Fhod3 Controls the Dendritic Spine Morphology of Specific Subpopulations of Pyramidal Neurons in the Mouse Cerebral Cortex

Abstract

Changes in the shape and size of the dendritic spines are critical for synaptic transmission. These morphological changes depend on dynamic assembly of the actin cytoskeleton and occur differently in various types of neurons. However, how the actin dynamics are regulated in a neuronal cell type-specific manner remains largely unknown. We show that Fhod3, a member of the formin family proteins that mediate F-actin assembly, controls the dendritic spine morphogenesis of specific subpopulations of cerebrocortical pyramidal neurons. Fhod3 is expressed specifically in excitatory pyramidal neurons within layers II/III and V of restricted areas of the mouse cerebral cortex. Immunohistochemical and biochemical analyses revealed the accumulation of Fhod3 in postsynaptic spines. Although targeted deletion of Fhod3 in the brain did not lead to any defects in the gross or histological appearance of the brain, the dendritic spines in pyramidal neurons within presumptive Fhod3-positive areas were morphologically abnormal. In primary cultures prepared from the Fhod3-depleted cortex, defects in spine morphology were only detected in Fhod3 promoter-active cells, a small population of pyramidal neurons, and not in Fhod3 promoter-negative pyramidal neurons. Thus, Fhod3 plays a crucial role in dendritic spine morphogenesis only in a specific population of pyramidal neurons in a cell type-specific manner.

actins, cytoskeleton, dendrites, formins, neurites, pyramidal cells, synapses, synaptogenesis

Introduction

The dendrites of pyramidal neurons are covered with numerous small pleomorphic protrusions called dendritic spines, which constitute the postsynaptic site for most excitatory synapses (DeFelipe and Fariñas 1992). During neuronal development and synaptic plasticity, dendritic spines undergo dynamic changes in shape and size (Huttenlocher and Dabholkar 1997; Zuo et al. 2005; Kasai et al. 2010; Yasuda 2017). Since their structural changes are tightly coupled to alterations in synaptic transmission, these modifications in the spine morphology are considered an important component of the molecular basis of learning and memory.

The morphological changes of dendritic spines are controlled spatially and temporally by rearrangement of the actin cytoskeleton. The assembly of F-actin in dendritic spines is regulated by multiple mechanisms, including those mediated via Rho-family small GTPases and their effectors (Saneyoshi and Hayashi 2012; Ba et al. 2013; Nishiyama and Yasuda 2015; Spence and Soderling 2015). The Arp2/3 complex, an actin nucleator activated downstream of Rac1 or Cdc42 signaling to create branched F-actin, has been shown to play important roles in the formation of F-actin in dendritic spines (Rotty et al. 2013; Spence et al. 2016). The activation of the Arp2/3 complex requires nucleation-promoting factors (NPFs), such as N-WASP and WAVE1 (Rotty et al. 2013). Ablation of NPFs leads to aberrations in spine morphology as well as behavioral abnormalities (Kim et al. 2006; Soderling et al. 2007), indicating that the Arp2/3 complex is essential for the maintenance and function of dendritic spines.

Formins, another group consisting of 15 mammalian members of actin nucleators that promote the formation of linear unbranched F-actin downstream of RhoA signaling (Goode and Eck 2007; Kühn and Geyer 2014), also seem to be involved in the morphogenesis of dendritic spines (Spence and Soderling 2015). In cultured hippocampal neurons, the formin mDia2 is required for dendritic spine development, especially to the initial step of elongation of the dendritic filopodia (Hotulainen et al. 2009). The analysis of cultured hippocampal neurons by super-resolution microscopy has shown that another formin, Fmnl2, is involved in spine head enlargement (Chazeau et al. 2014). The significance of formins in spinal morphogenesis is also evident from genetic studies, which have demonstrated that the loss of murine Fmn2 and deletion mutations of human Fmn2 are associated with memory loss and intellectual disability, respectively (Peleg et al. 2010; Almuqbil et al. 2013; Law et al. 2014; Agís-Balboa et al. 2017). Thus, several formins seem to participate in the morphogenesis of dendritic spines, thereby likely contributing to the plasticity of synaptic transmission. However, how the 15 formin family protein members share these tasks has remained elusive.

We have shown that Fhod3, a formin family protein that is expressed abundantly in the heart, plays an essential role in cardiogenesis and cardiac function (Kanaya et al. 2005; Taniguchi et al. 2009; Kan-o et al. 2012; Ushijima et al. 2018). In addition, Fhod3 is only transiently expressed in the neuroepithelium in the hindbrain during primary neurulation and plays a crucial role in neural plate morphogenesis (Sulistomo et al. 2019). In the present study, we examined the role of Fhod3 in the brain after birth. The expression of Fhod3 in the cerebral cortex became evident after birth. We demonstrated, by biochemical and immunofluorescent analyses, that Fhod3 accumulates in the dendritic spines of excitatory cortical neurons. The depletion of Fhod3 was found to lead to morphologic abnormalities in the dendritic spines both in vivo and in vitro. The defect in the spine morphology was only observed in Fhod3 promoter-active cells, a small population of pyramidal neurons, and not in Fhod3 promoter-negative pyramidal neurons. The present findings provide direct evidence that Fhod3 participates in the spine morphogenesis of cortical neurons in a cell type-specific manner.

Materials and Methods

Mice

Mice heterozygous for the constitutive null Fhod3 allele (Fhod3^+/−) were generated by replacement of exon 1 with lacZ as previously described (Kan-o et al. 2012). For deletion of Fhod3 in the central nervous system (CNS), homozygous Fhod3^flox/flox mice (Ushijima et al. 2018) were crossed with Fhod3^+/−; Nestin-Cre⁺ mice that were generated by crossing Fhod3^+/− mice (Kan-o et al. 2012) with Nestin-Cre transgenic mice (Matsumoto et al. 2011). Deletion of exon 18 of the floxed Fhod3 allele was confirmed by PCR analysis of genomic DNA as described previously (Ushijima et al. 2018).

All the experimental protocol was approved by the Animal Care and Use Committee of Miyazaki University (Permit Number: 2014-526-6). All mice were housed and maintained in a specific pathogen-free animal facility at University of Miyazaki, and all efforts were made to minimize the number of animals used and their suffering. All experiments were performed in strict accordance with the guidelines for Proper Conduct of Animal Experiments (Science Council of Japan) and the “Guide for the Care and Use of Laboratory Animals” published by the US National Institute of Health.

