All animal experiments were conducted in compliance with the European union and Belgian laws in accordance with the guidelines of the Ethical Committee Animal Experimentation of the KU Leuven (P267/2015). Mice were maintained on CD1 Swiss genetic background. The CD1 mice were crossed with transgenic Dlx5/6cre-IRES-eGFP mice strain, which labels telencephalic interneurons green (Dlx5/6-Cre-IRES-eGFP) (Stenman et al., 2003). The mice were kept in a 14/10-h light-dark cycle in a humidity- and temperature-controlled pathogen free animal unit. Timed matings were performed to obtain pregnant female mice, and a plug check was done every day early in the morning to confirm mating. The vaginal plug day was considered as E0.5.Dlx5/6-Cre-IRES-eGFP positive embryos were verified under a fluorescence binocular microscope (SteREO Discovery.V8; Zeiss; Oberkochen; Germany). Dissection of mouse brains was done in cold phosphate-buffered saline (PBS) and fixation in 4% w/v paraformaldehyde (PFA)/PBS for 16–24 h at 4°C. After fixation, samples were washed two times in PBS and then placed into a storage buffer (0.01% w/v thimerosal/PBS) to preserve the brains. In addition, mouse tail biopsy samples (~5 mm) were also collected for DNA extraction and genotyping of the embryos.

PCR reaction was done with a GoTAQ Polymerase (Promega). The used forward and reverse primer sequences are shown in Supplementary Table S1. Initial denaturation was performed at 94°C for 4 min. Cycling was done for 35 cycles with a denaturation at 94°C for 30 s, primer annealing at 58°C for 30 s and elongation at 72°C for 30 s. The final elongation was done for 7 min at 72°C and then cooled down to 4°C. The PCR product was visualized on a 1% agarose gel.

PCDH19 FL, ECDTM, ICD, and ICDΔNLS constructs were obtained from a plasmid containing the full-length murine sequence (PCDH19FL) and cloned into a second plasmid containing the green fluorescent protein (eGFP) behind a cytomegalovirus promoter. Primer sequences to generate the PCR products are shown in Supplementary Table S2.

PCDH19ICDeGFP and PCDH19ICDΔNLSeGFP were generated using tail PCR, and PCDH19FLeGFP and PCDH19ECDTMeGFP were generated using dovetail PCR. Finally, all PCDH19eGFP constructs were recloned into a pCAGGS plasmid, expressing IRES-TdTomato.

All the primers used to generate the PCDH19-V5 and PCDH19 KO mouse lines were ordered from IDT. These mouse lines were created in the lab of Bernhard Schermer.

A small ear biopsy was collected at weaning and used as well for identification of the mouse. Ear biopsy was deposited into a sterile Eppendorf tube and incubated with a 200 μl lysis buffer [1-M Tris-HCl (pH = 8.5), 0.5-M EDTA (pH = 8), 10% sodium dodecyl sulfate (SDS), and 5-M NaCl], containing 1:100 Proteinase K (10 μg/μl in 40% glycerol/nuclease free water) at 56°C overnight.

PCR reaction was done with a GoTAQ Polymerase. The used forward and reverse primer sequences are shown in Supplementary Table S4. Initial denaturation was performed at 94°C for 3 min. Cycling was done for 34 cycles with a denaturation at 94°C for 30 s, primer annealing at 60°C for 30 s, and elongation at 72°C for 1 min. The final elongation was done for 10 min at 72°C and then cooled down to 10°C. The PCR product was visualized on a 2% agarose gel. Primers are in Supplementary Table S4.

A 560 base pair fragment of the mouse PCDH19 gene exon1 was cloned into a pJET1.2 vector. In vitro transcription was done using 1 μg as a template of the linearized plasmid, using the T7 DIG RNA labeling kit (Roche, Sigma-Aldrich). Purification of the RNA was done using the Micro Bio-Spin™ P30 gel columns (Bio-rad) according to the manufacturer's instructions, and RNA concentration was measured using the SimpliNano spectrometer (Biochrom). All the ISH were done on 6-μm paraffin sections using an automated platform (Ventana Discovery, Roche). About 200 ng of dioxigenin labeled probe was diluted in RiboHybe (Roche) and vortexed prior to use. After deparaffination via heat, slides underwent pre-treatment with a citrate buffer (pH = 6.) at 95°C and with proteinase K at 37°C for 4 min. The probe in the aforementioned amount was added per slide, and denaturation was performed at 80°C. Next, hybridization was performed for 6 h at 65°C. Several washes had occurred before the anti-DIG antibody conjugated to alkaline phosphatase was added at the concentration of 1:1,000, followed by an incubation of 8 h with the substrate (BlueMap Detection Kit, Roche). Slides were manually dehydrated in a graded ethanol dilution and mounted with Eukitt (Sigma). Brightfield images were acquired using a LeicaDM6 B microscope connected to a digital CMOS camera (DMC2900, Leica) with the LAS Xsoftware suite (Leica). Further processing was done with Fiji and GIMP or Photoshop.

This protocol is an adaptation of the HCR3.0 protocol described in Choi et al. (2018). We refer to buffer compositions described in this protocol, and used the method described in Elagoz et al. (2022) for probe design. To ensure the probe could distinguish WT from KO, we used the ~1,000 bp PCDH19 sequence that was deleted in the PCDH19 KO mice as input for the probe generator. Off-target probes were identified using BlastN and were excluded from the probe set. DNA oligo pools from the designed probes were ordered from Integrated DNA Technologies (IDT), dissolved in UltraPureTM DNase/RNase-Free Distilled Water (Invitrogen) and stored at −20°C. HCR B2 amplifier labeled with Alexa Fluor^® 647 was ordered from Molecular Instruments. Components of all used buffers and solutions as well as the sequences for the DNA probe sets are listed in Supplementary Tables S5–S9.

