Rpsa Signaling Regulates Cortical Neuronal Morphogenesis via Its Ligand, PEDF, and Plasma Membrane Interaction Partner, Itga6

Sara M Blazejewski; Sarah A Bennison; Ngoc T Ha; Xiaonan Liu; Trevor H Smith; Kimberly J Dougherty; Kazuhito Toyo-Oka

doi:10.1093/cercor/bhab242

. 2021 Aug 4;32(4):770–795. doi: 10.1093/cercor/bhab242

Rpsa Signaling Regulates Cortical Neuronal Morphogenesis via Its Ligand, PEDF, and Plasma Membrane Interaction Partner, Itga6

Sara M Blazejewski ¹, Sarah A Bennison ², Ngoc T Ha ³, Xiaonan Liu ⁴, Trevor H Smith ⁵, Kimberly J Dougherty ⁶, Kazuhito Toyo-Oka ^7,^✉

PMCID: PMC8841558 PMID: 34347028

Abstract

Neuromorphological defects underlie neurodevelopmental disorders and functional defects. We identified a function for Rpsa in regulating neuromorphogenesis using in utero electroporation to knockdown Rpsa, resulting in apical dendrite misorientation, fewer/shorter extensions, and decreased spine density with altered spine morphology in upper neuronal layers and decreased arborization in upper/lower cortical layers. Rpsa knockdown disrupts multiple aspects of cortical development, including radial glial cell fiber morphology and neuronal layering. We investigated Rpsa’s ligand, PEDF, and interacting partner on the plasma membrane, Itga6. Rpsa, PEDF, and Itga6 knockdown cause similar phenotypes, with Rpsa and Itga6 overexpression rescuing morphological defects in PEDF-deficient neurons in vivo. Additionally, Itga6 overexpression increases and stabilizes Rpsa expression on the plasma membrane. GCaMP6s was used to functionally analyze Rpsa knockdown via ex vivo calcium imaging. Rpsa-deficient neurons showed less fluctuation in fluorescence intensity, suggesting defective subthreshold calcium signaling. The Serpinf1 gene coding for PEDF is localized at chromosome 17p13.3, which is deleted in patients with the neurodevelopmental disorder Miller–Dieker syndrome. Our study identifies a role for Rpsa in early cortical development and for PEDF-Rpsa-Itga6 signaling in neuromorphogenesis, thus implicating these molecules in the etiology of neurodevelopmental disorders like Miller–Dieker syndrome and identifying them as potential therapeutics.

Keywords: calcium imaging, dendrite formation, dendritic spines, in utero electroporation, ubiquitination

Introduction

Neuronal morphogenesis transforms an immature spherical neuron into a mature neuron with a complex structure. Investigation of the mechanisms that drive neuromorphogenesis has clear applications for improving treatments for neurodevelopmental disorders, as improper neurite formation and dendritic development have been strongly associated with intellectual disability and autism spectrum disorder (Tohgo et al. 1994; Parrish et al. 2007; Ammar et al. 2013; Van Maldergem et al. 2013; Wang et al. 2013). Inappropriate dendritic arborization may also impact inputs and signaling efficiency between the synapse and soma (Häusser et al. 2000; Hoshiba et al. 2016). To gain an understanding of the role that a signaling mechanism plays in regulating neuromorphogenesis, there are multiple developmental stages that must be investigated including dendrite formation, dendritic arborization, and spine formation. Defects at any of these stages of morphogenesis could directly impact subsequent stages, leading to multiple levels of defects. Cortical pyramidal neurons have one apical dendrite, which typically extends directly toward the cortical plate. Apical dendrite orientation is crucial in determining synaptic connectivity and is important for neuronal function, as misorientation of the apical dendrite could cause formation of aberrant connections (Ye et al. 2008). The density and morphology of dendritic spines is another important aspect of neuromorphogenesis. Dendritic spines are the site of over 90% of excitatory synapses in the central nervous system (Harris and Kater 1994). Spine morphology directly relates to synapse function and altered spine phenotypes can result from neurodevelopmental disorders (Fiala et al. 2002; Nimchinsky et al. 2002).

While neuronal morphogenesis is a critical aspect of cortical development, the earlier foundational stage of neurodifferentiation must also be considered. Division of radial glial cells (RGCs) is responsible for directly or indirectly generating most neurons in the brain (Paridaen and Huttner 2014). Asymmetrical RGC division produces an RGC and a neuron, either directly or via production of an intermediate progenitor cell (IPC) (Pontious et al. 2008). This occurs in the ventricular zone (VZ), following which the IPCs will migrate to the subventricular zone and divide to generate 2 neurons (Toyo-oka et al. 2014; Cornell and Toyo-Oka 2017). Newly generated neurons will migrate through the intermediate zone to the cortical plate using RGCs to facilitate their movement and form the cortical layers, with deeper cortical layers forming before the outer layers. An understanding of cortical layering is necessary to gain an appreciation for the functional anatomy of the brain. Each layer has distinctive inputs, outputs, and morphological characteristics. Thus, the morphogenesis of neurons in each layer may be differentially regulated.

Ribosomal protein SA (Rpsa) is also known as the 67-kDa laminin receptor, which is produced from maturation of a 37-kDa precursor protein via an unconfirmed mechanism that has been hypothesized to involve SUMOylation, heterodimerization with an unknown binding partner, homodimerization, or fatty acylation (Rao et al. 1989; Landowski et al. 1995; Buto et al. 1998; Digiacomo et al. 2015). Rpsa functions in a variety of roles including cell anchoring via laminins, ribosomal biogenesis, and chromatin and histone binding (DiGiacomo and Meruelo 2016). Such functional diversity is made possible by Rpsa’s presence in the plasma membrane, ribosomes, cytosol, and nucleus (DiGiacomo and Meruelo 2016). The 37-kDa precursor form is found in the nucleus and cytoplasm and associates with the 40S small subunit of ribosomes, while the 67-kDa form of Rpsa is a cell surface receptor that is embedded in the plasma membrane and may be concentrated in lipid rafts, suggesting its function as part of a signaling complex (Fujimura et al. 2005; Nelson et al. 2008; Ben-Shem et al. 2011; Malygin et al. 2011; Anger et al. 2013). A transmembrane domain for mature 67-kDa Rpsa has been proposed by multiple models (Castronovo et al. 1991; DiGiacomo and Meruelo 2016). Rpsa’s role as a laminin receptor has been extensively studied, revealing its importance in interacting with the extracellular matrix (Lesot et al. 1983; Malinoff and Wicha 1983; Rao et al. 1983; DiGiacomo and Meruelo 2016). Additionally, Rpsa initiates signaling for protection against cell death induced by serum withdrawal in Neuroscreen-1 cells upon treatment with laminin-1, demonstrating an important role for Rpsa in a neuron-like cell line (Gopalakrishna et al. 2018). However, the specific role of Rpsa in cortical development remains unknown.

Rpsa has been shown to directly bind the secreted glycoprotein pigment epithelium-derived factor (PEDF) by a yeast two-hybrid assay, confirming that Rpsa is a PEDF receptor (Bernard et al. 2009). PEDF has neurotrophic properties and protects neurons in a variety of regions throughout the central nervous system against excitotoxicity and oxidative damage (Tombran-Tink and Barnstable 2003). Recently, PEDF has been shown to promote axon regeneration and functional recovery in dorsal root ganglion neurons after spinal cord injury (Stevens et al. 2019). PEDF expression declines with age and is downregulated by more than 100-fold in aged human fibroblast as compared with young human fibroblast, suggesting an important role for PEDF in early development (Pignolo et al. 1993). Additionally, Serpinf1, coding for the PEDF protein, is encoded in a clinically relevant region of chromosome 17p13.3 known as the Miller–Dieker syndrome (MDS) critical region that is frequently deleted or duplicated in a variety of neurodevelopmental disorders (Shimojima et al. 2010; Blazejewski et al. 2018). Since rat cerebral cortical neurons are known to secrete PEDF, the relationship between PEDF and its receptor Rpsa in the developing cortex is of interest (Sanchez et al. 2012).

Binding of PEDF to Rpsa may involve a receptor complex that includes the integrin subunit α6 (Itga6), which binds to the β4 subunit to form a complete integrin. Co-localization of Rpsa and Itga6 on the plasma membrane may result from initial co-localization of Rpsa and Itga6 in cytoplasmic complexes, leading to trafficking to the membrane together (Romanov et al. 1994; DiGiacomo and Meruelo 2016). Furthermore, Rpsa and Itga6 expression may be co-regulated (Ardini et al. 1997). The Rpsa-Itga6 complex is hypothesized to have a role in regulating or stabilizing Rpsa’s interaction with laminin, suggesting that an interaction between Rpsa and Itga6 could be important for other signaling mechanisms like PEDF-Rpsa signaling (Ardini et al. 1997). However, the possibility of a PEDF-Rpsa-Itga6 signaling mechanism and whether this mechanism is important for cortical development remains completely unexplored.

We analyzed the functions of Rpsa in cortical neuronal morphogenesis by performing in utero electroporation at embryonic day (E)15.5 to induce knockdown (KD) of Rpsa in upper layer pyramidal neurons in the mouse cortex (Sehara et al. 2010; Matsui et al. 2011; Quiquempoix et al. 2018). We found that Rpsa-deficient cells show defects in apical dendrite orientation, initiation and elongation of dendrites, dendritic branching, and dendritic spine density and morphology in vivo. To determine if Rpsa has a role in regulating other neuronal layers, we performed in utero electroporation at E13.5 to target lower layer pyramidal neurons for Rpsa KD and observed that only dendritic arborization was disrupted. Additionally, Rpsa KD impacts RGC morphology and neuronal layering. Similar defects are observed following Rpsa KD, PEDF KD, and Itga6 KD at E15.5. Rpsa overexpression (OE) rescued morphological defects resulting from PEDF KD in vivo, suggesting that PEDF initiates Rpsa signaling to regulate neuromorphogenesis. Treatment of control primary cortical neurons with PEDF results in increased neurite length. However, PEDF treatment cannot rescue neurite length defects in Rpsa-deficient neurons, suggesting that neurite extension is primarily regulated by PEDF via Rpsa. Itga6 OE rescued dendrite formation deficits caused by PEDF KD. Itga6 OE also increases and stabilizes Rpsa expression on the plasma membrane by preventing ubiquitination of Rpsa, thus indicating an important role for Itga6 in this signaling mechanism. Additionally, we show that morphological changes associated with Rpsa KD impact subthreshold calcium activity. Our study investigates Rpsa as a key regulator of cortical neuromorphogenesis, with different roles in regulating the morphogenesis of neurons in at least 2 cortical layers and in regulating RGC fiber morphology implicating the PEDF-Rpsa-Itga6 signaling pathway in multiple aspects of cortical development and the etiology of neurodevelopmental disorders.

