Introduction
Serum response factor (SRF) is a transcription factor that is highly expressed in granular neurons in the dentate gyrus (DG) of the hippocampus (Etkin et al.
2006; Ramanan et al.
2005). It regulates the expression of many cytoskeletal genes and genes that are activated by neuronal stimulation (Benito et al.
2011; Etkin et al.
2006; Kalita et al.
2006; Kuzniewska et al.
2016; Losing et al.
2017; Miano et al.
2007; Parkitna et al.
2010; Ramanan et al.
2005; Treisman
1995). During brain development, SRF-regulated genes control the neuronal cell migration, neurite outgrowth and organization of the DG and the formation of mossy fiber circuitry (Alberti et al.
2005; Etkin et al.
2006; Knoll et al.
2006; Lu and Ramanan
2011; Scandaglia et al.
2015; Stritt and Knoll
2010). The early postnatal deletion of
Srf resulted in impairments in long-term potentiation and long-term depression and alterations of the formation of immediate memory of a novel context (Etkin et al.
2006; Ramanan et al.
2005). In contrast to early SRF knockout (KO), time-specific SRF deletion in adult excitatory neurons does not influence gross morphology of the hippocampus (Kuzniewska et al.
2016; Losing et al.
2017). Moreover, SRF deletion in the mature brain caused a very limited alteration of basal gene expression, restricted mostly to genes that encode the actin cytoskeleton (Kuzniewska et al.
2016; Losing et al.
2017; Parkitna et al.
2010). Nevertheless, these mice exhibit hyperactivity, a decrease in anxiety-like behavior, and epilepsy (Kuzniewska et al.
2016; Losing et al.
2017; Zimprich et al.
2017).
The DG is part of the hippocampal formation and mostly composed of granule cells (GCs) (Kempermann et al.
2004; Radic et al.
2017). The DG receives excitatory inputs from the entorhinal cortex, and GCs send excitatory outputs via mossy fibers to pyramidal cells in the Cornu Ammonis CA3 (Amaral et al.
2007). Alterations of hippocampal DG circuity have recently been implicated in the pathophysiological mechanisms of psychiatric disorders (Hagihara et al.
2013; Yu et al.
2014).
To investigate whether SRF controls the morphology and function of mature GCs, we employed a model of SRF depletion that was restricted to excitatory neurons (Kuzniewska et al.
2016). In the present study, we found that the ablation of SRF in adult GCs decreased β-actin levels and led to cofilin 1 inactivation through an increase in the phosphorylation at regulatory Ser3. The decrease in actin expression coincided with abnormal spine morphology and impairments in basal excitatory synaptic transmission in DG neurons. Moreover, the lack of SRF increased animal activity and impaired rodent-typical behaviors, such as digging, marble burying, and nesting. Altogether, our data indicate that SRF expressed in adult excitatory neurons plays a crucial role in maintaining actin expression and controlling the morphological and electrophysiological properties of granular neurons and innate animal behaviors.
Materials and methods
Animals
Conditional Srf KO mice (mutant mice;
Srff/fCaMKCreERT2) were generated as previously described (Kuzniewska et al.
2016). As a control, CreERT2-negative littermates were used (
Srff/f; control mice; wild type [WT]). The mice were bred on a C57BL/6J genetic background. Adult male and female
Srff/f and
Srff/fCaMKCreERT2 mice were intraperitoneally injected with 1 mg tamoxifen (TAM; cat. #T5648, Sigma) twice daily for 10 days, resulting in the translocation of Cre-recombinase to the nucleus. Experiments were performed at least 6 weeks after the TAM injection. The animals were bred in the Animal House of the Nencki Institute of Experimental Biology. The mice were housed in individual cages with a 12 h/12 h light/dark cycle, constant temperature (23–25 °C), and food and water available ad libitum. Both male and female mice were used for the experiments, except when marked otherwise. All of the experiments were performed by experimenters who were blind to mouse genotype. All of the work was conducted in accordance with the European Community Council Directive (86/609/EEC) and Animal Protection Act of Poland (Directive 2010/63/EU). The procedures were approved by the 1st Local Ethics Committee in Warsaw, Poland (permission no. 389/2012, 678/2015, and 144/2016).