Before starting the protocol, vibratome sections were placed in a 24-well plate (maximum, three sections per well). First, tissues were permeabilized with 600 μl of a permeabilization buffer (1% DMSO, 1% Triton-X in autoclaved PBS) for 2 h at 37°C in a humidified surrounding in the dark. Next, a permeabilization buffer was replaced with a 600 μl pre-warmed probe hybridization buffer, and tissue was incubated for 1 h at 37°C in a humidified surrounding in the dark. In the meantime, HCR probes were thawed on ice, spun, and a 12 nmol DNA probe was diluted in a 200 μl pre-warmed probe hybridization buffer (probe solution) per well or glass slide. After incubation, the probe hybridization buffer was replaced by 200 μl of probe solution. Tissue was incubated for 18 h at 37°C in a humidified surrounding in the dark.

The next day, excess probes were washed from the tissue in several wash steps (600 μl of each wash solution, see Table 1) at 37°C in a humidified surrounding in the dark.

To amplify the probe, the last wash solution was replaced by a 600 μl amplification buffer, and tissue was incubated for at least 30 min at RT in a humidified surrounding in the dark. Next, 9 pmol of both amplification hairpins (H1 and H2) per well or glass slide was pipetted into pcr-tubes, heated at 95°C for 90 s, immediately put on ice for 5 min, and incubated at RT for 30 min in the dark. Next, the hairpins were individually dissolved in the amplification buffer (100 μl per 9 pmol hairpin) before they were combined and mixed well (hairpin solution). The amplification buffer was replaced with 200 μl hairpin solution. Tissue was incubated for 16 h at RT in a humidified surrounding in the dark.

The following day excess hairpins were removed by washing tissue three times with 600 μl of 5X SSCT for 10 min. Next, sections were stained with DAPI and mounted.

A control condition in a well was included in which probe solution was replaced by a 300 μl probe hybridization buffer, and hairpin solution was replaced by a 300 μl amplification buffer. Thus, these sections were not treated with probes and hairpins to control for autofluorescence of the tissue.

Depending on the experiment, MGE, LGE and CGE was dissected out of 10 embryos at stage E13.5 and pooled per brain region (Figure 1J) or individual whole telencephalon (Figures 1E,J), ventral telencephalon (Figure 1J) or entire brain (Supplementary Figure 1F) were isolated. For protein isolation and quantification, the tissue lysates were homogenized in a 100 μl/sample ASBA buffer [1% Triton-X-100, 0.1% SDS, 1-mM EDTA, 50-mM NaCl, 20-mM Trizma base (pH = 7.5)] and 4 μl/sample cocktail protease inhibitor solution (Roche). The homogenization was performed mechanically with a drill for 5-x-10 s/sample and kept on ice. Afterwards, the brain samples were centrifuged for 5 min at 13.000 rpm at 4°C. Supernatants were collected, and the Qubit Protein Assay Kit (Invitrogen) was used to determine protein concentration of the brain lysates.

About 15 μg of the protein lysates was loaded on a precasted 4–12% Criterion XT Bis-Tris 26 well-gel of Biorad and blotted on a PVDF membrane using the Trans Turbo Blot from BioRad (7 min). Next, the membranes were dehydrated in Tris-saline for 5 min and incubated with a blocking buffer for the following 2 h. All the washing and incubation steps occurred at room temperature (RT) and placed on the shaker. After the 2-h incubation, the membrane was incubated with the PCDH19 primary antibody (Bethyl A304-468A) diluted 1/1,000 and left overnight to incubate. The next day, blots were rinsed 3 x 5 min with Tris-saline and incubated with the secondary antibody solution with HRP-labeled antibodies (GAR:Goat Anti-Rabbit) 1/10,000 in the blocking buffer for 30 min. Next, membranes were rinsed with Tris-saline (2 × 5 min) and incubated with Tris-Stock (1 × 5 min). After the Tris-Stock incubation, the Clarity western ECL substrate kit (BioRad) was used to reveal the blots. Imaging of the membranes occurred with the imaging system (BioRad).

The membranes were stripped using a Restore western blot stripping buffer of Thermo Scientific. Next, the membrane was rinsed again with Tris-saline (3 × 5 min) and incubated with the blocking buffer for 2 h. The membranes were incubated overnight with GADPH 1/1,000 (GAPDH Millipore cat nr: MAB374) in the blocking buffer. The next day, a HRP-labeled antibody (Goat Anti-Mouse) 1/25,000 solution was added and, afterwards, imaged with the BioRad imaging system.

