Myosin Va-dependent Transport of NMDA Receptors in Hippocampal Neurons

Gong, Ru; Qin, Linwei; Chen, Linlin; Wang, Ning; Bao, Yifei; Lu, Wei

doi:10.1007/s12264-023-01174-y

Myosin Va-dependent Transport of NMDA Receptors in Hippocampal Neurons

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Published: 30 January 2024

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Myosin Va-dependent Transport of NMDA Receptors in Hippocampal Neurons

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Ru Gong¹,
Linwei Qin¹,
Linlin Chen³,
Ning Wang³,
Yifei Bao¹ &
…
Wei Lu ORCID: orcid.org/0000-0001-5920-564X^1,2,3,4

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Abstract

N-methyl-D-aspartate receptor (NMDAR) trafficking is a key process in the regulation of synaptic efficacy and brain function. However, the molecular mechanism underlying the surface transport of NMDARs is largely unknown. Here we identified myosin Va (MyoVa) as the specific motor protein that traffics NMDARs in hippocampal neurons. We found that MyoVa associates with NMDARs through its cargo binding domain. This association was increased during NMDAR surface transport. Knockdown of MyoVa suppressed NMDAR transport. We further demonstrated that Ca²⁺/calmodulin-dependent protein kinase II (CaMKII) regulates NMDAR transport through its direct interaction with MyoVa. Furthermore, MyoVa employed Rab11 family-interacting protein 3 (Rab11/FIP3) as the adaptor proteins to couple themselves with NMDARs during their transport. Accordingly, the knockdown of FIP3 impairs hippocampal memory. Together, we conclude that in hippocampal neurons, MyoVa conducts active transport of NMDARs in a CaMKII-dependent manner.

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Introduction

The N-methyl-D-aspartate receptor (NMDAR) is a subtype of glutamate receptor important for postsynaptic response modulation and thus plays a crucial role in various brain functions [1]. The number of synaptic NMDAR components can be altered in response to electrophysiological inputs or sensory cues [1,2,3,4,5,6,7,8,9]. Transport of new NMDARs into dendritic spines increases the number of postsynaptic NMDARs, thereby facilitating the subsequent occurrence of synaptic plasticity and memory consolidation [1, 7, 10,11,12,13]. In another light, dysregulated NMDAR transport has been found in the pathophysiology of neural disorders such as Alzheimer’s disease and schizophrenia [10, 14, 15]. In contrast to extensive studies on the role of AMPAR (α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor) trafficking in the plasticity of AMPAR-mediated synaptic responses, only a few studies have documented the requirement of NMDAR transport for NMDAR-mediated synaptic plasticity.

After being released from the endoplasmic reticulum, the assembled NMDARs are sent via Golgi bodies to the neuronal surface [16, 17]. The journey from the soma to the dendritic spine requires members of the kinesin motor protein family, together with adaptor proteins, to transport NMDARs along microtubules in dendrites. Kinesin family 3B (KIF3B) and KIF17 are respectively responsible for the transport of NMDARs containing GluN2A and GluN2B subunits [10, 11, 13, 18]. Once they arrive at the base of the dendritic spine, NMDARs are believed to be released from KIF and transferred to myosin proteins that are associated with F-actin [19,20,21], which provides cytoskeletal support in dendritic spines.

Among the great family of motor proteins, unconventional class V myosins have been found to be associated with the active transport of diverse cargos along actin filaments [20, 22, 23], which makes them attractive candidates for the transport of synaptic proteins. Of the three class V myosins, only myosin Va (MyoVa) and myosin Vb (MyoVb) are present in the brain and possess features that allow them to serve as effective organelle motors within neurons [20, 22]. To date, it is unknown which type of myosin conducts NMDAR transport in spines. Considering the crucial role of NMDAR trafficking in both physiological brain functions and various neurological disorders, it is of great importance to uncover the specific motor protein and the process of regulation of NMDAR trafficking.

In this study, we specified that MyoVa, but not MyoVb, associates with NMDARs through its globular, cargo-binding domain in CA1 hippocampal neurons. This bond is fortified during NMDAR trafficking. Using a combination of biochemical, immunofluorescent, and electrophysiological measurements, we further demonstrated that MyoVa conducts a Ca2+/calmodulin-dependent protein kinase II (CaMKII)-dependent surface transport of NMDARs. This transport process requires Rab11 family-interacting protein 3 (Rab11/FIP3) as the adaptor proteins to couple NMDARs and MyoVa. Taken together, these results support MyoVa as the specific motor protein that conducts NMDAR transport, which is important for the formation of hippocampal memory.

Materials and Methods

Animals

All animal-related procedures were approved by the Experimental Animal Ethics Committee at Southeast University. Male and female Sprague−Dawley (Charles River, Beijing, China) rats were housed in a temperature-controlled (26°C) vivarium and maintained on a 12-h/12-h light/dark cycle (lights on from 07:00 to 19:00) with ad libitum access to food and water. Male and female rats, 14–16 days old, were used in electrophysiology and Western blotting. One-month-old male rats were used in behavioral testing experiments.