Materials and Methods

Mice

All animal experiments were performed under protocols approved by the Drexel University Animal Care and Use Committees and following the guidelines provided by the US National Institutes of Health. Institute of Cancer Research (ICR) mice were purchased from Taconic Inc. Embryonic day (E) 0.5 was defined as noon of the day the vaginal plug appeared. Females and males were used for in utero electroporation and primary cortical neuron culture.

Plasmids

CRISPR was chosen as the method to accomplish Rpsa KD, as opposed to shRNA, because we were not able to identify a specific shRNA target sequence. We tested Rpsa shRNA but observed off-target phenotypes that could not be rescued by shRNA-resistant Rpsa OE (data not shown). Thus, Rpsa KD via CRISPR was selected as the method to accomplish a specific KD. pCAG-eCas9-GFP-U6-gRNA was a gift from Jizhong Zou (Addgene plasmid # 79145; http://n2t.net/addgene:79145; RRID:Addgene_79 145). This plasmid contains high-fidelity eSpCas9 to reduce the off-target effects. gRNAs were designed using the web-based design tools, including CHOPCHOP (http://chopchop.cbu.uib.no/) and CRISPy-web (https://crispy.secondarymetabolites.org/#/input), and then cloned into pCAG-eCas9-GFP-U6-gRNA. Rpsa target gRNA sequence is ATCTACAAAAGGAAAAGTGA(CGG) (112-131 of mouse Rpsa). For calcium imaging, GFP in pCAG-eCas9-GFP-U6-control-gRNA and pCAG-eCas9-GFP-U6-Rpsa-gRNA was replaced into tdTomato by PCR. 6XHis-tagged Rpsa was cloned into pCAGItdTomato vector (pCAGItdTomato-Rpsa) as described previously (Wachi et al. 2016). CRISPR-resistant Rpsa OE plasmid was created by PCR. The sense and antisense primers containing the CRISPR target sequence with mutations were designed, and PCR was performed using PrimeSTAR GXL (Takara) with pCAGItdTomato-Rpsa. Ten mutations without amino acid change were introduced into primers. The sequence with mutation is ATTTATAAGCGCAAGTCAGA(TGG) where underlined nucleotides are mutated. The insertion of mutations was confirmed by sequencing. To create pCAGEN-GCaMP6s, pCAGEN and pGP-CMV-GCaMP6s were obtained from Addgene, and GCaMP6s fragment amplified by PCR was cloned into pCAGEN. pGP-CMV-GCaMP6s was a gift from Douglas Kim (Addgene plasmid # 40753; http://n2t.net/addgene:40753; RRID:Addgene_40 753) (Chen et al. 2013). pCAGEN was a gift from Connie Cepko (Addgene plasmid # 11160; http://n2t.net/addgene:11160; RRID:Addgene_11 160) (Matsuda and Cepko 2004). PEDF and Itga6 shRNAs were designed using the web-based tools described above and cloned into pSCV2-Venus plasmid as described previously (Hand and Polleux 2011; Wachi et al. 2016). The target sequence of mouse PEDF is GAACTTGACCATGATAGAA (849-867). The target sequence for mouse Itga6 shRNA is GACCAAAGACTCGATGTTT (1113-1131). Certified scramble shRNA (ACTACCGTTGTTATAGGTG) was also used as a negative control (Invitrogen). All plasmids used in this study were purified by NucleoBond Xtra purification kit (MACHEREY-NAGEL). pLenti-CMV-Itga6-Myc-DDK-P2A-Puro vector was purchased from Origene Technologies, Inc. (PS100092). Itga6-Myc-DDk fragment was amplified by PCR and cloned into pLV-CAG-P2A-mScarlet plasmid. HA-Ubiquitin was a gift from Edward Yeh (Addgene plasmid # 18712; http://n2t.net/addgene:18712; RRID:Addgene_18 712) (Kamitani et al. 1997).

Analysis of Genomic Alterations by CRISPR/Cas9

pCAG-eCas9-GFP-U6-Rpsa-gRNA plasmid was transfected into mouse Neuro-2a cells using PolyJet transfection reagent (SignaGen Laboratories), and genomic DNA was isolated and subjected to PCR to amplify the 433 bp fragment containing gRNA target sequence using Q5 High Fidelity DNA polymerase (NEB) and primers (GAATTC(EcoRI)/GAGTTCTAGTGTCAGAAGAAAAAAGATGAATTTTATTCC and GGATCC(BamHI)/AGCTTTAATAGTGTGCAGGGTCAGTCAG). PCR products were purified by PCR clean-up/Gel extraction kit (MACHEREY-NAGEL) and cloned into pBluescript SK (+). Plasmid DNA isolated from the transformed bacteria was sequenced.

Antibodies

The primary and secondary antibodies used in this research were as follows: 1:200 Anti-Rpsa (Santa Cruz, sc-376 295), 1:500 Anti-GAPDH (Proteintech, 60004-1-Ig), 1:2000 Anti-PEDF (Santa Cruz, sc-16 956), 1:500 Anti-His-Tag (Proteintech, 66005-1-Ig), 1:500 Anti-βIII tubulin (Thermo Scientific Pierce, 2G10), 1:1000 Anti-DYKDDDDK epitope (FLAG) tag (Thermo Scientific Pierce, MA1-91878), 1:500 Anti-HA (Thermo Scientific Pierce, 26183), 1:500 Anti-E-Cadherin (Cell Signaling Technology, #3195), 1:200 Anti-GFAP (DAKO – Fischer Scientific, z-0334), 1:5000 HRP-conjugated donkey-anti-mouse IgG, 1:5000 HRP-conjugated donkey-anti-rabbit IgG, 1:5000 HRP-conjugated donkey-anti-goat IgG, 1:200 Cy5-conjugated donkey-anti-goat IgG, and 1:200 TRITC-conjugated donkey-anti-mouse IgG (Jackson ImmunoResearch Laboratories, 715-025-151). To check the specificity of immunostainings, negative controls were included in which only secondary antibody, and no primary antibody, was used, so that autofluorescence could be accounted for (data not shown).

Histology and Immunohistochemistry

To analyze Rpsa expression and RGC morphology using GFAP as a marker, brains were dissected at E18.5 and fixed with paraformaldehyde/phosphate-buffered saline overnight at 4 °C. Fixed samples were cryoprotected by 25% sucrose/phosphate-buffered saline for 48 h at 4 °C and then embedded with O.C.T. compound (Sakura). Cryo-sections (30 μm thickness) were cut by cryostat (Microm HM505 N) and air dried. Sections were rinsed 3 times in Tris-buffered saline and treated with 0.2% Triton X-100/Tris-buffered saline for 10 min at room temperature, followed by blocking for 30 min in 5% bovine serum albumin/phosphate-buffered saline supplemented with 0.25% Tween-20 to prevent nonspecific binding. Primary antibodies were diluted in blocking buffer, and sections were incubated in primary antibody overnight at 4 °C. All secondary antibodies were diluted with blocking buffer and sections were incubated in secondary antibody for 30 min at room temperature. Sections were stained by 4',6-diamidino-2-phenylindole, dihydrochloride (DAPI, 600 nM) and embedded with 90% glycerol made with Phosphate-buffered saline.

To analyze neuronal morphology, brains were dissected at E14.5, E15.5, E16.5, and E17.5 or postnatal day (P) 3 or P15 and fixed with paraformaldehyde/phosphate-buffered saline overnight at 4 °C. Fixed samples were processed as described above. Cryo-sections (60 μm thickness) were cut and stained by DAPI.

Preparation and Culture of Primary Cortical Neurons

Primary mouse cortical neurons were harvested from embryonic mice at E15.5, prior to gliogenesis at E17.5, to allow for the exclusive culture of neurons (Qian et al. 2000). Neurons were plated on coverslips coated with PLL and laminin. For experiments in which transfection was necessary, neurons were transfected with the appropriate plasmid via nucleofection before plating. Neurons treated with recombinant PEDF (rPEDF) (Sino Biological, 50235-M08H) received 100 ng/mL rPEDF and were fixed at DIV2 after replating. To confirm the efficacy of the KD plasmids, neurons were fixed using 4% PFA at DIV 6 and stained with appropriate antibodies. The DIV 6 time-point was selected because neurons will have progressed through all stages of polarization by this time (Tahirovic and Bradke 2009). To test the expression of PEDF, Rpsa, and Itga6 at a later stage after dendritic spines have developed, primary cortical neurons prepared at E15.5 were fixed and analyzed at DIV14.

In Utero Electroporation

In utero electroporation was performed as previously described (Tabata and Nakajima 2001; Cornell et al. 2016; Wachi et al. 2016). Briefly, pregnant dams were anesthetized with Avertin and the uterine horn was exposed. Plasmids (1 μg) were then injected into the lateral ventricle of either E13.5 or E15.5 embryo brains. Electroporation at E13.5 will allow for transfection of RGCs that will differentiate into lower layer cortical neurons (Cánovas et al. 2015). Electroporation at E15.5 will allow for transfection of cortical pyramidal neurons in upper layers (Sehara et al. 2010; Matsui et al. 2011; Quiquempoix et al. 2018; Liu et al. 2021). Embryo heads were placed between electrodes with the positive anode angled toward the cortex and 3 electric pulses of 35 V were applied with 50 ms intervals by a CUY21 electroporator (Nepa GENE). The developmental timing of the procedure and the electrode angle, which determined which neurons incorporate plasmid, allowed neurons in the somatosensory cortex to be transfected (Taniguchi et al. 2012). Embryos were placed back into the uterus and allowed to develop uninterrupted until brain samples were collected at P3 or P15.

Analysis of Neuronal Morphology

All images were obtained using a confocal microscope (Leica SP8) and the experimenter was blind to the phenotype at the time of imaging.

Apical Dendrite Orientation at P3

The angle at which the apical dendrite extends with respect to the cortical plate at P3 was measured using the ImageJ software angle tool. A straight line perpendicular to the edge of the tissue was used as a reference point to start measuring the angle. Absolute values of the measured angles were used to calculate mean angles. Standard deviation projection images from z-projection photos produced from z-stack data were used for analysis.

Neuronal Morphology Analysis at P15

ImageJ software measuring features were used to analyze neuronal morphology of upper and lower layer cortical pyramidal neurons. Standard deviation projection images from z-projection photos produced from z-stack data were used for analysis. Sholl analysis was performed using the Sholl Analysis Plugin (Gosh Lab, UCSD) for ImageJ following the developer instructions.

For cells electroporated at E13.5 and analyzed at E15.5 or electroporated at E15.5 and analyzed at E17.5, the angle of cell division was measured using the ImageJ angle tool to measure spindle orientation of RGCs relative to the ventricular surface, which is a well-documented marker of the division angle (Théry et al. 2007; Siller and Doe 2009).

RGC process morphology in cells electroporated at E13.5 and analyzed at E14.5 or in cells electroporated at E15.5 and analyzed at E16.5 was assessed as previously described (Hartfuss et al. 2003). Briefly, radial processes reaching the pial surface were classified as long, radial processes ending below the cortical plate were classified as short, radial processes without a basally oriented process were classified as club-shaped, and radial processes terminating with a growth cone-like structure were classified as such.

