We first examined the expression of latrophilins in the cerebellum. The expression of latrophilins is developmentally regulated with peak expression observed at postnatal day P10, therefore we probed cerebellar sections at P10, P30 and P60 to cover latrophilin expression during synaptogenesis and in adult brain (Sando et al., 2019). Single molecule RNA fluorescent in situ hybridizations (smRNA-FISH) showed that the three latrophilin genes are differentially expressed in cerebellum, but in overlapping patterns (Figure 1). All latrophilins are expressed in cerebellar cortex; in addition, Lphn1, Lphn2 and to a lesser extent Lphn3 are expressed in deep cerebellar nuclei (Figure 1A). In the cerebellar cortex, Lphn1 is primarily expressed in granule cells, Lphn2 in Purkinje cells, and Lphn3 in Bergmann glia (that are also localized to the Purkinje cell layer) and in interneurons (Figure 1B and C). Quantification of Lphn RNA puncta in Purkinje cell bodies revealed that transcripts for all Lphn isoforms are found in Purkinje cells and that Lphn1 and Lphn2 puncta were more abundant than Lphn3 puncta (Figure 1—figure supplement 1).

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In situ hybridizations identify key cell types with high-level gene expression, but can overlook lower levels of expression that might still be significant. To probe the cell-type specific expression of latrophilins by an independent method, we crossed Lphn2 and Lphn3 double cKO mice (Sando et al., 2019; Anderson et al., 2017) with five different Cre-driver mouse lines: Nestin-Cre mice that express Cre recombinase in the precursor cells which generate all cerebellar cell types, L7/Pcp2-Cre mice that express Cre recombinase only in Purkinje cells, PV-Cre mice that express Cre recombinase in parvalbumin-expressing cerebellar cells (Purkinje cells, inhibitory interneurons, and neurons of the deep cerebellar nuclei), Math1/Atoh1-Cre mice that express Cre recombinase in granule cells, and GLAST/Slc1a3-CreER mice that express tamoxifen-inducible Cre recombinase in Bergmann glia (Figure 2A and B). For the GLAST-CreER mice, Cre activity was induced by intraperitoneal injections of 4-hydroxytamoxifen on postnatal days P10, P11, and P12. All mice were analyzed on day P35. Lphn2 and Lphn3 protein levels were measured in the cerebellum by quantitative immunoblotting, using antibodies to HA and to GFP since the cKO mice express Lphn2 with an mVenus tag (Anderson et al., 2017) and Lphn3 with an HA-epitope (Sando et al., 2019).

Figure 2

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Nestin-Cre nearly completely ablated Lphn2 and Lphn3 expression as expected (Figure 2C). L7-Cre had no significant effect on the levels of either Lphn2 or Lphn3, although there was a trend towards a decrease for Lphn2 (Figure 2D). This result seems to contradict the in situ data showing abundant expression of Lphn2 in Purkinje cells (Figure 1B and C). However, despite their remarkable size and prominence, the number of Purkinje cells is much lower than that of other cerebellar neurons (e.g., granule cells, interneurons, and deep cerebellar nuclei neurons), and the selective loss of Lphn2 and Lphn3 from Purkinje cells may have been occluded by the combined levels of these proteins in other neurons, in particular granule cells and deep cerebellar nuclei neurons. Consistent with this conclusion, both Lphn2 and Lphn3 were significantly decreased in PV-Cre mice that express Cre in deep cerebellar nuclei neurons and interneurons in addition to Purkinje cells, with Lphn2 expression reduced by nearly 80% (Figure 2E). Math-Cre, conversely, had no significant effect on the levels of Lphn2 and Lphn3, consistent with the fact that granule cells primarily express Lphn1 (Figure 2F). GLAST-Cre-mediated deletion of Lphn2 and Lphn3 in Bergmann glia, finally, did not detectably alter the levels of Lphn2, but dramatically decreased the levels of Lphn3, indicative of substantial Lphn3 expression in these astrocytes (Figure 2G).

