimulation of fibers should be also performed.

2. Baclofen still presynaptically potentiated EPSCs in KCTD KO mice. However, loss of KCTD12b impaired GABABR action probably via increasing release probability and occlusion. In contrast, lack of KCTD8 reduces release probability. But what about the effects of GABABR modulation? Is baclofen more powerful at MH synapses in KCTD8 KO?

3. The manuscript lacks to experimentally demonstrate that GABA-B receptor potentiation of ventral MH and the rostral interpeduncular nucleus synapses is physiologically relevant. This should be addressed by using physiologically salient repetitive stimulation in control and in the presence of GABA-B R antagonists. It would be also interesting to test the GABA-B R ligands on evoked EPSPs followed by IPSPs to see the comprehensive pharmacological effects on both excitation and inhibition. At present none of these important points are addressed and this reduces the impact of the data currently submitted.

[Editors’ note: further revisions were suggested prior to acceptance, as described below.]

Thank you for submitting your article "GABAB receptor auxiliary subunits modulate Cav2.3-mediated release from medial habenula terminals" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by John Huguenard as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Marco Capogna (Reviewer #2).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

The revised manuscript has been significantly improved by refocusing the study on the role for KCTDs on neurotransmitter release. Several additional experiments have been performed, including the comparisons of Cav2.3-dependence of synaptic transmission between the two distinct medial habenula to IPN inputs by electrical stimulation, which has been made possible by the establishment of thick-slice preparation. New data lend further support to the conclusion that KCTD8 and KCTD12b directly interact with Cav2.3 and regulate release from ventral MHb terminals independently of GABAB receptors, where effectively KCTD12b serves to restrain release while the recruitment of KCTD8 to the active zone potentiates release by promoting Cav2.3 currents. Prior to accepting the manuscript for publication, a number of points require attention.

Figure 6F-G: the EPSC waveforms of some rescued currents appear to be quite different than one would expect. Specifically, the LV-hSyn-KCTD8 current is quite prolonged, and may have two components to the rising phase. Additionally, the LV-hSyn-KCTD12b current appears to have a delayed onset. Are these examples representative of the population? If so, are these effects a result of over-expression, or more an indicator of the distinct roles of these two KCTD proteins?

In the introduction (Line 47 or Line 53-54), it would be clearer to readers who may not necessarily be familiar with KCTDs if the authors could briefly state the nature of the molecular difference especially between KCTD12 and KCTD12b.

Line 302-304. "The decrease in release probability by KCTD12b in the presence but not in the absence of KTCD8…" is confusing. Do the authors mean that the reason why the decreased PPR observed in KCTD12bKO could be recovered to control levels when KCTD8 is additionally absent (as observed in double KO mice), could be because the loss of KCTD12b in itself alters the localization of KCTD8 or Cav2.3? Perhaps this sentence should be rephrased.

https://doi.org/10.7554/eLife.68274.sa1

Author response

[Editors’ note: the authors resubmitted a revised version of the paper for consideration. What follows is the authors’ response to the first round of review.]

Reviewer #1:

This study examines presynaptic modulation of the medial habenula (MHb) to interpeduncular nucleus (IPN) connection, which plays a crucial role in aversion and addiction related behaviors. The authors focus on the special features of these synapses, namely the expression of R-type VGCC, Cav2.3 in presynaptic active zones and the robust presynaptic GABAB receptor (GBR)-mediated potentiation. In particular, they seek to determine the mechanism of GBR-dependent potentiation of release by considering the actions of the KCTD subfamily members that serve as auxiliary subunits of GBR.

The study reveals in presynaptic terminals of ventral MHb to rostral IPN connection in KCTD12b KO mice, a gain of function activity of KCTD8 in potentiating neurotransmitter release, likely via interacting with Cav2.3 to modulate its channel activity. It remains unclear what the normal functions of KCTD8 and KCT12b might be. For example, the lateral IPN has no detectable KCTD12b but expresses high levels of KCTD8 (Figure 2), yet its response to baclofen is opposite to the response detected in the rostral IPN in KCTD12b KO mice. Given that KO mice have been analyzed in the adult stage, whether the developmental compensatory effects could have contributed to the observed effects, requires attention also.

Altogether, the study provides an informative characterization of the ultrastructural localization of KCTD proteins, GBR and Cav2.3 at the synapse, and neurotransmitter release properties of synapses lacking KCTD8 and KCTD12b either singly or in combination. The paper is written clearly and the experiments have been carefully executed. However, further experiments and analysis are required to compellingly support the main conclusions and establish further insights into the mechanism by which KCTD8 and KCTD12 regulate basal release and clarify their contributions to GBR-dependent release.

We have re-structured our manuscript now focusing on the role of the KCTDs on Cav2.3-mediated basal release, and added further experiments and analysis to support our initial claims as follows.

Specific points

If there is a bi-directional regulation of neurotransmitter release by KCTDs in which KCTD8 potentiates whereas KCTD12b depresses, then one should test the effects of overexpression of KCTD8 and KCTD12b in WT animals to evaluate the competitive nature of the actions of KCTD8 and KCTD12b.