Dendritic Spine Morphology Analysis at P15

Spines of upper layer neurons were objectively characterized based on geometric characteristics. Spines longer than 1 μm were classified as filopodia, while spines shorter than 1 μm were classified as thin. Stubby and mushroom spines were classified based on morphological appearance. Spines with 2 heads were classified as branched. Photos of spines were taken at ×63 with ×5 zoom and standard deviation projection images were used for analysis.

Subcellular Fractionation

The subcellular fractionation kit (NBP2-47659) from Novus Biologicals was used to fractionate cells according to the protocol provided by the manufacturer. Briefly, COS-1 (Fig. 7A) or N-2a (Fig. 7B) cells were transfected with appropriate plasmids and grown to confluency (~90%) on 10 cm dishes. Cells were collected and the subcellular fractionation was performed by the addition of a series of buffers provided by the manufacturer and centrifugation steps as directed. The entire subcellular fractionation protocol was performed at 4 °C.

Itga6 OE increases and stabilizes Rpsa expression on the plasma membrane. (A) Western blot showing Rpsa and Itga6 expression in the membrane and cytosolic subcellular fractions after either empty backbone control plasmid OE, Rpsa OE, or Rpsa OE + Itga6 OE and subcellular fractionation. E-cadherin was used as a membrane fraction marker to show that the subcellular fractionation was effective. (B) Rpsa expression levels on the plasma membrane of N-2a cells. Subcellular fractionation was completed to isolate the membrane fraction. Fractionation was performed at 0, 3, 6, 12, 24, and 48 h following cycloheximide treatment. (C) Pull-down assay showing decreased ubiquitination in Rpsa OE + Itga6 OE cell lysate, as compared with Rpsa OE only lysate.

Pull-down Assay

Immunoprecipitation was performed as previously described (Toyo-oka et al. 2014; Cornell et al. 2016). Briefly, N-2a cells were transfected with either HA-Ubiquitin + His-Rpsa + FLAG-Control OE or HA-Ubiquitin + His-Rpsa + FLAG-Itga6 and grown to confluency (~90%) on 10 cm dishes. Cells were lysed by NP-40 lysis buffer: 1 M Tris pH 7.4, 5 M NaCl, 0.5 M EDTA, 20% NP-40, H₂O. Supernatant was immunoprecipitated by anti-6XHis antibody-conjugated agarose beads (Santa Cruz). After being thoroughly washed by washing buffer, immunoprecipitated proteins were subjected to western blot and blotted with an anti-HA antibody to detect ubiquitination levels. We also blotted with anti-His and anti-FLAG antibodies to confirm expression of Rpsa and Itga6, respectively.

Cycloheximide Treatment

For the analysis of Rpsa turnover rate on the plasma membrane, cells were treated with cycloheximide (1 mg/mL) 48 h after transfection. After 0, 3, 6, 12, 24, and 48 h, subcellular fractionation was performed as described above to isolate the cytosolic and membrane fractions.

Calcium Imaging of Live Brain Slices

Slice Preparation

Brain slices were prepared using a modified version of a previously described protocol (Wachi et al. 2016). Briefly, male and female mice (approximately P30) were decapitated and brains were quickly removed and placed in ice cold high sucrose artificial cerebrospinal fluid (ACSF) solution for slicing. Brains were embedded in 4% low-melting agarose and coronal cortical slices were generated (300 μm) with a vibrating microtome (VTS1000 Leica Microsystems). Slices were next incubated for 60 min at 37 °C in DMEM/F-12 imaging media without phenol red supplemented with 10% fetal bovine serum.

Imaging of Spontaneous Activity

The experimenter was blind to the phenotype at the time of the imaging. Slices were gently transferred into glass bottom 35 mm dishes (MatTek) for imaging. A membrane was placed on top of the slices to reduce movement during imaging. Time-lapse live imaging was performed using an inverted fluorescent microscope (Zeiss, Axio Observer Z1) with a ×20 objective. Images were captured every 50 ms for 1 min while the slices were maintained at 37 °C with a stage top incubator (Zeiss).

Calcium Imaging Data Analysis

All cells included in the analysis were double-positive for GCamp6s and either pCAG-eCas9-tdTomato-U6-control-gRNA or pCAG-eCas9-tdTomato-U6-Rpsa-gRNA. All image analysis was performed using Zen 2 Pro analysis software (Zeiss 2011). Circular regions of interest were placed on the cell soma. Baseline fluorescence (F₀) was obtained by averaging the fluorescence intensity inside the region of interest immediately before beginning the time course imaging. Images were captured every 50 ms for 1 min. Fluorescence intensity for the time course was measured by averaging all pixels in the region of interest at each frame of the imaging (Chen et al. 2013). Percent change in fluorescence (ΔF/F₀) was calculated as (F_measured − F₀)/F₀ for every frame of the time course.

Electrophysiological Recordings

Slice Preparation

Mice (approximately P30) were decapitated and brains were quickly removed and placed in ice cold sucrose solution containing the following (in mM): 87 NaCl, 75 sucrose, 25 glucose, 25 NaHCO₃, 1.25 NaH₂PO₄, 2.5 KCl, 0.25 CaCl₂, and 3.5 MgSO₄ for slicing. Coronal cortical slices were generated (300 μm) with a vibrating microtome (Leica Microsystems). Slices were next incubated for 30 min at 37 °C in recording ACSF containing the following (in mM): 111 NaCl, 3 KCl, 11 glucose, 25 NaHCO₃, 1.3 MgSO₄, 1.1 KH₂PO₄, and 2.5 CaCl₂. Slices were then maintained at room temperature in ACSF for at least 30 min before recording. Both slicing solutions and recording solutions were continuously aerated with 95%/5% CO₂/O₂.

Recordings

The experimenter was blind to the phenotype at the time of the recording. All recordings were performed at room temperature. Cortical neurons transfected with either the control CRISPR plasmid or the Rpsa CRISPR plasmid were visualized based on their fluorescence (tdTomato) with a ×63 objective lens on a BX51WI scope (Olympus) using LED illumination (X-cite). Patch electrodes (Harvard Apparatus) were pulled to tip resistances of 5–8 M'Ω using a multistage puller (Sutter Instruments) and were filled with K-gluconate-based intracellular solution, containing the following (in mM): 128 K-gluconate, 10 HEPES, 0.0001 CaCl₂, 1 glucose, 4 NaCl, 5 ATP, and 0.3 GTP. Data were collected with a Multiclamp 700B amplifier (Molecular Devices) and Clampex software (pClamp9, Molecular Devices). Signals were digitized at 20 kHz and filtered at 4 kHz.

Resting membrane potential of the neuron was recorded in current-clamp mode immediately after breaking in. For passive and active membrane properties, current- and voltage-clamp protocols were performed as previously described (Chen et al. 2013). Briefly, input resistance was measured from a series of hyperpolarizing steps in voltage clamp using Clampfit. Time constant, tau, was calculated from the standard exponential fit of hyperpolarizing steps in current clamp. Capacitance was calculated from the input resistance and the tau measurements. Rheobase was defined as the lowest current step (in 10 pA increments) that evoked an action potential in the neuron.

Quantification of Corrected Total Cell Fluorescence

To quantify immunofluorescence staining, we used the corrected total cell fluorescence (CTCF), which accounts for both the background fluorescence and the size of the cell being quantified. ImageJ was used to measure the area, mean gray value, and raw integrated density for both the background fluorescence and the fluorescence of the cell. CTCF was calculated using the following formula: raw integrated density of the cell – (area of the cell ^* mean gray value of background fluorescence).

Statistical Analysis

Quantitative data were subjected to statistical analysis using SPSS (IBM Analytics), Prism (GraphPad Software), and MATLAB (MathWorks). The data were analyzed by 2-tailed independent-samples t-tests, 1-way ANOVA, or 2-way ANOVA when appropriate. Values represented as mean ± SEM. Results were deemed statistically significant if the P value was <0.05. ^*, ^*^*, and ^*^*^* indicate P < 0.05, P < 0.01, and P < 0.001, respectively.

Results

Expression of Rpsa in the Developing Cerebral Cortex

The expression pattern of Rpsa has not been previously investigated in the developing brain. Therefore, it was unknown at which levels or developmental time-points Rpsa is expressed, which should be established when investigating the function of Rpsa during development. To quantify the expression of Rpsa protein levels in the developing cerebral cortex, we performed western blots using lysate of the mouse cerebral cortex from E13.5 to P7 samples (Fig. 1A,B). The Rpsa protein is moderately expressed in the developing cortex during early embryonic stages (E13.5) and then expression decreases but persists to at least P7. This suggests that Rpsa expression may be most important at earlier embryonic stages. Staining of primary cortical neurons confirmed the expression of Rpsa in neurons by staining with the neuronal marker, class III β-tubulin, showing that Rpsa is highly concentrated in the soma and proximal extensions but is not found in the nucleus or more distal extensions (Fig. 1C). We also analyzed the spatial distribution of the Rpsa protein by immunofluorescence using E18.5 brain sections (Fig. 1D). E18.5 was the earliest developmental time-point at which the formation of all cellular layers was complete (Molnár et al. 2006). Rpsa is expressed throughout all cortical layers and in the cytoplasm, but not the nucleus at E18.5. Therefore, functions of Rpsa that can be associated with these cellular localizations, such as interacting with the extracellular matrix, are likely more important at this time-point. Additionally, Rpsa appears to be expressed in the soma of RGCs, but not the glial fibers (Fig. 1E).

Rpsa KD in Upper Layer Neurons Is Associated with Apical Dendrite Misorientation at P3

To test the effects of Rpsa KD on one of the earliest stages of neuromorphogenesis, we analyzed apical dendrite morphology at P3. Extension of the apical dendrite is the first step of dendritic morphogenesis, and this is usually the only dendritic extension present at P3. KD efficiency of our Rpsa CRISPR construct was tested via western Blot and the KD efficiency was approximately 77% (Supplemental Fig. 1A,B). Genomic alteration by Rpsa CRISPR was mainly accomplished by deletions, but cases of single nucleotide additions were also observed (Supplemental Fig. 1C,D). To further confirm Rpsa KD, we performed immunostaining for Rpsa in primary cortical mouse neurons that were transfected with Rpsa CRISPR. Neurons positive for Rpsa CRISPR (43881.13 CTFC ±5156.70) had significantly decreased fluorescence intensity for Rpsa, as compared with untransfected controls (85504.51 CTFC ±9189.28) (Supplemental Fig. 1E,F).

