Knock-out of Tpm4.2/Actin Filaments Alters Neuronal Signaling, Neurite Outgrowth, and Behavioral Phenotypes in Mice

Sian Genoud; Chanchanok Chaichim; Rossana Rosa Porto; Tamara Tomanic; Holly Stefen; Esmeralda Paric; Soumalya Sarkar; Dasol Yoo; Wendi Gao; Edna C Hardeman; Peter W Gunning; Tim Karl; John Power; Thomas Fath

doi:10.1007/s12035-025-05259-9

. 2025 Aug 2;62(12):16316–16341. doi: 10.1007/s12035-025-05259-9

Knock-out of Tpm4.2/Actin Filaments Alters Neuronal Signaling, Neurite Outgrowth, and Behavioral Phenotypes in Mice

Sian Genoud ^1,^#, Chanchanok Chaichim ^2,^#, Rossana Rosa Porto ^3,^#, Tamara Tomanic ¹, Holly Stefen ¹, Esmeralda Paric ¹, Soumalya Sarkar ¹, Dasol Yoo ¹, Wendi Gao ², Edna C Hardeman ², Peter W Gunning ², Tim Karl ³, John Power ^2,^#, Thomas Fath ^1,^✉,^#

PMCID: PMC12559059 PMID: 40753314

Abstract

Tropomyosins (Tpm) are master regulators of actin dynamics through forming co-polymers with filamentous actin. Despite the well-understood function of muscle Tpms in the contractile apparatus of muscle cells, much less is known about the diverse physiological function of cytoplasmic Tpms in eukaryotic cells. Here, we investigated the role of the Tpm4.2 isoform in neuronal processes including signaling, neurite outgrowth, and receptor recycling using primary neurons from Tpm4.2 knock-out mice. Live imaging of calcium and electrophysiology data demonstrated increased frequency, yet reduced strength of single neuron spikes. Calcium imaging further showed an increase in neuronal networks. In vitro assays of Tpm4.2 knock-out neurons displayed impaired recycling of the AMPA neurotransmitter receptor subunit GluA1. Morphometric analysis of neurite growth showed increased dendritic complexity and altered dendritic spine morphology in Tpm4.2 knock-out primary neurons. Behavioral analysis of Tpm4.2 knock-out mice displayed heightened anxiety in the open field test, while the elevated plus maze displayed heightened anxiety only in females. Our study depicts the multi-faceted role of the Tpm4.2 isoform and its co-polymer F-actin population in neurons, with potential implications for better understanding diseases of the nervous system which involve actin cytoskeleton dysfunction.

Supplementary Information

The online version contains supplementary material available at 10.1007/s12035-025-05259-9.

Keywords: Tyopomyosin, Actin cytoskeleton, Neurons, Neuronal signaling

Introduction

Tropomyosins (Tpm) are helical coiled-coil dimers that polymerize head-to-tail along the actin filament and are regarded as a master regulator of actin dynamics in mammalian cells as they regulate the access of other actin-binding proteins [1]. The co-polymer nature of Tpms with filamentous actin (F-actin) is suggested to determine specific F-actin functions through influencing F-actin stability and promoting or inhibiting the activity of other actin-binding proteins [2, 3]. As Tpm isoforms are associated with molecularly distinct F-actin populations, Tpm isoforms can be used as a proxy to investigate the functional properties of F-actin populations [4]. In eukaryotic cells, over 40 different Tpm isoforms are found, arising by alternative splicing from four different genes. The expression of these isoforms has been found to be spatially and temporally regulated. Products from Tpm1, 3, and 4 are expressed in neuronal cells and have distinct spatial and temporal distributions related to their distinct functions [5]. Little is known regarding the physiological function of Tpm4 (also previously known as deltaTm) in non-muscle cells; however, Tpm4.2 is the only identified Tpm4 isoform in mice [6] and has been shown to enhance non-muscle myosin recruitment [7] and facilitate ER/Golgi trafficking [8].

Previous studies have identified Tpm4 isoforms expressed in neuron-like cells early in development, before a reduction in expression levels during maturation [9]. During neuronal differentiation, there is a shift from enrichment in axonal growth cone [10] to enrichment in the post-synaptic density [11], suggesting a functional change from a potential role in neurite outgrowth to a proposed role in synaptic plasticity of mature neurons [11, 12]. This is supported by evidence of increased neurites, filopodia, branch formation, and enlarged growth cone area in differentiating B35 neuroepithelial cells overexpressing Tpm4.2 [13]. Overexpression of Tpm4.2 is associated with an increase in phosphorylated (inactive) ADF (actin-depolymerizing factor)/cofilin. ADF/cofilin acts by severing actin filaments, therefore leading to increased actin stability and filament length [4, 13]. As the post-synaptic compartment contains a complex of scaffolding proteins, receptors, actin cytoskeleton proteins, adhesion, and signaling molecules, understanding the physiological function of Tpm4.2 in this compartment could provide essential insight into neuronal growth and synapse formation and signaling. In this study, we aim to further elucidate the physiological function of Tpm4.2 in neuronal development, maintenance, and signaling using primary neurons and behavioral testing in Tpm4.2 knock-out (Tpm4.2^−/−) mice [6].

Results

Expression Profile of Tpms in Mouse Brain

Immunoblotting was used to assess the brain region–specific expression of Tpm4.2 in the olfactory bulb, hippocampus, cerebellum, anterior cortex, and posterior cortex of 7-month-old wild-type (Tpm4.2^+/+) mice. Tpm4.2 was similarly expressed in all brain regions, with a trend towards higher expression levels found in the cerebellum, which did not reach significance. There was no significant difference in the relative abundance of Tpm4.2 between all brain regions (p > 0.05; Fig. 1). Relative abundance normalized to total protein is outlined in Table S1.

Fig. 1 — Tpm4.2 expression in the Tpm4.2^+/+ mouse brain. A Immunoblot results of sub-dissected brains from Tpm4.2^+/+ mice probed for Tpm4.2 (28 kDa) and normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH; 36 kDa). Olfactory bulb (OB), hippocampus (Hipp), cerebellum (CB), anterior cortex (CA), posterior cortex (CP). B Data are represented as min–max box plots with n = 4 per group. One-way ANOVA followed by Bonferroni’s multiple comparisons detected no significant difference between brain regions (p > 0.05). ns = not significant

To determine if any other Tpm isoforms compensate for Tpm4.2 deletion, immunoblotting was performed on brain tissue from 7-month-old Tpm4.2^+/+ and Tpm4.2^−/− mice. There were no significant differences in the relative expression of the Tpm3.1/2 isoforms in Tpm4.2^+/+ (1.051 ± 0.082) and Tpm4.2^−/− (1.078 ± 0.056), in total Tpm3 products between Tpm4.2^+/+ (0.17 ± 0.020) and Tpm4.2^−/− (0.22 ± 0.018), or in Tpm1.10/12 isoforms between Tpm4.2^+/+ (1.017 ± 0.054) and Tpm4.2^−/− (0.98 ± 0.036) mice (Fig. S1).

GluA1 Receptor Recycling in Tpm4.2^−/− Neurons

To assess whether Tpm4.2 plays a role in receptor recycling pathways, we performed an assay to induce receptor internalization by probing for surface and total (surface plus internalized) GluA1 receptors on dendrites of Tpm4.2^+/+ and Tpm4.2^−/− neurons following stimulation with glycine, N-methyl-D-aspartate (NMDA), and bicuculline (Fig. 2A, B). As expected, stimulated Tpm4.2^+/+ neurons showed a reduction in GluA1 receptors on the surface, compared with unstimulated neurons (Fig. 2C). NMDA-stimulated neurons had a 43.65% reduction in surface-total receptor ratio when compared with control extracellular solution (ECS) incubated neurons (p = 0.042). Glycine stimulation reduced this ratio by 63.21% (p = 0.0092), and bicuculline-stimulated neurons had a 49.84% reduction in surface-total receptors when compared with ECS (p = 0.011). Tpm4.2^−/− neurons had a much higher surface-total GluA1 receptor ratio when stimulated with NMDA (52.77% increase; p = 0.0035), glycine (66.45% increase; p = 0.0002), and bicuculline (44.79%; p = 0.012, Fig. 2C).

Fig. 2 — GluA1 receptor recycling in Tpm4.2^−/− neurons. A Representative immunocytochemistry images of surface and total GluA1 receptors from unstimulated 18 days in vitro primary hippocampal Tpm4.2^+/+ and Tpm4.2^−/− neurons in extracellular solution (ECS), compared with B surface and total GluA1 receptors on bicuculline-stimulated neurons. C A significant reduction in surface to total (surface plus internalized) ratio was observed in Tpm4.2^+/+ neurons when stimulated with NMDA and bicuculline (BIC) when compared with non-stimulated ECS Tpm4.2^+/+ neurons. A significant increase in surface to total ratio was also observed in NMDA and BIC-stimulated neurons of Tpm4.2^−/− neurons compared with their Tpm4.2.^+/+ controls reflective of a loss in GluA1 internalization. n = 10 dendrites per neuron, 10 neurons per condition × 4 biological replicates. Red boxes indicate area of dendrite measured. Data are represented as min–max violin plots with median and quartiles indicated by solid and dashed lines respectively. Scale bar = 5 µm; ns = not significant, *p < 0.05, **p < 0.005, ***p < 0.0005, ****p < 0.0001

Neuronal Activity in Tpm4.2^−/− Versus Tpm4.2^+/+ Neurons

Calcium Spike Analysis

Fluorescent calcium imaging was used to assess the role of Tpm4.2 in controlling neural network activity. Individual regions from Tpm4.2^+/+ and Tpm4.2^−/− neuronal cultures were imaged at 20 days in vitro (DIV) (Tpm4.2^−/−, n = 2094 neurons/26 FOV/4 preparations; Tpm4.2^+/+, n = 775 neurons/7 FOV/3 preparations). Neuronal density did not differ between groups (p > 0.05, Fig. S2).

Neuronal activity was primarily characterized by synchronized somatic calcium spikes (Fig. 3A(a), (b)). These synchronous events were on average 138% more frequent in the Tpm4.2^−/− (3.1 ± 0.3 Hz) than in Tpm4.2^+/+ (1.3 ± 0.5 Hz) neuronal cultures (unpaired t-test, t(31) = 2.816, p = 0.008; Fig. 3B). The amplitude of the synchronous events did not differ between Tpm4.2^−/− and Tpm4.2^+/+ cultures (Mann–Whitney U = 76, p = 0.53; Fig. 3C). Calcium spikes in individual neurons occurred 65% more frequently in the Tpm4.2^−/− (3.9 Hz) than in Tpm4.2^+/+ (2.3 Hz) neurons (nested t-test, t(31) = 2.251, p = 0.03; Fig. 3D). However, these individual spike peaks were ~ 60% smaller in Tpm4.2^−/− (0.08 ΔF/F) than Tpm4.2^+/+ (0.20 ΔF/F) neurons (nested t-test, t(31) = 2.845, p = 0.008; Fig. 3E). No differences were observed in the coefficient of variation of either the amplitude (nested t-test, t(31) = 1.037, p = 0.31) or inter-event interval (nested t-test, t(31) = 1.037, p = 0.31) of the calcium spikes.