We thank the reviewer for raising this important point. We do not think that KCTD12b is actively depressing release since it did not affect Cav2.3-mediated currents in vitro. We rather think that KCTD8 is enhancing release by increasing currents through Cav2.3 and KCTD12b may just compete with KCTD8 for binding to Cav2.3 and prevent the effect of KCTD8. To test the effect of overexpression of KCTD8 on basal release from ventral MHb terminals, we injected lentivirus expressing either EGFP or KCTD8-p2A-EGFP bilaterally into the MHb of WT mice (Figure 6F). PPR values recorded in rostral IPN neurons of EGFP control-injected animals remained unchanged (LV-EGFP: 2.17 ± 0.18, n = 17, 5 mice), whereas KCTD8-injected animals exhibited significantly lower PPR values (LV-KCTD8: 1.41 ± 0.14, n = 13, 3 mice; t28 = 3.20, P = 0.0034; unpaired t-test). These results are described on page 14, starting from line 288.

In order to address whether there may be developmental contribution of KCTD8 and KCTD12b, one should test whether acute knock-down produces the same effects and/or demonstrate the sufficiency by rescuing the mutant phenotype with exogenous overexpression in the adult.

To address this important point, we used lentiviral expression of KCTD12b in MHb neurons of KCTD12b KO mice. Lentiviral expression of KCTD12b-p2A-EGFP in KCTD12b KO mice significantly increased PPR values compared with EGFP control-injected animals (Figure 6G; KCTD12b KO EGFP Control: 1.38 ± 0.09, n = 11, 3 mice; PPR KCTD12b KO rescue: 2.18 ± 0.29, n = 14, 3 mice; t23 = 2.98, P = 0.0068, unpaired t-test), rescuing the mutant phenotype (no significant difference in PPR compared with wild-type mice shown in Figure 6A; WT: 2.17 ± 0.18, n = 17, 5 mice; KCTD12b KO rescue: 2.18 ± 0.29, n = 14, 3 mice; t29 = 0.40, P = 0.6925, unpaired t-test). These results are described on page 14, starting from line 293.

p. 4, middle paragraph: "KCTD subunits co-precipitate with release machinery proteins of the presynaptic active zone…" Please specify which active zone proteins?

We apologize for the lack of detail. We included a more detailed description of the findings from the affinity purifications in the cited paper. The updated sentence now reads “Based on proteomics studies, voltage-gated Ca2+ channels co-precipitate GBRs and their auxiliary KCTD subunits together with release machinery proteins of the presynaptic active zone, including SNAP-25, synaptotagmins, synaptobrevin-2, Munc13-1, syntaxins, RIM1 and synapsins, among others (Müller et al., 2010).” This sentence is found in the introduction section on page 4, lines 59 – 62.

In KCTD8 KO neurons, is there a compensatory recruitment of KCTD12b to the active zone, that is, the opposite of what takes place in KCTD12b KO neurons?

This is an important question. It is unclear whether a compensatory increase in KCTD12b occurs in KCTD8 KO mice. Our initial results from WT pre-embedding EM indicate that KCTD12b displays peak densities in the active zone whereas KCTD8 shows peak densities in the perisynaptic region. Therefore, although it may be possible to see a compensatory increase in KCTD12b in the active zone of KCTD8 KO mice, due to the weaker expression of KCTD8 in the WT active zone, such an increase would be expected to be smaller than the compensatory increase of KCTD8 seen in KCTD12b KO animals. Furthermore, the functional consequence of a compensatory increase in KCTD12b in the active zone in KCTD8 KO mice would not be detected in functional experiments, as KCTD12b did not alter Cav2.3 function in vitro and KCTD12b has no effects on PPR in the absence of KCTD8 (no significant difference between KCTD8 KO and KCTD8/12b double KO in Figure 6A). Our results suggest that the relatively mild reduction in release probability seen in KCTD8 KO mice resulted from removal of the comparatively low levels of KCTD8 from the WT active zone, rather than from a compensatory increase in KCTD12b. We added these thoughts to the discussion on page 21, starting from line 439.

Figure 2 – it would be useful to test for cross-labelling to evaluate any compensatory expression for the loss of particular KCDT subtype with another KCDT subtype at the level of light microscopy.

We were unable to detect significant differences in fluorescence intensities for KCTD8 in KCTD12b KO animals. This might have been due to potential postsynaptic expression of KCTD8 as well as difficulty to detect nanoscale changes in protein distribution on the level of light microscopy. Instead, to confirm the compensatory increase of KCTD8 in ventral MHb terminals in the absence of KCTD12b, we performed pre-embedding EM immunolabeling for KCTD8 in presynaptic terminals in rostral and lateral IPN of KCTD12b KO mice (Figure 7C). The distribution pattern of KCTD8 in the lateral IPN showed peak densities in the peri-synaptic region, as seen in WT (Figure 3C). In contrast in the rostral IPN, the density of KCTD8 particles in the active zone, but not in the peri- and extrasynaptic area, was significantly higher compared with lateral IPN synapses (main effect of presynaptic distribution: F4, 775 = 31.65, P < 0.0001; rostral AZ vs. lateral AZ: P < 0.0001; two-way ANOVA with Bonferroni post hoc test). This observation confirmed that KCTD8 compensates for the loss of KCTD12b selectively in the active zone of ventral MHb terminals. This was added to the Results section on page 16, starting from line 329.

Figure 6A. Regarding the representative plot for the double KO mice, it is worrisome that there is a drastic increase in the baseline prior to baclofen addition, such that it is difficult to interpret the rest of the data points.