Construction of Recombinant Plasmids

The cDNA fragment encoding mouse Fhod3 short variant of 1427 amino acids (previously designated as mFhos2S) was prepared and ligated to pEGFP-C1 (Clontech) or pEF-BOS as described previously (Kanaya et al. 2005; Taniguchi et al. 2009). The cDNA encoding mouse Fhod3 short variant carrying the I976A substitution (equivalent to I1127A in the cardiac Fhod3 long variant) was constructed by PCR using the cDNA encoding mouse Fhod3. The cDNA fragment encoding Lifeact, an actin-binding peptide comprising 17 amino acids (Riedl et al. 2008), was prepared by annealing of oligonucleotides and ligated to pmCherry-N1 (Clontech). The cDNA fragments encoding rat CaMKIIα carrying the T286D and K42R/T286D substitutions were constructed by PCR using the cDNA encoding rat CaMKIIα and ligated to the monomeric enhanced green fluorescent protein (pmEGFP) vector (Fujii et al. 2013). All constructs were sequenced for confirmation of their identities.

Primary Culture of Cortical Neurons and Transfection

Primary cultures of embryonic mice cortical neurons were prepared as described previously (Nemoto et al. 2014) with minor modifications. Briefly, primary cortical neurons from embryonic brains at embryonic day 17.5 were plated on coverslips coated with poly l-ornithine (Wako) in neurobasal plus medium (GIBCO) supplemented with 2% B27-plus (GIBCO) containing 2% Glutamax (GIBCO), and half of medium was replaced every 3–4 days. The neuronal cells were grown under 5% CO₂ for 21 days in an incubator maintained at 37 °C. All cell culture experiments included a B27-plus supplement, which is a standard component for neuronal cell culture. Transfection of cortical neurons with plasmid DNAs was performed using X-tremeGENE 9 (Roche) or Lipofectamine 2000 (Invitrogen), according to the manufacturer’s protocol.

Antibodies

Rabbit polyclonal antibodies for Fhod3 (anti-Fhod3-(650–802) and anti-Fhod3 C20) were prepared as described previously (Kanaya et al. 2005). The following antibodies were purchased: mouse anti-Brn2 clone B-2 (Santa Cruz); mouse anti-NeuN clone A60 (Chemicon); mouse anti-MAP2 clone HM-2 (Sigma Aldrich); mouse anti-PSD95 clone 6G6-1C9 (Novus); mouse anti-glutamate decarboxylase (GAD67) clone 1G10.2 (Sigma Aldrich); mouse anti-β-actin clone AC-15 (Sigma Aldrich); mouse anti-α-tubulin clone 10G10 (Wako); mouse anti-GAPDH clone 5A12 (Wako); mouse anti-GM130 clone 35/GM130 (BD Biosciences); mouse anti-β-galactosidase clone (Promega); rabbit anti-β-galactosidase (Invitrogen); rabbit anti-glutaminase (KGA/GAC; Proteintech); guinea pig anti-vesicular glutamate transporter 1 (vGlut1; Chemicon); Alexa Fluor 488, 555, or 657-conjugated F(ab’)₂ fragment of anti-rabbit IgG (Cell Signaling Technology); Alexa Fluor 488 or 555-conjugated F(ab’)₂ fragment of anti-mouse IgG (Cell Signaling Technology); and Alexa Fluor 555-conjugated anti-guinea pig IgG (Invitrogen). Nuclei were stained with Hoechst 33342 (Dojindo) or DAPI (Dojindo).

Immunoblot Analysis

Immunoblot analysis was performed as previously described (Ushijima et al. 2018). Briefly, organs of mice were homogenized and sonicated at 4 °C in a lysis buffer composed of 10% glycerol, 135 mM NaCl, 5 mM EDTA, and 20 mM Hepes, pH 7.4 containing Protease Inhibitor Cocktail (Sigma-Aldrich). The lysates were applied to SDS-PAGE and were transferred to a polyvinylidene difluoride membrane (Millipore). Protein transfer was immediately confirmed by Fastgreen staining (loading). The membrane was probed with the antibody, followed by development using ECL-plus (GE Healthcare) for visualization of the antibodies.

Immunofluorescence Staining

Immunofluorescence staining was performed as previously described (Sulistomo et al. 2019) with minor modification. The brain of mice were fixed overnight at 4 °C in 3.7% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) followed by cryoprotection at 4 °C in 30% sucrose and embedded in Colored Tissue Freezing Medium (Triangle Biomedical Sciences). The blocks were frozen and serial sections were obtained using a cryostat CM3050S (Leica Biosystems). Sections were washed with PBS containing 0.1% Triton X-100 and blocked with PBS containing 3% bovine serum albumin for 60 min at room temperature. Sections were labeled overnight at 4 °C with primary antibodies diluted in PBS containing 1% bovine serum albumin, washed with PBS, and then labeled for 3 h at 4 °C with fluorescently labeled secondary antibody mixture in the same buffer. Images were taken with Zeiss LSM700 confocal microscope or Leica TCS SP8 confocal microscope under the same conditions between Fhod3-null and wild-type mice.

Cultured neurons were fixed in 3.7% formaldehyde for 15 min, permeabilized with 0.1% Triton X-100 in PBS for 4 min, and blocked with PBS containing 3% bovine serum albumin for 60 min. Cells were labeled overnight at 4 °C with primary antibodies diluted in PBS containing 1% bovine serum albumin, washed with PBS, and then labeled for 3 h at 4 °C with fluorescently labeled secondary antibody mixture in the same buffer. Images were taken with Zeiss LSM700 confocal microscope or Leica TCS SP8 confocal microscope under the same conditions between Fhod3-null and wild-type mice.

X-Gal Staining

X-gal staining was performed as previously described (Sulistomo et al. 2019). The adult brain in Fhod3^+/+ or Fhod3^+/− mouse was sliced in 700 μm sagittal or 900 μm coronal sections by using microslicer. The sliced sections were fixed at 4 °C by immersion in PBS (137 mM NaCl, 2.68 mM KCl, 8.1 mM Na₂HPO₄, and 1.47 mM KH₂PO₄, pH 7.4) containing 1% formaldehyde, 0.2% glutaraldehyde, 0.02% Nonidet P-40, and 1 mM MgCl₂. Fixed brain sections were incubated at 37 °C in PBS containing 1 mg/ml X-gal, 5 mM K₃Fe(CN)₆, 5 mM K₄Fe(CN)₆, and 2 mM MgCl₂. In the case of immunostaining and X-gal staining using consecutive sagittal sections, brains were fixed overnight at 4 °C in 3.7% PFA in PBS and then immersed at 4 °C in 30% sucrose. Cryosections were performed at 5 μm thickness.