Since latrophilins are postsynaptic adhesion molecules and Purkinje cells are the postsynaptic target of almost all synapses in the cerebellar cortex, we focused our analyses of the cerebellar role of latrophilins on Purkinje cells.

First, we examined individual Lphn2 and Lphn3 cKO mice, examining parallel-fiber synapses as the most abundant cerebellar synapse. We analyzed mice obtained from crosses of Lphn2 and Lphn3 cKO mice with L7-Cre mice and compared Cre-negative homozygous Lphn2 or Lphn3 cKO mice to Cre-positive littermate mice to assess the effects of the Lphn deletion. To estimate parallel-fiber synapses morphologically, we stained cerebellar sections for vGluT1. Because parallel-fiber synapses are too abundant in the cerebellar cortex to resolve individual vGluT1-positive puncta by confocal microscopy, we measured the overall vGluT1 staining intensity, which depends on both the density and the size of parallel-fiber synapses (Zhang et al., 2015). We observed no significant difference in vGluT1 staining intensity between sections from control mice and mice lacking either Lphn2 or Lphn3 in the Purkinje cells (Figure 3A and E). Next, we cut acute slices from the mice and used whole-cell patch-clamp recordings to measure the input resistance and capacitance of Purkinje cells, but again detected no difference between control and Lphn2- or Lphn3-deficient Purkinje cells (Figure 3B and F). Finally, we monitored parallel-fiber EPSCs induced by extracellular stimulation in the upper molecular layer, and determined both the paired-pulse ratio of EPSCs in response to two stimuli separated by 50 ms and the overall synaptic strength quantified by the input-output relation and the rise slope of EPSCs (Figure 3C–D and G–H). Here, we also observed no differences between Purkinje cells expressing or lacking Lphn2 or Lphn3, suggesting that each isoform alone is not essential for proper parallel-fiber synapse numbers or parallel-fiber synaptic transmission.

Figure 3

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To address the potential for redundancy among Lphn2 and Lphn3, we generated double cKO mice in which both Lphn2 and Lphn3 are deleted in Purkinje cells by an L7-Cre allele. We stained cerebellar sections from littermate control and double cKO mice for vGluT1 to examine parallel-fiber synapses, for vGluT2 to assess climbing-fiber synapses and for vGAT to monitor inhibitory basket-cell and stellate-cell synapses (Figure 4). Strikingly, we found that the Lphn2/3 double-deficient sections displayed a ~ 30% decrease in vGluT1 staining intensity, suggesting a decrease in parallel-fiber synapses (Figure 4A). The density of vGluT2-positive puncta and of vGAT-positive puncta was unchanged (Figure 4B and C). The two types of excitatory inputs onto Purkinje cells can be distinguished anatomically as parallel fibers form synapses onto secondary and tertiary dendrites while climbing-fibers form synapses onto primary dendrites that are closer to the Purkinje cell soma. In support of a reduction of parallel-fiber but not climbing-fiber synapse number, we stained Lphn2/3 double-deficient sections for Homer1 and found that Homer1 puncta density was reduced on distal tertiary dendrites but not primary dendrites of Purkinje cells (Figure 4—figure supplement 1). In addition, we compared synaptic protein levels of cerebellar lysates from control and Lphn2/3 double-KO mice by immunoblot and found no difference in any of the proteins tested, including vGluT1 (Figure 4D). As discussed in our immunoblot analysis with the L7-Cre line in Figure 2D,a reduction in vGluT1 levels at Purkinje cell synapses in the cerebellar cortex may be obscured by abundant vGluT1 expression in other regions of the cerebellum. Overall, these results indicate that the double Lphn2/3 deletion may specifically alter parallel-fiber synapses, suggesting a redundant function of Lphn2 and Lphn3 in these synapses.