To investigate apical dendrite orientation, we performed in utero electroporation at E15.5 to induce a CRISPR/Cas9-mediated KD of Rpsa in upper layer cortical pyramidal neurons. The orientation of the apical dendrite is an important feature of neuronal morphology, since the angle at which the apical dendrite extends with respect to the cortical plate is important for neuronal connectivity (Demyanenko et al. 2004; Kvajo et al. 2008; Ye et al. 2008). Conducting this analysis at P3 allowed for easy identification of the apical dendrite, since neuronal morphology is relatively simple at this stage. We found that Rpsa KD cells (34.2° ± 3.61) had a significantly greater mean angle from the soma than the control CRISPR cells (14.1° ± 1.63) (Fig. 2A,B). The frequencies for degree of angle in Rpsa KD cells were more broadly distributed across a wider range from −30° to 30°, while angle measurements were closer to 0° and more narrowly distributed in control CRISPR cells (Fig. 2C). This suggests that Rpsa-deficient cells have apical dendrites that are inappropriately positioned at an abnormal angle, which may lead to formation of aberrant synapses. The Rpsa KD phenotype could be rescued by CRISPR-resistant wild-type Rpsa OE (11.8° ± 1.13), resulting in a phenotype comparable to the control CRISPR + control OE group (11.7° ± 1.25) (Fig. 2). The possibility of the phenotype being caused by an off-target effect of the CRISPR is minimal, since the Rpsa KD phenotype was able to be rescued by CRISPR-resistant Rpsa OE.

Rpsa KD of upper layer neurons is associated with apical dendrite misorientation at P3 that can be rescued by Rpsa OE. (A) Neurons transfected via *in utero* electroporation at E15.5, orientation of the apical dendrite measured using the angle at which the apical dendrite extends from the middle of the soma relative to the cortical plate, scale bar = 50 μm. Each representative photo is accompanied by multiple tracings. (B) One-way ANOVA showed a significant difference in mean angle of apical dendrite from soma (F(3,76) = 22.620, P < 0.0005). Tukey post hoc test showed degree of angle of apical dendrite from middle of soma is significantly greater for the Rpsa CRISPR group (30.4° ± 3.09) compared with control CRISPR (10.7° ± 1.20) (P < 0.0005). There was no significant difference between Rpsa CRISPR + Rpsa OE (10.8° ± 1.71) and control CRISPR + control OE groups (12.5° ± 1.49) (P = 0.924). For all groups, n = 21 cells from 3 mice. Data are represented as mean ± SEM. (C) Dendrograms show frequency distribution of angle measurement.

Additionally, we measured the length of the apical dendrite and found no significant difference between Rpsa KD (27.7 μm ± 1.4) and control CRISPR (27.5 μm ± 1.6) (Supplemental Fig. 2). This suggests that there is no defect in the early period of apical dendrite elongation.

Rpsa KD in Upper Layer Neurons Is Associated with Defects in Neuronal Morphogenesis at P15

To test whether Rpsa KD also affected later stages of dendrite morphogenesis, we analyzed dendritic morphology at P15 in upper layer cortical pyramidal neurons. A decrease in the number of dendrites extending from the soma suggests that defective dendrite initiation has occurred, while a decrease in dendrite length suggests a defect in elongation of dendrites. These early pivotal processes have implications for the remaining stages of neuronal morphogenesis, such as dendritic arborization. In utero electroporation was used to induce a CRISPR/Cas9-mediated KD of Rpsa at E15.5 and neuronal morphology was analyzed at P15, since neuritogenesis is well established by this time-point. All Rpsa-deficient neurons reached the cortical plate (CP) by P15, but the somas were more broadly distributed (Supplemental Fig. 3A–C). The Rpsa KD neurons had significantly fewer and shorter extensions with less complex branching as compared with the control CRISPR neurons (Fig. 3A). The mean number of dendrites from the soma in Rpsa-deficient cells was 2.9 ± 0.24, while the mean number of dendrites, including apical and basal dendrites, from the soma of control neurons was 4.2 ± 0.24 (Fig. 3B). The mean dendrite length in Rpsa-deficient cells was 20.9 μm ± 2.0 as compared with control neurons with a mean dendrite length of 49.5 μm ± 2.2 (Fig. 3C). Additionally, Sholl analysis of Rpsa-deficient neurons indicates that the Rpsa KD neurons have a statistically significant defect in dendritic branching at radial distances 20–75 μm (Fig. 3D). The Sholl profile is consistent with our analysis of dendrite number and length, since it also indicates that Rpsa-deficient cells have fewer and shorter dendrites than controls. All described morphological deficits from Rpsa KD were rescued by CRISPR-resistant Rpsa OE, making it unlikely that the observed phenotype was due to off-target effects (Fig. 3). This suggests that Rpsa signaling is crucial in regulating appropriate dendrite formation, during both initiation and elongation, in addition to dendritic arborization. Since the apical dendrite was correctly extended as observed at P3 (Supplemental Fig. 2), the defects in the extension of dendrites are mainly in basal dendrites.

Rpsa KD in Lower Layer Neurons Is Associated with Decreased Dendritic Branching and Disrupted Cell Layering at P15

To understand whether Rpsa-mediated signaling has a role on selected types of cortical neurons in a specific layer, we knocked down Rpsa via in utero electroporation at E13.5. Performing the KD at this time-point allowed for lower layer cortical pyramidal neurons to be electroporated. Morphological analysis at P15 showed that the number of dendrites and dendrite length did not significantly differ between controls and Rpsa-deficient cells; however, Rpsa KD resulted in a significant decrease in dendritic branching (Fig. 4A). The mean number of dendrites from the soma in Rpsa-deficient cells was 4.2 ± 0.30, while the mean number of dendrites, including apical and basal dendrites, from the soma of control neurons was 4.4 ± 0.21 (Fig. 4B). The mean dendrite length in Rpsa-deficient cells was 54.74 μm ± 5.2 as compared with control neurons with a mean dendrite length of 58.11 μm ± 3.8 (Fig. 4C). Sholl analysis revealed a significant decrease in dendritic branching at radial distance 25–70 μm (Fig. 4D). We also found that most Rpsa-deficient neurons reached near the CP by P15, but the somas were more broadly distributed (Supplemental Fig. 3D–F). These data indicate that Rpsa signaling has a significant role in mediating dendritic branching in at least 2 different cortical layers, while the role of Rpsa in regulating dendrite number and length may be layer specific. Also, our data implicate Rpsa in neuronal layering.

Rpsa KD of lower layer neurons is associated with decreased dendritic branching and disrupted cell layering at P15. (A) Neurons transfected via *in utero* electroporation at E13.5. Scale bars = 50 μm. (B) Two-tailed independent-samples t-test showed no significant difference between mean number of dendrites in Rpsa CRISPR neurons (4.2 ± 0.30) compared to control CRISPR neurons (4.4 ± 0.21), t(38) = 0.680, P = 0.51. For both groups, n = 20 cells. Data are represented as mean ± SEM. (C) Two-tailed independent-samples t-test showed no significant difference between mean dendrite length in Rpsa CRISPR neurons (54.74 ± 5.17) compared with control CRISPR neurons (58.11 ± 3.78), t(38) = 0.526, P = 0.60. For both groups, n = 20 cells. Data are represented as mean ± SEM. (D) Sholl analysis shows dramatic decrease in branching in Rpsa-deficient cells. Multiple unpaired t-tests show a significant decrease in branching for the Rpsa CRISPR group at radial distances 25–70 μm as compared with the control CRISPR group. For all groups, n = 20 cells analyzed from 3 mice.

Rpsa KD Disrupts RGC Process Morphology, but Results in No Significant Change in RGC Spindle Orientation at Multiple Time-points

Since Rpsa is expressed in the somas of RGCs, we analyzed whether Rpsa was involved in early cortical developmental steps. We analyzed the cells at E14.5 after electroporation at E13.5. We classified the morphology of the radial process as either long, short, club, or growth cone as previously described (Fig. 5A) (Hartfuss et al. 2003). We found that Rpsa-deficient RGCs showed an increase in growth cone radial processes that was significantly different than expected based on the controls (Fig. 5B,C). Additionally, we analyzed the cells electroporated at E15.5 at E16.5 and found that Rpsa-deficient RGCs showed an increase in long and growth cone radial processes that was significantly different than expected based on the controls (Fig. 5D,E). Neurons may be generated in the cortex via asymmetrical division of RGCs, which produces one neuron/IPCs and one RGC. Therefore, we analyzed the ratio of asymmetrically versus symmetrically dividing RGCs by measuring the angle of cleavage plan in the VZ or spindle orientation. Symmetrically dividing RGCs divide with a spindle orientation between 70 and 80°, while asymmetrically dividing RGCs divide at an angle of less than 70° (Fig. 5F) (Théry et al. 2007; Siller and Doe 2009). Control CRISPR cells transfected at E13.5 and analyzed at E15.5 divided mostly symmetrically with a mean spindle orientation of 72.01°, while Rpsa-deficient cells transfected at E13.5 and analyzed at E15.5 showed a slight decrease in cleavage angle with a mean spindle orientation of 68.37°, although this change was not significant (Fig. 5G). Both control CRISPR and Rpsa CRISPR cells transfected at E15.5 and analyzed at E17.7 divided mostly symmetrically with a mean spindle orientation of 74.03° and 73.92°, respectively (Fig. 5H). This analysis shows that Rpsa KD does not impact the cleavage angle of RGCs.

Rpsa KD disrupts RGC radial process morphology, but results in no significant change in RGC spindle orientation at multiple time-points. (A) Representative photos of RGC morphology. Blue arrow indicates long, red arrow indicates short, green arrow indicates club, and purple arrow indicates growth cone classifications. Scale bars = 50 μm. VZ, ventricular zone. Dashed line indicates the ventricular surface in photo (ii) and the upper edge of the tissue in photo (iii). (B) RGCs transfected via *in utero* electroporation at E13.5. Dashed line indicates the ventricular surface. Scale bars = 100 μm. (C) Chi-square test for goodness of fit indicated that the percentage of long (25.56%), short (30.00%), club (23.33%), and growth cone (21.11%) RGC morphology types observed in the Rpsa CRISPR group was significantly different than the percentage of long (26.67%), short (35.56%), club (33.33%), and growth cone (4.44%) RGC morphology types observed in the control CRISPR group, P < 0.005. For both groups, n = 90. (D) RGCs transfected via *in utero* electroporation at E15.5. Dashed line indicates the ventricular surface. Scale bars = 100 μm. (E) Chi-square test for goodness of fit indicated that the percentage of long (43.90%), short (21.95%), club (14.63%), and growth cone (19.51%) RGC morphology types observed in the Rpsa CRISPR group was significantly different than the percentage of long (37.40%), short (29.27%), club (25.20%), and growth cone (8.13%) RGC morphology types observed in the control CRISPR group, P < 0.005. For both groups, n = 123. (F) Examples of symmetrically or asymmetrically dividing RGCs in the VZ. Dotted lines indicate the angle of cell division as measured by the angle of cleavage with respect to the plane of the VZ or spindle orientation. (G) Two-tailed independent-samples t-test showed that the mean angle of cleavage plane in VZ for the Rpsa CRISPR group (68.37° ± 1.87) was not significantly different from the control CRISPR group (72.01° ± 1.56), t(38) = 1.493, P = 0.144. For both groups, n = 20 cells. Data are represented as mean ± SEM. (H) Two-tailed independent-samples t-test showed that the mean angle of cleavage plane in VZ for the Rpsa CRISPR group (73.92° ± 2.87) was not significantly different from the control CRISPR group (74.03° ± 2.86), t(38) = 0.027, P = 0.978. For both groups, n = 20 cells. Data are represented as mean ± SEM.