In the case of brains of embryos, brains were fixed by immersion for 3 h at 4 °C in PBS (137 mM NaCl, 2.68 mM KCl, 8.1 mM Na₂HPO₄, and 1.47 mM KH₂PO₄, pH 7.4) containing 1% formaldehyde, 0.2% glutaraldehyde, 0.02% Nonidet P-40, and 1 mM MgCl₂ and then immersed at 4 °C in 30% sucrose. Fixed brains were sliced by using Cryostat at 50 μm thickness. The all sliced sections were incubated at 37 °C in PBS containing 1 mg/ml X-gal, 5 mM K₃Fe(CN)₆, 5 mM K₄Fe(CN)₆, and 2 mM MgCl₂ followed by counter staining with 1% Orange G (WAKO) in 2% tungstophosphoric acid.

Histological Analysis

Histological analysis was performed as previously described (Ushijima et al. 2018). Briefly, mice were euthanized by cervical dislocation, and the brains were removed and fixed by immersion in PBS containing 3.7% formaldehyde. The fixed organs were dehydrated in graded ethanol solutions, embedded in paraffin, sectioned, and stained with hematoxylin and eosin. Images were acquired using BZ-9000 microscope (Keyence).

Golgi Staining and Quantification of Dendritic Protrusions

Brains from Fhod3 conditional knockout (cKO) mice (Fhod3^flox/−; Nestin-Cre⁺) and control mice (Fhod3^flox/+ and Fhod3^flox/+; Nestin-Cre⁺) at P35 were processed with the FD Rapid GolgiStain Kit (FD Neurotechnologies, Inc.), which is based on the staining principles previously described by Ramón-Moliner (1970) and Glaser and Van der Loos (1981). In brief, after the establishment of anesthesia, brains were immediately removed, rinsed in double distilled water, and then transferred to a mixture of equal volumes of solutions A and B provided in the kit. This mixture of solutions was replaced once after 24 h of immersion and stored at room temperature in darkness for 2–3 weeks. The embedded brains were then transferred into solution C provided in the kit and stored at room temperature for at least 1 week in the dark. Tissues were next rapidly frozen with dry ice and quickly embedded in Tissue Freezing Medium (Triangle Biomedical Sciences). Brains were cut into 150-μm-thick sections in the sagittal plane with a freezing cryostat. Sections were transferred onto gelatin-coated slides (FD Neurotechnologies, Inc.). Following air drying in the dark, sections were rinsed with distilled water, reacted in a developing mixed solution D and B provided in the kit, and dehydrated with 50%, 75%, 95%, and 100% ethanol, respectively. After defatting with xylene, the slides were mounted and coverslipped with Permount (FALMA).

Pyramidal neurons displaying a triangular soma with an apical dendrite were selected from the layer II/III of presumptive Fhod3-positive areas for tracing. Neurons were viewed and traced using a Zeiss LSM700 confocal microscope with a ×63 (oil immersion, NA 1.4) objective. Secondary or tertiary dendrites of pyramidal neurons and Purkinje cells were traced. All reconstructions and dendritic protrusion measurements were performed with the researcher blinded to genotype. The density and location of dendritic protrusions relative to the soma were measured using ImageJ (National Institutes of Health) with Fiji plugins. The statistics were conducted with a Student’s t-test and a factorial analysis of variance.

Quantification of Synaptic Puncta in the Cortex Sections

Quantification of synaptic puncta was performed by the method of Marzo et al. (2016) with modification. Briefly, the images of brain slices were taken with a ×63 objective lens of Leica TCS Sp8 confocal microscope. The threshold of image intensity was optimized, applied to all images, and punctate were counted using ImageJ/Fiji (National Institute of Health).

Preparation of Synaptosomal and PSD-Enriched Fractions

Mouse brain synaptosomal fractions were prepared by the method of Kamat et al. (2014). Briefly, the mouse cerebral cortex was dissected and suspended in 10 volumes of 0.32 M sucrose HEPES buffer (145 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 5 mM glucose, and 25 mM HEPES, pH 7.4) supplemented with Protease Inhibitor Cocktail. The tissue was homogenized using a Teflon glass homogenizer, and the homogenate was centrifuged at 600g for 10 min at 4 °C to separate the supernatant (S1) from the nuclei and large debris fraction. The S1 fraction was then diluted 1: 1 with 1.3 M sucrose in HEPES buffer, to yield a suspension at a final concentration of 0.8 M sucrose in HEPES buffer. This suspension was further centrifuged at 12 000g for 15 min to separate the supernatant (S2; cytosolic fraction) and the pellet (P2; crude synaptosomal fraction). The P2 fraction was washed twice with HEPES buffer to obtain the synaptosomal fraction. The washed synaptosomal fraction was solubilized with HEPES buffer containing 0.5% Triton X-100 at 4 °C for 20 min and then centrifuged at 20000g for 30 min to separate the triton-soluble and insoluble PSD-enriched fractions.

Transmission Electron Microscopic Analysis

Transmission electron microscopy of thin sections was performed as described previously (Sulistomo et al. 2019). Briefly, mice were deeply anesthetized with an intraperitoneal injection of pentobarbital and sevoflurane inhalation and systemically perfused with the fixative solution (2.0% PFA, 2.5% glutaraldehyde, and 0.1 M sodium cacodylate, pH 7.4). The dissected brains were immersed in the same fixative solution and then rinsed in PBS, postfixed in 1% osmium tetroxide, dehydrated in ethanol and propylene oxide, and embedded in epoxy resin. Thin sections stained with uranyl acetate and lead citrate were then examined with a HT7700 (Hitachi) transmission electron microscope. Mean synaptic densities were calculated by counting asymmetric synapses, which had clear synaptic vesicles with PSD.

Quantification of Dendritic Protrusions in Cultured Neurons

Quantification of dendritic protrusions was performed by the method of Hotulainen et al. (2009) with minor modification. Briefly, the images of transfected pyramidal neuron were taken with a ×63 objective lens of Leica TCS SP8 confocal microscope. Each dendritic spine was traced manually using ImageJ (National Institutes of Health) with Fiji plugins and categorized as filopodia, thin spine, mushroom spine, and stubby spine. Filopodia was defined as thin dendritic protrusion without a head. Thin spine was defined as long neck dendritic protrusion with <0.75-μm head diameter. Mushroom spine was defined as short neck dendritic protrusion with >0.75-μm head diameter. Stubby spine was defined as dendritic protrusion without a distinguishable neck. Dendrites were selected randomly and protrusions counted approximately a 50-μm dendrite length. Data were showed as density of spines or filopodia-like protrusions per micrometer dendrite length. The statistics were conducted with a Kruskal–Wallis test followed by Dunn’s post hoc test.