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We further tested the functional impact of the double Lphn2/3 deletion using whole-cell patch-clamp recordings from control and Lphn2/3 double KO Purkinje cells. Lphn2/3-deficient and control Purkinje cells exhibited the same capacitance and input resistance values, suggesting that the double deletion did not grossly impair the development of Purkinje cells (Figure 5A). Input-output measurements of parallel-fiber synaptic responses, however, uncovered an approximately 50% decrease in parallel-fiber synaptic strength, confirming the immunohistochemistry findings that suggest that parallel-fiber synapses are impaired by the double Lphn2/3 deletion (Figure 5B). This decrease is likely not due to a decrease in release probability, as demonstrated by a lack of change in paired-pulse ratios (Figure 5C). Analyses of climbing-fiber synaptic responses showed that the amplitude, paired-pulse ratio, and step number of climbing-fiber EPSCs (reflecting the number of climbing fibers innervating a Purkinje cell) were normal in Lphn2/3 double-deficient Purkinje cells (Figure 5D).

Figure 5

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Recordings of spontaneous mEPSCs performed in the presence of tetrodotoxin revealed no differences in the average mEPSC frequency between Lphn2/3 double-deficient and control Purkinje cells (Figure 5E–F). mEPSC rise times can be used to distinguish between events likely derived from parallel-fiber inputs that form synapses onto distal dendrites of Purkinje cells and events likely derived from climbing-fiber inputs that form synapse onto proximal Purkinje cell dendrites. The majority of mEPSCs with 10–90% rise times of less than 0.6 ms are derived from climbing fibers that are insensitive to Group III mGluR agonists, while the majority of mEPSCs with 10–90% rise times greater than 2 ms are derived from parallel-fiber synapses that are severely depressed in response to Group III mGluR agonists (Ichikawa et al., 2016). The frequency of mEPSCs with rise times less than 0.6 ms was unchanged between Lphn2/3 double cKO and control cells, whereas the frequency of mEPSCs with rise times greater than 2 ms was reduced by ~25%, a difference that was almost statistically significant (p=0.07). The frequency of mEPSCs with rise times between 0.6 ms and 2 ms, which reflects a mixed population of parallel-fiber- and climbing-fiber-derived events, was only slightly reduced in Lphn2/3 deficient Purkinje cells. A significant difference in the cumulative frequency distribution of mEPSC amplitudes was observed despite minimal difference in average mEPSC amplitude after the Lphn2/3 deletion (Figure 5G). The average 10–90% rise times of mEPSCs was significantly reduced in Lphn2/3 deficient Purkinje cells compared to control, however, the average rise times for both the <0.6 ms and >2 ms subgroups of mEPSCs were unchanged, suggesting that the reduction in average mEPSC rise time was the result of the altered proportion of parallel-fiber to climbing-fiber events after Lphn2/3 double KO rather than a change in intrinsic mEPSC properties of either synaptic input (Figure 5H). Recordings of spontaneous miniature inhibitory postsynaptic currents (mIPSCs) revealed no changes in mIPSC amplitude or frequency between control and Lphn2/3-deficient Purkinje cells (Figure 5I). Overall, these data indicate that the double deletion of both Lphn2 and Lphn3, as opposed to the single deletion of each latrophilin isoform individually, causes a selective and robust impairment of parallel-fiber synapses.