Electrophysiological Analysis

mEPSCs in Dissociated Cultures of Primary Tpm4.2^+/+ and Tpm4.2^−/− Neurons

To investigate whether depleting Tpm4.2 would reduce F-actin stability and therefore affect synaptic function, we recorded mEPSCs from cultured neurons, prepared from embryos of Tpm4.2^−/− mice, and Tpm4.2^+/+ controls (Fig. 4A). Tpm4.2^−/− cells had significantly reduced mEPSC frequency (Fig. 4B; Tpm4.2^+/+, 8.20 ± 1.26 Hz; Tpm4.2^−/−, 3.32 ± 0.66 Hz; Mann–Whitney test p = 0.002) and amplitude (Fig. 4F; Tpm4.2^+/+, 33.79 ± 2.69 pA; Tpm4.2^−/, 22.56 ± 1.53 pA; unpaired t-test t = 3.57, p = 0.001). Tpm4.2^−/− mEPSCs had increased rise time (Fig. 4G; Tpm4.2^+/+, 0.58 ± 0.04 ms; Tpm4.2^−/−, 0.70 ± 0.03 ms; unpaired t-test p = 0.02) and decay (Fig. 4H; Tpm4.2^+/+, 3.85 ± 0.32; Tpm4.2^−/−, 5.3 ± 0.29; unpaired t-test p = 0.002). We also noted that Tpm4.2^−/− neurons had a higher membrane resistance (Fig. 4D; Tpm4.2^+/+, 226.1 ± 27.88; Tpm4.2^−/−, 337 ± 39.77; Mann–Whitney test p = 0.03) and lower membrane capacitance (Fig. 4E; Tpm4.2^+/+, 105 ± 4.11 pF; Tpm4.2^−/−, 94.45 ± 2.62 pF; unpaired t-test t = 2.14, p = 0.04). Relative amplitude frequency was also significantly reduced (Fig. 4I), and relative inter-event interval frequency was significantly increased in Tpm4.2^−/− neurons (Fig. 4J).

Fig. 4 — Electrophysiology analysis of Tpm4.2^−/− neurons and acute brain slices. A Example recording traces for Tpm4.2^+/+ and B Tpm4.2^−/−. C Mean mEPSC frequency was significantly decreased in Tpm4.2^−/− neurons. D Tpm4.2^−/− neurons had a higher membrane resistance and E lower membrane capacitance than Tpm4.2^+/+ neurons. F Mean mEPSC amplitude was significantly decreased in Tpm4.2^−/− neurons. G Mean mEPSC rise time and H decay were increased in Tpm4.2^−/− neurons. I Cumulative probability histograms of mEPSC inter-event interval and J amplitude; 200 events sampled from each cell were significantly different in Tpm4.2^−/− neurons. n = 17 cells from 3 separate culture preparations. Measuring basal synaptic activity in Tpm4.2^−/− brain slices. K Locations of recording and stimulating electrodes in CA1 hippocampus. L Paired pulse ratio, a measure of synaptic release probability. M IO curve of fEPSP slope. N IO curve of fiber volley. O fEPSP slope plotted against fiber volley to show synaptic efficacy. Tpm4.2^+/+ n = 28 slices from 19 mice, Tpm4.2^−/− n = 31 slices from 19 mice. Data are represented as min–max violin plots with median and quartiles indicated by solid and dashed lines respectively. Significance was determined as *p < 0.05; **p < 0.01; and ****p < 0.0001. ns = not significant

Field Excitatory Post-synaptic Potentials (fPESPs) in Tpm4.2^+/+ and Tpm4.2^−/− Brain Slices. The Effect of Tpm4.2 Knock-out on Basal Synaptic Transmission

As the depletion of Tpm4.2 caused a significant reduction in mEPSC amplitude and frequency in dissociated primary neuron cultures, we examined the effect in acute brain slices prepared from Tpm4.2^−/− mice (Fig. 4K). An input–output (IO) curve was constructed to assess basal function. There was no significant difference between groups in baseline paired pulse ratio (PPR) (Fig. 4L, unpaired t-test p = 0.57), fEPSP slope (Fig. 4M; two-way repeated measures ANOVA interaction p = 0.96, genotype p = 0.87), fiber volley (Fig. 4N; two-way repeated measures ANOVA interaction p = 0.79, genotype p = 0.85), or synaptic efficacy (Fig. 4O).

The Effect of Tpm4.2 Knock-out on LTP

The effect of Tpm4.2 deletion on synaptic plasticity was assessed, using extracellular field potentials in brain slices. After fEPSP amplitude stabilization and induction of LTP (Fig. 5A), there was no change in the magnitude of LTP induced between groups, either at the early or late stages (Fig. 5B; two-way ANOVA interaction p = 0.52, genotype p = 0.69). The average level of stimulation chosen to run the protocol based on the IO curve was similar for both groups (Fig. 5C, D; Tpm4.2^+/+ = 12.93 ± 0.35 V; Tpm4.2^−/− = 12.63 ± 0.44 V; p = 0.62). PPR was measured throughout to check for presynaptic changes. We observed no significant differences in PPR at any point in the experiment (Fig. 5E; two-way RM ANOVA interaction p = 0.33, genotype p = 0.07).

Fig. 5 — The effect of Tpm4.2 knock-out on LTP (A–E) and LTD (F–J) in acute brain slices. A Plot of fEPSP over the course of the experiment, with each point representing the average from 2 min of recording for LTP. B Mean normalized fEPSP at first and last 10 min of recording after inducing LTP was not significantly different between Tpm4.2^+/+ (black) and Tpm4.2^−/− (red) cultures. C Example waveforms from baseline (blue), first 10 min after low frequency stimulation (red) and last 10 min of recording (green) for LTD. D Stimulation intensities used for LTP. E PPR at baseline and first and last 10 min after high-frequency stimulation was not significantly different between groups for LTP. F Plot of fEPSP over the course of the experiment, with each point representing the average from 2 min of recording for LTD. G Mean normalized fEPSP at first and last 10 min of recording after inducing LTD was not significantly different between Tpm4.2^+/+ (black) and Tpm4.2^−/− (red) cultures. H Example waveforms from baseline (blue), first 10 min after low frequency stimulation (red) and last 10 min of recording (green) for LTD. I Stimulation intensities used for LTD were the same as those used for LTP (D) experiments. J PPR at baseline and first and last 10 min after stimulation was not significantly different between groups for LTD. Tpm4.2^+/+ n = 19 slices from 11 mice, Tpm4.2^−/− n = 18 slices from 9 mice

The Effect of Tpm4.2 Knock-out on LTD

A stimulation level evoking approximately half of the maximal response was chosen. A stable baseline of at least 20 min was recorded, before 15 min of a 1-Hz paired pulse to induce LTD (Fig. 5F). No significant differences in the level of depression either at the early or late stage of recording were identified (Fig. 5G; two-way repeated measures ANOVA interaction p = 0.58, genotype p = 0.42). The stimulation level used for the experiments did not differ between the two groups (Fig. 5H, I; Tpm4.2^+/+ = 13.58 ± 0.31 V; Tpm4.2^−/− = 13.5 ± 0.38 V; unpaired t-test t = 0.16, p = 0.87). The PPR was also not significantly altered (Fig. 5J; two-way repeated measures ANOVA interaction p = 0.07, genotype p = 0.25).

Neurite Complexity of Tpm4.2 Knock-out Neurons

To investigate whether Tpm4.2 knock-out affects neuronal morphology, morphometric analysis was conducted to quantify neurite outgrowth in Tpm4.2^−/− compared with Tpm4.2^+/+ neurons (Figs. S3 and 6). Tpm4.2^−/− axons were significantly longer than axons from Tpm4.2^+/+ neurons (37.91% increase, p < 0.0001, Fig. 6A). There was no significant difference in the number of axon primary branches (p > 0.05, Fig. 6B) or axon secondary branches (p > 0.05, Fig. 6C) between Tpm4.2^+/+ and Tpm4.2^−/− neurons.

There was no significant difference in the number of dendritic trees in Tpm4.2^−/− neurons (p > 0.05, Fig. 6D). However, there was a significant increase in the length and complexity of Tpm4.2^−/− dendrites compared with Tpm4.2^+/+ neurons. Tpm4.2^−/− neurons had a higher mean total length of dendritic trees (52.62% increase, p < 0.0001; Fig. 6E), dendrite total length (56.68% increase, p < 0.0001, Fig. 6F), and number of primary branches (47.86% increase, p = 0.0012, Fig. 6G). The mean length of primary branches was unchanged (Fig. 6H). Tpm4.2^−/− neurons had a higher mean number of primary branches per dendritic tree (33.02% increase, p = 0.0004, Fig. 6I). Sholl analysis identified that this increased dendritic complexity occurred between 20 and 90 µm from the soma (p < 0.0001, Fig. 6J).

Spine Morphology in Tpm4.2^−/− Neurons

There was no significant difference in dendritic spine density between the Tpm4.2^−/− and Tpm4.2^+/+ neurons (Fig. 7A, B; Tpm4.2^+/+, 0.87 ± 0.03 µm⁻¹; Tpm4.2^−/−, 0.8 ± 0.03 µm⁻¹; unpaired t-test t = 2.51, p = 0.25). There was also no significant difference in the mean length (Fig. 7C; Tpm4.2^+/+, 0.71 ± 0.02 µm; Tpm4.2^−/−, 0.74 ± 0.02 µm; unpaired t-test t = 2.06, p = 0.29) or width (Fig. 7D; Tpm4.2^+/+, 0.39 ± 0.01 µm; Tpm4.2^−/−, 0.37 ± 0.01 µm; unpaired t-test t = 1.32, p = 0.19) of the dendritic spines. However, the cumulative probability histograms showed a shift towards longer (Fig. 7E; Kolmogorov–Smirnov p = 0.007) and thinner (Fig. 7F; Kolmogorov–Smirnov p < 0.0001). Additionally, there was a difference in the distribution of dendritic spine types (Fig. S4; two-way ANOVA interaction p = 0.03). Tukey post hoc multiple comparisons however did not detect any significant differences between groups (p > 0.05 for all); however, there was a trend towards a decrease in stubby dendritic spines in Tpm4.2^−/− neurons compared with Tpm4.2^+/+ (Fig. S4; 0.1661 vs 0.1212 ± 0.018, 36.64% reduction; p = 0.067).

Mouse Behavioral Testing

We performed a battery of behavioral tests on Tpm4.2^+/+ and Tpm4.2^−/− littermates that assessed anxiety, learning, and socialization to test the effect of Tpm4.2 on these cognitive aspects.

Open Field (OF)

Three-way repeated measures ANOVA revealed a significant main effect of “time” on the OF distance travelled (F(5,205) = 100.5, p < 0.001, Fig. 8A). All animals exhibited a decline in the distance travelled across time, with females displaying greater locomotion than males (main effect of “sex”: F(1,41) = 5.7, p = 0.022, Fig. 8A). A similar sex effect was also demonstrated in the cumulative distance travelled over the 30-min protocol (F(1,41) = 5.7, p = 0.022, Fig. 8B), and no discernible “genotype” effect on distance travelled was detected (F(1,41) = 0.625, p = 0.434; Fig. 8B). The rearing frequency was similar between sexes (F(1,41) = 1.5, p = 0.223, Fig. 8C) and genotypes (F(1,41) = 1.5, p = 0.231, Fig. 8C).