We apologize for unfortunate choice in the example time course for the double KO. Upon close inspection of the example time course in question, we realized that the baseline prior to baclofen was actually stable with the exception of the first minute of the recording (min 2 – 5 showed no tendency to ramp up). We therefore excluded the first 5 sweeps of this baseline recording. Using linear regression, we confirmed that the slope of the regression line over the remaining baseline points was not significantly deviating from 0 (P = 0.1318). We updated the corresponding relative values which were included in the graph, now moved to Figure 6—figure supplement 1B.

Does baclofen still potently increase EPSC amplitudes in KCTD8, KCTD12b KO or KCTD8/12b DKO mice if recordings are carried out in 4 mM extracellular Ca instead of 2.5 mM?

Although we did not test the effect of baclofen at higher Ca2+ concentrations in KCTD KO mice, we think it is a likely possibility that potentiation still occurs at increased external Ca2+ levels. However, since other reviewers felt the paper suffers from focusing too much on GBR-mediated potentiation instead of the role of KCTDs in basal release, we moved the baclofen experiments in KCTD KO mice from the main text to the supplement, and completely removed the analysis and discussion of alterations in potentiation time course by KCTDs and increased Ca2+ concentrations. We will deal with the mechanism of GBR-mediated potentiation in a separate paper.

Figure 7. To determine the nature of the increase in the active zone expression of KCTD8 in KCTD12b KO terminals, one should perform (1) the distance-dependent analysis of immunogold labelling for KCTD8 in KCTD12b KO preparation as shown for Figure 3, especially with respect to whether the profile of KCTD8 changes to what is shown for KCTD12b in rostral IPN, and (2) analysis at the level of light microscopy to assess whether there is an increase in the level of expression or KCTD8 in KCTD12b KO rostral IPN (i.e. the same comment as for Figure 2 above).

Please see our detailed answer to point 5. In short, we performed the suggested measurements of pre-embedding labeling and found a significant increase in KCTD8 densities in the AZ of rostral but not lateral terminals in KCTD12b KO mice.

Figure 8. The relative levels of surface expression of KCTD8 and KCTD12b in HEK cells needs to be quantified in order to fully interpret the preferential increase in Cav2.3-mediated currents in KCTD8-transfected cells.

To address this issue, we performed a plasma membrane purification experiment to determine the relative levels of surface expression of KCTD8 and KCTD12b in HEK cells. For this we transiently transfected HEK293T cells with the pore forming α-subunit (rat Cav2.3) together with auxiliary β3 and α2δ1 subunits in order to emulate experimental conditions that are as close as possible to the stable cell line used for the electrophysiology experiments in Figure 2F – H. These cells were then subjected to hypotonic lysis to separate cytosolic proteins from membrane proteins (a fraction that we called TM or “total membrane”). A fraction of the TM was then subjected to a plasma membrane purification protocol using a commercially available plasma membrane (PM) extraction kit from Abcam (ab 65400). We decided to work with this kit due to its extensive use in peer reviewed publications (so far it has been used in 85 papers – 10 of which were published between 2019 and 2020). This kit separates proteins of the plasma membrane from proteins located on other cellular membranes or the cytosol, using a 2-phase aqueous polymer method that takes advantage of differences in surface properties of the membranes rather than size and density. The PM fraction obtained after the purification as well as the TM fraction were subsequently solubilized in detergent-containing buffer (0.5% Nonidet P-40) and loaded on 10% SDS-PAGE gels. The presence of the KCTD proteins in the corresponding tissues was determined by Western Blotting using ECL detection of HRP-coupled antibodies against Flag-tagged KCTD8 (α-KCTD8 rabbit) and Flag-tagged KCTD12b (α-Flag mouse).

The following has been added to the Results section on page 8, starting from line 171.

“Plasma membrane preparations (see Methods) revealed that in HEK293T cells co-expressing Cav2.3 and KCTDs, both KCTD8 and KCTD12b are highly and similarly enriched in plasma membrane extracts, when compared to total membrane extracts [Figure 2—figure supplement 1; KCTD8: 10.7-fold (± 1.9), KCTD12b: 12.7-fold (± 2.6), no significant difference between KCTD8 and KCTD12b, P = 0.34, unpaired t-test]. We next performed co-IP experiments using total and plasma membrane extracts of transfected HEK293T cells and found that both KCTD8 and KCTD12b associate with Cav2.3 at the plasma membrane (Figure 2E).”

The direct interaction between KCTD8 and KCTD12b with Cav2.3 should be confirmed in the brain.

We agree that it would be ideal to show the direct binding of KCTDs to Cav2.3 in the brain. Unfortunately, due to technical limitations, we realized that we are unable to perform such an experiment for two main reasons:

– The IPN is relatively small (~1 x ~1 x ~0.7 mm) and weighs approximately 0.6 – 1 mg. Based on earlier Co-IP experiments that we performed in order to look at the binding of endogenous KCTDs to other interacting proteins, such as GBRs, we estimate that it would be necessary to pool tissue from over 100 mice. This constitutes an unreasonable number of animals to perform this experiment just once, and replicates might be required to consolidate findings.

– Alternatively, a co-IP could be performed from full brain lysates but this approach comes with the drawback that Cav2.3 is expressed in many brain regions whereas KCTD8 and KCTD12b show much more restricted expression. It would be theoretically possible to perform the pull down for the KCTDs, however, this runs into the problem that endogenous Cav2.3 is notoriously difficult to detect in lysates and even more challenging to detect in pull downs as co-IPs.