One day before the actual experiment, Millicell Cell Culture inserts with a pore size of 0.4 μm were coated with poly-L-Lysin (Sigma) and Laminin (Sigma). Coated inserts were placed in PBS into a 6-well plate overnight at 37°C in the cell culture incubator. On the next day, coating solution was removed, and inserts were placed on top of a slice culture medium. Dlx5/6-Cre-IRES-eGFP E13.5 brains were dissected in ice-cold L15++ (a Leibovitz‘s L15 medium, supplemented with glucose and Hepes). Subsequently, the dissected brains were embedded in 4% low-melting point agarose in L15++. Polymerization was obtained after 1 h on ice. Next, coronal slices of 300 μm were obtained with the vibratome and collected in L15++ media. Settings of the vibratome were the following: speed was less than 15, frequency was set at 60, and the amplitude at 0.6. Sectioned slices were placed on top of a membrane and placed on top of an agarose bed, previously prepared into the square well of a Petri dish square platinum electrode (CUY701P20E, Nepagene). Injection was performed under a binocular (Leica) into the MGE. For OE Plasmids, we injected at a concentration of 2 μg/μl. To generate the LOF, clustered regularly interspaced short palindromic repeats (CRISPR-Cas9) components were pre complexed to ribonucleoprotein (RNP) in an Opti-Mem medium for at least 20 min at room temperature injected at the amount of 380 ng for the guide ribonucleic acid (RNA) and 300 ng for the Cas9 protein. All LOF compounds were mixed with pCAGGS plasmid at a concentration of 1 μg/μl in order to trace the electroporated cells. To visualize the injection site, Fast Green was used at an amount of 1 μl per 33 μl. For the negative electrode, an agarose cylinder of 1% was attached to the cover square of platinum plate electrode (CUY701P20L, Nepagene). Electroporation was performed using the following settings: 5 pulses, 150V with a pulse length duration of 5 ms using 100-ms intervals. A BTX electroporator (ECM830, Harvard Apparatus) was used. Subsequently, after electroporation, electroporated sliced were placed on top of the coated inserts and placed on ice. Three slices with the same condition were placed on top of an insert. After all the inserts were covered with slices, the slices were cultivated in the cell culture incubator. After 2 days in vitro (div), 500 μl of the used slice culture media was removed and refreshed with new slice culture media. After 3 div slices were fixed in 4% PFA for at least 2 h, stained with DAPI and mounted on microscope slides with Mowiol and imaged with the confocal microscope (Olympus). Analysis was performed using ImageJ software, total amount of TdT positive cells was quantified within every slide side, and the proportion or cells reaching the cortex compared to the total amount was calculated.

Statistical significance was determined using a Kruskal Wallis multicomparison test with Dunn‘s post hoc test.

At E13.5 Dlx5/6-Cre-IRES-eGFP embryos were dissected from the mother in L15++ media. Embryos were kept on ice, while the skin and meninges were removed. Subsequently, the neocortex (NCTX) was open up from the top, superficially, in order to expose the GEs. Injection into the MGE was performed using approximately 10 injection sites per MGE. The same injection mixtures and amount of OE plasmids and CRISPR-Cas9 components as used for the slice electroporation (slice EP) were used. Subsequently, after injection in both MGEs, electroporation was performed using the following settings, 5 pulses, 50 V with a pulse length duration of 50 ms using 1-s intervals. Electrodes (CUY650P5 tweezer electrodes) connected to a BTX electroporator (Harvard Apparatus) were placed on each side of the embryonic head at the position of the ears. After electroporation embryos were kept at 4°C for 3 h in L15++. Meanwhile, the matrigel was thawed in ice. Matrigel was diluted in a 1:1 ratio with complete neuro basal media. After the incubation period, MGE was dissected from the embryos, and each MGE was cut into small pieces. Subsequently, these pieces were embedded carefully into the neurobasal media-diluted matrigel and placed as matrigel drops containing the piece into 35-mm glass bottom imaging (Ibidi) dishes. Once placed into the dish, the Matrigel containing explant was allowed to polymerize at 37°C. After polymerization, the explant was surrounded with complete neurobasal media and cultured in the cell culture incubator for 2 days. After 2 div explants were fixed with 4% PFA and imaged with the confocal microscope (Olympus). Stacks were projected with the Image J software, and analysis of migration from the explant was performed using Cell Profiler software (McQuin et al., 2018; Stirling et al., 2021). Statistical significance was determined using the Mann–Whitney U Test for the sample size of 2 and with the Kruskal–Wallis non-parametric test and the Dunn's post hoc test for bigger sample sizes.

The following cell profiler pipeline for analysis with the following modules was used. The z-projected image of the explant and the red fluorescence channel for the positive TdT cells were used as starting images for the images module. Metadata was extracted from the image file. The explant brightfield image and the TdT cell image were defined with names and types. We used “identify primary objects” as modules to detect the explant and the TdT neurons. The explant was defined with the Cell profiler as one primary object. All the TdT neurons cell bodies were identified as primary objects; the result was verified. Once all objects were identified, we used the two-module mask objects and relate objects. In the relate object module, the explant was set as a parent object and the neuron cell bodies as child objects. Using this module, distance between the parent object and all child objects was measured. Distances were obtained in pixels. Transforming of pixels into μm was achieved using the conversion factor stored in the .oib file using Image J. Finally, Graph Pad Prism 8.3.0 was used to determine the statistical significance.

Morphological analysis of MGE explant-derived neurons was performed using the SNT ImageJ plugin (Longair et al., 2011; Arshadi et al., 2021). A minimum of 4 explants obtained from at least two experimental replicates were analyzed per treatment group. Means per experimental group were compared with the Kruskal–Wallis non-parametric test and the Dunn's post hoc test for a sample size bigger than 2 and with the Mann–Whitney U test for the comparison between two samples. Polarity analysis was compared across experimental conditions using two-way repeated measures ANOVA with Holm-Sidak post hoc comparison. Neuronal category frequencies were compared between neuronal morphology and experimental groups using two-way repeated measures ANOVA with Holm-Sidak post hoc comparison.