When assessing anxiety behaviors, females spent less time (F(1,41) = 4.6, p = 0.037, Fig. 8D), yet had more entries (F(1,41) = 10.4, p = 0.003, Fig. 8E) in the open field center compared to their male counterparts. Notably, a main effect of “genotype” was evident for center entries (F(1,41) = 4.089, p = 0.049, Fig. 8E) and center distance ratio (F(1,41) = 4.614, p = 0.037, Fig. 8F), with Tpm4.2^−/− mice exhibiting fewer entries and reduced locomotion in the center of the open field, indicating heightened anxiety levels in Tpm4.2^−/− mice relative to Tpm4.2^+/+ regardless of sex (no “sex” by “genotype” interaction, p > 0.05).

Elevated Plus Maze (EPM)

Behaviors assessed using EPM appeared largely sex-specific, with male Tpm4.2^−/− mice demonstrating an anxiolytic-like phenotype, whereas female Tpm4.2^−/− mice had higher anxiety levels compared to the respective controls. Two-way ANOVA revealed significant “sex” by “genotype” interactions for time spent on open arms (F(1, 41) = 12.13, p = 0.001, Fig. 8G) and entries into open arms (F(1, 41) = 6.712, p = 0.012, Fig. 8H) as well as for the distance ratio exploring the open arms (F(1, 41) = 12.65, p = 0.001, Fig. 8I). Subsequent post hoc analysis splitting the data by sex revealed that female Tpm4.2^−/− mice spent less time in (t(20) = 3.269, p = 0.003, Fig. 8G) and exhibited fewer entries into (t(20) = 2.811, p = 0.010, Fig. 8H) the open arms than their respective controls, whereas no genotype differences were observed for these behaviors in male mice. Conversely, Tpm4.2^−/− males exhibited an increased locomotion ratio in the open arms compared to Tpm4.2^+/+ control males (t(21) = 2.102, p = 0.047, Fig. 8I), while female Tpm4.2^−/− mice displayed a decreased ratio distance in the open arms (t(20) = 3.037, p = 0.006, Fig. 8I). Additionally, a main effect of sex (F(1, 41) = 5.817, p = 0.020) and genotype (F(1, 41) = 5.017, p = 0.0301) was observed for the time spent in the center of the maze (Fig. 8J).

Social Preference Test (SPT)

A three-way RM ANOVA revealed a significant main effect of “chamber” for the nosing time in the sociability trial (F (1, 35) = 96.07, p < 0.001, no interactions, Fig. 9A)—all groups explored the A/J mice more than the empty chamber. Similar findings were observed using one-sample t-tests to analyze percentage exploration of each group compared to chance exploration (i.e., 50%). All animals, regardless of genotype or sex, showed a significant preference above chance levels for the side with the A/J mouse (p < 0.05 for all, Fig. S5A).

Fig. 9 — Social preference test in Tpm4.2^−/− mice. Sociability test behaviors—social preference and social novelty. Total time spent nosing chambers, containing A an A/J mouse versus no mouse (i.e., empty chamber) (sociability) or B a novel versus a familial A/J mouse (social novelty preference). Data are shown as mean ± SEM for Tpm4.2^+/+ (black) n = 25 mice and Tpm4.2.^−/− (red) n = 20 mice. Three-way RM ANOVA effects of “side” are indicated by *** (p < 0.001)

Three-way RM ANOVA also revealed a significant main effect of “chamber” for the nosing time in the social novelty preference test (F (1, 35) = 18.39, p < 0.001, no interactions, Fig. 9B). However, when using one-sample t-tests to also analyze percentage exploration compared to chance exploration, males of both genotypes preferred the new mice over the familiar ones (p > 0.05 for all, Fig. S5B), whereas no females developed a preference for the novel mouse (t(12) = 1.183, p = 0.091, Fig. S5B; t(8) = 0.477, p = 0.645, Fig. S5B). This is in line with our previous studies [14].

Novel Object Recognition Test (NORT)

A three-way RM ANOVA detected a significant main effect of “object” (F(1,38) = 8.129, p = 0.007) and “genotype” (F(1,38) = 6.466, p = 0.015), with the latter indicating that Tpm4.2^−/− mice exhibited lower levels of object exploration compared to Tpm4.2^+/+ across sex (Fig. 10A). Additionally, a significant “object” by “sex” by “genotype” triple interaction was observed (F(1,38) = 4.459, p = 0.041, Fig. 10A). Based on the triple interaction, data were split for corresponding factors. Subsequent paired t-tests elucidated an increased exploration of the new object by Tpm4.2^−/− males (t(10) = 3.407, p = 0.006, Fig. 4A), which was also trending for females Tpm4.2^+/+ mice (t(11) = 2.035, p = 0.066). Male Tpm4.2^+/+ controls (t(8) = 1.478, p = 0.177, Fig. 10) and Tpm4.2^−/−females (t(9) = 0.704, p = 0.499, Fig. 10) failed to show a preference for the novel object. To analyze percentage exploration compared to chance levels with one-sample t-tests, we observed a similar finding (Fig. S6). Male Tpm4.2^−/− mice had a preference for the novel object (t(10) = 3.644, p = 0.0045), whereas their respective Tpm4.2^+/+ controls exhibited no statistically significant difference (t(8) = 1.779, p = 0.113). Again, the female Tpm4.2^+/+ displayed a trend in nosing index (t(11) = 2.176, p = 0.052), while the Tpm4.2^−/− females explored both objects equally (t(9) = 0.216, p = 0.833).

Contextual and Cue Fear Conditioning (CFC)

During the conditioning phase, a three-way repeated measures ANOVA revealed a significant main effect of “time” (F(6, 246) = 93.80, p < 0.001, Fig. 10B), indicating an overall increase in freezing post foot shock across all animals. While there were no discernible effects of “sex” (F(1, 41) = 0.696, p = 0.409, Fig. 10B) or “genotype” (trend only: F(1,41) = 3.621, p = 0.064, Fig. 10B), interactions of “time” with “sex” (F(6, 246) = 5.736, p < 0.001, Fig. 10B) and with “genotype” (F(6, 246) = 3.142, p = 0.005, Fig. 10B) were evident. However, further examination using a two-way ANOVA split by “time” disclosed effects of “genotype” and “sex” only at minute 7 (sex: F(1, 42) = 7.069, p = 0.011, genotype: F(1, 42) = 6.466, p = 0.014, Fig. 10B). Tpm4.2^−/− mice exhibited heightened freezing in the final minute of the test following two foot shocks compared to Tpm4.2^+/+ controls (p < 0.05), and Tpm4.2^−/− females also displayed increased freezing compared to Tpm4.2^−/− males (p < 0.05). Total freezing time analysis suggested a trend for a “genotype” effect (F(1, 41) = 3.621, p = 0.064, Fig. 10C), yet no sex effect was evident (F(1, 41) = 0.6955, p = 0.4091, Fig. 10C).

In the context test, freezing increased over time as expected (“time”: F(6, 246) = 7.301, p < 0.001, Fig. 10D). No effects of “sex” (F(1, 41) = 1.132, p = 0.293, Fig. 10D) or “genotype” (F(1, 41) = 0.006, p = 0.937, Fig. 10D) and no interactions with “time” (all p’s > 0.05) were discerned. Two-way ANOVA for total freezing time during the context test revealed no significant differences either (p > 0.05 for all, Fig. 10E).

A three-way RM ANOVA for the cue trial demonstrated an overall effect of “time” (F(8, 328) = 70.95, p < 0.001, Fig. 10F), with all animals exhibiting increased freezing levels in response to the cue. No effects of “sex” (F(1, 41) = 0.038, p = 0.845, Fig. 10F) or “genotype” (F(1, 41) = 0.008, p = 0.931, Fig. 10F) were observed. Similar non-significant results were found for total freezing time during cue presentation (3rd to 7th minute; data not shown). An overall effect of “cue x baseline” (average freezing per minute during baseline versus cue freezing, Fig. 10G) was identified in the three-way repeated measures ANOVA (F(1, 42) = 357.7, p < 0.001, Fig. 10G), confirming that the freezing response increases at cue presentation, while no effects of “sex” (F(1, 42) = 0.0208, p = 0.885, Fig. 10G) or “genotype” (F(1, 42) = 0.032, p = 0.858, Fig. 10G) were evident or interfered with the overall freezing response to the cue (no interactions with “time,” p > 0.05 for all).

Prepulse Inhibition (PPI)

In examining the startle response, a three-way repeated measures ANOVA yielded a significant main effect of “startle intensity” (F(2, 82) = 122.9, p < 0.001, Fig. S7A), demonstrating that the startle response increased in tandem with the intensity across all groups. No significant effects were observed for sex (F(1, 41) = 3.408, p = 0.0721, Fig. S7A) or genotype (F(1, 41) = 2.797, p = 0.102, Fig. S7A), and no interactions with “startle intensity” were evident either (all p’s > 0.05). Similarly, analysis of habituation to the startle over time revealed an overall effect for the startle pulse blocks (F(2, 82) = 17.25, p < 0.001, Fig. S7B), indicating habituation across all groups. No significant differences were found for sex (F(1, 41) = 2.487, p = 0.122, Fig. S7B) or genotype (F(1, 41) = 2.27, p = 0.139, Fig. S7B). Finally, when analyzing the prepulse inhibition response, all animals demonstrated an elevated PPI response to increasing prepulse intensities (F(2, 82) = 347.2, p < 0.001, Fig. S7C) regardless of experimental condition (all other p’s > 0.05).

Discussion

The current study examined Tpm4.2 depletion-dependent behavioral phenotypes and cellular functions of Tpm4.2 in developing and mature neurons, using cellular assays in primary hippocampal neuron cultures and acute brain slices and a battery of cognitive test regimes. Here, we confirm previous reports [10] that Tpm4.2 is expressed throughout the brain and identified a role for Tpm4.2 in aspects of neuronal signaling, with increases in synchronous firing and firing frequency yet impaired neuronal signaling strength in Tpm4.2 knock-out neurons. Tpm4.2 knock-out mice exhibited a mild behavioral phenotype of anxiety which was predominantly sex-dependent. Dissociated neuronal cultures of Tpm4.2-depleted neurons demonstrated increased dendritic complexity, impaired receptor internalization, and increased neuronal network connectivity; however, more complex brain slices exhibited no significant changes in basal activity or alterations in either LTP or LTD. These data indicate a physiological role of Tpm4.2 in aspects of neuronal signaling, neurite growth regulation, and development, with no discernable effect on overall synaptic plasticity.

Using primary hippocampal neurons as a neuronal model, we confirmed a previously suggested role of Tpm4.2 in neurite outgrowth, providing a physiological function for the high expression of Tpm4.2 levels in dendritic growth cones of immature neurons [10]. During maturation, Tpm4.2 expression becomes more localized to the post-synapse [12], where we demonstrate a role for Tpm4.2 in signaling frequency, strength, and AMPAR recycling at the post-synapse. As Tpm forms co-polymers with F-actin, these data suggest specific roles for the corresponding F-actin subpopulations in normal neuronal health and signaling. One possible mechanism is increased cofilin activity in response to the lack of Tpm4.2 to inhibit cofilin severing of actin filaments.