Serum Response Factor Reporter Gene Activity

Serum response factor (SRF) activation by Fhod3 was estimated using a serum response element (SRE)-dependent reporter as described previously (Arimura et al. 2013) with minor modifications. Briefly, HEK293 cells (3.5 × 10⁴) were plated onto a 24-well plate and transfected using PEI MAX (Polysciences) with 67 ng of pEF-BOS-Fhod3, 13 ng of pmEGFP-rCaMKIIa, 20 ng of phRL-TK encoding for Renilla luciferase (Promega), and 400 ng of pSRE-Luc reporter plasmid (Clontech). After 16 h of transfection, the culture medium was changed to decrease the concentration of fetal calf serum from 10% to 0.5%. After culture for another 12 h, luciferase activity was determined using the Dual Luciferase Reporter Assay System Kit (Promega) with a luminometer (TD-20/20; Promega). SRE firefly luciferase counts were normalized to the activity of the internal control, Renilla luciferase, and calculated as fold transactivation in comparison with the counts from cells cotransfected with pEF-BOS-Fhod3-WT and the empty pmEGFP vector.

Real-Time PCR Analysis

Real-time PCR analysis was performed as described previously (Sanematsu et al. 2019) with minor modifications. Briefly, total RNAs were extracted from the cerebral cortex using TRIzol reagent (Thermo Fisher Scientific). Complementary DNAs were synthesized using Superscript First-Strand Synthesis System for reverse transcription-PCR (Invitrogen). Real-time PCR was performed using the TB Premix Ex Taq II (TAKARA) on StepOnePlus Real-Time PCR System (Thermo Fisher Scientific). Primers for Fmnl1, Fmnl2, Fmnl3, mDia1, mDia2, mDia3, Fhod1, Inf1, Fmn1, Fmn2, Daam1, Daam2, and Delphilin were according to the study by Rosado et al. (2014). Primers for Inf2 were according to the study by Subramanian et al. (2016). Primers for Fhod3 were as follows: forward 5′-AAGTTCCGCCTGGTAGTGAAGA-3′ and reverse 5′-TGTTGGACCAGGGTTTGACA-3′. Primers for GAPDH were described previously (Sanematsu et al. 2019). The relative abundance of mRNAs for formins was quantified using the standard curve method under the assumption that amplification efficiencies of the target formins and the reference GAPDH are the same. Only cycle threshold (C_t) values of <35 were used to compare the relative expression levels.

Analysis of Expression Patterns of Formins in the Mouse Brain

In situ hybridization (ISH) data and heat maps of ISH signal intensity for various formins were collected from Allen Mouse Brain Atlas (Available from: http://mouse.brain-map.org; Lein et al. 2007). Data were taken from C57BL/6J mice at P56.

Statistical Analysis

Data were shown as mean ± standard deviation. Statistical analysis was performed using GraphPad Prism 5.0 (GraphPad Software Inc., San Diego, CA). Two groups were compared by paired or unpaired Student’s t-test. Multiple groups were compared by analysis of variance followed by Tukey’s post hoc test, unless noted otherwise. The level of significance was set at P value <0.001, unless noted otherwise.

Results

Fhod3 Expression in the Brain

Two major forms of Fhod3 are expressed in the mouse organs in a tissue-specific manner; the long variant is strongly expressed in the heart, whereas the short variant is abundant in the brain (Fig. 1A; Kanaya et al. 2005). Although we have demonstrated that the cardiac Fhod3 long variant plays an essential role in the assembly and maintenance of myofibrils in the heart (Taniguchi et al. 2009; Kan-o et al. 2012; Ushijima et al. 2018), the role of the Fhod3 short variant in the brain has remained elusive. To address this question, we first examined the protein expression level of Fhod3 in various regions of the adult mouse brain. Immunoblotting revealed that the Fhod3 protein was prominently expressed in the cerebral cortex and also (but to a lesser extent) in the thalamus, hypothalamus, and olfactory bulb (Fig. 1B). On the other hand, in the hippocampus and cerebellum, Fhod3 was only marginally expressed.

Figure 1

Fhod3 expression in the adult mouse brain. (A, B) Proteins prepared from indicated tissues or brain regions of adult wild-type mice were analyzed by immunoblot with antibodies against Fhod3 (anti-Fhod3-C20), α-tubulin, or GAPDH. Positions for marker proteins are indicated in kDa. (C) The X-gal staining of sagittal (a–e and a’–e’) and coronal (f–l and f’–l’) sections of the adult brain of Fhod3^+/− (Fhod3^+/lacZ) and Fhod3^+/+ mice. Scale bars, 2 mm. (D) The layer-specific expression of Fhod3 in the adult brain. Consecutive sagittal sections of Fhod3^+/− (Fhod3^+/lacZ) adult mice were stained with either X-gal or immunohistochemically with anti-Brn2. The orientation of the images in the lower panels is shown as dotted boxes in upper panels. Scale bars, 1 mm (upper panels) and 100 μm (lower panels).

Distribution of Fhod3 in the Cerebral Cortex

We next examined the Fhod3 expression pattern in the adult brain by X-gal staining of heterozygous Fhod3^+/− mice, which carry one Fhod3 null allele replaced with lacZ but show no discernible phenotype (Kan-o et al. 2012). As shown in Figure 1C, the expression of Fhod3 was observed in the neocortex and thalamus but not in the hippocampus, consistent with the results of immunoblotting. In the neocortex, Fhod3 showed a region-specific distribution pattern: Intense signals were detected within a single layer of the medial prefrontal and motor cortices, whereas 2 separate layers of signals were observed in the somatosensory cortex. No specific signal was detected in the retrosplenial cortex.

To identify the layers of the cerebral cortex (which consists of 6 layers) in which Fhod3 is expressed, we stained consecutive sections of the brain from Fhod3^+/− mice with an antibody against Brn2, a marker for layers II/III and V (Molyneaux et al. 2007), and with X-gal for the detection of β-galactosidase activity. As shown in Figure 1D, Fhod3 was predominantly expressed within layer II/III in the somatomotor cortex and layers II/III and V in the somatosensory cortex. We further examined Fhod3 expression in the developing cortex and found that Fhod3 expression can be detected in the isocortex around birth and became prominent during postnatal corticogenesis (Supplementary Fig. S1).