Peptides

The interfering peptide Tat-MyoVa (4 μmol/L, YGRKKRRQRRR-PKPGHKRTDSTHSSNESEY) was designed based on the CaMKII binding sequence in MyoVa. The sequence of the scrambled peptides was YGRKKRRQRRR-YHSPSDGKSNTSHPTKERE, in which YGRKKRRQRRR represents the structural domain that mediates cell membrane conductance. Peptides were synthesized by ChinaPeptides Co. (Nanjing, China) and applied in artificial cerebrospinal fluid (ACSF) used in patch-clamp recording 20 min prior to the induction of long-term potentiation (LTP).

Virus

A sequence corresponding to 5344–5362 nucleotides (5’-GGT CTC TGT TTC ATT TAT C-3’) in rat MyoVa messenger RNA was used to inhibit endogenous rat MyoVa. A nonspecific scrambled short-hairpin RNA (shRNA) was treated as a control (5’-GGT TTC GTA CTT TCT CTT A-3’). The sequences were packaged into rAAV2/9 (Shanghai Sunbio Medical Biotechnology, Shanghai, China) or lentiviral particles (BrainVTA, Wuhan, China) tagged with mCherry. The flag-tagged wild-type (rescue) and 1725 mutant (Q1725A mut, the 1725 site is homologous to 1750 in humans) MyoVa gene resistant to knockdown (KD) were packaged into lentiviral particles by BrainVTA. The nucleotides used for endogenous rat FIP3 KD (5’-GCA TTC TGC TAC TTG CTA AAG-3’), and the nonspecific scrambled shRNA (5’-CGC TGA GTA CTT CGA AAT GTC-3’) were tagged with the enhanced green fluorescent protein (EGFP) and packaged into rAAV2/9 by Shanghai GeneChem (Shanghai, China).

In Vivo Stereotactic Injections

One-week-old rats used in electrophysiology experiments and 8-week-old rats used in behavior tests were prepared for stereotaxic injection [24, 25]. Rats were anesthetized with isoflurane and then immobilized on a stereotaxic apparatus. Small bilateral holes were drilled into the skull and then concentrated virus solution (rAAV2/9 expressing MyoVa or FIP3 KD shRNA) was injected either unilaterally into the CA1 region of 1-week-old (300 nL, bregma −2.0 mm; lateral ±2.5 mm; ventral ±2.4 mm) or bilaterally into 8-week-old rats (400 nL, bregma −3.5 mm; lateral ±2.8 mm; ventral ±2.6 mm) with a glass pipette attached to a microinjection pump (World Precision Instruments WPI, Sarasota, USA). After the virus injection, the needle was left in place for 10 additional min. Two weeks after the AAV injection, the rats were sacrificed for extracellular recordings, Western blot, or behavioral tests.

To verify virus expression in the CA1 region of the hippocampus, stereotaxic slices were obtained 2 weeks after virus injection. Rats were anesthetized with isoflurane and cardio-perfused. The brain was removed and fixed in 4% paraformaldehyde for 24 h and soaked in 40% sucrose for 2–3 days for dehydration. The 35-μm-thick hippocampal sections of the CA1 region were cut on a microtome cryostat (CM1950, Leica, Wetzlar, Germany) and then cover-slipped with mounting medium (Southern Biotech, Birmingham, USA) containing DAPI. Images were captured under an upright microscope (DM5000B, Leica) or laser scanning confocal microscope (LSM700, Zeiss, Oberkochen, Germany).

Acute Hippocampal Slice Preparation

Rats were anesthetized with isoflurane and rapidly decapitated. The isolated brain was quickly placed in cold ACSF containing (in mmol/L): 124 NaCl, 5 KCl, 1.2 KH₂PO₄, 1.3 MgSO₄, 2.4 CaCl₂, 26 NaHCO₃, and 10 glucose (pH 7.4). Then the hippocampal tissue was cut into 350 μm thick slices on a Leica vibratome. After incubation at 34°C for 1 h in oxygenated ACSF continuously ventilated with 95% O₂ and 5% CO₂, the slices were acclimated to room temperature for another 30 min. During recordings, slices were placed in a chamber filled with ACSF continuously ventilated with 95% O₂ and 5% CO₂.