RNA preparation and quantitative real-time polymerase chain reaction
Total RNA was isolated from the mouse hippocampus using the RNeasy Mini Kit (cat. #74104; Qiagen) as described by the manufacturer. DNA contamination was removed by digestion with DNase I (cat. #1023460, Qiagen). The RNA concentration was calculated from absorbance at 260 nm, and the 260/280 nm absorbance ratio determined the purity of RNA. RNA was reverse transcribed with SuperScript III or IV Reverse Transcriptase (cat. #18080-044, #18090050, Invitrogen) according to the manufacturer’s instructions. Quantitative real-time polymerase chain reaction (PCR) was performed using Fast TaqMan Master Mix (cat. #44456, Applied Biosystems) with an Applied Biosystems 7900HT Fast Real-Time PCR System using TaqMan probes (ThermoFisher). Fold changes in expression were determined using the ∆∆CT relative quantification method. The values were normalized to relative amounts of GAPDH.
Immunohistochemistry
The animals were perfused with 4% paraformaldehyde (PFA). Brains were then fixed in 4% PFA overnight at 4 °C and cut into 40-μm coronal slices. For SRF staining, brain slices were stained with anti-SRF antibody (1:500; cat. #sc-13029; Santa Cruz Biotechnology) overnight at 4 °C followed by avidin/biotin complex (cat. #PK-6100, Vector) and visualized using SIGMAFAST™ DAB (cat. #D0426, Sigma). For Nissl staining, the brain sections were stained with 0.1% cresyl violet solution and 3% acetic acid for 5 min, washed, dehydrated, cleared in xylene, and coverslipped.
Western blot
Twenty micrograms of protein extracts were run on polyacrylamide gels (cat. #4569033, BioRad). The standard procedure of Western blot was performed using anti-Ser 3-phospho cofilin 1 and cofilin 1 (cat. #3313, #5175, Cell Signaling), and β-actin (cat. #A1978, Sigma) antibodies. To ensure equal total proteins level, blots were reprobed with tubulin (cat. #T9026, Sigma) antibodies. For detection, the chemiluminescent method was used. To quantify individual bands, a scan of photographic films was analyzed by densitometry with GeneTools Software (Syngene).
Electrophysiology
Whole-cell patch-clamp recordings were carried out in voltage-clamp mode. Mice (male and female) were anesthetized with isoflurane and decapitated. Brain slices (coronal, 250 μm thick) prepared using Leica VT 1200S vibratome in ice-cold NMDG solution (135 mM NMDG, 1.2 mM KH2PO4, 1 mM KCl, 1.5 mM MgCl2·6H2O, 0.5 mM CaCl2·2H2O, 20 mM choline bicarbonate, 10 mM d-glucose, saturated with carbogen—95% O2, 5% CO2) were then transferred to the artificial cerebrospinal fluid (ACSF) solution (119 mM NaCl, 2.5 mM KCl, 1.3 mM MgCl2·6H2O, 1 mM NaH2PO4, 26 mM NaHCO3, 20 mM d-glucose, 2.5 mM CaCl2·2H2O, saturated with carbogen), incubated for 12 min at 31 °C and then at least 1 h at room temperature. During recordings, slices were held in a recording chamber perfused with carbogenated ACSF solution supplemented with 100 μM picrotoxin and 0.5 μM tetrodotoxin and heated up to 31 °C. Granule cells of the upper blade of dentate gyrus were identified visually. Borosilicate glass capillaries (4–7 MΩ resistance) used for patch-clamp recordings were filled with Cs-based internal solution (130 mM Cs-gluconate, 20 mM HEPES, 0.4 mM EGTA, 3 mM TEA-Cl, 4 mM Na2ATP, 0.3 mM NaGTP, 4 mM QX-314Cl; osmolarity: 285–290 mOsm, pH = 7.0–7.1). Data were acquired using custom algorithms in Igor Pro (Wavemetrics) with an NPI amplifier and digitized at 10 kHz with an ITC-18 InstruTECH/HEKA. Recorded currents were filtered at 2 kHz. Series and input resistances were monitored throughout the experiment. To measure miniature excitatory postsynaptic currents (mEPSCs) 10- to 20-min-long voltage-clamp recordings were collected. Miniature events were analyzed using the MiniAnalysis software (Synaptosoft). mEPSCs amplitude detection threshold was set to 7 pA. All mini events automatically detected by the software were verified visually by the experimenter.