Given the selective high-level expression of Lphn3 in Bergmann glia in the cerebellum we observed by in situ hybridization analyses (Figures 1 and 2) that is consistent with single-cell RNAseq data (Saunders et al., 2018; Zeisel et al., 2018; Zhang et al., 2014; Koirala and Corfas, 2010), we asked whether Lphn3 might have a synaptic organizing function in these cells. This question was prompted by our unpublished finding that the expression of neuroligin-3, another postsynaptic adhesion molecule, in Bergmann glia appears to be essential for the function of parallel-fiber synapses (Zhang, B. and Südhof, T.C., submitted). Thus, we used Lphn3 cKO mice crossed with GLAST-CreER mice to delete Lphn3 from astrocytic Bergmann glia in the cerebellum, employing daily 4-OHT injections from age P10-P12 to induce Cre recombination (Figure 6A). This manipulation caused a major decrease in Lphn3 protein levels as determined by immunoblot in total cerebellum (Figure 2G). Consistent with this observation, we observed a similar decrease in Lphn3 staining intensity (visualized by the HA-epitope knock-in) in the cerebellar cortex after deletion of Lphn3 in Bergmann glia (Figure 6B). These results indicate that the majority of Lphn3 in cerebellar cortex is expressed by Bergmann glia, and that despite the functional importance of Lphn3 in Purkinje cells as described above, the Lphn3 levels in Purkinje cells are much lower than those in Bergmann glia.

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We first examined markers for excitatory parallel-fiber and climbing-fiber synapses as well as for inhibitory synapses to determine if Lphn3 deletion from Bergmann glia impairs synapses of the cerebellar cortex (Figure 6C–E). As above, we quantified parallel-fiber synapses by overall vGluT1 staining intensity because the density of parallel-fiber synapses is too high to enable resolution of individual puncta by confocal microscopy. The resulting vGluT1 staining intensity reflects a combination of parallel-fiber synapse density and size (Zhang et al., 2015). The deletion of Lphn3 from Bergmann glia had no effect on this parameter (Figure 6C). We then quantified the density, staining intensity, and size of vGluT2-positive puncta that represent climbing-fiber synapses. We observed no change in climbing-fiber density, but a significant decrease in their staining intensity and size, hinting at a possible impairment (Figure 6D). Moreover, we investigated inhibitory synapses both in the upper molecular and the Purkinje cell layer that are presumably derived from stellate cells and basket cells, respectively, but again observed no significant changes (Figure 6E).

We next asked whether the deletion of Lphn3 from Bergmann glia might change the presence and composition of Bergmann glia processes in the cerebellar cortex. We used immunocytochemistry to stain the cerebellar cortex of littermate-control homozygous Lphn3 cKO mice containing or lacking a GLAST-Cre allele for three Bergmann glia-specific markers: the glutamate transporters GLAST and GLT-1, and the glutamate receptor GluA1 (Figure 6F; Rothstein et al., 1994; Bergles et al., 1997; Saab et al., 2012). For every glial marker, the staining pattern intensity were indistinguishable between sections from control mice and mice in which Lphn3 had been deleted from Bergmann glia postnatally, suggesting that Bergmann glia were unaffected by Lphn3 deletion (Figure 6F).

These experiments suggest that the Lphn3 deletion from Bergmann glia does not cause a major change in the structure and synaptic composition of the cerebellar cortex. To independently confirm this conclusion, we analyzed the cerebellum from control mice and mice with a deletion of Lphn3 from Bergmann glia by quantitative immunoblotting for a series of synaptic marker proteins (Figure 6G). We observed no changes in any synaptic protein, in particular not in vGluT1 and vGluT2 that are specific for parallel-fiber and climbing-fiber synapses, respectively, except for a decrease in the levels of vGAT that is specific for inhibitory synapses, and a trend towards a decrease for the scaffolding proteins CASK and gephyrin. These results confirm that the deletion of Lphn3 from Bergmann glia, the major cell type in cerebellum in which Lphn3 is expressed, does not cause a major change in the structure and composition of the cerebellar cortex.