PEDF KD and Itga6 KD result in neuromorphological defects in upper layer neurons similar to Rpsa KD

Since PEDF is a known ligand of Rpsa that has neurotrophic properties and is encoded in the clinically relevant Miller-Dieker Syndrome critical region of chromosome 17p13.3, we investigated whether PEDF binding was responsible for initiating Rpsa signaling to regulate neuronal morphogenesis in upper layer cortical pyramidal cells. We chose to further analyze upper layer neurons because Rpsa KD produced more significant morphological effects on upper cortical layers, as opposed to lower cortical layers. We used PEDF shRNA in combination with in utero electroporation to conduct an initial analysis of PEDF deficient neurons in vivo. KD efficiency of the PEDF shRNA was tested via Western Blot and the knockdown efficiency was approximately 84% (Suppl. Fig. 1A and B). Also, we performed immunostaining for PEDF in primary cortical neurons that were transfected with PEDF shRNA. Neurons positive for PEDF shRNA (11,735.13 CTFC ± 2,019.42) had significantly decreased fluorescence intensity for PEDF, as compared with untransfected controls (25,563.82 CTFC ± 6,629.78) (Suppl. Fig. 1 G and H). All PEDF deficient neurons reached the CP by P15 but showed broader distribution of neurons in the CP compared to the control, like that seen following Rpsa KD (Suppl. Fig. 3 A-C). The initial analysis of dendritic morphology revealed that PEDF deficient neurons showed similar defects to Rpsa deficient neurons, with PEDF KD neurons having significantly fewer and shorter extensions with less complex branching as compared to the scramble shRNA control neurons (Fig. 3 and 6A). The mean number of dendrites from the soma in PEDF deficient cells was 2.6 ± 0.21, while the mean number of dendrites from the soma in control neurons was 4.4 ± 0.25 (Fig. 6B). The mean dendrite length in PEDF deficient cells was 27.0 μm ± 2.6 as compared to control neurons with a mean dendrite length of 50.9 μm ± 3.6 (Fig. 6C). These data indicate that PEDF is important for the initiation and elongation of dendrites. Additionally, Sholl analysis of PEDF deficient neurons indicates that these neurons have a statistically significant defect in dendritic branching at radial distances 20-70 μm (Fig. 6D). The Sholl profile is consistent with our analysis of dendrite number and length, since it also indicates that PEDF deficient cells have fewer and shorter dendrites than controls. PEDF has known roles in axon regeneration (Liu et al. 2016; Stevens et al. 2019; Vigneswara et al. 2013; Vigneswara et al. 2015). Therefore, we analyzed axon bundle length in vivo following PEDF KD and found no significant difference between the mean axon bundle length in PEDF deficient cells, which was 5.4mm ± 0.41, and the mean axon bundle length in control neurons of 5.0mm ± 0.41 (Suppl. Fig. 4).

To determine whether these similar phenotypes could be a result of PEDF and Rpsa functioning in the same signaling pathway, we overexpressed Rpsa in PEDF-deficient cells. Rpsa OE rescued morphological defects in the initiation and elongation of dendrites after PEDF KD in vivo (Fig. 6A). The mean number of dendrites from the soma in PEDF KD + Rpsa OE neurons was 4.2 ± 0.17, which was not significantly different from the mean of 4.35 ± 0.30 for the double transfected control neurons (scramble shRNA + control OE) (Fig. 6B). The mean dendrite length in PEDF KD + Rpsa OE neurons was 44.3 μm ± 3.6, which was not significantly different from the double transfected control neurons (scramble shRNA + control OE) with a mean dendrite length of 44.3 μm ± 3.3 (Fig. 6C). These data indicate that PEDF is involved in Rpsa-mediated initiation and elongation of dendrites. Additionally, Sholl analysis of PEDF KD + Rpsa OE neurons indicates that Rpsa OE was able to compensate for the defect in branching complexity caused by PEDF KD and resulted in an increase in branching (Fig. 6D). Taken together with the fact that PEDF is a ligand of Rpsa, these results suggest that PEDF binding initiates Rpsa signaling to regulate neuronal morphogenesis. To understand more fully how Rpsa regulates neuromorphogenesis, we overexpressed Rpsa alone (Fig. 6A). The mean number of dendrites from the soma in Rpsa OE cells was 5.3 ± 0.26, while the mean number of dendrites from the soma in control neurons was 4.4 ± 0.25 (Fig. 6B). The mean dendrite length in Rpsa OE cells was 76.6 μm ± 6.3 as compared with control neurons with a mean dendrite length of 50.9 μm ± 3.6 (Fig. 6C). Sholl analysis of Rpsa OE neurons did not reveal any significant difference in dendritic branching (Fig. 6D).

Rpsa is known to bind to Itga6 on the plasma membrane (Ardini et al. 1997). To determine if Itga6 is also involved in PEDF-Rpsa signaling, we used Itga6 shRNA in combination with in utero electroporation at E15.5 to conduct an initial analysis of Itga6-deficient upper layer cortical pyramidal neurons in vivo. KD efficiency of the Itga6 shRNA was tested via western blot and the KD efficiency was approximately 79% (Supplemental Fig. 1A,B). In addition, we performed cell staining for Itga6 in primary cortical neurons that were transfected with Itga6 shRNA. Neurons positive for Itga6 shRNA (3947.66 CTFC ±819.61) had significantly decreased fluorescence intensity for Itga6, as compared with untransfected controls (21478.55 CTFC ±6285.73) (Supplemental Fig. 1I,J). Itga6-deficient neurons did not show any deficits in layering with all Itga6 KD neurons reaching the CP by P15 (Supplemental Fig. 3A–C). The analysis revealed that Itga6-deficient neurons showed similar defects to Rpsa- and PEDF-deficient neurons in neuronal morphogenesis (Figs 3 and 6A). The mean number of dendrites from the soma in Itga6-deficient cells was 2.5 ± 0.24, while the mean number of dendrites from the soma in control neurons was 4.4 ± 0.25 (Fig. 6B). The mean dendrite length in Itga6-deficient cells was 26.2 μm ± 3.3 as compared with control neurons with a mean dendrite length of 50.9 μm ± 3.6 (Fig. 6C). These data indicate that Itga6 is a key protein for the initiation and elongation of dendrites. Additionally, Sholl analysis of Itga6-deficient neurons indicates that these neurons have a statistically significant defect in dendritic branching at radial distances 20–70 μm, similar to the defects observed in neurons deficient in Rpsa and PEDF (Figs 3D and 6D).

To determine whether Itga6 is involved in PEDF signaling, we overexpressed Itga6 in PEDF-deficient cells. We found that Itga6 OE in PEDF-deficient neurons can rescue the morphological defects associated with PEDF KD (Fig. 6A). This rescue phenotype was like the control (scramble shRNA + control OE) and PEDF shRNA + Rpsa OE phenotypes. Thus, these results suggest that Itga6 is involved in a PEDF-Rpsa signaling pathway. The mean number of dendrites from the soma in PEDF KD + Itga6 OE neurons was 4.1 ± 0.29, which was not significantly different from the mean of 4.35 ± 0.30 for the double transfected control neurons (scramble shRNA + control OE) (Fig. 6B). The mean dendrite length in PEDF KD + Itga6 OE neurons was 44.8 μm ± 3.4, which was not significantly different from the double transfected control neurons (scramble shRNA + control OE) with a mean dendrite length of 44.3 μm ± 3.3 (Fig. 6C). This indicates that Itga6 is involved both in the initiation and elongation of dendrites mediated by PEDF and Rpsa. Additionally, Sholl analysis of PEDF KD + Itga6 OE neurons indicates that the branching complexity is restored (Fig. 6D). These results suggest that Itga6, a known binding partner of Rpsa, is important for PEDF-Rpsa signaling to regulate dendrite formation during initiation and elongation stages, and dendritic branching.

To further investigate the role of PEDF in neuromorphogenesis and Rpsa signaling, we treated control CRISPR and Rpsa CRIPSR transfected primary cortical neurons with a recombinant PEDF (rPEDF) (Fig. 6E). For mean number of neurites, Rpsa KD (3.1 ± 0.19) decreased the mean number of neurites observed in vitro, but the decrease was not statistically significant (Fig. 6F). The treatment of control cells with rPEDF (4.5 ± 0.38) did not cause significant change in mean neurite number as compared with untreated control cells (4.0 ± 0.23). However, rPEDF treatment of Rpsa KD cells (4.9 ± 0.38) resulted in increased neurite number (Fig. 6F). For mean length of neurites likely to become dendrites, Rpsa KD (29.83 μm ± 2.29) caused a significant decrease (Fig. 6G). The treatment of control cells with rPEDF (127.2 μm ± 11.5) significantly increased the mean length of neurites likely to become dendrites as compared with untreated control cells (58.7 μm ± 4.2). Interestingly, rPEDF treatment was unable to rescue this defect in Rpsa KD cells (40.8 μm ± 3.16) (Fig. 6G). For mean length of neurites likely to become axons, Rpsa KD (70.5 μm ± 6.96) caused a significant decrease (Fig. 6H). The treatment of control cells with rPEDF (340.4 μm ± 34.5) significantly increased the mean length of neurites likely to become axons as compared with untreated control cells (158.0 μm ± 17.0). Again, rPEDF treatment failed to rescue this defect in Rpsa KD cells (128.6 μm ± 13.1) (Fig. 6H). For dendritic branching in untreated cells, Rpsa KD resulted in a statistically significant decrease in dendritic branching at radial distance 25–100 μm and 110 μm (Fig. 6I). The treatment of control cells with rPEDF significantly increased dendritic branching at radial distance 50–150 μm. Treatment with rPEDF was able to increase dendritic branching in Rpsa-deficient cells at radial distances 20–40 μm and 55 μm (Fig. 6I). These data suggest that neurite extension is predominantly regulated by PEDF via Rpsa, but neurite initiation and arborization are also regulated by a non-Rpsa-mediated mechanism.