Electrophysiology

Patch pipettes were made with a horizontal microelectrode puller (P-1000, Sutter Instruments, Novato, USA). The average resistance was 3–5 MΩ. To evoke field excitatory postsynaptic potentials (fEPSPs) in CA1, a bipolar stimulating electrode (FHC, Bowdoin, USA) was placed in the stratum radiatum to stimulate Schaffer commissural projections at 0.05 Hz. The resistance of the glass electrode filled with 2 mmol/L NaCl to record fEPSP was 3–5 MΩ. NMDAR-mediated fEPSPs were recorded in ACSF with a low Mg²⁺ (0.25 mmol/L) concentration. 10 μmol/L NBQX (2,3-dihydroxy-6-nitro-7-sulfamoylbenzo quinoxaline-2,3-dione) was added to the recording solution to block AMPAR-mediated excitatory synaptic currents (EPSCs). The EPSCs of NMDAR were obtained by whole-cell recording with a patch pipette (4–6 MΩ) containing (in mmol/L) 135 K-gluconate, 5 KCl, 2 NaCl, 0.2 EGTA, 10 HEPES, 5 ATP, and 5 QX-314 (pH 7.4). Cells were held at −65 mV in the low concentration of Mg²⁺ (0.25 mmol/L) ACSF. 10 μmol/L BMI (bicuculline methiodide) and 10 μmol/L NBQX were applied in the recording solution for the blockade of GABA_A and AMPA receptors, respectively. NMDAR-mediated miniature EPSCs (mEPSCs) were recorded at −65 mV in Mg²⁺-free ACSF containing NBQX (10 μmol/L), BMI (5 μmol/L), and tetrodotoxin (TTX, 1 μmol/L). NMDAR-dependent LTP was induced by theta-burst stimulation (TBS, 3 times 5 bursts of 5 Hz, each burst containing 5 stimuli at 100 Hz, with a 30 s interburst interval, for references see [26]) or incubation with the protein kinase C (PKC) agonists PMA (phorbol myristate acetate, 0.5 μmol/L, for references see [27,28,29]) for 10 min. NMDAR-dependent LTD (long-term depression) was induced by low-frequency stimulation containing 900 stimuli at 1 Hz [30]. Individual experiments were normalized to the baseline to generate a summary graph, and three consecutive reactions were averaged for 1-min plots. The magnitudes of LTP or LTD were calculated based on the averaged fEPSP or EPSC values in the last 10 min of recording after induction. Clampex 10.4 and Clampfit 10.4 (Axon Instruments, San Jose, USA) were used for data recording and analysis, respectively.

Primary Hippocampal Cell Culture

Primary hippocampal cultures were prepared from E18–19 rats as described previously [31, 32]. Hippocampi were dissected and digested with 0.125% trypsin (Gibco, CA, USA) diluted in FBS (fetal bovine serum) for 8 min at 37°C, then the reaction was quenched with 10% FBS (Gibco) diluted by Dulbecco’s modified Eagle’s medium. Neurons were plated on coverslips coated with 0.1 mg/mL poly-D-lysine (Gibco) at a density of 50,000–80,000 cells per cm² (for imaging studies) or 300,000 cells per cm² (for Western blots). Neurons were grown in Neurobasal medium (Gibco) supplemented with B-27 (Gibco) containing glutamine (Gibco), β-mercaptoethanol (Gibco), and 1× penicillin-streptomycin (Gibco). Cultures were kept in a humid 5% CO₂ tissue culture incubator at 37°C. Half fresh medium was exchanged every 3 days until experiments. Neurons were infected with MyoVa-shRNA or MyoVa scrambled-shRNA at 7 days in vitro (DIV) by adding rAAV2/9-packaged viruses into wells with the final virus titer at 10⁹–10¹⁰ genome copies/mL. For the expression of MyoVa or Q1725A mutant MyoVa in neurons, lentivirus was added at DIV7.

Western Blotting

The hippocampal slices were homogenized in cold homogenization buffer (pH 7.4) containing (in mmol/L) 320 sucrose, 1 HEPES, 1 MgCl₂, 1 NaHCO₃, 20 sodium pyrophosphate, 20 β-phosphoglycerol, 0.2 dithiothreitol, 1 EDTA, 1 EGTA, 50 NaF, and 1 Na₃VO₄. Protease inhibitors and phosphatase inhibitors were also added, which included 1 mmol/L phenyl-methylsulfonyl fluoride (PMSF), 1 mmol/L p-nitrophenyl phosphate, 5 µg/mL aprotinin, 5 µg/mL leupeptin, 5 µg/mL pepstatin A, and 16 µg/mL benzamidine. The sample was centrifuged at 1,000× g for 10 min. The supernatant was used for total protein analysis. To obtain the Triton insoluble fraction (TIF) [33, 34], the supernatant was centrifuged at 3,000× g for 15 min and the resulting pellet was resuspended in 8 mL of hypotonic buffer with protease inhibitors, followed by centrifugation at 100,000× g for 1 h. The resulting pellet was resuspended and centrifuged for 1 h at 100,000× g in 8 mL of buffer containing 75 mmol/L KCl and 1% Triton X-100. The final pellet was homogenized three times in 20 mmol/L HEPES, and the resulting fraction was referred to as the TIF. The concentration of total or TIF proteins was determined with the Bradford protein assay kit (Thermo Fisher, Waltham, USA). Equal amounts of proteins were denatured by 5× SDS (sodium dodecyl sulfate)-loading buffer and then separated by SDS-PAGE using precast 8% gels. Proteins on the gels were transferred to PVDF (hydrophobic polyvinylidene fluoride) membranes (BioRad, Hercules, USA) and blocked in 3% BSA (bovine serum albumin) in TBST solution for 2 h at room temperature. Then, the membranes were incubated with primary antibodies diluted in 1% BSA in TBST solution at 4°C overnight. After washing 5 times in TBST, the membranes were incubated with the corresponding secondary antibody diluted in 1% BSA in TBST solution for 1 h at room temperature. The membranes were then washed another 5 times. Results were visualized using an imaging system (Tanon, Shanghai, China) and analyzed by ImageJ. The major primary antibodies were: MyoVa (Sigma, St. Louis, USA, HPA001356), GluN2A (Millipore, Burlington, USA, AB1555P), GluN1 (rabbit: Abclonal, Wuhan, China, A11699; mouse: Abcam, Cambridge, UK, ab134308), GluA1 (Millipore, MAB2263), GluN2B (Cell Signaling Technology, Boston, USA, 14544), Rab11 (Invitrogen, Carlsbad, USA, 71-5300), FIP3 (Rockland, Philadelphia, USA, 600-401-994), PSD95 (Millipore, MABN68), MyoVb (Sigma, HPA040902), CaMKII (Santa Cruz Biotechnology, Inc., Dallas, USA, sc-13141), GST (glutathione-S-transferase, Genscript, Nanjing, China, A00866-100). The secondary antibodies were: goat anti-rabbit IgG-HRP (horseradish peroxidase, Thermo Fisher Scientific, Waltham, USA, A16110) and goat anti-mouse IgG-HRP (Jackson ImmunoResearch, West Grove, USA, 115-035-003). For visualization of the signal, the Super Signal West Pico Chemiluminescent Substrate Kit (Thermo Fisher Scientific, 34580) was used according to the manufacturer’s instructions.