In a final set of experiments we explored the possibility that the Lphn3 deletion from Bergmann glia may impair cerebellar synaptic functions even though the overall synaptic architecture of the cerebellum appears to be largely unaffected by the Lphn3 deletion. Using whole-cell patch-clamp recordings from Purkinje cells in acute cerebellar slices, we confirmed by measurements of the capacitance and input resistance that the Lphn3 deletion in Bergmann glia has no major effect on the basic properties of Purkinje cells (Figure 7A). Evoked climbing-fiber synaptic responses were unchanged in either amplitude, paired-pulse ratio or number of ‘steps’ (reflecting the number of climbing fibers innervating a Purkinje cell) after Lphn3 deletion from Bergmann glia (Figure 7B). Measurements of spontaneous miniature EPSCs and IPSCs (performed in the presence of tetrodotoxin) showed that the frequency and amplitude of mEPSCs and of mIPSCs were also unchanged (Figure 7C–D). Thus, despite high levels of Lphn3 expression in Bergmann glia, Lphn3 is not essential in this type of astrocyte for any of the major synaptic components of the cerebellar cortex.

Figure 7

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Lphn2 and Lphn3 conditional knockout (cKO) and knock-in lines were generated as described (Sando et al., 2019; Anderson et al., 2017). Other mouse lines used in this paper were L7/Pcp2-Cre (Barski et al., 2000; JAX #004146), PV-Cre (JAX #017320), Math1/Atoh1-Cre (Matei et al., 2005; JAX #011104), GLAST/Slc1a3-CreERT2 (JAX #012586), Ai14 (JAX #007914) and Nestin-Cre (Tronche et al., 1999, JAX #003771). All mice used were on a CD57BL/6 background. All procedures conformed to the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals and were approved by the Stanford University Administrative Panel on Laboratory Animal Care. All analyses of Lphn KOs were performed with male and female littermate-controlled Cre-positive and Cre-negative mice that were homozygous for single or double Lphn cKOs.

Wild type CD-1 mice were euthanized with isofluorane at P10, P30 and P60 followed by transcardial perfusion with ice cold PBS. Brains were quickly dissected and embedded in Optimal Cutting Temperature (OCT) solution on dry ice. 16 µm thick sections were cut using a Leica CM3050-S cryostat, mounted directly on to Superfrost Plus slides and stored at −80°C until use. Single-molecule FISH for Lphn1 (Cat# 319331), Lphn2 (Cat# 319341-C2) and Lphn3 (Cat# 317481-C3) mRNA was performed using the multiplex RNAscope platform (Advanced Cell Diagnostics) according to manufacturer instructions for fresh-frozen sections.

Immunoblotting was performed as described previously (Zhang et al., 2015). Littermate-controlled mice at P35 were anesthetized and cerebella were dissected out, then homogenized in lysis buffer containing (in mM): 20 Tris-HCl, pH 7.5, 100 NaCl, 4 KCl, 2 MgCl2, 2 CaCl2, 1% Triton X-100, 1x protease inhibitor cocktail (Roche) at 4°C. The samples were then centrifuged for 20 mins at 20,000 g to remove insoluble materials at 4°C. 20 μg of protein lysate was run on 4–20% SDS-polyacrylamide gels then transferred onto nitrocellulose membranes on a semi-dry transfer apparatus (BioRad Trans-Blot Turbo). Membranes were blocked in 5% milk powder in 0.05% TBS-Tween for 1 hr at room temperature (RT). Primary antibodies were diluted in blocking buffer and incubated overnight at 4°C. Membranes were then washed four times for 10 min with 0.05% TBS-Tween then incubated with fluorescence-labeled secondary antibodies (donkey anti-rabbit IR Dye 680CW, 1:10,000; donkey anti-mouse IR Dye 800CW, 1:10,000; donkey anti-guinea pig IR Dye 680CW, 1:10,000; all from LI-COR Biosciences). Signals were visualized with an Odyssey Infrared Imager and Odyssey software (LI-COR Biosciences). Total intensity values were calculated with Odyssey software and quantification was performed by normalizing protein abundance to actin first then to Cre-negative control samples. Antibodies used: anti-HA, 1:5000, Covance, MMS-101P; anti-GFP, 1:5000, rabbit, Invitrogen, A11122; anti-ACTB, 1:10000, mouse, Sigma, AC-15; anti-Tuj1, 1:5000, mouse, Covance, MMS-435P; anti-synapsin 1, 1:5000, mouse, Sysy, 106011; anti-synapsin 1, 1:5000, rabbit, Yenzym, clone YZ6079; anti-CASK, 1:5000, mouse, Neuromab, K56A/50; anti-vGluT1, 1:5000, guinea pig, Millipore, AB5905; anti-vGluT2, 1:5000, guinea pig, Millipore, AB2251; anti-vGaT, 1:5000, rabbit, Millipore, AB5062; anti-GAD67, 1:5000, mouse, Millipore, MAB5406; anti-Homer1, 1:5000, rabbit, Sysy, 160003; anti-Homer1, 1:5000, rabbit, Yenzym, YZ6081; anti-PSD95, 1:5000, mouse, Sysy, 124011; anti-gephyrin, 1:5000, mouse, Neuromab, L106/83; anti-collybistin, 1:5000, mouse, Neuromab, L120/12; anti-GABARA1, 1:5000, mouse, Neuromab, N95/35; anti-calbindin, 1:5000, mouse, Sigma, C9848.