To determine whether Tpm4.2 is involved in cellular processes of learning and memory through long-term potentiation (LTP) and long-term depression (LTD), we analyzed receptor recycling processes, calcium signaling, and mEPSCs in dissociated primary hippocampal neurons as a neuronal model and fEPSPs in acute slices to provide a more complex biological system more representative of the human brain. We observed a reduced internalization of the GluA1 subunit component of AMPA receptors in Tpm4.2^−/− neurons stimulated with NMDA, bicuculline, and glycine, indicating the potential for overstimulation and excitotoxicity in Tpm4.2-depleted neurons. Selective modifications of post-synaptic AMPA receptors play a key role in the cellular mechanism for learning and memory [15]. The regulation of AMPAR is essential as it mediates most of the fast excitatory neurotransmission, and the abundance of AMPAR at the surface of excitatory synapses dictates the strength of responses to excitatory stimulation [16]. The AMPAR subunit GluA1 mediates activity-dependent changes in excitatory synaptic transmission between neurons [17]. Internalization of receptors following stimulation is a necessary process to prevent overstimulation. NMDA-mediated receptor internalization and LTD were achieved through nonselective activation of both synaptic and extrasynaptic NMDA receptors as previously described [18, 19]. Prolonged exposure to a high concentration of bicuculline and glycine was also used to induce LTD through increased receptor internalization as previously described [20, 21]. In neurons lacking Tpm4.2, however, GluA1 receptor subunit internalization was impaired, indicating a physiological function of Tpm4.2 in receptor recycling processes and therefore in mediating aspects of neuronal excitation. Diminished GluA1 internalization and the resulting increase in surface GluA1 receptors can have substantial effects on neuronal excitability and are proposed to contribute to the increased neuronal firing frequency observed in live imaging of calcium levels in Tpm4.2^−/− neurons in the current study.

Despite evidence that the actin cytoskeleton plays a key role in processes of endocytosis, including endosomal retrieval and recycling [22], the precise role of actin in AMPAR trafficking is not fully understood. As F-actin is abundant in dendritic spines, it is suggested to play a physiological role in regulating various aspects of the post-synapse—including receptor trafficking. Actin-depolymerizing drugs such as latrunculin reduce GluA1-containing AMPARs in dendritic spines [23] and reduce surface expression at synapses [24], while jasplakinolide, an F-actin-stabilizing drug, prevents AMPAR internalization [25]. In addition, there are reports of AMPAR pools that are regulated by actin processes and others that are unaffected, suggesting distinct actin subpopulations, some of which affect AMPAR recycling processes and some that are not involved in this process. As Tpms form co-polymers with F-actin and have previously been suggested to play a role in bulk endocytosis [26], we hypothesized that Tpms are involved in processes of receptor internalization in an isoform-specific manner. Here, we report a role for Tpm4.2 and its associated F-actin subpopulation in AMPAR recycling with impaired GluA1 internalization in Tpm4.2^−/− neurons. The concurrent increase in abundance of excitatory surface receptors ultimately increases the likelihood of further activation and hyperexcitability in Tpm4.2^−/− neurons—highlighting the important link between spine dynamics and synaptic strength.

Dissociated Tpm4.2^−/− neuron cultures showed increases in both synchronous and single-cell firing frequency when compared with Tpm4.2^+/+ neurons. However, the calcium response was diminished. The reduced calcium response may reflect a reduction in the number of APs generated within the underlying calcium spike or differences in ion-channel activity. The reduced mEPSC amplitude and frequency in Tpm4.2^−/− neurons (Fig. 4A–J) suggest a role for Tpm4.2 in maintaining synaptic strength. The decreased strength of individual synapses may attenuate the depolarization induced by network activity and the number of APs generated by a network event. The observed impairment in GluA1 receptor recycling (Fig. 2C) is consistent with this idea. It is feasible to suggest that Tpm4.2 depletion also affects the recycling of other receptors, including ion channels, which could account for the observed differences in passive membrane properties and mEPSC kinetics.

Despite observing an altered synaptic activity in Tpm4.2^−/− primary hippocampal neurons, we did not observe any significant differences in basal synaptic function and plasticity of Tpm4.2^−/− brain slices. Further, slice-specific factors such as network inhibition and glial buffering could explain discrepancies between the dissociated cultures and ex vivo brain slices. Another possible factor underlying this discrepancy is that brain slice experiments are performed in animals that are 6–8 weeks old, while dissociated cultures are harvested from embryonic mouse brains, and experiments are performed at 20DIV when neurons are deemed mature based on synapse formation and neuronal signaling [27]. It is possible that Tpm4.2 is only critical for synaptic activity during development, or that adult brains develop other compensatory mechanisms to account for the loss of Tpm4.2. Differences in dissociated neuron culture and acute brain slice experiments could also be attributed to differences in model complexity, with acute brain slices also containing other cells that would also be depleted of Tpm4.2. Tripartite synapses, for example, contain neurons and astrocytes, and as astrocytes exchange information with the synaptic neuronal elements and respond to synaptic activity, they could potentially compensate for the loss of neuronal synaptic activity in Tpm4.2^−/− neurons and mask this neuronal phenotype. Future studies could explore this through generating mice with neuron-specific Tpm4.2 knock-out.

Calcium amplitude was lower in the Tpm4.1^−/− neurons, consistent with reduced mEPSC amplitude. The enhanced frequency of calcium events could be due to compensatory changes in intrinsic excitability which act to normalize average firing across the network. We also observe differences in membrane resistance which may result in increased propensity for Tpm4.2^−/− to generate action potentials.

An additional contributor to neuronal signaling probability and strength is the morphology of the neuron and its neurites. Tpm4.2-depleted neurons demonstrated increased total axonal length and increased dendritic complexity early in development, with increases in the number of primary branches per tree, dendrite length, and number of interactions near the soma. The observed increased dendrite length, branching, and complexity in Tpm4^−/− neurons could strongly influence synapse dispersal along a dendrite [28], which in turn affect post-synaptic strength and excitability of neurons. On the other hand, with neurite arborization strongly influenced by extracellular signals and synaptic activity [29], increased dendrite branching and complexity may be a result of hyperexcitable Tpm4.2^−/− neurons with increased firing frequency. Further, these increases in dendrite complexity could contribute to the increased network connectivity displayed in live neuron imaging of calcium in Tpm4.2^−/− neurons.

On the molecular level, the effect of Tpm4.2 depletion on neurite growth may be associated with its property of cooperatively binding with tropomodulin (Tmod) to filamentous actin. Tropomyosins binding enhances tropomodulin’s capping activity of actin filament pointed ends [30], assisting in the stabilization of filaments and inhibiting their disassembly and turnover [31]. Tropomodulins also have isoform-specific effects, with Tmod3 not demonstrated to affect neurite outgrowth, while Tmod1 and Tmod2 are reported to increase dendritic complexity [32], and both Tmod1 and Tmod2 require both of their actin-binding sites to regulate dendritic morphology and dendritic spine shape [33]. Tpm4.2 may interact with Tmod1 or Tmod2, and therefore, Tpm4.2 depletion may affect this interaction and result in the observed increase in dendrite branching complexity of Tpm4.2^−/− neurons [34]. Neurite outgrowth is therefore proposed to be tightly regulated by numerous components of the actin cytoskeleton, with an isoform-specific function of Tpm4.2 in dendritic outgrowth and complexity.

Dendritic spine expression and morphology can also strongly influence neuron excitability and signaling strength. Dendritic spines are highly dynamic structures that can grow, change shape, and be eliminated through neuronal development, maturation, and maintenance. Understanding their morphological properties in different neuron populations can provide insight into their functional differences, with their activity-dependent plasticity directly affecting neuronal signaling, learning, and memory processes. Tpm4.2 knock-out neurons exhibit longer, thinner dendritic spines when compared with Tpm4.2^+/+ neurons. Thin spines are reported to form more transient and weaker synapses [35], and are associated with stress [36], as are longer spines [37]. These longer, thinner spines may account for the weakened synaptic activity observed in the dissociated primary hippocampal Tpm4.2^−/− neurons. Long, thin spines are immature spines which then transform into mushroom or stubby spines following LTP or LTD respectively [38, 39]. Here, we show that subtle alterations in normal, healthy dendritic spine dynamics of single cells can have substantial effects on neuronal signaling and learning and associative memory processes such as those we observe in Tpm4.2^−/− neurons. These neuronal changes, however, may be masked or mitigated by other processes in more complex brain tissues, including the interaction of glial cells and, in particular, the role of astrocytes in regulating neuronal activity and plasticity.

Neuronal hyperexcitability was observed in Tpm4.2^−/− neurons, which likely arose due to the increased dendritic complexity and alterations in excitatory surface receptors. These changes, in addition to alterations in spine morphology, can result in more aberrant neuronal connections within the hippocampus and neighboring regions [40], depicted in the current study with increased neuronal connectivity in Tpm4.2^−/− neurons. Hyperexcitable neurons and increased neuronal connections are both implicated in producing anxiety-like behaviors [41]. Here, we observed a heightened anxiety phenotype demonstrated in Tpm4.2^−/− mice, particularly in female mice. As increasing evidence is implicating hippocampal hyperexcitability with anxiety-like behaviors [41], it is important to further investigate Tpm4.2 expression and regulation in the context of models of anxiety, which could uncover some potential mechanisms underlying anxiety-like behaviors. This is of particular clinical relevance, as previous studies silencing other actin cytoskeleton-associated proteins, including protein kinase A [42] and SAP90/PSD95-associated protein [43], have demonstrated an anxiety-like phenotype in mice. As the Tpm4.2 was deleted throughout the entire brain in our model, it is important to consider the involvement of other brain regions in the production of the observed anxiety-like phenotype, specifically other limbic regions such as the amygdala and nucleus accumbens [44]. Overall, there was no overt genotype-dependent impact on social or novel object recognition in Tpm4.2^−/− mice. The anxiety phenotype in Tpm4.2^−/− mice that predominantly affects females suggests that further studies should more closely investigate sex differences in not only Tpm4.2 but also other actin cytoskeleton proteins and their functions.

Notably, knock-out of Tpm4 causes a rare form of macrothrombocytopenia in humans and mice [6]. However, in the current study, circulating platelet levels are unlikely to impact the brain and the resulting behavioral phenotype. Although we have not observed any compensation for the lack of Tpm4.2 by other Tpm isoforms in brain tissue, we cannot exclude the possibility that other neuronally expressed Tpms could contribute to the observed phenotypes by functional compensation.

Conclusion

In conclusion, our study sheds light on the role of Tpm4 in neuronal development, signaling, and maintenance, highlighting its important regulation in the healthy brain. The observations regarding heightened neuronal firing and neuronal networks producing anxiety-like phenotypes could implicate the dysregulation of Tpm4.2 in anxiety disorders. Future research is required to further dissect the molecular interactions of Tpm4.2 that drive the observed functional changes in early developing and mature neurons. Further, the observation of impaired recognition memory warrants further investigation of a role for Tpm4.2 in memory disorders such as dementia.

Materials and Methods

Generation of Tpm4.2^−/− Mice

Tpm4.2^−/− mice were designed as a homozygous knock-out of exon 1b of Tpm4.2 using the C57BL/6 J mouse strain as described in [6]. Mice were genotyped by using isopropanol-precipitated DNA from tail biopsies as a template for polymerase chain reaction (PCR), using the following primers Tpm4.2 knock-out forward primer: 5′-GTGACCTCATGGGCCTGAC-3′ and Tpm4.2 knock-out reverse primer: 5′-GGACGAAAAGTGGGATCG-3′. Deletion of Tpm4.2 protein was further confirmed by Western blotting in previous studies [6] and in Fig. S8. All procedures involving animals were approved by the UNSW and Macquarie University Animal Care and Ethics Committees and conducted in accordance with national and international guidelines.