Due to these challenges, we designed an alternative experiment to test the interaction of Cav2.3 with KCTDs in MHb terminals. Specifically, we tested whether antibodies bound to KCTD8 might interfere with labeling of Cav2.3 by steric hindrance in KCTD12b KO mice, where KCTD8 is strongly localized to the active zone. We hypothesized that the compensatory increase in KCTD8 in the AZ might lead to increased binding of KCTD8 to Cav2.3 and, therefore, pre-incubation with anti-KCTD8 antibody might reduce the ability of the anti-Cav2.3 antibody to bind to Cav2.3, resulting in lower labeling densities. In rostral IPN synapses, we found that pre-incubation significantly reduced Cav2.3 labeling densities in the AZ but not the peri- or extrasynaptic regions (Figure 7D; main effect of synaptic distribution: F4, 880 = 31.42, P < 0.0001; control AZ vs. pre-incubation AZ: P = 0.0006; two-way ANOVA with Bonferroni post hoc test). In the lateral IPN, pre-incubation slightly but significantly reduced Cav2.3 labeling density only in the peri-synaptic region (Figure 7E; main effect of synaptic distribution: F4, 890 = 39.01; control <50 nm vs. pre-incubation <50 nm: P = 0.0320). The peri-synaptic region is the location exhibiting peak expression for KCTD8 in dorsal MHb terminals. The significant reduction in Cav2.3 labeling density at this position therefore validates our steric hindrance approach.

These results indicate a close association between KCTD8 and Cav2.3 in situ, if not their direct binding. This point was added to the Results section on page 16, starting from line 340, and discussed on page 20, staring from line 417.

Reviewer #2:

This manuscript aims to examine the mechanism of the unusual potentiating effect of baclofen acting on presynaptic GABAB receptors in the MHb-IPN pathway. I was expecting an examination of underlying second messenger pathways, but this was not the route taken, as the authors investigating whether there is a role for the KCTD proteins which interact with GABAB receptors.

My impression of this manuscript is that by trying to understand the potentiating effect of baclofen on the synapses in question, the authors have uncovered a very interesting effect of KCTD8 potentiating CaV2.3, and an interplay between KCTD8 and KCTD12b, shown with several techniques including very nice EM images and analysis, which however is completely separate from the effect of baclofen acting on GABAB receptors.

In trying nevertheless to combine the two stories, the authors have overcomplicated the message of their study, in my view. I would prefer to see the results separated, with the KCTD study stripped out and potentially published here, as it is novel and well performed. Otherwise it seems to me the manuscript is over-complex and the very interesting message about the KCTDs is difficult to extract.

We appreciate very much this advice and fully agree with the reviewer’s suggestion. Accordingly, we have re-structured our manuscript mostly focusing on the role of KCTDs on the Cav2.3-mediated basal neurotransmission in the MHb-IPN pathway. We will deal the mechanism of GABAB receptor-mediated potentiation in a separate paper.

1. Line 99 SNX482 is not "selective". The authors are using it at 1 µM. It may be relatively selective between calcium channels, but it also blocks certain K channels in the same concentration range, in fact 500 nM was used in slices to block A-type K currents (Kimm and Bean, 2014). It also affects other Ca channels and Na channels at concentrations below 1µM (Newcomb et al., 1998). How sure are the authors this blockade of other channels is not affecting their study ?

Thank you for highlighting this important point. We agree that the concentration that we used in our acute slice preparation is higher than the concentrations used in the papers cited by the reviewer, which used either cell culture (Kimm and Bean, 2014) or oocyte preparations (Newcomb et al. 1998). Although slices were cut in the paper by Kimm and Bean 2014, they were only used to extract substantia nigra cells to prepare primary neuronal cultures, not for pharmacological experiments. An important consideration, in this respect, is the fact that SNX-482 is a peptide which may easily adhere to surfaces such as biological membranes, thereby potentially reducing its effective concentration considerably, particularly in acute brain slices. Use of SNX-482 in cell culture or oocytes naturally does not run into such issues as the pharmacological targets are readily exposed to the extracellular solution containing the drug. To reduce loss of SNX-482 as much as possible, we added 0.1 % BSA to the ACSF but it is still likely that the effective SNX-482 concentration inside the acute slice at the site of active terminals was considerably lower than 1 µM.

In addition, a block of A-type K+ channels would be expected to increase membrane excitability and/or action potential half-width, both of which would result in increased neurotransmitter release. However, we did not observe increases in EPSC amplitudes in any of the SNX experiments, suggesting that the observed decrease in EPSC amplitudes by the application of SNX-482 was not mediated by an inhibition of A-type K+ channels.

To test whether the decrease in EPSC amplitude caused by the application of SNX-482 resulted from a block of voltage-gated Na+ or K+ channels, we measured action potential properties in ventral MHb neurons in ACSF containing 0.1% BSA and in ACSF containing 0.1% BSA + 1 µM SNX-482. We defined the action potential peak time as the time between the threshold potential and the action potential peak. Action potential full-width at half maximum was measured as the time between the rising and falling phase of the action potential at half of the maximal action potential amplitude. SNX-482 did not significantly affect action potential peak times or full-widths at half maximum in ventral MHb neurons (Figure 1—figure supplement 1A; peak times: control: 0.86 ± 0.03 ms, n = 3; SNX-482: 0.93 ± 0.05 ms, n = 4; t5 = 1.083, P = 0.3282; full-widths: control 1.08 ± 0.07 ms, n = 3; SNX-482: 1.06 ± 0.03 ms, n = 4, t5 = 0.3852, P = 0.7159; unpaired t-test), suggesting that the reduction of neurotransmitter release by SNX-482 was not caused by a blockade of voltage-gated Na+ or K+ channels. This part was added to the Results section on page 6, starting from line 107.