Analysis of dendritic spine morphology
Serum response factor KO and WT mice (male and female
n = 8) were anesthetized and transcardially perfused with phosphate-buffered saline (PBS) and with 1.5% paraformaldehyde (PFA) at room temperature. The brains were placed in 1.5% PFA for 20 min for postfix and then transferred to ice-cold PBS for at least 20 min. Next, these brains were cut on a vibratome into 130-µm-thick slices and placed in PBS at room temperature for 1 h. Gene gun technique was used to label sections with tungsten particles (Bio-Rad, Hercules, CA, USA) coated with lipophilic dye DiI (1,1ʹ-dioctadecyl-3,3,3ʹ,3ʹ-tetramethyl indocarbocyanine perchlorate; d-3911, cat. #D282, ThermoFisher). The slices were incubated in PBS at the room temperature for about 3 h and next in 1.5% PFA in 4 °C overnight, which enables diffusion of the dye into neuronal processes and allows to visualize dendritic spines. Z-stacks of confocal images of the seven dendrites per animal from the middle molecular layer of the upper blade of dorsal DG were acquired with 561-nm laser line with Zeiss LSM780 confocal system. Neurons of immature morphology (very sparse dendritic spines) were excluded from the analysis. The semiautomatic SpineMagick! software was used to measure and analyze dendritic spines by obtaining the virtual skeletons (Ruszczycki et al.
2012). We used a scale-free parameter (the length/width ratio) which reflect the spine shape (Michaluk et al.
2011). Length/width ratio was calculated and plotted using a logarithmic scale. The density of spines was calculated as a number of spines per 1 μm of dendrite length. Spines’ shapes were divided into clusters and then sorted into two groups: “long” and “mushroom and stubby” spines using custom scripts (Jasinska et al.
2016).
Behavioral tests
Open field The apparatus had a wooden floor (50 cm × 50 cm) surrounded by walls (34 cm hight). Animals’ behavior was monitored by a video camera placed above the center of the apparatus. Adult females (WT n = 12, SRF KO n = 12, about 6 months old or older) were put individually in one corner of the open field facing the wall and were allowed to explore freely for 15 min. The floor of the apparatus was cleaned with 5% ethanol after each session. Data were analyzed using EthoVision 3.1 System (Noldus Information Technology), total distance traveled was counted.
Species-typical behaviors All species-typical behaviors tests were performed on the same group of animals (WT n = 10, SRF KO n = 7; adult female and male, about 6 months old or older), starting from overnight nesting test and then marble burying and digging.
Nesting Mice were housed individually in their home cages with standard bedding for at least 1 week. After that time paper towel (divided into six pieces 9 cm × 9 cm each) was placed in the middle of each cage and left overnight to assess the nest building ability of the mice. The following scoring system was used: (1) paper towel was mostly untouched (> 90%) and was left in the middle of the cage; (2) flat pad-shaped nest, mostly moved to a corner; (3) more complex nest with biting the towel but not gathered in one place; (4) nest with shredded paper to form a cup; (5) perfect nest, paper towel torn up to form a crater, where walls are higher than mouse body height. Nest building is spontaneous home cage behavior of mice. It indicates healthy functioning and well-being of an animal (Jirkof
2014; Moretti et al.
2005). It is also interpreted by some authors as social behavior (Crawley
2012).
Marble burying Clean cage was filled 5 cm deep with bedding, tamped down to make even and flat surface. The same 12 glass marbles (15 mm in diameter) were placed in a regular pattern in two rows on bedding surface. Mice were placed individually into the experimental cage for 30 min. After that time, mice were taken to their home cages, and the number of marbles buried to at least two-thirds of their depth with bedding was counted. The same cage was re-used for all tested animals.
Digging Clean cage was prepared with the flat surface of wooden, 5-cm-deep bedding. Mice were placed individually in the experimental cage. The duration of the test was 3 min and latency to start digging, a number of digging bouts and total duration of digging were recorded and analyzed. Coordinated movements of force and/or hindlimbs which displace the wooden bedding defined the digging. The new bedding was used for all tested animals. Marble burying measures spontaneous digging behavior, which is typical for mice and is dependent on hippocampal function (Deacon
2006; Deacon and Rawlins
2005).
Statistical analyses
To compare the distributions, Shapiro–Wilk normality test was performed. To test the differences between two groups, unpaired t test or Mann–Whitney test (nonparametric) was used. When required repeated-measures ANOVA was performed, followed by Bonferroni’s multiple comparisons post hoc test. To compare the cumulative distributions of mEPSCs amplitudes and frequencies, the Kolmogorov–Smirnov test was used. The number of animals and neurons which were used for analysis is provided in figures legend. Data on the graphs are expressed as cumulative probability or means ± standard errors of the means (SEM). The difference between the experimental groups was considered as significant when p < 0.05. Results were analyzed in GraphPad Prism software.