Fhod3 Is Expressed in Excitatory Neurons in the Cerebral Cortex

The cerebral cortex contains 2 fundamentally different cell types, that is, neuronal and glial cells. We then examined which type of cells express Fhod3 by double immunostaining for β-galactosidase, which showed a diffuse nuclear and/or a punctate cytoplasmic pattern, and several marker proteins (Fig. 2). Double immunostaining for a neuronal-specific nuclear protein (NeuN; Mullen et al. 1992) and β-galactosidase revealed that Fhod3 is expressed in neuronal cells (Fig. 2A). Neurons in the cerebral cortex are further divided into 2 major classes: glutamatergic neurons characterized by a typical pyramidal morphology and GABAergic inhibitory interneurons that exhibit a nonpyramidal morphology (DeFelipe and Fariñas 1992; Molyneaux et al. 2007). As shown in Figure 2B, double immunostaining for β-galactosidase and glutaminase, an enzyme that synthesizes glutamate, only detected β-galactosidase in glutaminase-positive excitatory neurons, even though some β-galactosidase-positive cells showed a punctate cytoplasmic pattern. In contrast, double immunostaining for β-galactosidase and glutamate decarboxylase, an enzyme that synthesizes GABA, only detected β-galactosidase in glutamate carboxylase-negative neurons and not in glutamate carboxylase-positive inhibitory neurons (Fig. 2C). Thus, Fhod3 is expressed specifically in glutamatergic pyramidal neurons in the cerebral cortex.

Figure 2

The expression of Fhod3 in excitatory neurons in the motor cortex. (A–C) Confocal fluorescence micrographs of layer II/III of the motor cortex of Fhod3^+/− (Fhod3^+/lacZ) adult mice. Sections of adult cerebral cortex were stained with the anti-β-galactosidase (green) and the anti-NeuN (A), or anti-glutaminase (B), or glutamate decarboxylase (C) (red) antibodies, followed by Hoechst (blue) staining. Scale bars, 5 μm.

Fhod3 Is Localized in Dendritic Spines and Postsynaptic Densities

Several formins have been shown to be localized in the dendritic spines and involved in the formation and maturation of their morphological changes. Thus, in order to investigate whether Fhod3 is localized in the dendritic spines, we examined the subcellular localization of the Fhod3 protein using a biochemical fractionation method. As shown in Figure 3A, Fhod3 protein was found in the synaptosome and enriched in the triton-insoluble PSD-enriched fraction. To confirm the ability of Fhod3 to localize into the dendritic spines, we transfected primary cortical neurons with Fhod3 cDNA. Neurons in primary culture develop axonal and dendritic arbors and form functional synaptic connections. During this process, filopodia-like protrusions extend from developing dendrites and evolve into dendritic spines. As shown in Figure 3B, EGFP-tagged Fhod3 accumulates in these dendritic protrusions (filopodia and spines). Thus, Fhod3 protein seems to accumulate in postsynaptic dendritic filopodia and spines.

Figure 3

The accumulation of Fhod3 in postsynaptic spines. (A) Biochemical fractionation of the mouse cerebral cortex. Equal amount of proteins (10 μg) from fractions, including initial homogenate (whole lysate), synaptosomal fraction (Synaptosome), and the triton-soluble (Tx-soluble) and insoluble (Tx-insoluble) PSD-enriched fractions from a synaptosomal fractionation of mouse cerebral cortex, were analyzed by immunoblot with antibodies against Fhod3 (anti-Fhod3-C20), PSD-95, β-actin, and GAPDH. (B) Representative confocal images of cultured cortical neurons expressing GFP-Fhod3 (green) and mCherry (red) at 14 days in vitro (DIV). Ratiometric images (GFP-Fhod3 vs. mCherry) are also shown. Scale bar, 5 μm.

Conditional Deletion of Fhod3 in the CNS

To address the role of Fhod3 in the physiological context, we conditionally deleted Fhod3 from the entire CNS by crossing Fhod3^flox mice (Ushijima et al. 2018) with Nestin-Cre mice. The resultant Fhod3 cKO mice (Fhod3^flox/−; Nestin-Cre⁺) were born in the expected Mendelian ratio and developed normally. The protein expression of Fhod3 in the whole brain of Fhod3 cKO mice was reduced to ~10% of that of wild-type mice, as demonstrated by immunoblotting of mouse brain tissue (Fig. 4A). On the other hand, the size and organization of the brain appeared indistinguishable from wild-type mice (Fig. 4B). Using the brain of Fhod3 cKO mice, we performed immunofluorescence staining of Fhod3 (Fig. 4C). Although the Fhod3 antibody used cross-reacted with some cytoarchitectural components, including the Golgi apparatus (Supplementary Fig. S2), the accumulation of immunoreactive signals for endogenous Fhod3 was specifically detected in layer II/III of control mice (Fhod3^flox/+) but not in layer II/III of Fhod3 cKO mice (Fhod3^flox/−; Nestin-Cre⁺). As shown in Figure 4D, Fhod3 immunoreactivity was detected not only in cell bodies but also in the surrounding regions within the layer II/III, in a punctate pattern. These Fhod3 puncta were partially colocalized with puncta of vGlut1 and PSD-95 (pre- and postsynaptic markers of excitatory synapses, respectively; Fig. 4E), supporting the presence of Fhod3 in dendritic spines that carry excitatory synapses. Thus, the Nestin-Cre-mediated depletion of Fhod3 from the entire CNS does not seem to exert a significant gross anatomical change in mice, despite the almost complete absence of Fhod3 protein.

Figure 4

Genetic deletion of Fhod3 in the mouse CNS. (A) Proteins of lysates prepared from the brain of Fhod3 cKO mice (Fhod3^flox/−; Nestin-Cre⁺) and control mice (Fhod3^flox/+ and Fhod3^flox/+; Nestin-Cre⁺) at P35–45 were analyzed by immunoblot with the anti-Fhod3 and anti-α-tubulin antibodies. Positions for marker proteins are indicated in kDa. (B) Histological analyses of the brain of Fhod3 cKO and control littermate mice. Paraffin-embedded sections of brains were stained with hematoxylin and eosin. Scale bars, 1 mm. (C, D) Confocal fluorescence micrographs of the cerebral cortex of Fhod3 cKO and control littermate mice. Sections of adult cerebral cortex at the motor area were stained with the following anti-Fhod3-(650–802) (red) and the anti-Brn2 (green) antibodies and DAPI (blue). Scale bars, 5 μm. (E) Colocalization of Fhod3 with excitatory synaptic markers in vivo. Sections of the control adult cerebral cortex layer II/III at the motor area were stained with the anti-vGlut1 (red), anti-PSD-95 (green), and anti-Fhod3-(650–802) (blue) antibodies. Scale bars, 10 μm.