2. Line 104. You cannot say the two MHb-IPN pathways have different sensitivities to SNX482, as no dose response curve has been performed, and the difference between the pathways at the single dose studied may indicate the involvement of other Ca channels.

We apologize for the poor wording. We removed this sentence.

3. Similarly, in the Discussion, line 310 "We confirmed that both dorsal MHb-IPN and ventral MHb-IPN pathways require Cav2.3 for transmitter release", is not completely correct, as how is release occurring for the ~50% SNX482-insensitive lateral epscs (Figure 1) ?

We changed this sentence to: “We confirmed that Cav2.3 is involved in mediating transmitter release at both MHb-IPN pathways, but only release at the ventral pathway exclusively relies on Cav2.3.” page 17, starting from line 364.

4. Figure 8A needs correct labelling of the panels, with MW markers. I assume the top panel is KCTDs, but this is unclear as both CaV2.3 and KCTDs have Flag tags, which is an unusual way of performing the experiment. The entire blot should be shown for the top panel, as blotting with Flag would show both KCTDs and CaV2.3.

We apologize for this oversight and added MW markers to the co-IP blots, now moved to Figure 2. Unfortunately, due to substantial differences in molecular weight, KCTDs and Cav2.3 cannot be run on the same gels because the high molecular weight of Cav2.3 requires a less dense gel which would not allow for adequate band separation of the significantly smaller KCTDs, while in gels with higher density that allow us to better visualize the size differences between the individual KCTDs, large molecules like Cav2.3 get trapped at the pore openings. Furthermore, since KCTDs are expressed stronger than Cav2.3, two separate gels were necessary to avoid overexposure of KCTD bands.

5. If there is co-ip between CaV2.3 and KCTD8/12b, should there be actual co-localization observed in the EM images? Can this be measured?.

The reviewer raises a very important consideration. In freeze-fracture replica labeling, proteins that are closely associated with the membrane, such as transmembrane proteins or membrane-anchored proteins, get trapped in a carbon layer and remain there even after all remaining tissue is digested by SDS. Therefore, KCTDs, which are cytosolic proteins, are undetectable in replica unless they are covalently bound by chemical fixation to another membrane–integrated protein and getting trapped together in the carbon. As we readily detect them at high densities in the active zone, it very likely that the vast majority of detected KCTDs in replica are actually bound to either GBRs or Cav2.3.

Another important aspect is that it is difficult to label multiple proteins that are bound to each other in a complex because a) the protein complex might mask epitopes and b) primary/secondary antibodies with gold conjugates might hamper the binding of additional antibodies to another protein in the same complex in a process called steric hindrance.

Nevertheless, we made use of this steric hindrance effect to confirm the co-localization of KCTD8 and Cav2.3 in the active zone. We found that pre-incubation with anti-KCTD8 antibodies in KCTD12b KO mice led to a reduction in Cav2.3 labeling densities in the active zone of only ventral but not lateral MHb terminals. Although this does not prove that the two proteins are directly bound to each other, it suggests that they are in close proximity (<16 nm, two times IgG size of 8 nm) to each other. The following part was added to the Results section on page 16, starting from line 340:

“In order to examine further the spatial proximity of Cav2.3 to KCTD8, we tested whether steric hindrance of antibodies bound to KCTD8 might interfere with labeling of Cav2.3. We hypothesized that the compensatory increase in KCTD8 in the AZ might lead to increased binding of KCTD8 to Cav2.3 and, therefore, pre-incubation with anti-KCTD8 antibody might reduce Cav2.3 labeling in the AZ. In rostral IPN synapses, we found that pre-incubation significantly reduced Cav2.3 labeling densities in the AZ but not the peri- or extrasynaptic regions (Figure 7D; main effect of synaptic distribution: F4, 880 = 31.42, P < 0.0001; control AZ vs. pre-incubation AZ: P = 0.0006; two-way ANOVA with Bonferroni post hoc test). In the lateral IPN, pre-incubation significantly reduced Cav2.3 labeling density only in the peri-synaptic region (Figure 7E; main effect of synaptic distribution: F4, 890 = 39.01; control <50 nm vs. pre-incubation <50 nm: P = 0.0320).”

6. In Figure 8c, how selective is the augmentation with KCTD8 and CaV2.3, ie does it also occur for any other Ca channels? It is possible that a low level of other presynaptic Ca channels might be also augmented by KCTD8 in these terminals, increasing Ca2+ entry.

This is an interesting question and we do not have a definite answer to it. However, we did not observe any EPSC responses in rostral IPN neurons in Cav2.3 KO mice, suggesting that Cav2.3 is the sole Ca2+ channel mediating release at this pathway. Although the increase in release probability at ventral MHb terminals is likely caused by a direct effect of KCTD8 on Cav2.3, it is unknown whether other Ca2+ channel subtypes involved in transmitter release from dorsal MHb terminals are equally affected by KCTD8. We added this to the discussion on page 20, starting from line 435.