Discussion
The present study assessed morphological, electrophysiological, and behavioral effects of SRF deficiency in adult excitatory granular neurons. The depletion of SRF in adult neurons leds to the direct downregulation of mRNAs of two actin-encoding genes and the functional inhibition of cofilin 1. Although alterations of actins and actin-binding protein did not alter gross hippocampal architecture, changes in dendritic spine morphology and basal synaptic transmission in granular neurons of the DG were observed. Moreover, SRF KO animals exhibited hyperactivity and impairments in behaviors that are typical for small rodents, recognized as hippocampus-dependent tasks. Altogether, our results suggest that basal SRF expression in adult DG neurons is engaged in the regulation of structural plasticity, basal synaptic transmission, and species-typical behaviors.
Alterations of spine morphology without gross changes in hippocampal architecture were the most prominent phenotype we identified in animals with a decrease in SRF expression in adult neurons. At the molecular level, we observed a decrease in the expression of neuronal actins and an increase in the phosphorylation of the actin-depolymerization factor cofilin 1. The actin cytoskeleton in neurons is composed of two distinct isoforms, non-muscle γ-actin and β-actin. Both actin genes are SRF targets and are downregulated in DG neurons in SRF KO animals (Miano et al.
2007; Sun et al.
2006). The present results confirmed earlier findings. The expression of both β-actin and γ-actin is impaired upon SRF ablation. We essentially generated the double knockdown of brain-expressed actin isoforms in
Srf mouse mutants.
We also showed that the depletion of SRF in adult neurons coincided with an increase in cofilin 1 phosphorylation at Ser3 (inactivation), suggesting that in wild-type cells, SRF is involved in cofilin activation. Cofilin 1 activity could be inhibited by increased phosphorylation by LIM kinases (LIMKs) or decreased activity of phosphatases like slingshot (SSH) family of protein phosphatases (Mizuno
2013). In MRTFs-deficient neurons (myocardin-related transcription factors; SRF coactivators), a decrease in Pctaire1 kinase activity was shown to stimulate the Pak1-Limk cascade to promote cofilin phosphorylation (Mokalled et al.
2010). Pctaire1 (Cdk16) gene is a direct target of SRF/MRTF (Mokalled et al.
2010). Moreover, the decrease in actin levels observed in SRF-deficient neurons might impair slingshot activation, which is stimulated by association with actin filaments (Kurita et al.
2008; Liu et al.
2007; Nagata-Ohashi et al.
2004). Both mechanisms of increased cofilin 1 phosphorylation observed in SRF-deficient neurons are possible. Collectively, our data are consistent with a previous study that reported higher levels of phosphorylated cofilin 1 in SRF- or MRTFs -depleted neurons (Alberti et al.
2005; Beck et al.
2012; Mokalled et al.
2010; Zimprich et al.
2017).
In the present study, we observed a sharp reduction of total actin levels in SRF-depleted neurons. Interestingly, SRF deletion in non-neuronal cells resulted in the downregulation of β-actin and a decrease in the ratio of polymerized F-actin to G-actin (Randrianarison-Huetz et al.
2018; Schratt et al.
2002; Taylor et al.
2014). Thus, changes in spine morphology in SRF-depleted neurons could be attributable to both lower total actin expression and alterations of the regulation of actin polymerization. However, further quantification of the F-actin-to-G-actin ratio in neurons is needed. The link between SRF and actin is multifaceted. Actin is a direct target of SRF and controls SRF-dependent gene expression (Sotiropoulos et al.
1999). The conversion of monomeric G-actin to polymeric F-actin results in the activation of SRF-dependent transcription that is regulated by the translocation of MRTFs (Kalita et al.
2006; Kaneda et al.
2018) from the cytoplasm to the nucleus (Miralles et al.
2003; Posern et al.
2002,
2004). Moreover, nuclear actin dynamics were also shown to regulate the activity of SRF/MRTFs (Baarlink et al.
2013; Ho et al.
2013; Lundquist et al.
2014; Vartiainen et al.
2007). Thus, there is a feedback loop between SRF and actin, in which the activity of SRF regulates actin levels, the dynamics of which influence the transcriptional activity of SRF.
Actin cytoskeleton remodeling, stabilization, or depolymerization is crucial for dendritic spine morphology and synaptic function (Ethell and Pasquale
2005; Honkura et al.
2008; Matus
2000). Modifications of the number of neuronal connections and the size and shape of dendritic spines underlie synaptic plasticity (Borczyk et al.
2019; Holtmaat and Svoboda
2009; Kasai et al.