Characterization of Tpm4.2^−/− Mice

Perfusion and Tissue Collection

To confirm the complete knock-out of Tpm4.2, Tpm4.2^−/− and Tpm4.2^+/+ control mice were euthanized and transcardially perfused with 1 × phosphate buffered saline (PBS) and their brains harvested. Brains were sagittally sectioned into two halves along the midline, and one-half was immediately snap frozen in liquid nitrogen and later stored at − 80 °C for downstream molecular studies.

Western Blot

Snap frozen half brain tissues were lysed with ice-cold radioimmunoprecipitation assay (RIPA) lysis and extraction buffer (20 mM Tris pH 8.0, 1% Nonidet P-40, 0.25% sodium deoxycholate, 0.1% sodium dodecyl sulfate, 150 mM sodium chloride, 5 mM ethylenediaminetetraacetic acid, 30 mM sodium fluoride, 60 mM β-glycerophosphate, 20 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM dithiothreitol, 1 × protease inhibitor, and Milli-Q water). In brief, tissues were thawed upon addition of RIPA buffer followed by sonication at 1 pulse/s with 20% amp in a probe sonicator for 30 s and then centrifuged at 14,000 rpm for 10 min at 4 °C. The supernatants were collected, and bicinchoninic acid assay was performed for protein quantification using Pierce BCA Protein Assay Kit (Thermo Scientific, Cat No. 23225). 12.5 µg or 10 µg of total protein from brain tissue lysates was loaded onto 12.5% SDS–polyacrylamide gels for brain region–specific expression and Tpm isoform compensation analysis, respectively. Electrophoresis was run at 80 V for 20 min through the 4% stacking gel, before 120 V for 2.5 h through the 12.5% resolving gel. Resolved proteins were then transferred onto methanol activated polyvinylidene difluoride membrane (Merck Millpore) using a Bio-Rad Trans-blot Turbo transfer system and chilled transfer buffer containing 10% Tris–Glycine SDS buffer (10 ×), 20% methanol in Milli-Q water. Membranes were then blocked with 5% skim milk in Tris-buffered saline with 0.1% Tween 20 (TBST) on the rocker for 2 h at room temperature followed by incubation with primary antibodies diluted in blocking buffer (rabbit polyclonal anti-tropomyosin 4.2 (Gift from Peter Gunning, 1:2000) for 3 h in 2.5% blocking buffer at room temperature; mouse anti-Tpm3.1/2 (2G10.2, 1:250 Gift from Peter Gunning); mouse anti-total Tpm3 (CG3, 1:250); and mouse anti-Tpm1.10/12 (clone 5–54, 1:250) in 5% blocking buffer and overnight incubation at 4 °C on rocker) [45]. After four washes with TBST (5 min each at room temperature), the blots were incubated with the respective secondary antibodies (1:5000 in their respective blocking buffer) for 1.5 h at room temperature on rocker before washing again four times with TBST (5 min each). Next, the blots were developed with Crescendo substrate (Millipore) and visualized using ChemiDoc imaging system (Bio-Rad). All blots were subsequently probed with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) housekeeping gene (Merck-Millipore; Cat No. MAB374) (1:5000 for Tpm4.2 blot and 1:1000 for all other blots) and imaged to obtain the corresponding loading controls. The western blots were quantified using ImageJ software (NIH). Uncropped Western blot images are provided in Figs. S8, S9 and S10.

All values are expressed as mean ± SEM (standard error of mean) and were subjected to unpaired t-test using GraphPad Prism (Version 8.3). Significance (if found) was indicated as p < 0.05.

Primary Culture of Mouse Hippocampal Neurons

Mouse primary hippocampal neurons were cultured from embryonal day 16.5 (E16.5) from Tpm4.2^+/+ and Tpm4.2^−/− mice with C57BL/6 J background as previously described [46]. In brief, brains were removed from E16.5 mouse embryos and placed in Hanks buffered salt solution (HBSS; Sigma). Meninges were removed and hippocampi extracted using microclippers. A 1:100 dilution of Trypsin (Sigma-Aldrich) was then added to hippocampi and incubated at 37 °C for 20 min. Deoxyribonuclease I 1:100 final concentration of 0.1 mg/mL (Sigma) was added for 30 s before washing twice with 10 mL of DMEM (Life Technologies, Berkley, CA, USA) with 10% fetal bovine serum (FBS, Thermo Fisher Scientific) to remove DNAse I. Hippocampi were then dissociated by trituration with 1 mL DMEM/10% FBS using fire-polished, serum-coated Pasteur pipettes. Cells for receptor recycling assay and calcium imaging of live neurons were then plated on poly-D-lysine (PDL; Sigma)-coated 1.5-mm glass coverslips at a density of 70,000 cells per well in a 24-well plate and incubated at 37 °C and 5% CO₂ for 2 h in DMEM/10% FBS. Media were then changed to Neurobasal medium (NBM; Neurobasal, Life Technologies; supplemented with 2% B27, Life Technologies, and 0.25% GlutaMAX, Invitrogen).

Receptor Internalization Assay

Neuron Treatment

Primary hippocampal neurons were seeded at 70,000 per well on 1.5 mm, PDL-coated glass coverslips and incubated in NBM at 37 °C with 5% CO₂ for 18 days. An assay to induce receptor internalization was performed as previously described [47] with the following modifications—80% of culture media was removed and neurons either left untreated, incubated with extracellular solution (ECS; 150 mM NaCl, 2 mM CaCl₂, 5 mM KCL, 10 mM HEPES (pH 7.4), and 30 mM glucose in dH₂O), 25 μM NMDA in ECS, 100 M glycine in ECS, or 25 M bicuculline in ECS for 30 min at 37 °C with 5% CO₂. Incubation solutions were then aspirated and replaced with original cell culture medium and re-incubated for 60 min before fixing for 5 min in 4% PFA.

Surface and Total GluA1 Receptor Probing

Half of the neurons were then probed for surface GluA1 receptors only by blocking for 1 h at RT in blocking buffer (2% FBS (Sigma) in PBS) followed by incubation for 60 min with mouse N-terminal anti-GluA1 (1:250 in BB; Merck-Millipore MAB2263) diluted in blocking buffer. Surface-probed neurons are then washed in PBS and fixed with 4% PFA for 5 min, permeabilized with 0.1% Triton-X in PBS for 5 min at RT with PBS wash steps in between each. The remaining half of the neurons are probed for total GluA1 receptors by permeabilizing with 0.1% Triton-X in PBS for 5 min at RT, washing in PBS, blocking for 60 min in BB before incubation with 1:150 mouse anti-GluA1 at RT for 60 min. All neurons are then washed, blocked for 60 min in blocking solution, and incubated with Chicken anti-MAP2 (1:500 in blocking buffer; Abcam ab5392) for 4 °C overnight. All neurons are then washed and incubated for 60 min with donkey anti-mouse AlexaFluor-555 (1:500 Life Technologies in BB) for GluA1, goat anti-chicken AlexaFluor-647 (1:500; Life Technologies in BB) for MAP2. Secondary antibodies are then washed off in PBS before incubation with phalloidin 488 (1:100 in PBS, Thermo Fisher Scientific) and DAPI (1:1000, Life Technologies) for 20 min at RT. Neurons are then washed in PBS, dipped in H₂O, and mounted onto glass slides with Fluoromount G (Thermo Fisher Scientific).

Imaging and Analysis Receptor Internalization

Slides were imaged on a Zeiss AxioImager, using a 63 × oil objective. Neurons were stitched, exported as uncompressed.Tif files, and analyzed on FIJI (ImageJ). Relative intensity was measured for three dendrites per neuron and ten neurons per condition, maintaining a consistent distance away from the soma (5 M) and measurement area (15 µM²) and normalized to background intensity. Ratios of surface to total GluA1 intensity were measured, and the ratio was compared between groups using GraphPad Prism software (version 10.2).

Plasmid Production and Cloning

For calcium imaging of live neurons, we used a the green fluorescent protein (GFP)–based GCaMP sensor protein with fast kinetics jGCaMP7f as previously characterized [48] and inserted it into an AAV vector containing a PhP.B capsid, human synapsin promoter, and woodchuck hepatitis virus post-transcriptional regulatory element (WPRE), flanked by AAV inverted terminal repeats (ITRs). NEBuilder Hifi DNA Assembly Master Mix was used to clone plasmids (E2621L, New England Biolabs, MA, USA). DNA fragments were amplified by PCR, and 0.06 pmol was combined with 0.06 pmol of digested vector fragments, 5 µL of water, and 5 µL of NEBuilder DNA Assembly Master Mix. The mixture was incubated for 1 h at 50 °C before 5 µL was added into 50 µL of One Shot Stbl3 Chemically Competent E. coli (C737303, Thermo Fisher Scientific, MA, USA) and incubated on ice for 30 min, heat-shocked at 42 °C for 45 s, and then placed back on ice for a further 2 min. Transformed bacteria were then mixed with 200 µL of SOC Outgrowth Medium (B9035, New England Biolabs, MA, USA), incubated with 300-rpm shaking for 1 h at 37 °C, and plated on agar plates with 1:1000 ampicillin. Plates were then incubated at 37 °C overnight.

For each construct, a pipette tip was used to pick five distinct colonies from each agar plate and added to LB broth (0.5% yeast [Sigma-Aldrich, MO, USA], 1% tryptone [G-Biosciences, New Delhi, India] and 1% NaCl [Sigma-Aldrich, MO, USA]). DNA was then extracted from bacterial culture using Wizard Plus SV Miniprep Purification System (Promega, WI, USA), as per manufacturer’s protocol. In brief, the culture was pelleted for 5 min, before resuspension in 250 µL of cell resuspension solution. Ten microliters of alkaline protease solution was then added and the mixture incubated for 5 min at room temperature before neutralization with 350 µL of neutralization solution. The mixture was centrifuged (10 min at room temperature, 21,000 g) and the supernatant transferred to the spin column placed in a collection tube and centrifuged (21,000 g for 1 min at room temperature). Flowthrough was discarded, and the bound DNA was washed with 750 µL of wash solution spun (centrifuged for 1 min at room temperature, 21,000 g). The wash step was repeated with 250 µL of wash solution. The spin column was transferred to a sterile 1.5-mL microcentrifuge tube, and the DNA was eluted by incubating the spin column membrane with 50 µL of nuclease-free water for 2 min at room temperature, followed by centrifuging for 1 min at room temperature.

PureLink HiPure Plasmid Maxiprep Kit (Thermo Fisher Scientific, MA, USA) was used to obtain larger DNA amounts. Bacteria were grown in 4 mL of LB Broth (1:1000 ampicillin) for 8 h, before expansion to 250 mL LB Broth (1:1000 ampicillin) overnight. Overnight culture was spun at room temperature for 10 min at 4000 g and the pellet resuspended in 10 mL of resuspension buffer with RNase A. Ten milliliters of lysis buffer was then added to the cells, mixed by inversion, and incubated for 5 min at room temperature. Ten milliliters of precipitation buffer was then added, and the mixture centrifuged at 12,000 g for 30 min at room temperature. Supernatant was loaded onto the previously equilibrated column and drained by gravity flow. The column was washed with 60 mL of wash buffer and drained by gravity flow. Fifteen milliliters of elution buffer was then added to the column to elute DNA using gravity flow. The DNA was mixed well with 10.5 mL of isopropanol and centrifuged at 12,000 g for 30 min at 4 °C. Supernatant was removed, and the pellet resuspended in 5 mL of 70% ethanol, followed by centrifugation at 12,000 g for 10 min at 4 °C. Supernatant was discarded, and the pellet was left to air-dry for 10 min. The DNA was resuspended in 300 µL of nuclease-free water and used for adeno-associated virus production.