N2A cells were grown in DMEM high glucose supplemented with 2-mM L-glutamine, 1% penicillin/streptamycin, and 10% fetal bovine serum albumine (Neuro2A media). For confocal microscopy, we used glass bottom dishes (Ibidi) and pre coated them with geltrex at least 3 h prior seeding; for the rest of applications, we used 6-well plates. One day prior, transfection cells were seeded to a density of 5 × 10⁵ (a 6-well plate) and 5 × 10⁴ (Ibididish) and left to adhere. On the day of the transfection, the cells were checked for adherence and confluency under the microscope. Transfection was performed using Lipofectamine 3000 (Thermo Fisher Scientific). Briefly, Lipofectamine 3000 reagent was diluted in Opti-Mem using the higher amount suggested by the manufacturer for 6-well or 24-well (Ibidi dish) and vortexed and spun down. About 5 μg of plasmid DNA was diluted in Opti-MEM, and, subsequently, the 2 μl of P3000 reagent per 1 μg DNA was added to generate the master mix. The mastermix was spun down and mixed 1:1 with the prior diluted lipofectamine in Opti-Mem. The mixture was spun down and incubated for 30 min at room temperature. After the incubation, mixture was added to one well/dish drop by drop, and the cells were given an easy shake to mix the reagent better into their media. The cells were incubated at 37 °C in an incubator. Transfection efficiency was controlled after 24 and 48 h under a fluorescent microscope.

About 500 μg of protein per condition was used to start the pull down. The Miltenyi μM ACSGFP isolation kit (130-091-125) was used. About 50 μg GFP beads were incubated with lysate, containing 500 μg protein for 1 h in the cold room with overhead shaking. A μMACS column and magnet were placed in the cold room. About 200 μl of an ice cold lysis buffer containing the proteinase inhibitor cocktail was used to preclear the columns. Whole lysate containing the beads was loaded on the magnetic column. Beads were washed two times with a wash buffer 1 [50 mM Tris (pH = 7.5), 150-mM NaCl, 5% glycerol, and 0.05% Triton-X100]. Then, the beads were washed three times with wash buffer 2 [50-mM Tris (pH = 7.5), 150-mM NaCl]. Elution was conducted via adding 20 μl of a 95°C pre-heated elution buffer to the column and incubated at room temperature for 5 min. Subsequently, 50 μl was further eluted through the column, and analysis was done by Western Blot.

For western blot detection after the GFP pull down, proteins were heat denatured in a mixture of sample-reducing agent 10 × (NOVEX, NP0009), and loading dye LDS Sample Buffer 4 × (NOVEX, NP0007) was as well added to the protein. All amounts were calculated for a volume of 20 μl and boiled at 70°C for 10 min. Separation was performed on a precast NuPAGE 4–12% Bis-Tris gel [Invitrogen, (WG1403BX10) with NuPAGE MOPS SDS Running Buffer (NP0001) and immune blotted to a nitrocellulose membrane (Transfer Pack, Midi format, 0.2-μm nitrocellulose, Bio-Rad, 170-4159) using a Trans Blot Turbo system (Bio-Rad)]. Standard protein detection was performed using mouse anti-GFP antibody -HRP (1:5,000; Biorad), rabbit anti-PCDH19 (1:500 Millipore). After 2-h blocking in 5% w/v non-fat dry milk/TBST (a blocking buffer) at RT, o/n incubation at 4°C in a primary antibody diluted in the blocking buffer, and washing in TBST, transfer membranes were incubated during 45 min in HRP-conjugated anti-rabbit secondary antibodies (Bio-Rad) diluted 1:10,000 in a WB buffer. Protein bands were visualized with a ChemiDoc MP imaging system (Bio-Rad) after incubation in ECL substrate (Thermo Fisher Scientific).

To perform our apoptosis analyses, we trypsinated the Neuro2A cells and stopped the enzyme with media-containing serum. In addition, we kept all the washes to not loose dead cells. After centrifugation, the supernatant was discarded, and cell pellets were resuspended in PBS and transferred into micro centrifuge tubes. In order to have a positive control for early and late apoptosis, the untransfected cell condition was used. Half of the cells were killed intentionally for 15 min at 75°C. These cells were then mixed with the rest of untransfected cells in equal parts and spun down. Once the positive apoptosis control was made, all the cell suspensions were spun down and resuspended into a 100 μl Annexin V-binding buffer (BD Biosciences). Annexin V (V450, 560506 BDBiosciences) and 7-aminoactinomycin D (7AAD) (559925 BD Biosciences) were added to analyze for early and late apoptosis, excluding the single-stain controls. The single-stain controls were the positive apoptosis control either incubated with Annexin V or 7AAD. Incubation was performed for 15 min at room temperature in the dark. After the incubation, an Annexin V-binding buffer was added, and cells were passed through tubes with a 35-mm strainer (VWR) and analyzed with the SH800 Cell Sorter (Sony).

N2A cells images were acquired using the 60X magnification with oil immersion of the confocal microscope (an Olympus FV1000 microscope) and taking fluorescent z-stacks with a depth of 1–2 μm, a speed of 4 μs/pix and a pixel size of at least 1024 × 1024 pixels. Whole z-depth of the DAPI-channel was covered.