Fhod3 Depletion Causes Abnormal Spine Morphology and Synapse Loss in the Cerebral Cortex

To investigate the physiological role of Fhod3 in in vivo spine morphology, we performed Golgi staining and analyzed the spiny pyramidal cells within layer II/III of the medial prefrontal and motor cortices, where Fhod3 is highly expressed. There were no apparent differences in dendritic patterns between Fhod3 cKO (Fhod3^flox/−; Nestin-Cre⁺) and control mice. However, there were clear differences in the spine morphology on the apical dendrites between the Fhod3 cKO and control mice (Fig. 5A). In cKO mice, the number of filopodia was significantly increased, whereas the number of spines was decreased (Fig. 5B–D). Similar results were also observed in the protrusions on basal dendrites (Fig. 5E–H). In contrast, there was no significant difference in the dendritic spine morphology of Purkinje cells (Fig. 5I–L), where Daam1, another member of the formin family, plays a critical role in the formation of dendritic spines (Kawabata Galbraith et al. 2018). Thus, Fhod3 depletion led to dendritic spine malformation specifically in the cerebral cortex.

Figure 5

Effects of Fhod3 deletion on the spine morphology in vivo. (A, E, I) Representative Golgi-stained images of apical (A) and basal (E) dendrites of pyramidal neurons within layer II/III of the motor cortex and dendrites of Purkinje cells in the cerebellum (I) from Fhod3 cKO mice (Fhod3^flox/−; Nestin-Cre⁺) and control mice (Fhod3^flox/+ and Fhod3^flox/+; Nestin-Cre⁺). Scale bars, 5 μm. Quantification of density of dendritic protrusions of apical (B–D) and basal (F–H) dendrites of pyramidal neurons within layer II/III of the motor cortex and dendrites of Purkinje cells in the cerebellum (J–L). The density of protrusions was calculated as the number of protrusions on secondly or tertiary dendrites from Fhod3^flox/+ mice (apical: n = 22 dendritic protrusions from 18 neurons; basal: n = 23 dendritic protrusions from 19 neurons; Purkinje: n = 30 dendritic protrusions from 18 cells), Fhod3^flox/+; Nestin-Cre⁺ mice (apical: n = 19 dendritic protrusions from 14 neurons; basal: n = 21 dendritic protrusions from 16 neurons; Purkinje: n = 30 dendritic protrusions from 19 cells), and Fhod3^flox/−; Nestin-Cre⁺ mice (apical: n = 25 dendritic protrusions from 17 neurons; basal: n = 18 dendritic protrusions from 16 neurons; Purkinje: n = 30 dendritic protrusions from 18 cells) from 3 independent mice of each genotype. Values are means ± SD. ^*P < 0.001.

To examine the effect of Fhod3 deletion on synaptic formation, we performed immunofluorescence staining of synaptic markers within layer II/III of the motor cortices. In control mice, some PSD-95 puncta were colocalized with vGlut1 (Fig. 6A, upper panels), indicating formation of excitatory synapses (Marzo et al. 2016). In contrast, Fhod3 cKO mice (Fhod3^flox/−; Nestin-Cre⁺) exhibited fewer synapses with a decreased number of PSD-95 puncta (Fig. 6A, lower panels and Fig. 6B). In the retrosplenial cortex where Fhod3 is not expressed, the number of PSD-95 puncta was not reduced (Supplementary Fig. S3). As shown by a biochemical fractionation method, the amount of PSD-95 in the triton-insoluble fraction from a synaptosomal fraction of the motor cortex was significantly decreased in Fhod3 cKO mice (Supplementary Fig. S4). Transmission electron microscopic analysis also showed loss of synaptic formation in Fhod3 cKO mice (Fig. 6C,D). It thus seems likely that depletion of Fhod3 affects the spine morphology and results in synaptic defect.

Figure 6

Effects of Fhod3 deletion on the synaptic formation in vivo. (A) Confocal fluorescence micrographs of the cerebral cortex of control (Fhod3^flox/+) and Fhod3 cKO (Fhod3^flox/−; Nestin-Cre⁺) mice. Sagittal sections of adult cerebral cortex layer II/III at the motor area were stained with the anti-vGlut1 (red) and anti-PSD-95 (green) antibodies and DAPI (blue). Scale bars, 5 μm. (B) Quantification of PSD-95 puncta per μm² in the layer II/III motor cortex. Data were obtained from Fhod3^flox/+ mice (n = 5) and Fhod3^flox/−; Nestin-Cre⁺ mice (n = 4) from 4 independent experiments. Three to four fields were taken per brain slice per mouse (total calculated area in motor cortex = 809470.15 μm² from 37 fields). Values are means ± SD. (C) Electron microscopic analysis of synapses in layer II/III of the medial prefrontal cortex of Fhod3 cKO and control mice. Scale bars, 500 nm. (D) Quantification of synapse number in images shown in (C). Asymmetric synapses identified by an electron-dense region associated with clear synaptic vesicles (arrowhead in C) were counted. Data were obtained from Fhod3^flox/+ mice (n = 3) and Fhod3^flox/−; Nestin-Cre⁺ mice (n = 3) at the age of 4–6 weeks. Six fields were taken per mouse (total calculated area in motor cortex = 810 μm² per mouse). Values are means ± SD.

Fhod3 Depletion Causes Abnormal Spine Morphology in a Specific Population of Pyramidal Neurons

To further investigate the functional role of Fhod3 in the spine morphology, we prepared primary cultures of cerebrocortical neurons from Fhod3 cKO mice (Fhod3^flox/−; Nestin-Cre⁺). However, we were not able to distinguish Fhod3-positive neurons from Fhod3-negative neurons in the dissociated culture, since the lacZ reporter gene was removed in addition to exon 18 of the Fhod3 gene by Cre-mediated recombination in the brain of Fhod3 cKO mice. Consequently, we failed to detect Fhod3 depletion-dependent changes in the spine morphology. We therefore adopted an alternative strategy: We prepared primary cerebrocortical neurons from Fhod3^{−/−Tg(αMHC-Fhod3)} embryos in which Fhod3 was systemically depleted and transgenically expressed only in the heart (Kan-o et al. 2012). This embryo expresses β-galactosidase both in the heart and brain under the control of endogenous Fhod3 promoter. Although this embryo showed exencephaly, as previously reported (Kan-o et al. 2012), we were able to identify the cerebrocortical region in which β-galactosidase was positive in a layer pattern (Fig. S5) and therefore successfully dissected the cerebral cortex from the extracranially herniated brain.