Co-immunoprecipitation (Co-IP)

Total hippocampal lysates were incubated with primary antibodies in the immunoprecipitation buffer at 4°C by end-over-end spinning. The immunoprecipitation buffer (pH 7.4) contained 0.05 mol/L HEPES, 0.15 mol/L NaCl, 10% glycerol, 0.5% Nonidet P-40, 1% TritonX-100, 1 mmol/L EGTA, 1 mmol/L EDTA, 1 mmol/L Na₃VO₄, and 1 mmol/L PMSF. After equilibration, the Protein A/G agarose (Thermo Fisher Scientific) was pipetted into the solution and mixed at 4°C overnight. After 5 washes with 1% TritonX-100 buffer, the bound proteins were eluted by boiling with 2× SDS buffer for 8 min, separated by SDS-PAGE, and analyzed by immunoblotting.

Glutathione S-transferase (GST) Pull-down Assays

The GST constructs of the rat MyoVa globular tails (amino acids 1,430–1,830) and medial tail (amino acids 1,110–1,440) were generated by Genscript as described in previous studies (Correia et al., 2008; Costa et al., 1999). Equal amounts of total proteins from hippocampal slices or cells were mixed with glutathione resin (Genscript), together with either GST-MyoVa globular tails, medial tail sequence fusion proteins, or GST alone at 4°C overnight via end-over-end spinning. The beads were processed by centrifugation and washing 5 times with PBS (phosphate-buffered saline) containing 0.1% Triton X-100. The conjugated proteins were eluted by boiling in 2× SDS sample buffer for 10 min, separated by SDS-PAGE, and analyzed by immunoblotting.

Surface Biotinylation Assay

The surface biotinylation assay was conducted to detect the expression of NMDAR proteins on the plasma membrane, as described in the Pierce Cell Surface Protein Biotinylation and Isolation Kit (Pierce, Thermo Fisher). At DIV16, hippocampal neurons infected with the virus were treated with or without PMA for 10 min and then were incubated with 1 mg/mL EZ-Link™ Sulfo-NHS-SS-Biotin at 4°C for 45 min. After the labeling solution was removed from the wells, neurons were washed three times with ice-cold TBS and harvested by gentle scraping. Neurons were then transferred into the lysis buffer containing protease inhibitors and centrifuged at 15,000× g for 10 min at 4°C. Supernatants containing equal amounts of protein were incubated with NeutrAvidin™ agarose in a column overnight at 4°C to capture biotinylated (surface) proteins via an end-over-end mixing rotator. After 5 washes with the wash buffer in the kit, the captured surface proteins were eluted from the biotin-NeutrAvidin agarose by incubation with dithiothreitol (10 mmol/L) in elution buffer for 30 min at room temperature on an end-over-end rotator. The eluted proteins, representing the cell surface proteins, were collected by column centrifugation. The samples were then separated using SDS-PAGE and imaged by Western blotting.