7. If this large augmentation of CaV2.3 currents by KCTD8 plays a physiological role should the epsc's in the KCTD8 KO mice be reduced? I cannot see this basic result, although I note the release probability is indeed reduced.

This is an important question. It is generally difficult to quantify absolute amplitudes of electrically evoked EPSCs as they depend largely on the stimulation intensity. Therefore, we used the variance mean analysis to show the reduced release probability in KCTD8 KO mice. An important indication that KCTD8 functionally increases synaptic strength was the observation that viral overexpression of KCTD8 in WT ventral MHb neurons significantly lowered PPR values compared with EGFP controls. This result suggests that KCTD8 and KCTD12b may be in a state of dynamic balance. Creating an imbalance appears to result in alterations in synaptic strength. The new data showing the effect of viral overexpression of KCTD8 on PPR was added to the Results section on page 14, starting from line 288 and discussed on page 20, starting from line 427.

8. Following on from point 6, in Discussion line 329, has the 1997 study by Ludwig et al. showing no α1B, A, C mRNA in the habenula been replicated anywhere? The technique used in this early study is not very sensitive, are there other more recent studies using single cell RNAseq for example?

Unfortunately, we did not find a more recent study investigating the expression of different Cav2 α subunits in MHb neurons. However, according to the Allen Brain Atlas, Cav2.1 and Cav2.2 are not expressed in MHb neurons. We included this in the supplement as Figure 8—figure supplement 1 and added this information to the discussion on page 18, starting from line 382.

9. Discussion line 330 "Although this exclusivity has been described anatomically (Parajuli et al., 2012)" is not correct as it gives the impression other Ca channels were examined in this study, whereas this is not the case.

We changed this sentence to: “Although the predominant presynaptic localization of Cav2.3 immunoreactivity was previously reported (Parajuli et al. 2012), the functional implications of this localization were not determined.” page 18, starting from line 385.

10. Discussion line 331: SNX482 is here described as specific, which is incorrect.

We removed the word “specific”.

Reviewer #3:

The manuscript attempts to elucidate the mechanisms underlying the potentiation of synaptic transmission between the ventral medial habenula (MH) and the rostral interpeduncular nucleus mediated by presynaptic GABA-B receptors. The topic is interesting especially because it is very unusual that GABA-B receptors gate an enhancement of neurotransmission, since usually when they are activated, they depress the release of neurotransmitter.

The manuscript contains some high quality data, for example, freeze-fracture replica immunolabelling of presynaptic proteins in habenular terminals. However, I also found some technical problems, (e.g., see comment #1 below). Overall, the manuscript is potentially interesting and covers an original topic. However, the data currently submitted fail to answer the two major questions, namely the underlying mechanisms and the physiological relevance of GABA-B receptor-mediated modulation. A lot of efforts are devoted in characterizing key protein expression in various K+ channel tetramerization domain-containing (KCTD) proteins. However, I doubt that this strategy paid off and really shed light on the mechanisms underlying GABA-B receptor mediated modulation of the synapses studied.

We agree with the reviewer that we failed to elucidate the mechanisms of GABA-B receptor-mediated modulation in our previous manuscript. Also, according to suggestion by the reviewer #2 (please see above), we have re-structured our manuscript mostly focusing on the role of KCTDs on the Cav2.3-mediated basal neurotransmission in the MHb-IPN pathway. We will deal with the mechanism of GABAB receptor-mediated potentiation in a separate paper.

1. Data on the action of SNX-482 on evoked EPSCs need refinement. The presynaptic fibers of the fasciculus retroflexus were electrically stimulated and recordings were made in the rostral IPN. In contrast, stimulation of dorsal fibers were obtained via optogenetic stimulation and responses were recorded in the lateral IPN. The effect of SNX-482 was stronger at the former versus the latter synapses. However, this result is at odds with similar Cav2.3 densities in the active zones of dorsal and ventral MHb terminals. The Authors discussed this issue, and I acknowledged this mis-match in their data. Nevertheless, the problem remains: a quantitative comparison between drug modulation of synaptic responses evoked by electrical versus optogenetic stimulation is a poor choice. Amongst various problems, several recent papers report the non-physiological high release probability often caused by optogenetic stimulation of presynaptic fibers that contrasts with the electrical stimulation method. This discrepancy would affect the results obtained with SNX-482. This set of experiments should be repeated by using more uniform experimental conditions in the two synaptic pathways.

We agree that using two separate ways of stimulation for the two pathways was not ideal. We initially used optogenetics to study lateral IPN inputs because it was challenging to keep the fiber tract intact all the way to the lateral IPN. This is due to the fact that although MHb axons project linearly from the MHb to the IPN, they enter the IPN from the anterior side. Thus, to solve this problem, we have newly established thick-slice preparations (1 mm) cut at a 54° angle, which keep the most anterior portion of the IPN intact within the slice and allow reliable stimulation of the fasciculus retroflexus (FR). In this preparation we stimulated FR just below the MHb, 2 – 3 mm distal to the IPN, and could obtain EPSC responses in both rostral and lateral IPN neurons (see Figure 1—figure supplement 1B – D). We used a 10 Hz stimulus train rather than a single stimulus in order to not miss potentially weaker responses due to the absence of Cav2.3. The following was added to the Results section on page 6 starting from line 112:

“To corroborate the involvement of Cav2.3 in release from MHb terminals, we next performed electrical stimulation of the MHb-derived fiber tract, the fasciculus retroflexus (FR), in Cav2.3 KO mice and recorded EPSCs in rostral and lateral IPN neurons. To obtain maximally intact FR, slices were cut at 1 mm thickness which allowed for the stimulation of the fiber tract at a distance of 2 – 3 mm from the IPN (Figure 1H, I). […] Overall, these results suggest that ventral MHb terminals rely exclusively on Cav2.3 for release, whereas dorsal MHb terminals rely on Cav2.3 and additional Ca2+ channels.”