The primary cultures prepared from the cerebral cortex of Fhod3^{−/−Tg(αMHC-Fhod3)} embryos lack Fhod3 protein, whereas those from control Fhod3^+/− embryos express it (Fig. 7A). Using these primary cultures, we compared the dendritic spine morphology of β-galactosidase-positive neurons (i.e., Fhod3 promoter-active cells). As shown in Supplementary Figure S6, the β-galactosidase-positive neurons prepared from control Fhod3^+/− embryos indeed express Fhod3 protein. The β-galactosidase-positive neurons showed normal spiny dendrites with accumulation of PSD-95 (Fig. 7B, upper panels), vGlut1, and Fhod3 (Supplementary Fig. S7). In contrast, the β-galactosidase-positive neurons from Fhod3^{−/−Tg(αMHC-Fhod3)} embryos showed aberrant dendritic spine morphology (Fig. 7B, lower panels): Elongated filopodia-like protrusions were frequently observed, suggesting the involvement of Fhod3 in the expansion of the spine heads. A quantitative analysis revealed a significant increase in the number of dendritic filopodia, whereas the number of matured spines, such as stubby and mushroom-like, was decreased (Fig. 7C). The dendritic protrusion density and dendritic diameter were not significantly changed (Fig. 7D). Notably, these morphological changes were not observed in β-galactosidase-negative neurons (i.e., Fhod3 promoter-negative cells) prepared from the extracranially herniated brain of Fhod3^{−/−Tg(αMHC-Fhod3)} embryos (Fig. 7C and Supplementary Fig. S8), indicating that brain herniation per se does not lead to abnormal spine morphology. These results suggest that Fhod3 regulates spine head expansion in a specific population of pyramidal neurons in a cell type-specific manner.

Figure 7

Effects of Fhod3 deletion on the spine morphology in vitro. (A) Proteins of lysates of primary culture of cortex neurons prepared from Fhod3^{−/−Tg(αMHC-Fhod3)} embryos and control Fhod3^+/− embryos were analyzed by immunoblot with the anti-Fhod3 (C-20) and anti-α-tubulin antibodies. Positions for marker proteins are indicated in kDa. (B) Cortical neurons prepared from Fhod3^{−/−Tg(αMHC-Fhod3)} embryos and control Fhod3^+/− embryos were transfected at DIV 20 with Lifeact-mCherry and cultured for 24 h. Cells were fixed and double stained with the anti-PSD-95 (green) and anti-β-galactosidase (blue) antibodies. Scale bars, 10 μm. (C, D) Quantification of density of dendritic protrusions including all protrusions with different morphologies. The density of protrusions was calculated as the number of protrusions on dendrites of β-galactosidase-positive (shown in B) and -negative (shown in Supplementary Fig. S8) neurons from Fhod3^{−/−Tg(αMHC-Fhod3)} embryos (n = 627 dendritic protrusions from 7 neurons and n = 608 dendritic protrusions from 18 neurons, respectively) and β-galactosidase-positive (shown in B) and -negative (shown in Supplementary Fig. S8) neurons from control Fhod3^+/− embryos (n = 262 dendritic protrusions from 4 neurons and n = 320 dendritic protrusions from 9 neurons, respectively) from more than 3 independent cultures prepared from different mice. Values are means ± SD. ^*P < 0.001.

CaMKII Kinase Activity Facilitates Fhod3-Mediated F-Actin Assembly

We finally explored whether and how Fhod3 is regulated in excitatory neurons. In dendritic spines of excitatory neurons, Ca²⁺/calmodulin-dependent protein kinase II (CaMKII) plays pivotal roles in the activity-dependent changes in structure and function; CaMKII becomes autophosphorylated at Thr 286 as a result of Ca²⁺ influx through the NMDA-type glutamate receptor and then mediates morphological changes in spines via activation of Rho family GTPases and their effectors (Saneyoshi and Hayashi 2012; Hell 2014; Takemoto-Kimura et al. 2017). We tested whether CaMKII commits to the regulation of Fhod3-mediated actin assembly. To quantitatively assess the actin-assembling activity of Fhod3, we performed a reporter assay using a luciferase reporter plasmid under the control of SRF. It is established that activation of SRF correlates with in vivo actin polymerization status: The SRF coactivator MRTF associates with G-actin in the resting state but dissociates during actin polymerization to directly interact with SRF, resulting in transcriptional activation (Fig. 8A) (Miralles et al. 2003; Olson and Nordheim 2010). Actin polymerization mediated by formin family proteins including Fhod3 leads to the activation of SRF-dependent transcription (Copeland and Treisman 2002; Miralles et al. 2003; Arimura et al. 2013). As shown in Figure 8B, the wild-type Fhod3 short variant (brain type) exhibited the ability to induce SRF-dependent transcriptional activation when compared with Fhod3-IA, a defective mutant in binding to actin (Taniguchi et al. 2009). The wild-type Fhod3 activity was enhanced by coexpression of a constitutively active, calcium-independent form of CaMKII (CaMKII-T286D; Fong and Soderling 1990), but not by that of kinase-dead double mutant (CaMKII-K42R/T286D; Rongo and Kaplan 1999). Since the Fhod3-IA activity in the presence of CaMKII-T286D was low and comparable to that of CaMKII-K42R/T286D, it is suggested that the kinase activity of CaMKII induces SRF activation in a manner dependent on actin assembling activity of Fhod3. Thus, it seems possible that Fhod3-mediated actin assembly is regulated in a manner dependent on CaMKII during the activity-dependent morphological changes in spines.