Immunofluorescence Labeling and Analysis

Immunofluorescence was used to analyze NMDAR trafficking in primary hippocampal cultures as previously described [31, 35]. The GluN1 expressed on the surface of the neuronal plasma membrane or in the intracellular compartment were labeled with the corresponding antibodies under membrane-impermeable or membrane-permeable conditions, respectively. Surface GluN1 was labeled by incubating hippocampal neurons with mouse GluN1 antibodies (Abcam) for 40 min at room temperature, followed by 5 washes with PBS, fixation in ice-cold parafix solution (4% paraformaldehyde, 4% sucrose in PBS) for 15 min, and 5 washes with PBS. Neurons were then incubated with Alexa fluor 488-conjugated secondary antibody (Invitrogen) for 1 h at room temperature. Following permeabilization (0.3% Triton X-100 in PBS) and blocking (10% normal donkey serum, 1 h) at room temperature, total GluN1 was labeled with rabbit GluN1 antibodies (ABclonal) overnight at 4°C. After incubation with Alexa fluor 647 secondary antibodies (Abcam) for 1 h at room temperature and 5 washes with PBS, sections were then mounted on glass slides coverslipped with Fluoromount-G (SouthernBiotech). A 63× oil-immersion objective on a Zeiss LSM900 confocal microscope was used to capture images. Consecutive optical sections were collected at 0.35 μm intervals to create the maximum projection image. Fluorescence intensities from 3 independent areas per neuron were measured using Fiji software (National Institutes of Health, West Grove, USA) for surface (Alexa fluor 488-conjugated secondary antibody) and total GluN1 (Alexa fluor 647 secondary antibodies). Data are presented as surface/total GluN1 ratios.

For co-localization analysis of GluN1 and MyoVa (GluN1/MyoVa), GluN1/PSD95, GluN1/FIP3, or CaMKII/MyoVa, hippocampal neurons at DIV 16–18 were fixed and permeabilized as above and then blocked with 10% normal donkey serum for 1 h at room temperature. Primary antibodies labeling pairs of proteins (GluN1/MyoVa, GluN1/PSD95, GluN1/FIP3, or CaMKII/MyoVa) were incubated overnight at 4°C. Following 5 washes, corresponding secondary antibodies were incubated for 2 h at room temperature. After 5 washes, the coverslips were mounted on glass slides. Images were acquired with a 63 × oil immersion objective on a Zeiss LSM900 confocal microscope under the same exposure, gain, and intensity conditions and analyzed with Fiji software (National Institutes of Health) using the same parameters.

Behavioral Tests

Fear Conditioning

Before the beginning of the experiments, rats were handled for 5 days. The training and test procedures for both contextual and trace fear conditioning were performed as previously described [36]. On the first day, the rats were acclimatized for 12 min in a training room with metal walls and a stainless-steel grid floor. On day 2, rats were allowed to explore the chamber for 3 min before the onset of six consecutive training blocks, each consisting of a 20 s baseline, a 20 s, 2 kHz, 80 dB tone (conditioned stimulus), an 18 s trace interval of silence, followed by a 0.8 mA foot shock persisting for 2 s (unconditioned stimulus) and lastly an intertrial interval (ITI) persisting for 40 s. Memory tests were applied on the third day. For contextual memory recall, rats were first placed in the training chamber for 3 min to assess contextual fear responses. For tone-cued memory recall, rats were placed in a novel chamber with walls and floors made of acrylic panels, in which they were allowed free exploration for 3 min and then presented with four testing blocks, each consisting of a 20 s baseline, a tone (20 s, 2 KHz, 80 dB), and a 60 s ITI. The chamber was cleaned with 10% ethanol after each rat was tested. The Freeze Frame (Actimetrics, Wilmette, USA) software for automatic motion detection was used to quantify the percentage of time the animal spent in freezing behaviors.

Novel Object Recognition (NOR) Task

Rats were handled for 5 days and habituated to an open-field box (100 cm × 100 cm × 35 cm) for 3 days. The training and test procedures were executed as described [37]. Rats were allowed to explore two novel objects for 10 min during training sessions on the first day, and the time for rats to explore each object was recorded. In the test sessions on day 2, rats were positioned in the same box to explore two objects for 10 min, one of which was the familiar object, and the other was replaced by a novel object. To avoid the influence of olfactory cues, the objects were thoroughly cleaned with 10% ethanol after exploration by the rats. Recognition memory was measured by a preference index using Anymaze (Stoelting, Wood Dale, USA), the ratio of the time spent exploring either of the two objects (training session) or the new object (test session) to the total time spent exploring the two objects.

Novel Place Preference (NPP) Task

Rats were handled and habituated as described above before training. During the training phase, rats were placed in the open-field box to explore two identical objects located in the corners of the box for 10 min. On day 2, one copy of the objects was placed at a different location, whereas the other one was placed in the same location as during the training trials. Recognition memory was measured by taking the ratio of the time spent exploring the relocated object to the total time spent exploring both objects using Anymaze.

Temporal Order Memory Task

Rats were handled and habituated as described above before training. Experiments were performed following protocols previously described [38]. Rats were placed in an experimental apparatus to explore each set of three objects (referred to as A–A, B–B, and C–C) for 10 min with a 3-min inter-session interval. The rats were given a 3-min timeout after the third set of objects and then were returned to the apparatus where a third copy of object A and a third copy of object C were placed at opposite ends of the box. The preference for object A versus object C in a 10-min period was measured by Anymaze (Stoelting). The overall activity level for the temporal order task was calculated as the ratio of the time spent exploring object A to the total time spent exploring objects A and C.