Itga6 OE Increases and Stabilizes Rpsa Expression on the Plasma Membrane by Preventing Ubiquitination of Rpsa

Rpsa and Itga6 expression have been shown to be co-regulated and Rpsa and Itga6 co-localize in the cytosol, possibly leading to their being trafficked together to the plasma membrane (Romanov et al. 1994; Ardini et al. 1997). We tested whether Itga6 could be increasing Rpsa expression on the plasma membrane, which would allow PEDF-Rpsa signaling to occur more efficiently. We transfected 6XHis-Rpsa (only Rpsa OE) and 6XHis-Rpsa + FLAG-Itga6 (both Rpsa and Itga6 OE) into COS-1 cells and performed subcellular fraction to isolate the cytosolic and membrane fractions. Then, Rpsa expression level in the 6XHis-Rpsa + FLAG-Itga6 group was analyzed and compared with the control OE group and Rpsa OE only group. We found that Rpsa expression on the plasma membrane was increased with Itga6 OE, as compared with when only Rpsa was overexpressed (Fig. 7A). This suggests that Itga6 could be contributing to PEDF-Rpsa signaling to regulate neuronal morphology by increasing the amount of Rpsa available on the plasma membrane to participate in signaling and may explain the similar neuronal morphology phenotypes following Rpsa KD, PEDF KD, and Itga6 KD. We confirmed isolation of the membrane fraction by blotting with the plasma membrane marker E-cadherin.

Since Itga6 OE was shown to increase the amount of Rpsa present in the membrane, we next tested if Itga6 OE could increase the time spent by Rpsa in the membrane before internalization or degradation. We transfected Neuro2a (N-2a) mouse neuroblastoma cells with either 6XHis-Rpsa (only Rpsa OE) or 6XHis-Rpsa + FLAG-Itga6 (both Rpsa and Itga6 OE) and 48 h after transfection treated the cells with cycloheximide, which inhibits protein synthesis. Cells were then subject to subcellular fractionation to isolate the membrane fraction at 0, 3, 6, 12, 24, and 48 h following cycloheximide treatment. Western blot for 6XHis-Rpsa using anti-His antibody was completed on the membrane fraction revealing that Rpsa remained present in the membrane for dramatically longer following cycloheximide treatment when both Rpsa and Itga6 were overexpressed, as compared with when only Rpsa was overexpressed (Fig. 7B). This suggests that Itga6 stabilizes Rpsa in the membrane.

To determine the mechanism by which Itga6 increases and stabilizes Rpsa expression on the plasma membrane, we transfected N-2a cells with HA-ubiquitin and either 6XHis-Rpsa (only Rpsa OE) or 6XHis-Rpsa + FLAG-Itga6 (both Rpsa and Itga6 OE). Ubiquitin has already been shown to regulate the presence of Rpsa at the plasma membrane (Kim et al. 2012). Pull-down assay was done 48 h after transfection to isolate 6XHis-Rpsa. Western blot using anti-HA antibody showed decreased ubiquitination when both Itga6 and Rpsa were overexpressed, as compared with when only Rpsa was overexpressed (Fig. 7C). This suggests that Itga6 stabilizes the expression of Rpsa on the plasma membrane by preventing its ubiquitination, which would lead to internalization or degradation.

Rpsa KD and PEDF KD, but Not Itga6 KD, Cause a Decrease in Overall Spine Density and a Change in Spine Morphology in Upper Layer Neurons

We further investigated the neuromorphological changes observed after Rpsa KD, PEDF KD, and Itga6 KD by determining if there was a change in dendritic spine density and morphology, since these factors rely on the previous steps of dendrite morphogenesis occurring with fidelity and can impact synaptic function. We used in utero electroporation at E15.5 to transfect the relevant plasmids and analyzed spines at P15 in upper layer cortical pyramidal neurons (Fig. 8A). We observed a decrease in overall spine density after Rpsa KD (0.28 ± 0.04) and PEDF KD (0.21 ± 0.04), as compared with the control CRISPR (0.70 ± 0.05) and scramble shRNA (0.97 ± 0.09) controls, respectively (Fig. 8B). The Rpsa KD and PEDF KD groups also showed significant differences in spine morphology (Fig. 8C). There was a significant decrease in mushroom spines following both Rpsa KD (5.88%) and PEDF KD (6.98%) as compared with control CRISPR (55.56%) and scramble shRNA (60.10%), respectively. Thin spines were significantly increased after Rpsa KD (66.18%), while thin spines were significantly decreased following PEDF KD (34.88%) as compared with control CRISPR (22.22%) and scramble shRNA (20.21%), respectively. Rpsa KD also resulted in a significant decrease in stubby spines (10.29%), as compared with control CRISPR (18.06%), and a significant increase in filopodia spines (16.18%) as compared with control CRISPR (2.78%). This indicates that Rpsa KD results in a clear shift toward a more immature spine morphology, while PEDF KD follows this same trend. Interestingly, Itga6 KD (1.01 ± 0.10) resulted in a spine density phenotype like that of the scramble shRNA control group (Fig. 8B). The Itga6 KD and scramble shRNA groups also showed similar spine morphology phenotypes, with no significant differences in spine morphology between these groups (Fig. 8C). This suggests that PEDF-Rpsa signaling regulates spine density in addition to morphology, while Itga6 may not be involved in the regulation of spine formation by PEDF-Rpsa signaling.

Rpsa KD and PEDF KD, but not Itga6 KD, cause a decrease in overall spine density and a change in spine morphology. (A) Neurons transfected via *in utero* electroporation at E15.5, images taken with ×63 objective and ×5 zoom to visualize spines, scale bars = 5 μm. (B) One-way ANOVA determined there was a statistically significant difference (F(6,127) = 23.321, P < 0.0005). Tukey post hoc test revealed that mean spine density per μm was statistically significantly lower for Rpsa CRISPR (0.28 ± 0.04, n = 21) as compared with control CRISPR (0.70 ± 0.05, n = 12) (P = 0.001) and mean spine density per μm was statistically significantly lower for PEDF shRNA (0.21 ± 0.04, n = 20) as compared with scramble shRNA (0.97 ± 0.09, n = 20) (P < 0.0005). However, the Itga6 shRNA group (1.01 ± 0.10, n = 17) did not show a significant difference compared with the scramble shRNA group (P = 0.999). The PEDF shRNA + Rpsa OE group (0.99 ± 0.07, n = 21) was not significantly different from the scramble shRNA + control OE group (0.77 ± 0.08, n = 20) (P = 0.230). Data are represented as mean ± SEM. For each group, spines were analyzed from 3 mice. (C) Graph showing differences in percentage of total spines with filopodia, thin, stubby, mushroom, or branched morphology for each experimental group. Statistical analyses were done using raw spine density per μm data. Branched spines were significantly increased in the PEDF shRNA + Rpsa OE group (0.0381 ± 0.01, n = 21), as compared with scramble shRNA + control OE (0.0050 ± 0.01, n = 20), t(408) = −2.295, P < 0.0005. Mushroom spines were significantly decreased in the Rpsa CRISPR (0.0190 ± 0.01, n = 21, P < 0.0005) and PEDF shRNA (0.0150 ± 0.01, n = 20, P < 0.0005) groups as compared with control CRISPR (0.3333 ± 0.04, n = 12) and scramble shRNA (0.5800 ± 0.03, n = 20), respectively, while mushroom spines were significantly increased in the PEDF shRNA + Rpsa OE (0.4619 ± 0.03, n = 21, P = 0.004) group compared with the scramble shRNA + control OE (0.3850 ± 0.03, n = 20). Stubby spines were significantly decreased in the Rpsa CRISPR (0.0333 ± 0.01, n = 21, P < 0.0005) compared with control CRISPR (0.1083 ± 0.03, n = 12), t(328) = 2.770. Thin spines were significantly increased in the Rpsa CRISPR group (0.2143 ± 0.03, n = 21) as compared with control CRISPR (0.1333 ± 0.03, n = 12) (t(328) = −1.826, P < 0.0005), while thin spines were significantly decreased in the PEDF shRNA group (0.0750 ± 0.02, n = 20, P = 0.002), as compared with scramble shRNA (0.1950 ± 0.03, n = 20) (F(2,567) = 6.507, P = 0.002). Filopodia spines were significantly increased in the Rpsa CRISPR (0.0524 ± 0.02, n = 21, P = 0.001) and PEDF shRNA + Rpsa OE (0.1000 ± 0.02, n = 21) groups, as compared with the control CRISPR (0.0167 ± 0.01, n = 12, P < 0.0005) and scramble shRNA + control OE (0.0150 ± 0.01, n = 20), respectively. (D) Primary cortical neurons transfected with the plasmid coding for YFP cultured until DIV14 and stained for Rpsa. Scale bar = 5 μm. (E) Transfected primary cortical neurons cultured until DIV14 and stained for PEDF. Scale bar = 5 μm. (F) Transfected primary cortical neurons cultured until DIV14 and stained for Itga6. Scale bar = 5 μm.

To determine whether Rpsa OE could rescue spine defects after PEDF KD, we used in utero electroporation at E15.5 to overexpress Rpsa in PEDF-deficient cells (Fig. 8A). The PEDF shRNA + Rpsa OE group (0.99 ± 0.07) showed an overall spine density that was comparable to the scramble shRNA + control OE group (0.77 ± 0.08) (Fig. 8B). The PEDF shRNA + Rpsa OE group showed a significant increase in branched (3.88%), mushroom (47.09%), and filopodia (10.19%) spines compared with the scramble shRNA + control OE group (branched: 0.65%, mushroom: 50.00%, filopodia: 1.95%) (Fig. 8C). This rescue shows an increase in branched and mushroom spines, which indicate a mature morphology, while also showing an increase in filopodia spines, which are considered immature. Thus, Rpsa OE can rescue spine density after PEDF KD, and it is suggested that Rpsa could accelerate the creation of new spines.

To investigate the expression of Rpsa, PEDF, and Itga6 at the spines, we transfected cortical neurons with a plasmid coding for YFP and cultured primary cortical neurons until DIV14 which is approximately when spine formation occurs and early spines are already present (Srivastava et al. 2011). We also stained for either Rpsa, PEDF, or Itga6 and confirmed that Rpsa, PEDF, and Itga6 are expressed in mature neurons at DIV14 (Fig. 8D–F). We observed that Rpsa and PEDF were stained in dendrites and spines. However, it appears that there is no or marginal expression of Itga6 in dendrites or spines (Fig. 8F).

Calcium Imaging of Brain Slices Shows Subthreshold Functional Difference in Rpsa KD Upper Layer Neurons

We next investigated whether the Rpsa KD associated decrease in dendrite number and length in upper layer cortical pyramidal neurons, as well as reduced dendritic arborization and change in spine density and morphology, could be linked to functional deficits, since neuronal morphology is a critical factor in determining the extent of inputs received by the neuron. To investigate function in Rpsa-deficient neurons, we conducted calcium imaging using live brain slices after in utero electroporation at E15.5 collected from male and female mice (approximately P30). The calcium indicator GCaMP6s was used to image neurons double positive for Rpsa or control CRISPR (tdTomato) and GCaMP6s (GFP). GCaMP6s has a relatively high sensitivity and is a well-validated calcium indicator (Chen et al. 2013). Rpsa-deficient neurons had a significantly lower mean difference between maximum and minimum peaks (6.7 ± 1.8), as compared with control neurons (12.8 ± 1.1) when spontaneous calcium activity was measured (Fig. 9A), indicating that the Rpsa-deficient neurons show significantly decreased calcium signaling. We found that Rpsa KD neurons showed almost no fluctuation in fluorescence intensity, while the control CRISPR neurons showed a dramatically greater change in fluorescence intensity (Fig. 9B). Since few action potentials were recorded, these differences in fluorescence intensity indicate that Rpsa KD neurons have a defect in calcium signaling at a subthreshold level. Calcium signaling directly relates to neuronal function by contributing to transmission of the depolarizing signal, synaptic signaling processes, neuronal energy metabolism, and neurotransmission (Brini et al. 2014).