For nuclear-cytoplasmic distribution calculations, a single z-slice was manually selected from the provided z-stack in order to have the most central view of the cell through the nucleus and to have the highest number of cells in focus. These images were split in individual channels [DAPI (blue), GFP (green) and TexRed (red)]. Images were segmented and analyzed using Cell Profiler using a custom-made pipeline. Briefly, the cells were segmented by identifying individual nuclei through the DAPI channel using a global Otsu threshold method. Cells edges were identified using a propagation algorithm using the fluorescence channel that would best describe the totality of the cell, leading to a one-to-one correspondence between each nucleus and the respective cell. Nuclear areas were defined by the DAPI stain and the cytoplasmic areas were obtained by subtracting the nuclear areas to the identified total cell areas. Shrinking of 1 pixel of nuclei was used to better define the cytoplasmic areas. After segmentation, the average fluorescence intensity was calculated for each cell in both the nucleus and cytoplasm-identified objects. Image analysis with CellProfiler 2.2.0 and R Studio 3.6.1 was conducted by Marco Dalla Vecchia. Images with cells in a clearly dying state or with many saturated pixels were discarded from the analysis.

Z max projection was made of the explant images. The area of the explant was defined by the Dlx5/6-Cre-IRES-eGFP IN in the GFP channel. The same area was applied to the TdT channel, and the threshold was adjusted before measurement. Intensity measurements were performed with Fiji is just ImageJ and statistical analysis performed with Graph Pad Prism 8.3.

Different studies have shown that Pcdh19 is expressed in the developing mouse cortex and hippocampus within pyramidal neurons (Kim et al., 2007; Fujitani et al., 2017; Pederick et al., 2018; Gerosa et al., 2022). Only a few groups have investigated Pcdh19 expression in inhibitory neurons (Bassani et al., 2018; Serratto et al., 2020; Galindo-Riera et al., 2021). We hypothesized that proper migration of inhibitory neurons to the cortex could be affected in PCDH19-CE. To address this question, we first assessed Pcdh19 expression in developing cortical interneurons neurons by means of in situ hybridization (ISH). GABAergic cortical interneuron migration occurs from E12-E19 in the developing mouse brain from the ganglionic eminence to the NCTX (Guo and Anton, 2014). We, therefore, studied Pcdh19 mRNA at E13.5 and E16.5 in the ganglionic eminence and NCTX in mouse embryonic brain slices (Figures 1A–D). At E13.5, Pcdh19 is clearly expressed in the cortical plate and the ventricular zone within the NCTX, and in the mantle zone (MZ), the dorsal lateral ganglionic eminence (LGE) ventricular zone (VZ), and the lateral cortical stream (LCS) of the ventral telencephalon. Caudally, Pcdh19 mRNA could be detected throughout the CGE, thalamus (TH), and hypothalamus (HYP) (Figure 1B). At E16.5, Pcdh19 expression became less prominent in the cortical plate (CP) than at E13.5 (Figure 1C) but appeared more restricted, which could be the prospective layer 5, where Pcdh19 expression has been described postnatally (Galindo-Riera et al., 2021) (Figure 1C). In the ventral telencephalon, it was still visible within the LCS. Caudally, expression was high in the limbic system within the TH, amygdala (AM) and HYP (Figure 1D). To investigate the temporal dynamics of protein production of PCDH19, we performed western blot (WB) on lysates of whole telencephalon at E13.5, 15.5, 17.5, and P7 and quantified the relative amount of PCDH19 at each stage (Figure 1E, Supplementary Figure 1A for WB). This analysis showed a significant increase in PCDH19 production at late embryonic stages and at P7, in line with the described roles of PCDH19 at the synapse. To further explore the expression of Pcdh19 in embryonic cortical interneurons, we made use of the Dlx5/6-Cre-IRES-eGFP mouse line that labels migrating cortical interneurons during embryogenesis (Stenman et al., 2003). We focused our analysis on E13.5, given many assays in this manuscript use this time point, and interneuron migration is easy to visualize at this stage. We revealed Pcdh19 expression by hybridization chain reaction, which enables co-localization and semi-quantitative evaluation of expression in situ. The HCR signal appears dotty, and each dot represents at least one mRNA molecule. The number of dots can be used as a proxy for the expression level. This analysis confirmed the colorimetric ISH data shown in Figures 1A,B, and indicated that some, but not all interneurons migrating through the cortex, expressed Pcdh19 (Figures 1F,H, Supplementary Figure 1F). Pcdh19 was also expressed in the ventricular zone of the MGE, while it was nearly absent from the mantle zone (Figure 1G). The marginal zone stream of interneurons tends to express Pcdh19 at a low level in most cells (Figure 1I, Supplementary Figure 1G). We complemented this analysis with a study at the protein level, performing WB on lysates from the whole ventral telencephalon, telencephalon and distinct parts of the ganglionic eminence at E13.5. A strong band around 126 kDa, which is the predicted MW of PCDH19, indicated that PCDH19 was produced in the MGE, CGE, and LGE (Figure 1J). Taken together, these results show that PCDH19 was present in the main regions that generate cortical inhibitory neurons at the assessed time points, and in variable levels in individual interneurons migrating in the cortex.