Production of Adeno-Associated Virus for Neuron Transduction

GCamp7f-eGFP, Tpm4.2-mRuby2, and Tpm4.2-IRES-mRuby2 were packaged into PHP.B capsids, and adeno-associated viruses were produced as previously described [47]. In brief, HEK293T cells were grown to 70–80% confluency in DMEM/10% FBS before replacement with Iscove-modified Dulbecco medium (Sigma) + 5% FBS 3 h prior to transfection with polyethyleneimine-Max (PEI-max, Polysciences) containing pF0delta06 and AAV-PHP.B plasmid with rep and cap sequences.

Calcium Imaging of Live Neurons

Primary hippocampal neurons for calcium imaging of live neurons were seeded at 70,000 on 1.5-mm PDL-coated glass coverslips. Neurons were transduced with 1 × 10¹² vg/coverslip of AAV-PHP.B jGCamp7f and incubated at 37 °C with 5% CO₂ for 20 DIV. At each timepoint, cells were imaged at 37 °C with 5% CO₂ on an Axio Observer 7 Live cell imager (Carl Zeiss, Germany) at 5 × magnification for 5 min with 500-ms intervals.

A somatic region of interest (ROI) was drawn for each active neuron in FIJI [49], and the fluorescent time series for each ROI within the field of view (FOV) was saved as a table. Time series were analyzed using custom MATLAB (version R2024b, MathWorks) routines adapted from previously published in vitro calcium image analysis pipelines [50, 51]. Normalized fluorescence intensity (D_F/F) was extracted using the getDFF function in CaPTure [51] with tau [2 50]. Peaks and their locations were identified using the MATLAB findpeaks function (“MinPeakHeight” = 0.01 D_F/F, “MinPeakDistance” = 4 timepoints). To quantify the synchronous activity, the normalized fluorescence intensity for each ROI within a FOV was averaged and the peaks of the average intensity timeseries were extracted as described above. For each cell, we quantified the average calcium event amplitude and event frequency. These values were then aggregated by FOV. To account for the hierarchical structure of the data (multiple cells nested within each FOV), we performed a nested t-test using GraphPad Prism (GraphPad Software, La Jolla, USA), treating FOVs as the unit of replication and cells as nested observations. This approach allowed us to assess group-level differences in calcium responses while accounting for the non-independence of cells within the same FOV.

Electrophysiology

Whole Cell Patch Clamping of Primary Neurons in Dissociated Cultures

Primary hippocampal neurons were seeded at 70,000/well in a 24-well plate onto 1.5-mm PDL-coated coverslips and incubated in NBM at 37 °C and 5% CO₂. At 17 and 18 DIV, coverslips were transferred to the recording bath on a Leica DM IL inverted microscope, which was perfused with extracellular solution (110 mM NaCl, 10 mM HEPES, 10 mM glucose, 2 mM CaCl₂, 0.8 mM MgCl2, 5 mM KCl) at room temperature using a peristaltic pump (Minipuls 3, Gilson, France). After a cell was patched, the perfusion was switched to a separate 20-mL aliquot of 0.5 µM TTX and 100 µM picrotoxin. Patch electrodes were made from glass capillaries 1.2 mm OD, 0.94 ID, 100 length (Harvard Apparatus), pulled using a Narishige Model PC-10 microelectrode puller to a tip resistance of 3–5 MΩ. Electrodes were filled with internal solution (110 mM cesium methane sulfonate, 8 mM NaCl, 10 mM HEPES, 2 mM Mg₂ATP, 0.3 mM Na₃GTP, 0.1 mM spermine tetrahydrochloride, 7 mM phosphocreatine, 10 mM EGTA, 5 mM CsCl) with 50 µM Alexa Fluor 594, filtered through a 0.22-µm syringe-driven filter unit (Millex). Recordings of mEPSCs were made at a holding potential of − 70 mV with an Axopatch 200B amplifier, filtered at 2 kHz, digitized at 5 kHz with a Digidata 1440 A, and saved with Clampex 10.2 (Molecular Devices, USA).

mEPSC Analysis in Primary Neurons of Dissociated Cultures

mEPSCs were detected and measured using Axograph (Sydney, Australia). A notch filter (49.9–50.1 Hz) was applied, and an event template (maximum 0.5-ms rise time, 3-ms minimum decay time) was used to detect mEPSC [52]. Events outside of 5–150 pA were excluded, and detected events were manually verified. Event amplitude and inter-event interval were measured. Either the first 1000 events or 5 min of activity were analyzed from each cell, as there was a large variability in activity frequency.

Changes of Field Potential Recording in Brain Slices

All animal studies were carried out in accordance with the New South Wales Animal Research Act and Regulation and approved by the Animal Ethics Committee of UNSW Sydney under ethics protocol numbers 14/113A and 18/37B. Mice were housed in a temperature-controlled facility (22–24 °C) on a 12-h light–dark cycle. For all experiments, mice of both sexes were used at 6–8 weeks old. Data collection was performed blind to genotype and interspersed so no more than two animals of the same genotype were used on consecutive days.

Mice were anesthetized by open-drop exposure to isoflurane in an induction chamber before decapitation, and the brain was removed and placed in ice-cold modified artificial cerebrospinal fluid (ACSF; 124 mM sucrose, 62.6 mM NaCl, 2.5 mM KCl, 26 mM NaHCO₃, 1.2 mM NaH₂PO₄, 10 mM glucose, 0.5 mM CaCl2, and 3.3 mM MgCl₂). The brain was hemisected and immersed in cold modified ACSF. Horizontal slices (400 µm) were cut using a vibratome (model VT1200, Leica, Wetzlar, Germany) at room temperature. The solution containing the brain slices was continuously infused with 95% O₂/5% CO₂. Slices were left to recover for at least 1 h before recording. Slices were used within 7 h of cutting, corresponding to the optimal period of slice health [53].

Slices were transferred individually to the tissue recording system (Kerr Scientific Instruments, Christchurch, New Zealand) and continuously perfused with standard ACSF at room temperature. A bipolar, Teflon-coated tungsten stimulating electrode (Kerr Scientific Instruments) was placed in the stratum radiatum, aligned to the end of the dentate gyrus, and the recording electrode was placed approximately 800 µm from the stimulating electrode. Stimuli were delivered via an isolated stimulator (model DS2, Digitimer, Hertfordshire, England, or A-M Systems, Model 2200, WA, USA), triggered through a Powerlab (model 4/2ST, AD Instruments, Sydney, Australia) or multifunctional data acquisition card (National Instruments NI PCI-6221, USA or Data Translation DT9816; USA). Field potentials were amplified at 100 × using a KSI Tissue Recording System Amplifier (Kerr Scientific Instruments, Christchurch, New Zealand) and digitized with the Powerlab or multifunctional data acquisition card. Traces were acquired using Scope (AD Instruments, Sydney, Australia), AxoGraph X (Axograph Scientific) or custom software. To determine the optimal electrode positions, stimuli (15–20 V 100 µs) were delivered every 8–10 s, and electrodes were lowered until an extracellular field excitatory synaptic potential (fEPSP) was observed. The depth of the recording and stimulating electrodes was then adjusted to produce the largest response. After finding the optimal electrode position, the stimulus frequency was reduced to once every 30 s and left to stabilize for 10 min. A stimulus response curve was then conducted by varying the stimulus intensity from 5 to 70 V. Slices were discarded if the maximum fEPSP amplitude was below 0.6 mV, as the small response was considered to be a marker of poor health. The stimulus intensity eliciting 50% of the maximum amplitude was identified, and field potentials were evoked in pairs with a 50-ms interval at this stimulus intensity at 30-s intervals. After obtaining a stable baseline for 20–30 min, LTP was induced with two bursts of high-frequency stimulation (100 Hz 1 s). For LTD experiments, LTD was induced with 900 paired pulses (50-ms interval) at 1 Hz. Responses were then recorded for the following 60 min.

Electrophysiological data were analyzed offline using AxoGraph (Sydney, Australia). Measures included the fiber volley amplitude, the fEPSP slope, and the paired pulse ratio. Fiber volley amplitude was defined as the amplitude of the negative peak preceding the fEPSP. The fiber volley amplitude is indicative of the number of presynaptic axons activated by stimulation. The fEPSP slope was defined as the maximum slope during the initial 2.5 ms after the fiber volley. This was used to measure excitatory activity. Slope was measured instead of amplitude, as the amplitude can be contaminated by the population spike at higher stimulus amplitudes. The paired pulse ratio was calculated as the second fEPSP slope divided by the first, to give a measure of presynaptic transmitter release probability [54].

Statistical Analysis for Electrophysiology

Statistical comparisons were performed using GraphPad Prism (GraphPad Software, La Jolla, USA). The D’Agostino and Pearson test was used to test data sets for normality. Differences between groups were tested using repeated measures ANOVA, paired t-test, unpaired t-test, Mann–Whitney test, or Kolmogorov–Smirnov test as indicated with α = 0.05. Data are presented as mean ± standard error of the mean. Data collection and analysis were performed blind to the genotype of the animals.

Neurite Outgrowth

Neuron Plating and Transfection for Neurite Outgrowth Analysis

For neurite growth experiments, primary hippocampal Tpm4.2^+/+ and Tpm4.2^−/− neurons were seeded at 70,000 cells/well in a 24-well plate containing PDL-coated 12-mm glass coverslips and cultured in 1 mL per well in complete NBM at 35 °C and 5% CO₂.

For neurite tracing experiments, pEGFP-C1 plasmid (Clontech) was used for control neurons. Tpm4.2^+/+ and Tpm4.2^−/− neurons were transfected with pEGFP-C1 at 2 DIV with Lipofectamine 3000 (Thermo Fisher Scientific, MA, USA) by making a transfection mixture containing 0.5 µg of DNA and 1 µL of Lipofectamine 3000 in a total of 100-µL NBM per well of a 24-well plate. The transfection mixture was incubated with the cells for 90 min at 37 °C and 5% CO₂ in 50% of the initial culture media volume (while the other 50% was collected before transfection procedure and kept at 37 °C). After incubation, the media was aspirated, and conditioned media were added to the cells.

Immunocytochemistry for Neurite Outgrowth Analysis

Two days after transfection at 4 DIV, neurons were fixed with 4% paraformaldehyde (PFA) at room temperature for 15 min, washed in PBS, and permeabilized in 0.1% Triton-X (Sigma-Aldrich) for 5 min at room temperature. Neurons were then washed and blocked in BB (2% FBS in PBS) at room temperature for 1 h before incubation in the following primary antibodies diluted in BB-mouse anti-Tau1 (1:500 Millipore ab3420) and chicken anti-B3-tubulin (1:250, Millipore ab9354) overnight at 4°. The following day, primary antibody was washed off in PBS, and secondary antibodies donkey anti-mouse Alexa-555 and goat anti-chicken Alexa-647 (both 1:500, Life technologies) were diluted in BB and placed on neurons for 1 h at room temperature. Neurons were washed in PBS and mounted on glass microscope slides with Prolong Gold antifade reagent with DAPI (Life technologies).