Open-field Tests

Open-field tests were used to test general behavior and locomotor activity as previously described [39]. Rats were paced in an open-field box (100 cm × 100 cm × 35 cm) and left to freely move for 3 min. Between tests for each rat, the test chamber was cleaned with 10% ethanol. Motion trails were recorded by video tracking through which the total distance, mean speed, and time mobile were measured and analyzed using the Anymaze system.

Quantification and Statistical Analysis

Data in the bar graphs are presented as the mean ± SEM and statistical analysis of the data was calculated by GraphPad software. Student’s t-test was applied to compare differences between the two groups. One-way ANOVA (analysis of variance) was applied to compare the differences between more than two groups. Two-way repeated measures ANOVA was used to assess how data from different groups evolved over time, that is, how animal freezing rates changed over time. The association between the two variables was evaluated by Pearson’s correlation coefficient. When P <0.05 (*P <0.05, **P <0.01), the difference was considered significant.

Results

Increased Association Between MyoVa and NMDARs During NMDAR Transport

To investigate whether MyoVa or MyoVb is involved in the transport of NMDARs, we first examined the possible association between MyoVa or MyoVb with NMDARs. We performed standard co-IP experiments on hippocampal homogenates with antibodies to GluN1, MyoVa, and MyoVb. As shown in Fig. 1A, MyoVa, but not MyoVb, was specifically immunoprecipitated by an anti-GluN1 antibody. Consistent with this, GluN1 and GluN2A were specifically immunoprecipitated by MyoVa (Fig. 1B), but not by MyoVb antibody (Fig. 1C). This selectivity for myosin association also applied to GluN2B (Fig. S1A).

To confirm the association between MyoVa and NMDARs, we used GST pull-downs targeting different domains of the MyoVa C terminus in both cultured hippocampal neurons (Fig. 1D) and hippocampal slices (Fig. 1E). We found that the globular tail of MyoVa (cargo-binding domain) was associated with both GluN1 and GluN2A NMDAR subunits in hippocampal neurons, whereas the plain GST did not. These results indicate that MyoVa associates with NMDARs in hippocampal neurons under basal conditions.

Activation of PKC rapidly transports NMDARs to the cell surface by phosphorylating GluN1 [29, 40, 41]. To further examine a possible alteration in the MyoVa-NMDAR association during NMDAR transport, we carried out co-IP analyses on hippocampal homogenates upon activation of PKC by phorbol myristate acetate (PMA, 0.5 μmol/L). Consistent with the previous studies [27, 28], perfusing brain slices with PMA for 10 min elicited enhancement of the surface expression of NMDARs at the postsynaptic site (Fig. S1B, C). As a control, the total amount of protein expression was not altered (Fig. S1D, E). More importantly, MyoVa showed a dramatic enhancement in its association with NMDARs, revealed by experiments with antibodies to MyoVa (GluN2A, 1.59 ± 0.16, P <0.05; GluN1, 1.44 ± 0.12, P <0.05; n = 6; Fig. 2A, B) and GluN1 (MyoVa, 1.52 ± 0.18, P <0.05; n = 7; Fig. 2C, D). By contrast, no NMDAR association with MyoVb was detected after PKC activation, reiterating the involvement of MyoVa rather than MyoVb in NMDAR transport (Fig. 2C). Moreover, we failed to detect any association between MyoVa and GABA receptors under control or PMA conditions (Fig. S1F).

To further confirm the enhancement in MyoVa-NMDAR association during NMDAR transport, we performed immunolabeling analyses of the endogenous proteins in dendrites from cultured hippocampal neurons. As shown in Fig. 2E, MyoVa displayed a partial colocalization with GluN1 under basal conditions. PMA treatment substantially enhanced the colocalization value (control: n = 59 neurons, 0.58 ± 0.01; PMA: n = 71 neurons, 0.61 ± 0.01, P <0.01; Fig. 2E, F). Consistent with the co-IP experiment (Figs. 1A, C and 2C), the Pearson coefficient for MyoVb/GluN1 was significantly lower than MyoVa/GluN1, indicating that GluN1 increased its association with MyoVa rather than with MyoVb (Fig. S1G, H). Taken together, these data indicate that MyoVa increases its association with NMDARs during NMDAR transport.

MyoVa KD Affects NMDAR Transport

To investigate whether MyoVa is required for NMDAR transport, we synthesized effective shRNAs consisting of a sequence previously demonstrated to KD endogenous MyoVa (Figs 3A and S2A) [42]. The shRNAs were packed into lentivirus that infects hippocampal cells. The efficiency of MyoVa KD was ensured by quantitative densitometry of immunoblots (MyoVa in KD, 0.24 ± 0.03, n = 6; P <0.01; Fig. 3B, C; 0.20 ± 0.04, P <0.01; n = 4; Fig. S2B, C). By contrast, the level of MyoVa was not affected by lentivirus containing scrambled shRNAs (scrambled, 0.96 ± 0.05, P >0.05; n = 4; Fig. S2B, C). Moreover, MyoVa KD did not alter the overall expression of MyoVb, GluN1, and other important synaptic proteins (Fig. S2B, C).