To complement our analysis of endogenous PCDH19 production, we generated two mouse lines: a C-terminus-tagged PCDH19-V5 mouse line, which would allow detection of PCDH19 with higher sensitivity, as the V5 antibody has been verified in more studies; a PCDH19 KO mouse line, which served as a negative control for the HCR analysis (Supplementary Figures 1B,C). The PCDH19 KO mouse line was designed to have a large deletion of 1,186 bp in the first exon 1. This deletion leads to a frameshift from AA176 onwards and a truncation of the protein at AA189. We could not observe any HCR signal nor any protein production in this knockout model (Supplementary Figures 1C,D), also not on any other height, ruling out alternative start exons that might generate a residual intracellular domain. Western Blot analysis of brain lysates showed the production of PCDH19-V5 with a stronger band around 126 kDA at E13.5, E14.5, and postnatal Day (P)7 (Supplementary Figure 1E). The multiple bands at the full-length size could represent different known PCDH19 mouse isoforms (Hunt et al., 2018) or post translational modifications. In the embryonic and P7 samples, but not in the adult brain, an additional band was detected at 50 kDA, the predicted MW of the PCDH19 intracellular domain. This fragment might have been generated by enzymatic cleavage, similar to some other PCDHs, as has been shown recently (Pancho et al., 2020; Gerosa et al., 2022). Besides a weak band around 70 kDa, no bands were observed in the wild-type control, showing the specificity of the V5 antibody. Our data thus suggested that, in the developing brain, PCDH19 appeared in different forms, including a cleaved version.

Cleavage of the ICD of transmembrane proteins is often followed by nuclear translocation, driven by a nuclear localization signal. We used distinct prediction tools to search for nuclear localization signals within the PCDH19 sequence (Figure 2A). Three NLS sequences of the longest PCDH19 isoform (Q80TF3) were predicted, yet scored differently in the prediction program NLStradamus (Supplementary Figure 2A) (Nguyen Ba et al., 2009).

To further study the subcellular localization of distinct PCDH19 subdomains, we generated C-terminally tagged overexpression constructs of subdomains of PCDH19 in bicistronic TdTomato expression plasmids (pCAGGS-PCDH19-eGFP-IRES-TdTomato): a full-length version: PCDH19FL; extracellular and transmembrane domains: PCDH19ECDTM, an intracellular domain: PCDH19ICD and intracellular domains without a predicted nuclear localization signal (NLS): PCDH19ICDΔNLS (Figure 2B). PCDH19ICDΔNLS was lacking the predicted NLS with the highest score and analysis with NLStradamus showed loss of most nuclear localization activity (Supplementary Figure 2A). The constructs were validated by expressing them in Neuro 2a cells (Supplementary Figure 2C). Western Blot analysis indicated a proper production of the GFP-tagged domain mutants (Supplementary Figure 2B). PCDH19 FL overexpression resulted in accumulation of tagged PCDH19 around the nuclear membrane. Overexpression of PCDH19ECDTM resulted in accumulation of PCDH19 at the membrane of the cell. PCDH19ICD localized to the nucleus, while PCDH19ICDΔNLS could be found distributed over the whole cell and less prominent in the nucleus. Therefore, removing the NLS in this construct seems to inhibit nuclear translocation at least partially. Indeed, when we quantified the nuclear-cytoplasmic ratio (Supplementary Figure 2D), we found that PCDH19ICD localized preferentially in the nucleus, while this preference was lost upon removal of the NLS. To investigate whether overexpression in the developing mouse forebrain yields similar results, we performed slice electroporation and visualized subcellular distribution of GFP-tagged PCDH19. A similar subcellular distribution could be observed: PCDH19ECDTM was mainly found at the membrane. PCDH19FL was at the membrane, strongly again surrounding the nucleus but could also be identified in the nucleoli. PCDH19ICD localized to the nucleus while PCDH19ICDΔNLS was distributed all over the cell (Figure 2C). Taken together, our data show that PCDH19ICD translocated to the nucleus, probably using the predicted NLS. In addition, these constructs now allowed us to study the role of PCDH19 at the membrane and distinguish it from a potential role in the nucleus in different mosaic overexpression paradigms.

As PCDH19-CE only affects women that display cellular mosaicism of PCDH19, the imbalance in PCDH19 dosage at cell-cell contact sites might be a driver of the phenotype. We decided to model this mosaic imbalance by altering PCDH19 levels in some cells only by electroporation. In order to follow up the survival and quality of the slices, we used Dlx5/6-Cre-IRES-eGFP organotypic brain slices in which proper interneuron migration of non-targeted cells was a proxy for healthy slices. A PCDH19 subdomain-expressing plasmid or the empty TdTomato control plasmid was electroporated in the MGE. This allowed us to study the effect of overdosing distinct domains of PCDH19 on the migrational behavior of MGE cortical IN after 3 days in organotypic culture (Figures 3A–C). Representative images of organotypic brain slices for the endogenous fluorescence Dlx5/6ireseGFP (Figure 3A) and the electroporated fluorescence TdTomato (Figure 3B) after 3DIV are shown for all conditions, and the cortical field is depicted by the dotted line (Figures 3A,B). Quantification of the percentage of TdT positive neurons within the cortical field compared to the whole brain slice showed significantly reduced cortical IN migration for PCDH19ECDTM and PCDH19FL compared to the control condition (Figure 3D, Kruskal–Wallis test with the Dunn's post hoc test). Stunted neuronal morphology could be observed after PCDH19FL (Figure 3B) as well as after PCDH19 ECDTM overexpression, suggesting that increasing the PCDH19 extracellular amount could affect cell morphology or survival. In contrast, increasing the dosage of the intracellular domain of PCDH19 did not impact IN migration nor morphology. As we hypothesized that an imbalance in PCDH19 dosage between migrating interneurons and their environment would influence migration, one could argue that non-targeted migrating interneurons might be affected as well when neighboring, co-migrating interneurons would express an excessive amount of PCDH19. We therefore investigated the migration of all eGFP-labeled neurons, and although we saw a reduction, this effect was not significant (Supplementary Figure 3, Kruskal–Wallis test), potentially because only a fraction of interneurons was targeted. Our data thus indicated that overdosing in especially the extracellular part of PCDH19 hampered tangential migration of interneurons to the neocortex.