Imaging of Neurite Outgrowth

Tpm4.2^+/+ and Tpm4.2^−/− mouse primary hippocampal neurons transfected with pEGFP-C1 were imaged using an Axio Imager upright fluorescent microscope fitted with a monochrome camera, with EC-Planachromatic Neofluar 40 × magnification objective (NA 0.75, WD 0.71 mm, Air immersion). Fluorescent illumination was obtained with a Xenon HXP lamp. The fluorescent filter sets used were BP450-490/BS495/BP500-550 (FS#38) for 488-nm channel, BP533-558/BS570/BP570-640 (FS#43) for 555-nm channel, and BP625-655/BS660/BP665-715 (FS#50) for 647-nm channel. At least 20 transfected neurons were imaged per single coverslip, and four coverslips per single experimental group were imaged per one biological replicate (80 imaged neurons per biological replicate in total—3 biological replicates). Images are representative of all three biological replicates.

Morphological and Statistical Analysis for Dendritic Spines

The axonal compartment of Tpm4.2^+/+ and Tpm4.2^−/− neurons, transfected with pEGFP-C1 plasmid, was identified as tau1-positive/β3-tubulin-positive. The dendritic compartment was verified as tau1-negative/β3-tubulin-positive. For each experimental group, ~ 20 neurons (~ 5 neurons per coverslip) from 80 imaged neurons per biological replicate were chosen, using a random number generator (RNG). The images were processed in ImageJ (v.2.1.0). The morphological analysis of neurons was performed using the semiautomated approach in Neurolucida software (MBF Bioscience, VT, USA, v2019.1.1) to outline soma, axons, and dendrites. AutoNeuron workflow was used to initiate automatic tracing, followed by manual corrections, labeling, and branch ordering. To quantify traced axonal and dendritic compartments in Neurolucida, branched structure analysis and centrifugal Sholl analysis options in the Neurolucida Explorer package were used. Statistical analysis was performed in GraphPad Prism software (version 9.1.2). To determine if experimental groups have a normal Gaussian distribution, we performed Anderson–Darling, D’Agostino-Pearson, Shapiro–Wilk, and Kolmogorov–Smirnov normality tests. After these tests showed no Gaussian distribution within the experimental groups, the significance was determined with a non-parametric Mann–Whitney U test for Tpm4.2^+/+ and Tpm4.2^−/− neurons transfected with pEGFP-C1.

DiI Injection for Dendritic Spine Analysis

Cells were fixed in 4% PFA for 15 min at room temperature and washed with PBS. The fixed coverslip was placed in a bath of PBS on a Leica DMIL microscope. A 1% solution of 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI) in ethanol was loaded into a sharp microelectrode. The tip of the microelectrode was maneuvered into the target cell soma and left for 5 min to allow the dye to enter the cell. Sometimes, pressure was added to expel the DiI solution. Cells were randomly selected for injection.

Approximately ten cells per coverslip were injected. The coverslips were then left in 1 × PBS at 4 °C overnight, or at room temperature for 5 h to allow the DiI to diffuse into neuronal processes. The coverslips were then mounted with either Fluoromount or Prolong Gold and then sealed with nail polish the next morning before imaging.

Imaging of Dendritic Spine Analysis

Immunofluorescence images were taken using an LSM 710 confocal microscope (Zeiss) with a 63 × 1.4 NA oil immersion objective. DiI was excited with a 561-nm DPSS laser and emission captured at 519–673 nm. For dendritic spine morphology analysis, two 20-µm-length segments of secondary dendrites, 50–75 µm away from the cell soma, were selected from each transfected neuron. A Z-stack image with a 0.21-µm interval and 0.04 × 0.04 µm pixel size was acquired of each selected dendrite.

Deconvolution was performed on images using the DeconvolutionLab2 plugin in ImageJ [55]. Images were processed with five iterations of the Richardson-Lucy algorithm using a point spread function generated by Diffraction PSF 3D plugin (Optinav).

Spine Quantification

To investigate whether Tpm4.2 deletion affects spine morphology, primary neuronal cultures were prepared from Tpm4.2^+/+ and Tpm4.2^−/− mice, fixed at 17–18 DIV, stained with DiI to assist labeling of entire cell membranes, and imaged. Fifty-one cells were analyzed for the Tpm4.2^+/+ group, and 57 cells were analyzed for the Tpm4.2^−/− group, from 3 separate culture preparations.

Spine morphology was analyzed according to Risher et al. [56]. Z-stack images were imported to RECONSTRUCT [57]. The length of the dendrite and the length and widths of spines were manually traced and measured. Measurements were copied to an Excel spreadsheet to calculate the spine density, average protrusion width, and average protrusion length for each cell that was imaged. Data were analyzed, and graphs were generated in GraphPad Prism. Data are given as mean ± SEM, unless otherwise indicated.

Mouse Housing for Behavioral Analysis

Male and female Tpm4.2^−/− (n = 20) and Tpm^+/+ mice (n = 25) littermates with C57BL/6 J background were transported at least 2 weeks prior to testing for habituation from Macquarie University and housed on arrival in the animal facility at the School of Medicine, Western Sydney University (Campbelltown Campus, Australia). Mice were housed in littermate groups separated for sex but not standardized for genotype to avoid the need to individually house test mice. Adult A/JArc mice were used in the social preference test to trigger explorative behavior of test mice. All animals were group housed (2–3/cage) in individually ventilated cages (Type Mouse Version 1; Airlaw, Smithfield, Australia; air change: 90–120 times per hour averaged; passive exhaust ventilation system), containing corn cob bedding, a mouse igloo (Bioserv, Frenchtown, USA), and a crinkle nest to provide nest building opportunities (Crink-l’Nest, Tecniplast, Australia). Mice were kept under 12:12 h (light phase: 0900–2100 h with white light at an illumination of 124 lx; dark phase: 2100–0900 h with red light at an illumination of less than 2 lx) and food and water ad libitum, with a temperature between 22 and 24 °C and humidity between 40 and 60 RH. Mice were 7 months old at the start of behavioral experiments. Research and animal care procedures were approved by the Western Sydney University Animal Care and Ethics Committee (approval number: A12918) and were in accordance with the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes.

Behavioral Phenotyping

For habituation purposes, all test animals were transported to the testing room 30 min prior to behavioral testing, and all experiments were performed within the first 6 h of the light phase, with an inter-test interval of at least 48 h. All test equipment was cleaned after each trial with 80% ethanol solution. Tests were carried out in the following order: open field, novel object recognition, social preference test, elevated plus maze, prepulse inhibition, and fear conditioning. The sample size for each experimental test condition was n = 10–12.

Open field (OF)

Locomotor activity, exploration, and anxiety-like behaviors were measured in the OF and were conducted as previously published from our laboratory [58, 59]. Mice were placed into the right corner of the OF chamber (43 × 43 cm; Activity Monitor, Med Associates Inc., Fairfax, USA) and allowed to explore the arena freely for 30 min. The arena was divided into a central and peripheral zone (MED software coordinates 3/3, 3/13, 13/3/, 13/13) with the central zone being a more aversive, anxiety-inducing zone of the open field [60]. Software settings for the detection of locomotion were box size, 3; ambulatory trigger, 2; resting delay, 1000 ms; and resolution, 100 ms. Time, horizontal (distance travelled), and vertical activity (rearing) in central and peripheral zones were measured by the chambers’ infrared photo beams. The ratio of central to total distance travelled (distance ratio) and time (time ratio) spent in the central area of the OF were taken as measures of anxiety.

Novel Object Recognition Test (NORT)

The distinction between familiar and unfamiliar objects is an index of recognition memory and is measured by the innate preference of rodents for novel over familiar objects [61]. The protocol was adapted from a previous publication from our group [58]. The apparatus for NORT was a gray Perspex arena (35 × 35 × 30 cm), and the protocol ran over two consecutive days. On the first day, mice were habituated twice to the test chamber for 10 min and allowed to freely explore the empty arena, with an intertrial interval of 2 h. On the second day, mice were placed in the NORT arena with two identical objects (2 DUPLO® elephants or 2 blocks of DUPLO®) and allowed to explore the objects for 10 min. After a 15-min intertrial interval, mice were replaced in the arena with one familiar object and one novel object (LEGO® elephant + block of LEGO®) for 10 min. Object exploration was scored when mice exhibited nosing behavior towards the objects (i.e., when the mouse directed its snout towards an object at a distance of < 1 cm). Object recognition was reported as INDEX (time spent nosing the novel object expressed as a percentage of the time spent nosing the novel + familiar objects); the behavior was manually scored using ANY-maze™.

Social Preference Test (SPT)

The SPT test was used to assess sociability and social novelty preference (i.e., social recognition memory) in test mice and was conducted as per a previous publication from our group [58]. The apparatus consists of three chambers, i.e., a central chamber (around 9 cm × 18 cm × 20 cm) and two outer chambers (around 16 cm × 18 cm × 20 cm). The dividing walls were made of clear Plexiglas, with square passages, 4 cm high and 4 cm wide. One circular cage (i.e., mouse enclosure) was placed into each outer chamber. The mouse enclosures were 15 cm in height with a diameter of 7 cm and bars spaced 0.5 cm apart to allow nose contact between mice but prevent fighting. The chambers and enclosures were cleaned in-between trials (intertrial interval of 2 min), and fresh bedding was added prior to each mouse. During the habituation trial, mice were placed individually in the central chamber and allowed to freely explore the apparatus and the two empty enclosures for 5 min. For the sociability test, an unfamiliar adult age-matched A/J mouse was placed in one of the two enclosures, while the experimental mouse was enclosed in the central chamber. Then, the test mouse was allowed to explore all three chambers for 10 min. Finally, test animals were observed in a 10-min social recognition test. For this, a second, unfamiliar A/J mouse was placed in the previously empty chamber so that the test mouse had the choice to explore either the familiar A/J mouse (from the previous trial) or the novel, unfamiliar mouse. AnyMazeTM tracking software was used to determine the time spent in the different chambers, number of entries, and distance travelled. In addition, time spent sniffing the opponent (i.e., A/J mouse) was recorded manually (i.e., snout of test mouse within the enclosure containing the opponent mouse or < 1 cm away from enclosure).

Elevated Plus Maze (EPM)

The EPM represents the natural conflict between the tendency of mice to explore a novel environment and the tendency to avoid a brightly lit open areas [62]. The behavior is also influenced by thigmotaxis and the fear of heights. The EPM was in the shape of a “ + ”, with the four arms extending from a central platform and raised 1 m above the floor. Two alternate arms were dark and enclosed (30.5 cm × 6.5 cm, sidewall height 18.5 cm) while two alternate arms were open (30.5 cm × 6.5 cm, no sidewalls, illumination 40 lx) with a central platform connecting the arms (6 cm × 6 cm). The mouse was placed onto the center field of the EPM (faced to a closed arm) and was allowed to freely explore the maze for 5 min. AnyMazeTM tracking software was used to determine the time spent in open/closed arms, the number of entries, and the distance travelled by the test mice. Anxiety was measured by the time spent on open arms as well as the ratio of open arm entries and distance (compared to total number of entries/total distance travelled). These parameters are inversely related to anxiety. The number of total arm entries/total distance travelled reflects general motor activity (locomotion).