Furthermore, a serious comparison of the efficacy of SNX-482 should rely on testing at least three different drug concentrations for each synapse. Furthermore, normalization assuring that approximatively an equal number of presynaptic fibers are stimulated at each synaptic connection should be employed. What ever methods of presynaptic stimulation will be used, optical or electrical, test of selective stimulation of fibers should be also performed.

As described in our response to the reviewer’s initial concern (1.), we tested the effect of SNX-482 using fiber tract electrical stimulation in WT mice and directly compared EPSC responses in rostral and lateral IPN neurons in the same slice of Cav2.3 KO mice using electrical fiber tract stimulation. Stimulation selectivity was confirmed by application of baclofen in WT slices (Figure 1—figure supplement 1C, D), which potentiated EPSCs when recording from rostral IPN neurons and inhibited EPSC when recording from lateral IPN neurons. We also compared the time courses of SNX-482-mediated EPSC reductions in lateral IPN for electrical and optogenetic stimulation in Figure 1L, confirming no significant difference between types of stimulation and no effect of Conotoxin application on EPSC amplitudes. This point was also added to the Results section on page 7, starting from line 136.

2. Baclofen still presynaptically potentiated EPSCs in KCTD KO mice. However, loss of KCTD12b impaired GABABR action probably via increasing release probability and occlusion. In contrast, lack of KCTD8 reduces release probability. But what about the effects of GABABR modulation? Is baclofen more powerful at MH synapses in KCTD8 KO?

We did not find a significant difference in baclofen-mediated potentiation between WT and KCTD8 KO mice, suggesting that the relatively minor reduction in release probability in KCTD8 KO mice did not affect GBR-mediated enhancement of release. Since the reviewers felt that our manuscript focused too much on GBR-mediated enhancement instead of the effect of KCTDs on basal release, we removed discussion and analysis of the potential connection between basal release and baclofen-mediated enhancement. We now only mention that GBR-mediated potentiation was still intact in all KCTD KO mice and moved the corresponding result to the supplement (Figure 6—figure supplement 1).

3. The manuscript lacks to experimentally demonstrate that GABA-B receptor potentiation of ventral MH and the rostral interpeduncular nucleus synapses is physiologically relevant. This should be addressed by using physiologically salient repetitive stimulation in control and in the presence of GABA-B R antagonists. It would be also interesting to test the GABA-B R ligands on evoked EPSPs followed by IPSPs to see the comprehensive pharmacological effects on both excitation and inhibition. At present none of these important points are addressed and this reduces the impact of the data currently submitted.

We were unfortunately unable to unravel the exact mechanism underlying the GBR-mediated enhancement of release in our original manuscript but instead found a surprising involvement of KCTDs with Cav2.3 that modulate basal release. As the reviewers suggested, we therefore shifted our focus away from potential GBR mechanisms, and now mostly focus on the role of KCTDs in basal release.

[Editors’ note: what follows is the authors’ response to the second round of review.]

The revised manuscript has been significantly improved by refocusing the study on the role for KCTDs on neurotransmitter release. Several additional experiments have been performed, including the comparisons of Cav2.3-dependence of synaptic transmission between the two distinct medial habenula to IPN inputs by electrical stimulation, which has been made possible by the establishment of thick-slice preparation. New data lend further support to the conclusion that KCTD8 and KCTD12b directly interact with Cav2.3 and regulate release from ventral MHb terminals independently of GABAB receptors, where effectively KCTD12b serves to restrain release while the recruitment of KCTD8 to the active zone potentiates release by promoting Cav2.3 currents. Prior to accepting the manuscript for publication, a number of points require attention.

Figure 6F-G: the EPSC waveforms of some rescued currents appear to be quite different than one would expect. Specifically, the LV-hSyn-KCTD8 current is quite prolonged, and may have two components to the rising phase.

We agree that the LV-hSyn-KCTD8 trace appears slightly broader than the control trace. Upon re-examination of the other recordings in this group, we did not consistently observe broader responses in KCTD8 overexpression experiments and therefore, replaced the corresponding example trace with one showing a more representative decay time.

Additionally, the LV-hSyn-KCTD12b current appears to have a delayed onset. Are these examples representative of the population? If so, are these effects a result of over-expression, or more an indicator of the distinct roles of these two KCTD proteins?

Concerning the slightly delayed onset in the KCTD12b rescue trace: This delay might be influenced by the position of the stimulating electrode relative to the recorded neuron. Inside the IPN, habenula-derived axons can be unmyelinated and therefore responses may occur slightly faster or slower depending on this distance. Although we try to keep recording conditions similar for all experiments, random biological variation cannot be completely controlled. Upon re-examination of the other recordings, we did not consistently observe delayed responses in LV-hSyn-KCTD12b injected mice, suggesting a KCTD-independent effect, and we replaced the example trace with one showing a more representative response onset.

In the introduction (Line 47 or Line 53-54), it would be clearer to readers who may not necessarily be familiar with KCTDs if the authors could briefly state the nature of the molecular difference especially between KCTD12 and KCTD12b.