Figure 8

Effects of CaMKII on the Fhod3-mediated F-actin assembly. (A) The basic principle of the measurement of Fhod3-mediated F-actin assembling activity using SRF reporter activity. See text for details. (B) HEK293 cells cotransfected with pSRE-Luc, phRLTK, pEF-BOS encoding wild-type (WT) Fhod3 or mutant Fhod3 carrying I976A substitution (IA), and pmEGFP-CaMKII-T286D or -K42R/T286D. Luciferase activity was normalized for transfection efficiency with Renilla luciferase activity. Fold activation was calculated as compared with normalized luciferase activity in cells cotransfected with pEF-BOS-Fhod3-WT and the empty pmEGFP vector, which was arbitrarily defined as 1.00. Each graph represents the mean ± SD of data from 3 independent transfection experiments.

Discussion

In the present study, we identified the critical role of Fhod3 in dendritic spine morphogenesis in the cerebral cortex. Unlike other neuronal formins that have previously been characterized, the expression of Fhod3 was hardly detected in the hippocampus but found to be abundant in the cerebral cortex. Fhod3 was expressed specifically in the excitatory pyramidal neurons in layers II/III and V of the cortex and was localized to the postsynaptic dendritic spines, thereby promoting spine head enlargement by regulating F-actin assembly. The Fhod3-mediated changes in the spine morphology only occurred in a specific population of pyramidal neurons and seemed to contribute to subtype-specific regulation of the spine morphology. To our knowledge, this is the first study showing that a mammalian formin contributes to spine morphogenesis of the cortex neurons in a neuronal subtype-specific manner.

Pyramidal neurons display a great diversity in their distribution, dendritic structure, projection pattern, and excitability (Hausser et al. 2000; Spruston 2008). However, the mechanism whereby dendritic spines change their shape and size in pyramidal neurons has been recognized as a one common to various neurons; in the currently accepted model, there are no differences between neuron subtypes. This is because the large majority of studies on actin regulators in dendritic spines have been carried out using hippocampal neurons (Ackermann and Matus 2003; Honkura et al. 2008; Gu et al. 2010; Bosch et al. 2014). Furthermore, the role of formins in dendritic spines has mainly been investigated using hippocampal neurons (Salomon et al. 2008; Hotulainen et al. 2009; Chazeau et al. 2014; Law et al. 2014). However, the expression patterns of actin-regulating proteins are not uniform across brain regions. It is therefore possible that mechanisms are variable depending on subtypes of neurons: The mechanism through which cerebrocortical neurons regulate spine morphogenesis may differ from that of hippocampal neurons. With regard to the formins, there are 15 mammalian members that have a distinct expression pattern (Krainer et al. 2013; Dutta and Maiti 2015; Kawabata Galbraith and Kengaku 2019). In the cerebral cortex, several members of formins are widely expressed but distributed in unique patterns (Supplementary Fig. S9). Intriguingly, some formins distribute in a mutually complementary manner not only at the level of brain regions but also across cortex layers. The biochemical property of actin assembly of each formin is unique and distinguishable (Courtemanche 2018). The present study demonstrates that Fhod3 is only expressed in a certain population of cortical neurons in layers II/III and V, indicating the possibility that Fhod3 is involved in a subtype-specific mechanism of regulation of the actin dynamics in dendritic spines.

During neuronal development, dendritic protrusions dynamically change their morphology; upon synaptic contact with the axon, thin dendritic filopodia can transform into more stable mushroom spines (Ziv and Smith 1996; Yoshihara et al. 2009; Hotulainen and Hoogenraad 2010). These complex steps seem to be achieved by different regulators. The formin mDia2 induces the elongation of dendritic filopodia by promoting actin polymerization at the filopodium tip (Hotulainen et al. 2009). Following the elongation of dendritic filopodia, mDia2-induced straight actin filament polymerization seems to be switched to the Arp2/3 complex-based polymerization of branched actin networks. In contrast to the study by Hotulainen et al., our present findings show that the formin Fhod3 is involved in the transition from filopodia to mushroom by promoting expansion of spine head. Since exogenously expressed Fmnl2, another formin family member, can localize at the tips of finger-like protrusions in mature spines and seems to contribute to spine enlargement (Chazeau et al. 2014), Fhod3 may function in a similar manner.

Modifications of spine morphology also occur in response to synaptic activation. This activity-dependent modification—enlargement or shrinkage of spines during long-term potentiation (LTP) or depression—is associated with learning and memory. At the initial phase of LTP, cofilin-mediated depolymerization of actin filaments is required for the rapid reorganization of the actin cytoskeleton (Gu et al. 2010; Bosch et al. 2014). The concomitant polymerization of actin seems to be achieved by CaMKII-mediated activation of various types of regulators including Arp2/3 complex, although the precise and comprehensive mechanism of regulation remains elusive. Our present findings implicate Fhod3 as one of the possible candidates for regulators of the activity-dependent reorganization of actin filaments in spines. Since learning and memory are based on synaptic communication across a broad network of various types of neurons, understanding the factors involved in the cell type-specific regulation of spine morphology is important. Future studies are awaited to elucidate the detailed molecular mechanism of Fhod3-mediated F-actin assembly in dendritic spines of a particular subset of neurons during neuronal development and synaptic plasticity.

Notes

We thank Drs Hideki Nishitoh (University of Miyazaki) and Naoya Murao (University of Miyazaki) for technical advice and helpful discussion. Dr Fumiyuki Sanematsu (University of Miyazaki) for advice on microscopic analysis, Yumiko Nomura (University of Miyazaki) for advice on histological analysis, Ami Inayoshi (University of Miyazaki), and Asami Akiyama (University of Miyazaki) for secretarial assistance. We also appreciate the technical support received from the Frontier Science Research Center, University of Miyazaki. Conflict of interest: The authors declare that they have no conflicts of interest with the contents of this article.

Funding

This work was supported in part by Grants-in-aid for Scientific Research (C) (JP18K06701 to T.N., JP19K07355 to R.T., JP19K11793 to Y.K.) from the Japan Society for the Promotion of Science; a Grant-in-aid for Research Activity Start-up (JP19K23830 to H.W.S.) from the Japan Society for the Promotion of Science; a Grant-in-aid for Scientific Research on Innovative Areas, “Platform of Advanced Animal Model Support,” (to R.T.) from the Ministry of Education, Culture, Sports, Science, and Technology; a grant from the Japan Research Institute of Industrial Science (Fukuyama) (to R.T.); a grant from Kobayashi Foundation (to R.T. and H.W.S.); a grant from TERUMO Life Science Foundation (to R.T.); the joint research program of the Biosignal Research Center, Kobe University (to R.T.); and the President’s Strategic Priority Budget of the University of Miyazaki (to R.T.).

Author notes

Hikmawan Wahyu Sulistomo and Takayuki Nemoto contributed equally to this work.