To further study the effect of excess PCDH19 on neuronal migration and morphology, we investigated PCDH19 overexpressing neurons in cultured MGE explants. Interneurons generated from these explants migrate out over the course of hours, during which they detach from the explant and make plenty short-term contacts with migrating cells in their vicinity. Besides revealing the impact of PCDH19 gain-of-function on individual cells and their migration capacity, this assay can also uncover disturbed cell-cell adhesion. E13.5 Dlx5/6-Cre-IRES-eGFP MGE explants were electroporated with the different PCDH19 subdomain expressing and control constructs, allowing the detection of electroporated (GFP + TdT +) vs. non-electroporated (GFP + TdT–) cells (Figure 4A). Next, we measured the total migration distance after 2DIV. Example images are shown for explant per experimental condition (Figures 4B–F). Statistical analysis with the Kruskal–Wallis non-parametric test and the Dunn's post hoc test showed a significant reduction in the spread of PCDH19FL-expressing Ins compared to the TdT control condition (p < 0.0001) (Figure 4G). Binned distribution analysis indicated that PCDH19FL-overexpressing IN distribution is closer to the explant than the rest of the conditions; nevertheless, no particular bin-specific means were found to be significantly changed (mixed-model ANOVA) (Figure 4H).

Similarly, as in the brain slice, we asked whether we would see a non-cell autonomous effect and, therefore, also analyzed non-targeted eGFP-positive neurons (Supplementary Figure 4). Also here, statistical analysis with the Kruskal–Wallis non-parametric test and the Dunn's post hoc test displayed a significant reduction in the spread of eGFP + TdT – neurons in the PCDH19FL condition (p < 0.0001) compared to the TdT control, and, unexpectedly, a significantly larger spread of the eGFP + TdT – neurons in the PCDH19ICD condition (p < 0.0001) (Supplementary Figure 4F). Statistical analysis per bin was also not significant for the non-cell autonomous effect of PCDH19 overexpression (Supplementary Figure 4G).

To further address the previous observation of the stunted neuronal morphology in the brain slices, we wondered whether upon overexpression of the different subdomains neuronal morphology was affected. Each neuron was assessed depending on the morphology of the leading process into “no process”, “unbranched process”, “single-branch process”, and into “multi-branch process”, and then the percentage of total neurons per experimental condition per explant was calculated per phenotype (Figure 4I). Most neurons had an unbranched process (50–70%), which fits to the previous observed morphology in the explant assay (Mitsogiannis et al., 2021). Overexpression of PCDH19FL compared to the control TdT-control did not result in any significant observation. Since total migration distance was reduced upon PCDH19FL overexpression, we also decided to look into the topological morphological characteristics of the neuronal processes. Assessment of the total cable length, a measure for neurite outgrowth, resulted in significant differences for PCDH19FL, PCDH19ECDTM, as well as PCDH19ICDΔNLS, compared to the control condition (Figure 4J, the Kruskal–Wallis non-parametric test, and the Dunn's post hoc test). Primary branch length and average branch length were only significantly affected upon overexpression of PCDH19ECDTM compared to the control condition (Figures 4K,L, the Kruskal–Wallis non parametric test, and the Dunn's post hoc test). Thus, the observed reduced migration distance upon PCDH19FL overexpression cannot be explained only by an affected neuronal morphology.

The observed impact on cortical IN migration and total migration distance could not be explained by aberrant cell morphology of the electroporated neurons only. Therefore, we sought to investigate whether cell survival was affected. First, we tested our hypothesis in the Neuro2A neuronal cell line. After 48 h of transfection with the PCDH19 or control constructs, apoptosis was measured using flow cytometry. AnnexinV was used to detect early apoptotic cells and 7AAD to detect late apoptotic cells within transfected and untransfected cells after overexpression. Flow cytometry example plots show transfected cells (TdT +) and early apoptotic cells (Figure 5A, the upper row) and late apoptotic cells (Figure 5A, the bottom row). Quantification of targeted cells results in more early and late apoptotic cells upon PCDH19FL overexpression compared to the control condition (Figure 5B, the left panel, Welch ANOVA, and the Dunnet's post hoc test) and in more late apoptotic cells upon PCDH19ECDTM and PCDH19FL compared to the control condition (Figure 5B, the right panel, Welch ANOVA, and the Dunnet's post hoc test). Quantification of early and late apoptotic cells in the untargeted cells resulted in non-significant differences, suggesting that the cells were not suffering because of the procedure, or that apoptosis was also induced by a non-cell-autonomous effect (Figure 5C, left and right panels). Therefore, the observed effect is specific for the PCDH19 overexpressing cell population, but only when the extracellular domain is present.