Immunohistochemistry was performed as previously described (Zhang et al., 2015). Mice (P21 for L7-Cre, P35 for GLAST-CreER KOs) were anesthetized with isoflurane, perfused with phosphate buffered saline (PBS) to clear blood then with 4% paraformaldehyde (PFA) in 0.1M PBS to fix tissues. Cerebella were dissected out and post-fixed in 4% PFA overnight at 4°C. Cerebella were then cryoprotected in 30% sucrose in 0.1 M PBS for a minimum of 48 hr at 4°C. Sagittal cerebellar sections (40 µm) were collected at −20°C with a cryostat (Leica CM1050). Sections were washed with PBS and incubated in blocking buffer (0.2% Triton X-100% and 5% goat serum in PBS) for 1 hr at RT under gentle agitation, followed by incubation for 24 hr at 4°C with primary antibodies diluted in blocking buffer (antibodies used: anti-vGluT1, 1:1000, guinea pig, Millipore, AB5905; anti-vGluT2, 1:1000, guinea pig, Millipore, AB2251; anti-vGaT, 1:1000, rabbit, Millipore, AB5062; anti-calbindin, 1:2000, mouse, Sigma, C9848; anti-GLAST, 1:500, rabbit, Abcam, ab416s; anti-GLT-1, 1:500, rabbit, Abcam, ab41621; anti-GluA1, 1:500, rabbit, Sysy, 182003; anti-HA, 1:1000, mouse, Covance, MMS-101P; anti-Homer1, 1:1000, rabbit, Millipore, ABN37; anti-MAP2, 1:1000, chicken, Encor Bioscience, CPCA-MAP2). Sections were washed 4 times for 10 min each in PBS then incubated in secondary antibodies diluted in blocking buffer for 1 hr at RT (1:1000, Alexa 488 or 546, Invitrogen). Sections were washed as described previously then mounted onto positively-charged microscope slides and covered in DAPI-containing mounting media (Vectashield, Vector Labs). Serial confocal z-stack images (1.5 μm intervals for 40 μm at 1024 × 1024 resolution) were acquired using a Nikon confocal microscope (A1Rsi) with a 60x oil objective (PlanApo, NA1.4). All staining procedures and acquisition parameters were kept constant within each experiment. Images of cerebellum molecular layer were taken from the midpoint of lobule IV/V. Image backgrounds were normalized, and immunoreactive elements were analyzed with Nikon analysis software (object size range 0.25–4.0 μm²).