Fear Conditioning (FC)

Fear conditioning (FC) is a type of associative learning task in which mice learn to associate a particular neutral conditional stimulus (CS) with an aversive unconditional stimulus (US) and show a conditional response [63]. The FC protocol used here evaluates both cued and contextual conditioning to assess amygdala-dependent and hippocampal-dependent fear-associated memory respectively. On day 1 (training), animals were placed into the test chamber (fear conditioning chambers from MED Associates Inc.) with vanilla scent cue presented in the chamber and lights on. For the first 2 min, mice were left undisturbed and could explore and habituate to the environment. After 2 min, the conditioned stimulus (CS: 30-s duration, 80-dB tone stimulus) was paired with a co-terminating unconditioned stimulus (US: electric foot shock of 0.4 mA for 2-s duration) twice with an inter-pairing interval of 120 s. The test mouse was returned to its home cage 120 s after the second CS–US pairing. On day 2 (context test), the mouse was returned to the testing chamber with the scent cue and light present, but no sound, for a total of 7 min. On day 3 (cue test), mice were placed in an altered context (e.g., no scent cue present, and a black wall in triangle was placed into the chamber, but lights are still on). Following the first 120 s, during which no auditory stimulus was presented (pre-CS), the CS was then presented continuously for 5 min. The experiment was then terminated after another 120 s without CS. In all trials, the percentage freezing response (the absence of all but respiratory movement) per 1-min block was automatically measured using SOF-843 video freeze (MED Associates Inc.), with a freezing threshold of 10. Total averages were calculated for each day, as well as the average freezing per minute in the cue test, during baseline testing in the first 2 min versus cue freezing in minutes 3 to 7.

Prepulse Inhibition (PPI)

Prepulse inhibition (PPI), an operational measure of sensorimotor gating, is the attenuation of the startle response by a non-startling stimulus (prepulse) presented 30–500 ms before the startling stimulus (startle pulse). Startle reactivity can be measured using SR-LAB startle chambers (San Diego Instruments, San Diego, USA) where the startle response intensity of rodents (whole body flinch amplitude) can be measured using a piezoelectric accelerometer. As previously described, animals were habituated to the startle chambers and the test enclosures twice a day for 10 min on two consecutive days (with an intertrial interval of 2 h). On the third day, the PPI test was carried out and consisted of a 5-min acclimatization period to a 70-dB background noise, followed by 97 trials presented in a pseudorandom order to test the acoustic startle response (ASR) and PPI, 5 × 70 dB trials (background noise), 5 × 100 dB trials, 15 × 120 dB trials (for ASR), and 72 PPI trials comprising six sets of a prepulse of either 74, 82, or 86 dB presented 32, 64, 128, or 256 ms (variable interstimulus interval; ISI) prior to a startle pulse of 120 dB. The intertrial interval varied randomly between 10 and 20 s. Responses to each trial were calculated as the average mean amplitude detected by the accelerometer. The startle response was calculated as the mean amplitude across all 15 startle trials, and percentage PPI (% PPI) was calculated as [(mean startle response (120 dB) – PPI response)/mean startle response (120 dB)] × 100. For acoustic startle habituation, blocks of the acoustic response to 120-dB startle pulses presented at the beginning, in the middle, and at the end of the PPI protocol (i.e., averaged across five trials each) were used to determine the effect of “startle block” [14].

Statistical Analysis

Behavioral data were analyzed using GraphPad PRISM 9.4.0. Three-way repeated measures (RM) or two-way analysis of variance (ANOVA) were conducted for the within-subjects factors “time” (OF, FC), “chamber” (SPT), and “object” (NORT) and the between-subjects factors “sex” and “genotype.” If ANOVA interactions were detected, data were split for the corresponding factor and paired t-tests were used to follow these up and identify differences between individual groups. In addition, single-sample t-tests against chance levels (i.e., 50%) were used for the SPT and NORT exploration index to test for preference of mice for mouse/novel mouse /novel object. Data is presented as mean ± standard error of the mean (SEM), and F-values and degrees of freedom are presented for ANOVAs. Differences were regarded as statistically significant if p < 0.05; trends were mentioned where p = 0.05–0.06. Significant “genotype” effects are indicated by “#” (# p < 0.05, ## p < 0.01, and ### p < 0.001), “sex” effects by “ + ” (+ p < 0.05, + + p < 0.01, and + + + p < 0.001), “repeated measures” effects by “&” (& p < 0.001), and significant interactions by “^” (^ p < 0.05, ^^ p < 0.01, and ^^^ p < 0.001). Significant t-test results against chance level (i.e., 50%, NORT/SPT) and paired t-test results are indicated by “*” (*p < 0.05).

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOCX 3.64 MB)^{(3.6MB, docx)}

Acknowledgements

We acknowledge and thank the staff at the Central Animal Facility (CAF) at Macquarie University and the staff from the Laboratory Operations team in the Faculty of Medicine Health and Human Sciences at Macquarie University. We would also like to thank the Prof Yazi Ke (AAV core, Dementia Research Centre, Macquarie University) for AAV production.

Author Contributions

Conceptualization: TF, JP and TK; Methodology: TF, SG, CC, RRP, TT, HS, EP, DY and SS. Formal analysis and investigation: SG, CC, RRP, TT, SS, WG, JP and TF; Writing—original draft preparation: SG, CC and RRP; Writing—review and editing: SG, RRP, CC, JP and TF; Funding acquisition: TF, TK, JP, RRP, TT, SS and PG; Resources: TF, PWG, TK, JP and ECH; Supervision: TF, JP and TK. All authors read and approved the final manuscript.

Funding

Open Access funding enabled and organized by CAUL and its Member Institutions This work was supported by funding from the National Health and Medical Research Council (NHMRC) (grant# APP1083209 and grant# APP200660) and the Australian Research Council (ARC) (grant #DP180101473) to TF. RRP is supported by the Ainsworth Medical Research Innovation Fund (AMRIF) as well as Dementia Australia. TK is supported by a Drug Development Grant from FightMND, NSW Health, as well as the AMRIF. SS has been supported by an International Macquarie Research Excellence Scholarship (iMQRES) from Macquarie University.

Data Availability

Data is provided within the manuscript or supplementary information files.

Declarations

Ethics Approval

Approval was obtained from the animal ethics committee of Macquarie University. The procedures used in this study adhere to the tenets of the Declaration of Helsinki.

Competing interests

P.W.G. and E.C.H. are directors and shareholders of TroBio Therapeutics Pty Ltd., a company that is commercializing anti-tropomyosin drugs for the treatment of cancer. Their laboratories receive funding from TroBio to evaluate anti-tropomyosin drug candidates.

Footnotes

Publisher's Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Sian Genoud, Chanchanok Chaichim and Rossana Rosa Porto contributed equally to this work.

John Power and Thomas Fath contributed equally to this work.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary file1 (DOCX 3.64 MB)^{(3.6MB, docx)}

Data Availability Statement

Data is provided within the manuscript or supplementary information files.

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[CR34] 34.Fath T, Fischer RS, Dehmelt L, Halpain S, Fowler VM (2011) Tropomodulins are negative regulators of neurite outgrowth. Eur J Cell Biol 90(4):291–300. 10.1016/j.ejcb.2010.10.014 [DOI] [PMC free article] [PubMed] [Google Scholar]

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[CR39] 39.Scofield MD, Heinsbroek JA, Gipson CD, Kupchik YM, Spencer S, Smith AC, Roberts-Wolfe D, Kalivas PW (2016) The nucleus accumbens: mechanisms of addiction across drug classes reflect the importance of glutamate homeostasis. Pharmacol Rev 68(3):816–871. 10.1124/pr.116.012484 [DOI] [PMC free article] [PubMed] [Google Scholar]

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[CR41] 41.Ghasemi M, Navidhamidi M, Rezaei F, Azizikia A, Mehranfard N (2022) Anxiety and hippocampal neuronal activity: relationship and potential mechanisms. Cogn Affect Behav Neurosci 22(3):431–449. 10.3758/s13415-021-00973-y [DOI] [PubMed] [Google Scholar]

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PERMALINK

Knock-out of Tpm4.2/Actin Filaments Alters Neuronal Signaling, Neurite Outgrowth, and Behavioral Phenotypes in Mice

Sian Genoud

Chanchanok Chaichim

Rossana Rosa Porto

Tamara Tomanic

Holly Stefen

Esmeralda Paric

Soumalya Sarkar

Dasol Yoo

Wendi Gao

Edna C Hardeman

Peter W Gunning

Tim Karl

John Power

Thomas Fath

Abstract

Supplementary Information

Introduction

Results

Expression Profile of Tpms in Mouse Brain

Fig. 1.

GluA1 Receptor Recycling in Tpm4.2−/− Neurons

Fig. 2.

Neuronal Activity in Tpm4.2−/− Versus Tpm4.2+/+ Neurons

Calcium Spike Analysis

Fig. 3.

Electrophysiological Analysis

mEPSCs in Dissociated Cultures of Primary Tpm4.2+/+ and Tpm4.2−/− Neurons

Fig. 4.

Field Excitatory Post-synaptic Potentials (fPESPs) in Tpm4.2+/+ and Tpm4.2−/− Brain Slices. The Effect of Tpm4.2 Knock-out on Basal Synaptic Transmission

The Effect of Tpm4.2 Knock-out on LTP

Fig. 5.

The Effect of Tpm4.2 Knock-out on LTD

Neurite Complexity of Tpm4.2 Knock-out Neurons

Fig. 6.

Spine Morphology in Tpm4.2−/− Neurons

Fig. 7.

Mouse Behavioral Testing

Open Field (OF)

Fig. 8.

Elevated Plus Maze (EPM)

Social Preference Test (SPT)

Fig. 9.

Novel Object Recognition Test (NORT)

Fig. 10.

Contextual and Cue Fear Conditioning (CFC)

Prepulse Inhibition (PPI)

Discussion

Conclusion

Materials and Methods

Generation of Tpm4.2−/− Mice

Characterization of Tpm4.2−/− Mice

Perfusion and Tissue Collection

Western Blot

Primary Culture of Mouse Hippocampal Neurons

Receptor Internalization Assay

Neuron Treatment

Surface and Total GluA1 Receptor Probing

Imaging and Analysis Receptor Internalization

Plasmid Production and Cloning

Production of Adeno-Associated Virus for Neuron Transduction

Calcium Imaging of Live Neurons

Electrophysiology

Whole Cell Patch Clamping of Primary Neurons in Dissociated Cultures

mEPSC Analysis in Primary Neurons of Dissociated Cultures

Changes of Field Potential Recording in Brain Slices

Statistical Analysis for Electrophysiology

Neurite Outgrowth

Neuron Plating and Transfection for Neurite Outgrowth Analysis

Immunocytochemistry for Neurite Outgrowth Analysis

Imaging of Neurite Outgrowth

Morphological and Statistical Analysis for Dendritic Spines

DiI Injection for Dendritic Spine Analysis

Imaging of Dendritic Spine Analysis

Spine Quantification

Mouse Housing for Behavioral Analysis

Behavioral Phenotyping

Open field (OF)

Novel Object Recognition Test (NORT)

GluA1 Receptor Recycling in Tpm4.2^−/− Neurons

Neuronal Activity in Tpm4.2^−/− Versus Tpm4.2^+/+ Neurons

mEPSCs in Dissociated Cultures of Primary Tpm4.2^+/+ and Tpm4.2^−/− Neurons

Field Excitatory Post-synaptic Potentials (fPESPs) in Tpm4.2^+/+ and Tpm4.2^−/− Brain Slices. The Effect of Tpm4.2 Knock-out on Basal Synaptic Transmission

Spine Morphology in Tpm4.2^−/− Neurons

Generation of Tpm4.2^−/− Mice

Characterization of Tpm4.2^−/− Mice