To examine the possible role of MyoVa during the surface targeting of NMDARs, we next applied a surface biotinylation assay as well as a Western blot to primary hippocampal neurons with MyoVa KD. As previously reported [27, 28], PKC activation caused a rapid and substantial increase in surface rather than total GluN1 and GluN2A expression (GluN1, 1.67 ± 0.08, P <0.01; GluN2A, 1.56 ± 0.08, P <0.01; n = 6; Fig. 3D, E). This enhancement in surface NMDAR expression, however, was absent in MyoVa KD neurons (GluN1, 0.94 ± 0.05, P >0.05; GluN2A, 1.08 ± 0.05, P >0.05; n = 6). The enhanced levels of surface NMDARs were not affected by lentiviruses containing scrambled shRNAs which were used as controls (GluN1, 1.49 ± 0.17, P <0.05; GluN2A, 1.53 ± 0.12, P <0.01; n = 6; Fig. S2D, E).

We next used an immunofluorescence assay in cultured hippocampal neurons to confirm the role of MyoVa in the regulation of NMDAR surface expression [31, 35]. NMDARs expressed on the membrane surface were labeled with antibodies against the N-terminal extracellular epitope of GluN1 under membrane-impermeable conditions. The total NMDARs were labeled with antibodies against the C-terminal intracellular epitope of GluN1 under membrane-permeable conditions. The amount of surface NMDARs was determined by quantifying the intensity of GluN1 at the membrane. Consistent with previous surface biotinylation data, we detected a substantial increase in the surface level of GluN1 shortly after PMA treatment (0.5 μmol/L) (control: n = 19 neurons, 1.00 ± 0.07; PMA: n = 20 neurons, 1.43 ± 0.06, P <0.01; Fig. 3F, G). This enhancement in surface GluN1 was absent in MyoVa KD neurons (MyoVa KD: n = 21 neurons, 0.95 ± 0.04, P >0.05; MyoVa KD-PMA: n = 21 neurons, 1.01 ± 0.05, P >0.05). As a control, the enhanced level of surface GluN1 was not affected by lentivirus containing scrambled shRNAs (scrambled: n = 20 neurons, 0.98 ± 0.03, P >0.05; scrambled-PMA: n = 19 neurons, 1.44 ± 0.08, P <0.01).

To investigate whether the effect of MyoVa on the surface NMDAR level also applied to the NMDARs specifically located at postsynaptic sites, we quantified the colocalization of NMDARs with PSD-95 (postsynaptic density 95), a postsynaptic marker (Fig. S2F, G). Consistent with the above results on surface NMDAR expression, we found that PMA treatment (0.5 μmol/L) led to an enhancement of the NMDAR/PSD-95 colocalization (control: n = 20, 1.00 ± 0.05; PMA: n = 20, 1.43 ± 0.04, P <0.01). This enhancement, however, was absent in cultured hippocampal neurons with MyoVa KD (MyoVa KD: n = 18, 1.02 ± 0.05, P >0.05; MyoVa KD-PMA: n = 20, 0.98 ± 0.05, P >0.05). As a control, neurons injected with scrambled shRNAs still displayed enhanced NMDAR/PSD-95 colocalization (scrambled: n = 18, 1.06 ± 0.06, P >0.05; scrambled-PMA: n = 13, 1.46 ± 0.04, P <0.01).

To further validate the above results, we performed rescue experiments by overexpressing full-length MyoVa resistant to the knockdown effect in cultured MyoVa KD neurons (MyoVa in rescue, 1.08 ± 0.04, P >0.05; n = 6; Fig. 3B, C). We found that MyoVa rescue completely reversed the deficiency in PMA-induced surface GluN1 enhancement, as evaluated by both biotinylations (GluN1, 1.39 ± 0.09, P <0.01; GluN2A, 1.42 ± 0.08, P <0.01; n = 6; Fig. 3D, E) and immunofluorescence assays (rescue: n = 21 neurons, 1.00 ± 0.04, P >0.05; rescue-PMA, n = 22 neurons, 1.35 ± 0.04, P <0.01; Fig. 3F, G), suggesting that MyoVa is important for PMA-induced enhancement in both surface and postsynaptic expression of NMDARs in hippocampal neurons.

The above results were obtained from cultured neurons prepared from the hippocampus of embryonic rats. These cultures, however, are likely not able to fully mimic mature synapses in situ. Therefore, we next determined whether MyoVa is also important for NMDAR transport in standard acute hippocampal slices when local circuits are partially maintained. One-week-old rats were stereotactically injected with AAV containing shRNAs into hippocampal CA1 (Fig. 4A) [24, 25], which suppressed the expression of MyoVa as revealed by Western blotting assays (MyoVa, 0.28 ± 0.05, P <0.01; n = 4; Fig. 4B, C). Injected rats were sacrificed 2 weeks later for fEPSP recording in the visually identified CA1 area in hippocampal slices.