Electrophysiological recordings from cerebellum were performed as previously described (Zhang et al., 2015; Caillard et al., 2000; Llano et al., 1991). Mice at age P21-25 or P35-40 were anesthetized with isofurane and decapitated. Cerebella were rapidly extracted and transferred into ice-cold low-Ca2+ aCSF containing (in mM): 125 NaCl, 2.5 KCl, 3 MgCl2, 0.1 CaCl2, 25 glucose, 1.25 NaH2PO4, 0.4 ascorbic acid, three myo-inositol, 2 Na-pyruvate, and 25 NaHCO3; pH was adjusted to 7.4 by continuous gassing with carbogen. Sagittal slices (250 µm) were sectioned from the vermis on a vibratome (Leica VT 1200S) and transferred to oxygenated aCSF at RT containing (in mM): 1 MgCl2, 2 CaCl2 instead of 3 MgCl2, 0.1 CaCl2 as described earlier. Slices were incubated for >1 hr before recording. Purkinje cells were visually identified using infrared DIC imaging with a 40 × water immersion objective. Whole-cell recordings from Purkinje cells in cerebellar lobules IV/V (voltage-clamped at −70 mV) were performed at RT with borosilicate glass pipettes (2–3 MΩ) pulled with a vertical micropipette puller (PC-10, Narishige). Internal pipette solutions contained (in mM): for all EPSCs, 140 Cs-gluconate, 5 CsCl, 2 MgCl2, 0.5 EGTA, 2 Na-ATP, 0.5 Na-GTP (pH 7.3, adjusted with CsOH); for IPSCs, 145 CsCl, 2 MgCl2, 0.5 EGTA, 2 Na-ATP, 0.5 NaGTP (pH 7.3, adjusted with CsOH). For CF-EPSC and PF-EPSC recordings, the aCSF bath solution contained 50 µM picrotoxin and 10 µM APV; miniature EPSC recordings also included 1 µM TTX. For mIPSC recordings, the aCSF bath solution contained 10 µM CNQX, 10 µM APV and 1 µM TTX. Focal square pulse stimuli (duration 50 μs, amplitude 5–15 V or 200-600mA) were applied with a bipolar stimulation electrode (FHC, Bowdoinham, ME) connected to a Model 2100 Isolated Pulse Stimulator (A-M Systems, Inc) synchronized with the Clampfit 10 data acquisition software (Molecular Devices), with currents recorded by a Multiclamp 700B amplifier (Molecular Devices). Climbing fibers were stimulated in the granule cell layer around Purkinje cells (~100 µm) and identified by their characteristic all-or-none response and paired-pulse depression at a 50 ms inter-stimulus interval (ISI). The number of steps of climbing-fiber EPSCs was defined as the maximal number of climbing-fiber EPSC responses with different amplitudes recorded with different stimulus strengths at different stimulation sites in the granule cell layer. Parallel fibers were stimulated in the distal molecular layer (~200 μm from the target Purkinje cell) and identified by paired-pulse facilitation at 50 ms ISI. Only Purkinje cells with a series resistance <12 MΩ (not compensated; monitored before and after experiments by applying 10 ms, −5 mV voltage pulses) were used for analyses. Amplitude and PPR of evoked EPSCs were averaged from 5 to 10 traces. mEPSCs and mIPSCs were thresholded at 5 pA and hand-picked and analyzed using Clampfit 10.7 (Molecular Devices). Whole-cell currents were recorded at 10 kHz and filtered with a Bessel filter at 4 kHz.

All data are shown as means ± SEM, with statistical significance (*p<0.05, **p<0.01, ***p<0.001) determined by Student’s t-test, one-way analysis of variance (ANOVA), two-way ANOVA or Kolmogorov-Smirnov (K-S) tests. Non-significant results (p>0.05) are not specifically identified.

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

We thank Dr. R Sando for advice on mouse genetics. This study was supported by a grant from the NIH. RSZ was supported by the Lucille P Markey Basic Biomedical Research Fellowship from Stanford University. K L-A is supported by EMBO long-term fellowship (ALTF 803–2017).

Animal experimentation: All animal procedures conformed to the National Institute of Health Guidelines for the Care and Use of Laboratory Animals and were approved by the Stanford University Administrative Panel on Laboratory Animal Care (protocol #21589).

© 2020, Zhang et al.

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