Calcium imaging shows subthreshold functional difference in Rpsa KD neurons. (A) Two-tailed independent-samples t-test showed the mean difference between maximum and minimum peaks was significantly greater in the control (12.79 ± 1.06, n = 22 cells from 3 mice) compared with Rpsa KD (6.73 ± 1.80, n = 19 cells from 3 mice), t(39) = 2.992, P = 0.035. Data are represented as mean ± SEM. (B) Representative % change in fluorescence (ΔF/F₀) for representative individual neuron in mouse brain slice. Single action potentials are defined as ΔF/F₀ ≥ 23% ± 3.2%. The left panels were taken over 20 s and the right panels show a 5-s section of each representative trace, with the y-axis measuring 8 units to provide a zoomed in view. (C) Electrophysiology recordings confirm low spontaneous firing rate of control CRISPR and Rpsa CRISPR neurons. Representative traces of spontaneous activity of 3 control CRISPR and Rpsa CRISPR neurons at resting membrane potential in current-clamp mode. n = 9 cells from 3 control CRISPR mice and n = 7 cells from 2 Rpsa CRISPR mice.

To complement our calcium imaging data, we conducted whole-cell electrophysiological recordings in mice (approximately P30) after knocking down Rpsa by CRISPR by in utero electroporation at E15.5. In agreement with our calcium imaging results, electrophysiological recordings showed that control and Rpsa KD neurons do not fire action potentials spontaneously at rest (Fig. 9C). However, both control and Rpsa-deficient neurons fired action potentials upon stimulation (Supplemental Fig. 5). Intrinsic membrane properties were comparable between groups and only differed significantly in capacitance, indicating a difference in cell size (Supplemental Fig. 5 and Table 1). Taken together, this corroborates our calcium imaging results by confirming the low spontaneous firing rate in both control and Rpsa KD cells and showing that Rpsa KD neurons are healthy and able to fire action potentials.

Table 1.

Comparison of membrane properties between control CRISPR and Rpsa CRISPR

	Vm (mV)	Resistance (MΩ)	τ (ms)	C (pF)	Rheobase (pA)	Spike height (mV)	Overshoot (mV)	Spike threshold (mV)
Control CRISPR (n = 9)	−67 ± 1.2	60 ± 4.9	14 ± 1.2	251 ± 30.2	141 ± 10.5	84 ± 1.8	50 ± 1.2	−34 ± 1.1
Rpsa CRISPR (n = 7)	−64 ± 0.9	50 ± 5.1	18 ± 2.6	369 ± 35.8	149 ± 33.3	81 ± 1.7	48 ± 2.4	−32 ± 1.2

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Data are represented as mean ± SEM. Vm, membrane potential; τ, membrane time constant; C, capacitance.

Discussion

Our study reports a role for the key signaling molecule Rpsa in regulating multiple functionally relevant stages of neuronal morphogenesis in upper and lower cortical layers. Additionally, Rpsa KD disrupts RGC fiber morphology but does not impact neuronal differentiation via asymmetrical division of RGCs. We found that PEDF, Rpsa’s ligand, and Itga6, Rpsa’s plasma membrane binding partner, are essential components of the signaling pathway that regulates neuromorphogenesis. Additionally, we found Itga6 increases and stabilizes Rpsa expression on the plasma membrane by preventing the ubiquitination of Rpsa. We have begun to clarify the PEDF-Rpsa-Itga6 signaling mechanism, which is involved in the coordination of multiple stages of cortical neuromorphogenesis resulting in a functional neuron.

Rpsa is present at early embryonic stages in the somas of RGCs (Fig. 1). This expression pattern suggested a role for Rpsa in regulating RGC division. However, we show that Rpsa KD does not impact RGC division, but rather RGC fiber morphology (Fig. 5). These data indicate that Rpsa could be regulating RGC fiber morphology indirectly, possibly by regulating downstream cytoskeletal elements or expression of other proteins that are crucial for fiber morphology, as Rpsa has been shown to be required for pre-rRNA processing (Griffin et al. 2018).

Orientation of the apical dendrite may lead to improper positioning and formation of inappropriate synaptic connections. Rpsa-deficient neurons show aberrant orientation of the apical dendrite at the proximal dendrite region at P3 (Fig. 2). However, the distal region of apical dendrites turns into the proper direction to the marginal zone (Fig. 2A). This implies that Rpsa affects the initial step of dendrite extension. Future studies including time-lapse live imaging will help elucidate the precise function of Rpsa in dendrite behavior during the initial stages of dendrite extension and orientation.

Later stages of neuromorphogenesis were dramatically impacted by Rpsa KD, as indicated by the deficits observed in upper cortical layers at P15. The decrease in number of dendrites and dendrite length suggests a deficit in the initiation and elongation of dendrites, respectively. The decrease in dendrite length seen at P15 (Fig. 3) was not observed in apical dendrite length at P3 (Supplemental Fig. 2), indicating the importance of Rpsa in basal dendrite formation.

To better understand the role of Rpsa in overall cortical development, we analyzed pyramidal neurons in upper and lower layer neurons and determined that Rpsa differentially regulates the morphogenesis of neurons in different cortical layers. Neurons in lower layers connect to other neurons within the cortex to integrate information, while neurons in upper layers project more widely to subcortical targets that are involved in behavior (Gerfen et al. 2018). The differential regulation of neuromorphogenesis via Rpsa between upper and lower layer neurons may serve to create some of these functional differences. Additionally, Rpsa KD changes RGC fiber morphology, which may be related to the neuronal layering phenotypes observed in Rpsa KD neurons (Supplemental Fig. 3). It appears that the effect of Rpsa KD in neuronal differentiation is moderate, as Rpsa deficiency did not result in changes to the number of neurons/IPCs generated via asymmetrical RGC division.

We next wanted to investigate the mechanism of Rpsa signaling. Before advancing to rescue experiments that would investigate the role of PEDF and Itga6 in Rpsa signaling, we first confirmed that KD of PEDF and Itga6 alone would produce morphological defects similar to those seen in Rpsa-deficient neurons. We also characterized the layering of neurons following Rpsa, PEDF, and Itga6 KD and found all neurons reached the CP (Supplemental Fig. 3). While Rpsa- and PEDF-deficient neurons did appear to be distributed more broadly within the cellular layer, the phenotype observed is not consistent with a severe defect in neuronal migration. Interestingly, we also found that Rpsa KD in lower layer neurons resulted in sparse distribution. This suggests that PEDF-Rpsa signaling is also involved in neuronal migration, which will be further analyzed in the future.

Our in vitro rPEDF rescue experiment (Fig. 6E–I) and genetic manipulation of the Rpsa signaling pathway in vivo using various rescue experiments allowed us to provide convincing evidence that PEDF is the ligand responsible for initiating Rpsa signaling to regulate neuronal morphogenesis and that a Rpsa-Itga6 signaling complex is important for dendrite formation (Fig. 10A). Our rPEDF rescue experiment revealed that rPEDF treatment of Rpsa-deficient neurons could not rescue the defects in neurite length both in the axon and dendrites. This strongly suggests that PEDF regulates neurite extension via Rpsa. In contrast, rPEDF could increase the number of neurites to a normal level and rescued arborization. Since we found that Rpsa KD in vivo resulted in defects in neurite initiation and arborization, Rpsa is involved in neurite initiation and arborization, but PEDF can also regulates these through a non-Rpsa-mediated mechanism, such as through another one of its receptors. Although we observed differences between Rpsa OE neurons and controls in vivo, this does not discredit our rationale regarding our rescue experiments. Endogenous PEDF would be present at typical levels in this in vivo experiment, thus our Rpsa OE phenotype may be reflecting increased activation of the Rpsa pathway via PEDF in this situation.

Schematic of Rpsa/PEDF/Itga6 signaling regulating neuronal morphogenesis. (A) Under normal conditions Itga6, which is likely bound to a β4 subunit to function as part of a complete integrin, is co-localized on the plasma membrane with Rpsa. Both Rpsa and Itga6 facilitate the binding of PEDF to Rpsa, which results in signaling to regulate proper neuronal morphogenesis (top left). Following Rpsa KD, PEDF KD, or Itga6 KD, the signaling mechanism initiated by Rpsa is attenuated, resulting in severe defects in morphology (top right). KD of the upstream ligand PEDF can be compensated for by increasing its downstream receptor Rpsa, to cause increase of Rpsa-Itga6-Itgb4 complex, thus resulting in normal level of signal intensity and normal morphogenesis. The increase in Rpsa-Itga6-Itgb4 complex can also be accomplished by Itga6 OE (bottom). (B) Itga6 increases both the expression level and time spent by Rpsa in the plasma membrane by preventing ubiquitination of Rpsa. Created with BioRender.com.

PEDF is secreted by rat cerebral cortical neurons, suggesting that mouse cortical neurons secrete PEDF since cell staining indicates PEDF expression in mouse cortical neurons (Supplemental Fig. 6A; Sanchez et al. 2012). The dramatic morphological defects seen after PEDF KD via in utero electroporation suggest that autocrine PEDF-Rpsa signaling is a major pathway in developing cortical neurons. Because PEDF is a secreted protein, neighboring cells may be secreting PEDF that binds Rpsa receptors present on neurons transfected with PEDF shRNA to initiate a paracrine signaling pathway. However, paracrine PEDF-Rpsa signaling in developing cortical neurons appears to be less efficient than the autocrine pathway, since severe defects are seen after PEDF KD and paracrine signaling cannot compensate for autocrine signaling. In our rescue experiment where Rpsa is overexpressed in PEDF-deficient cells, there is more Rpsa present on the membrane, which increases paracrine signaling efficiency by accelerating the opportunity for PEDF secreted by other cells to bind to an Rpsa receptor and compensate for the PEDF KD in the transfected cell. PEDF may also have autocrine and paracrine effects on motor neurons (Bilak et al. 1999).

Our rescue experiments showed that both Itga6 and Rpsa OE can correct defects in dendrite formation in PEDF-deficient neurons, with the Sholl profiles for these groups indicating an increase in complexity of dendritic branching as compared with the PEDF shRNA group (Fig. 6D). However, the PEDF shRNA + Rpsa OE group showed even more complex branching than its control. Rpsa OE could have resulted in a more complex branching phenotype, since Rpsa is further downstream than PEDF in the proposed signaling pathway. Thus, OE at a more downstream level in the signaling pathway could have overstimulated the mechanism responsible for regulating branching, leading to an increase in branching complexity.