2010). Alterations of spine morphology underlie many neuropsychiatric disorders, indicating their functional importance (Glausier and Lewis
2013; Penzes et al.
2011; Phillips and Pozzo-Miller
2015). Spine parameters, such as the dimension of the spine head or neck, determine spine motility and stability (Nimchinsky et al.
2002). Mushroom-shaped spines are generally considered mature and stable. In contrast, thin spines with a long neck are highly plastic (Bourne and Harris
2007). In the present study, we found that SRF depletion in granular neurons shifted the spine population toward long spines with a less mature phenotype. Thus, dendritic spines grew but did not form enlarged heads, which are characteristic of mature mushroom spines. In addition to spine morphology, we observed deficits in basal synaptic transmission in SRF-depleted neurons. The lower amplitude (mostly mEPSCs of small and medium amplitudes) and frequency of mEPSCs that were observed in DG neurons that lacked SRF could represent either a change in release probability or postsynaptic processes, such as the loss of functional synapses.
Using several behavioral tasks, we showed that animals that lacked SRF protein in adult forebrain neurons built poorer nests, performed fewer digging bouts, exhibited a reduction of burying behavior compared with their WT littermates. In agreement with other studies, mice lacking SRF were significantly more active than control animals in the open field test (Johnson et al.
2011; Parkitna et al.
2010; Zimprich et al.
2017). Similar alterations in innate behaviors and activity were observed in animals with hippocampal lesions (Deacon et al.
2002). Deficits in nest building, digging and marble burying, in SRF KO animals, were also present in the animal models with smaller volume of the DG and abnormal spine morphology of DG neurons (Kondratiuk et al.
2013,
2017) and in mice with deficits in DGs adult neurogenesis (Jedynak et al.
2012). Although behaviors observed in SRF KO animals may reflect dysfunction of the dentate gyrus, we cannot exclude the possibility that SRF deletion in other brain structures impacts the observed phenotype. Interestingly, marble burying and nest building tests are interpreted by some authors as a measure of repetitive or social behaviors, respectively, were found to be altered in several murine models of psychiatric disorders, such as schizophrenia and autism spectrum disorders (Crawley
2012; Deacon
2006; Deacon et al.
2002; Jirkof
2014).
Loss-of-function of the
ACTB gene in humans is associated with developmental delays or intellectual disability (Cuvertino et al.
2017; Palumbo et al.
2017). Moreover, a single-nucleotide polymorphism (SNP) of the
ACTB gene that caused missense mutations was shown to be linked with several neurological phenotypes, including global developmental delay, intellectual disability, cognitive impairment, abnormal aggressive and impulsive behavior, attention deficit, hyperactivity, and autism spectrum disorders (based on DECIPHER database;
http://www.decipher.sanger.ac.uk/, accessed 2018). An increase in hyperactivity and deficits in species-typical behaviors in SRF KO animals also resemble the phenotype of mice with the brain-specific deletion of β-actin, potentially linking some of the observed phenotypes in SRF-deficient neurons to the specific deletion of β-actin (Cheever et al.
2012). However, further work is needed to determine whether the specific depletion of β-actin and γ-actin in adult neurons mimics the phenotype that we observed in SRF KO animals. Overall, the present data support the notion that such behaviors as nest building, marble burying, and locomotor activity require intact hippocampal circuity.
Several lines of evidence suggest a role for SRF in brain pathology, such as epileptogenesis, cocaine-induced gene expression, and cocaine-induced dendritic spine formation (Cahill et al.
2017; Kuzniewska et al.
2016; Losing et al.
2017; Parkitna et al.
2010). Data also link SNPs of the
Srf and
Mrtfs genes to brain diseases. In humans, SNPs in CArG box (SRF transcription factor binding site), that disrupt SRF binding, were linked with neurological disorders, such as bipolar disorder, amyotrophic lateral sclerosis, and Alzheimer’s disease (Benson et al.
2011). Moreover, SNPs of the
Mrtfs genes were associated with neurodevelopmental disorders, such as schizophrenia and autism spectrum disorders (Holt et al.
2010; Luo et al.
2015; Wang et al.
2016).
Altogether, our data indicate that the adult deletion of SRF in DG neurons alters the expression of actins, dendritic spine morphology, and mouse behavior, showing that SRF expression is necessary to maintain proper brain circuitry. To our knowledge, this is the first report that demonstrates that SRF regulates the adult structural plasticity of intact, adult hippocampal neurons in vivo. Furthermore, the present results link aberrant spine morphology to changes in glutamatergic synaptic transmission and behavior.
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