Next, we wondered whether we could address apoptosis in a more physiological model like the MGE explant. More cells can be targeted than in the slice electroporation with this approach, and, also, cellular resolution is better achieved with the assay than with the slice electroporation. We approximated the cell number of targeted cells via the TdT signal, again using the background of the Dlx5/6-Cre-IRES-eGFP mouse line. We reasoned that, if overexpression would affect cell viability and survival, we might lose more targeted cells and, hence, TdT expression, whereas overall GFP expression should stay constant. We measured the ratio between the TdT+ and GFP+ signals, whereby we normalized measurements by the explant area as shapes were diverse between measured conditions, and repeated the experiment several times to account for differences in electroporation efficiencies between experiments (Figure 5D). Our data consistently showed significantly lower ratios for PCDH19ECDTM and PCDH19FL compared to the control condition (Figure 5E, the Kruskal–Wallis non parametric test, and the Dunn's post hoc test), which could indicate that overexpression of these plasmids triggers neuronal loss. Taken together, our data suggest that an additional explanation for the observed impact on cortical interneuron migration could rely on increased neuronal cell death.

Another way to model mosaic imbalance in PCDH19 dosage at cell-cell contact sites is to deplete PCDH19 levels in some cells only. We applied a CRISPR RNP electroporation approach to downregulate PCDH19. Two guides targeting Exon 1 were designed (P1, P2) (Supplementary Figure 5A) and validated in vitro on a PCDH19FLeGFPiresTdT construct expressed in Neuro2A cells. Both guides were able to significantly reduce the percentage of GFP-positive cells on the total number of transfected cells (Supplementary Figures 5B,C).

To address migration characteristics at the cellular level, we again used MGE tissue derived from Dlx5/6-Cre-IRES-eGFP mice. Embryonic brains were injected and electroporated with ribonuclear particles containing a guide targeting Pcdh19 and Cas9 protein (referred to as PCDH19KO RNPs) or Cas9 protein alone (control), and a TdT-expressing plasmid in the MGE at E13.5. Subsequently, MGE pieces were cultured in Matrigel for 2 days (DIV) (Figure 6A). At 2 DIV, total migration distance was measured from the edge to the neuronal soma (Figure 6B). Figures 6C,D show representative examples of TdT-labeled neurons for each condition images. Migration distance was measured for more than 200 neurons in the PCDH19KO and control condition. Statistical comparison with the Mann–Whitney U test resulted in a slight significant decrease (p < 0.05) of total migration distance upon loss of function of Pcdh19 (Figure 6E). Binned distribution analysis of the migration distance suggested that, in the PCDH19KO condition, more neurons are distributed closer to the explant; however, no bin-specific means were found to significantly differ (Multiple Mann–Whitney tests with FDR for multiple corrections) (Figure 6F). Also, for this analysis, we addressed the non-cell autonomous effect of the PCDH19KO condition on eGFP+TdT– neurons and detected a significant decrease (p < 0.001) compared to the control condition (the Mann–Whitney U test) (Supplementary Figure 6). We further investigated whether the direction of migration could be affected in the PCDH19KO-electroporated neurons. To this aim, we classified the leading neurite direction, showing away from the explant, toward the explant or when the direction could not be specified in both categories, parallel to the radial migration. The total amount of neurons within these three classifications was counted in 4 explants for each experimental condition. Statistical analysis indicated that more leading neurites were pointed toward the explant in the PCDH19KO condition (two-way repeated measures ANOVA with Holm–Sidak post hoc comparison, p < 0.05) (Figure 6G). Overall, about 70% of the counted neurons pointed their main neurite away from the explant in both conditions. During interneuron migration, extension of the leading process is followed by branching as the neuron senses the environment, and, eventually, nucleokinesis. Microtubule and actin cytoskeleton remodeling events govern these dynamic processes (Bellion et al., 2005; Métin et al., 2006; Guo and Anton, 2014). A second reason for hampered migration could be disturbed by branch elongation and secondary branch formation. Hence, we measured primary branch length (Figure 6H), average branch length (Figure 6I), and cable length (Figure 6J) for more than 200 neurons in each condition, yielding no difference between the conditions.

In addition, we also assessed cell survival in a similar manner as described above (Figure 5D). Reducing the level of PCDH19 did not affect cell survival; we observed a lower number of targeted cells only in the control condition; however, this difference remained statistically non-significant (the Mann–Whitney U test) (Supplementary Figure 7).

Collectively, these results indicated that, similar to PCDH19 overexpression, loss of PCDH19 in MGE-derived IN mildly reduced the total migration distance in MGE explants, which might be explained in part by a disturbed migration polarity. However, the cell-cell variation in the effectiveness of the knockout, as well as the potential off-target effects, was difficult to measure in this explant model.

To further study the impact of PCDH19 imbalance and its impact on cortical interneuron migration, we modeled the mosaic loss by crossing the PCDH19 knockout mouse line heterozygotes with Dlx5/6-Cre-IRES-eGFP mice in order to obtain control (PCDH19+/+; Dlx5/6-Cre-IRES-eGFP), heterozygotes (PCDH19+/–; Dlx5/6-Cre-IRES-eGFP), and PCDH19 KO (PCDH19–/–; Dlx5/6-Cre-IRES-eGFP) mice in which VT-derived interneurons would be eGFP labeled. We studied the proportion of labeled cells in the cortex at E13.5 as a proxy for cell migration, similar to the analysis performed on the slices (Figure 7A). In doing so, we found that, at this early stage, we could detect a mild but significant reduction in migration in the heterozygote mice, but not in the homozygous KOs [Figure 7B, (^*p = 0.0135, the Kruskal–Wallis test, and the Dunn's post hoc test)]. These data suggest that, also, in vivo, creating an imbalance in the dosages of PCDH19 in interneurons and neurons hampers the overall interneuron migration to the cortex.