In accordance with this suggestion, we rewrote the second paragraph of the introduction:

“Four KCTD subunits, KCTD8, 12, 12b and 16, serve as auxiliary GBR subunits (Schwenk et al., 2010). KCTD subunits bind as hetero- and homo-pentamers to GBRs and modulate their signaling kinetics (Fritzius et al. 2017; Zheng et al., 2019). Structurally, KCTD subunits are composed of an N-terminal T1 domain and an H1 domain. […] However, the functional consequences of these interactions and whether other voltage-gated Ca2+ channels interact with KCTDs remain unknown.”

Line 302-304. "The decrease in release probability by KCTD12b in the presence but not in the absence of KTCD8…" is confusing. Do the authors mean that the reason why the decreased PPR observed in KCTD12bKO could be recovered to control levels when KCTD8 is additionally absent (as observed in double KO mice), could be because the loss of KCTD12b in itself alters the localization of KCTD8 or Cav2.3? Perhaps this sentence should be rephrased.

We apologize for the confusing wording, we rephrased the sentence as follows: “The decreased PPR in the absence of KCTD12b recovered to control levels when KCTD8 was additionally absent suggesting alterations in either Cav2.3 or KCTD8 localization in the active zone of MHb terminals in KCTD12b KO mice.”

https://doi.org/10.7554/eLife.68274.sa2

Article and author information

Author details

  1. Pradeep Bhandari

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Data curation, Formal analysis, Investigation, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  2. David Vandael

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Data curation, Formal analysis, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
  3. Diego Fernández-Fernández

    Department of Biomedicine, University of Basel, Basel, Switzerland
    Contribution
    Data curation, Formal analysis, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-1431-3705
  4. Thorsten Fritzius

    Department of Biomedicine, University of Basel, Basel, Switzerland
    Contribution
    Data curation, Formal analysis, Investigation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3597-6623
  5. David Kleindienst

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Software, Formal analysis, Writing - review and editing
    Competing interests
    No competing interests declared
  6. Cihan Önal

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Data curation, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2771-2011
  7. Jacqueline Montanaro

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
  8. Martin Gassmann

    Department of Biomedicine, University of Basel, Basel, Switzerland
    Contribution
    Resources, Writing - review and editing
    Competing interests
    No competing interests declared
  9. Peter Jonas

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Resources, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5001-4804
  10. Akos Kulik

    1. Institute of Physiology II, Faculty of Medicine, Freiburg, Germany
    2. BIOSS Centre for Biological Signalling Studies, University of Freiburg, Freiburg, Germany
    Contribution
    Resources, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared
  11. Bernhard Bettler

    Department of Biomedicine, University of Basel, Basel, Switzerland
    Contribution
    Resources, Supervision, Funding acquisition, Writing - review and editing
    Competing interests
    No competing interests declared
  12. Ryuichi Shigemoto

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    ryuichi.shigemoto@ist.ac.at
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8761-9444
  13. Peter Koppensteiner

    Institute of Science and Technology Austria (IST Austria), Klosterneuburg, Austria
    Contribution
    Conceptualization, Data curation, Formal analysis, Investigation, Writing - original draft, Writing - review and editing
    For correspondence
    peter.koppensteiner@ist.ac.at
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3509-1948

Funding

H2020 European Research Council (694539)

  • Ryuichi Shigemoto

H2020 European Research Council (692692)

  • Peter Jonas

Schweizerischer Nationalfonds zur Förderung der Wissenschaftlichen Forschung (31003A‐172881)

  • Bernhard Bettler

H2020 Marie Skłodowska-Curie Actions (665385)

  • Cihan Önal

Deutsche Forschungsgemeinschaft (For2143)

  • Akos Kulik

Deutsche Forschungsgemeinschaft (BIOSS-2)

  • Akos Kulik

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are grateful to Akari Hagiwara and Toshihisa Ohtsuka for CAST antibody, and Masahiko Watanabe for neurexin antibody. We thank David Adams for kindly providing the stable Cav2.3 cell line. Cav2.3 KO mice were kindly provided by Tsutomu Tanabe. This project has received funding from the European Research Council (ERC) and European Commission (EC), under the European Union’s Horizon 2020 research and innovation programme (ERC grant agreement no. 694539 to Ryuichi Shigemoto, no. 692692 to Peter Jonas, and the Marie Skłodowska-Curie grant agreement no. 665385 to Cihan Önal), the Swiss National Science Foundation Grant 31003A‐172881 to Bernhard Bettler and Deutsche Forschungsgemeinschaft (For 2143) and BIOSS-2 to Akos Kulik.

Ethics

Animal experimentation: All experiments were performed in accordance with the license approved by the Austrian Federal Ministry of Science and Research (Animal license number: BMWFW-66.018/0012-WF/V/3b/2016) and the Austrian and EU animal laws.

Senior Editor

  1. John R Huguenard, Stanford University School of Medicine, United States

Reviewing Editor

  1. Yukiko Goda, RIKEN, Japan

Reviewer

  1. Marco Capogna, University of Aarhus, Denmark

Publication history

  1. Received: March 10, 2021
  2. Accepted: April 28, 2021
  3. Accepted Manuscript published: April 29, 2021 (version 1)
  4. Version of Record published: May 14, 2021 (version 2)

Copyright

© 2021, Bhandari et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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