The similarities between the Rpsa OE and Itga6 OE rescue phenotypes may be explained by our data, indicating that an increase in Itga6 expression increases Rpsa localization and stabilization on the plasma membrane, where Rpsa must be located to interact with its extracellular ligand PEDF (Fig. 10A). Itga6 prevents ubiquitination of Rpsa (Fig. 10B). Rpsa has a total of 6 ubiquitination sites: Lys11, Lys40, Lys52, Lys57, Lys89, and Lys166. While Lys89 may be located in a computer-predicted transmembrane domain at residues 86-101 and Lys166 is likely to be included in the extracellular laminin binding domain, Lys11, Lys40, Lys52, and Lys57 are located inside the cell and are likely targets for ubiquitination (Castronovo et al. 1991). Itga6 could potentially bind Rpsa such that Itga6 blocks the ubiquitination site on Rpsa that regulates its degradation. Itga6 is expressed in some of the same regions of the cortex as Rpsa, including the CP (Supplemental Fig. 6B). Previous studies indicate that ubiquitin E3 ligase Nedd4 is primarily responsible for labeling Rpsa for ubiquitin-mediated internalization (Kim et al. 2012). Nedd4 is strongly and ubiquitously expressed in the newborn brain and its expression decreases as development progresses, suggesting that Nedd4 is particularly important in the developing neocortex (Kawabe et al. 2010). Thus, Nedd4 is likely involved in PEDF-Rpsa-Itga6 signaling, and the mechanisms of Rpsa ubiquitination by Nedd4 should be analyzed in the future.

Interestingly, Itga6-deficient neurons did not display any defects in dendritic spine density or morphology, suggesting that the role of Itga6 in regulating neuromorphogenesis via Rpsa signaling may be limited to dendrite formation. The absence of a dendritic spine phenotype following Itga6 KD may be explained by the lack of expression of Itga6 in dendrites and spines, while Rpsa and PEDF were found to be present at dendrites and spines (Fig. 8D–F). These changes in spine phenotypes and Rpsa’s presence at the synapse suggested that Rpsa KD may impact neuronal function.

We conducted calcium imaging on live brain slices and detected a difference between Rpsa KD cells and control cells in subthreshold calcium signaling. The GCaMP6s calcium indicator was used, which has a relatively high sensitivity and has commonly been used to image relatively low cytoplasmic calcium levels (Chen et al. 2013). Previous studies have focused on calcium signaling in active dendritic spines. However, the amount of calcium that enters via these synaptic mechanisms may be very small, requiring that the signal be greatly amplified by intracellular calcium release to be detected (Emptage et al. 1999). Thus, imaging of the soma, as done in our study, allows for changes in subthreshold calcium signaling to be detected by measuring calcium release from internal stores in the soma that amplify signals. Subthreshold changes in calcium signaling could be the result of the decrease in dendritic surface area seen in Rpsa-deficient neurons. This morphological change may lead to fewer numbers of NMDA channels, NMDA and AMPA receptors, and/or voltage-gated calcium channels being present, which are crucial for regulating calcium levels in neurons (Magee et al. 1995; Koester and Sakmann 1998; Schiller et al. 1998; Yuste et al. 1999, 2000; Kovalchuk et al. 2000). Furthermore, the defects in spine density and morphology in Rpsa-deficient neurons likely play a significant role as dendritic spines compartmentalize calcium and spine morphology directly relates to calcium kinetics, since the thickness of the spine neck impacts the speed of calcium diffusion into dendrites (Yuste et al. 2000).

Single action potentials are defined as ΔF/F₀ ≥ 23% ± 3.2%. Thus, few action potentials were recorded by our calcium imaging. However, this is not surprising, since pyramidal cells can have a low spontaneous firing rate (Petersen and Crochet 2013). Our electrophysiology recordings confirm the minimal percent change in fluorescence detected by our calcium imaging by showing that inputs do not summate to a level that would cause action potentials for either the Rpsa KD or control group. This indicates that the specific cortical pyramidal neurons that were transfected via in utero electroporation have a low spontaneous firing rate (Fig. 9C). Thus, we should not expect to see many action potentials recorded in our calcium imaging experiments. Taken together, these data suggest that Rpsa KD causes a functional difference in calcium signaling at subthreshold levels in cortical pyramidal neurons with a low spontaneous firing rate. Additionally, our electrophysiology recordings indicate that control CRISPR neurons and Rpsa CRISPR neurons display similar intrinsic membrane properties (Supplemental Fig. 5 and Table 1). This is important to note because the 37-kDa Rpsa precursor is a component of the 40S ribosome and is involved in pre-RNA processing, thus it could be suggested that the morphological defects in Rpsa-deficient cells may be attributed to a poor overall state of the neurons (Tohgo et al. 1994; O'Donohue et al. 2010). However, our electrophysiology results suggest that Rpsa-deficient cells remain healthy, despite the phenotypic changes associated with Rpsa KD (Supplemental Fig. 5 and Table 1).

These results are of clinical interest because the gene coding for the PEDF protein, Serpinf1, is in the chromosome 17p13.3 region, which is often deleted causing MDS or duplicated causing 17p13.3 microduplication syndrome. There is no doubt that neuronal migration defects are a main cause of MDS, but the pathogenesis of MDS has not been completely clarified in detail. MDS has a complex etiology because it caused by a microdeletion that could include more than 26 genes, including Serpinf1. While most of the genes deleted in MDS patients have not been investigated, previous studies have analyzed some genes, including Pafah1b1 (Lis1), Ywhae (14-3-3ε), and Crk in the MDS critical region (Izumi et al. 2007; Sreenath Nagamani et al. 2009; Hyon et al. 2011; Cornell et al. 2016; Henry et al. 2016; Blazejewski et al. 2018). However, these studies focused on the gene functions in neuronal migration and neural activity, since most MDS patients suffer from epilepsy. The role of neuronal morphogenesis in MDS etiology remains uncertain. PEDF deficiency may not be the main cause of MDS, but it could contribute to MDS pathogenesis. Thus, our findings will provide new insights into the mechanisms underlying MDS.

In conclusion, Rpsa signaling mediates functionally relevant aspects of cortical neuronal morphogenesis including apical dendrite orientation, initiation and elongation of dendrites, dendritic branching, and dendritic spine density and morphology. This Rpsa signaling mechanism is initiated by binding of its ligand PEDF, while Itga6 promotes and stabilizes the expression of Rpsa on the membrane. Future studies should focus on investigating the mechanism by which Rpsa KD impacts the synapse and elucidating the downstream targets of Rpsa during cortical neuronal morphogenesis. The PI3K and MAPK signaling pathways have been shown to be downstream of Rpsa and Itga6 in promoting pancreatic cancer invasion and metastasis (Wu et al. 2019). This indicates that PI3K and MAPK are potentially downstream of PEDF-Rpsa-Itga6 signaling during cortical development. Furthermore, Nedd4, which is known to ubiquitinate Rpsa, has been implicated in branching and regulation of neurite growth by acting as a downstream target of PI3K/PTEN-mTORC1 (DiAntonio 2010; Hsia et al. 2014; Kim et al. 2012). Future studies should determine if these signaling mechanisms are downstream effectors of PEDF-Rpsa-Itga6-mediated regulation of cortical neuromorphogenesis. Further investigation of this pathway will advance our understanding of neuronal morphogenesis in normal brain development as well as increase our knowledge of neurodevelopmental disorders, such as MDS.

Supplementary Material

Supplemental_Figures_and_Legends_Cerebral_Cortex_Revision_Final_bhab242

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Contributor Information

Sara M Blazejewski, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Sarah A Bennison, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Ngoc T Ha, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Xiaonan Liu, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Trevor H Smith, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Kimberly J Dougherty, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Kazuhito Toyo-Oka, Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA 19129, USA.

Funding

National Institute of Neurological Disorders and Stroke (R01NS096098 to K.T., R01NS095366 to K.J.D., and F31NS113404 to S.M.B.). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Notes

We thank Dr Bryan W. Luikart for his technical advice and support. We are also grateful to Drs Peter Baas, Elias Spiliotis, Eric Olson, and Itzhak Fischer for their reading the manuscript and comments. Conflict of Interest: The authors have declared that no competing interests exist.

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PERMALINK

Rpsa Signaling Regulates Cortical Neuronal Morphogenesis via Its Ligand, PEDF, and Plasma Membrane Interaction Partner, Itga6

Sara M Blazejewski

Sarah A Bennison

Ngoc T Ha

Xiaonan Liu

Trevor H Smith

Kimberly J Dougherty

Kazuhito Toyo-Oka

Abstract

Introduction

Materials and Methods

Mice

Plasmids

Analysis of Genomic Alterations by CRISPR/Cas9

Antibodies

Histology and Immunohistochemistry

Preparation and Culture of Primary Cortical Neurons

In Utero Electroporation

Analysis of Neuronal Morphology

Apical Dendrite Orientation at P3

Neuronal Morphology Analysis at P15

Dendritic Spine Morphology Analysis at P15

Subcellular Fractionation

Figure 7 .

Pull-down Assay

Cycloheximide Treatment

Calcium Imaging of Live Brain Slices

Slice Preparation

Imaging of Spontaneous Activity

Calcium Imaging Data Analysis

Electrophysiological Recordings

Slice Preparation

Recordings

Quantification of Corrected Total Cell Fluorescence

Statistical Analysis

Results

Expression of Rpsa in the Developing Cerebral Cortex

Figure 1 .

Rpsa KD in Upper Layer Neurons Is Associated with Apical Dendrite Misorientation at P3

Figure 2 .

Rpsa KD in Upper Layer Neurons Is Associated with Defects in Neuronal Morphogenesis at P15

Figure 3 .

Rpsa KD in Lower Layer Neurons Is Associated with Decreased Dendritic Branching and Disrupted Cell Layering at P15

Figure 4 .

Rpsa KD Disrupts RGC Process Morphology, but Results in No Significant Change in RGC Spindle Orientation at Multiple Time-points

Figure 5 .

PEDF KD and Itga6 KD result in neuromorphological defects in upper layer neurons similar to Rpsa KD

Figure 6 .

Figure 6 .

Itga6 OE Increases and Stabilizes Rpsa Expression on the Plasma Membrane by Preventing Ubiquitination of Rpsa

Rpsa KD and PEDF KD, but Not Itga6 KD, Cause a Decrease in Overall Spine Density and a Change in Spine Morphology in Upper Layer Neurons

Figure 8 .

Calcium Imaging of Brain Slices Shows Subthreshold Functional Difference in Rpsa KD Upper Layer Neurons

Figure 9 .

Table 1.

Discussion

Figure 10 .

Supplementary Material

Contributor Information

Funding

Notes

References

Associated Data

Supplementary Materials

ACTIONS

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