Postsynaptic NMDAR transport has been reported as an important molecular mechanism underlying the LTP of NMDAR-mediated synaptic responses [9, 12, 43, 44]. We thus examined whether MyoVa KD could affect the LTP of NMDAR-mediated fEPSPs in hippocampal slices. In acute slices prepared from the hippocampus injected with AAV targeting MyoVa, the LTP of NMDA fEPSPs was absent, no matter whether it was induced by PMA (0.5 μmol/L; control: 1.52 ± 0.11, n = 8; MyoVa KD: 0.97 ± 0.02, n = 7, P <0.05; Fig. 4D, F) [28, 29] or TBS (control: 1.53 ± 0.09, n = 9; MyoVa KD: 0.98 ± 0.02, n = 8, P <0.01; Fig. 4G, I) [12, 26]. By contrast, the LTP of NMDA fEPSPs was intact in slices injected with scrambled shRNAs (PMA: 1.59 ± 0.13, n = 7, P >0.05; TBS: 1.59 ± 0.09, n = 8, P >0.05). Moreover, the blockade of LTP by MyoVa KD was reversed by the expression of shRNA-insensitive full-length MyoVa (PMA: 1.53 ± 0.18, n = 6, P >0.05; TBS: 1.52 ± 0.16, n = 8, P >0.05). In contrast, the expression of full-length MyoVb failed to restore the impaired NMDAR LTP (Fig. S3A, F), further indicating that MyoVb does not participate in NMDAR transport.

To determine whether the impaired NMDAR LTP was caused by altered basal excitatory synaptic transmission, we measured NMDAR mEPSCs and found both their amplitude and frequency were unaffected (Fig. S3G–I). Moreover, the NMDAR LTD was normal in MyoVa KD slices (Fig. S3J–L), which suggests that MyoVa mainly contributes to NMDAR delivery to the postsynaptic membrane rather than an LTD-related change in surface mobility. These results indicate that MyoVa KD blocks the LTP of NMDA fEPSPs by interrupting postsynaptic NMDAR transport.

Taken together, our results suggest that MyoVa is essential for postsynaptic NMDAR transport.

CaMKII Regulates NMDAR Transport via MyoVa

MyoVa is a Ca²⁺-sensitive motor protein, as both its conformation and actin-activated ATPase activity can be regulated by micromolar Ca²⁺ [23, 45,46,47,48,49]. On the other hand, activation of NMDARs elicits Ca²⁺ influx and Ca²⁺ elevation in spines [50,51,52], which could lead to multiple downstream cascade events including the activation of various protein kinases. CaMKII is a downstream effector that is activated by Ca²⁺/CaM. Importantly, CaMKII has been reported to be associated with MyoVa [53, 54]. Interestingly, the CaMKII-MyoVa association increased during PMA treatment, which also induced NMDAR transport (Fig. S5A–D). We thus investigated whether CaMKII plays a role in regulating the MyoVa-NMDAR association and NMDAR transport.

Co-IP of MyoVa with NMDAR subunit GluN1 was applied to hippocampal homogenates. As expected, the MyoVa-GluN1 association was enhanced after PMA treatment (0.5 μmol/L) (Fig. 5A, B). This enhancement, however, was absent when the antagonist of CaMKII, AIP (autocamtide inhibitory peptide, 1 μmol/L), was applied (PMA, 1.50 ± 0.13, P <0.05; PMA + AIP, 1.01 ± 0.09, P >0.05; AIP, 0.99 ± 0.01, P >0.05; n = 4; Fig. 5A, B). These results suggest that CaMKII regulates the interaction of MyoVa with NMDARs during NMDAR transport.

CaMKII-dependent regulation of MyoVa-NMDAR points to CaMKII’s potential role in NMDAR transport. To test this possibility, we applied a Western blotting assay to the TIF preparation, which roughly represents postsynaptic components [31, 33, 34]. As shown in Fig. S4, PMA-induced enhancement in postsynaptic NMDAR expression was reduced by AIP treatment (Fig. S4A, B). By contrast, AIP alone failed to have any effect on postsynaptic GluN1 expression (Fig. S4A, B).

The role of CaMKII in NMDAR transport was further examined using whole-cell patch-clamp recording. Both PMA and TBS-induced LTP of NMDAR EPSCs, which have been proposed to be mediated by postsynaptic NMDAR transport, were completely abolished by perfusing acute hippocampal slices with AIP (1 μmol/L; Fig. S4C–H). Taken together with the above biochemical data, our results support the notion that CaMKII plays an important role in NMDAR transport.

Under basal conditions, CaMKII associates with MyoVa, with a consensus phosphorylation site close to this binding site [53]. Notably, this CaMKII-MyoVa association was enhanced during PMA-induced NMDAR transport, as revealed by both co-IP (Fig. S5A, D) and immunofluorescence assays (Fig. S5E, F). We speculate that CaMKII regulates NMDAR transport by increasing its association with MyoVa for subsequent switching MyoVa to an active state capable of binding cargo. To test this speculation, we design a cell-permeable short peptide based on the conservative CaMKII binding domain of MyoVa (Tat-MyoVa) that is centered around the proximal tail region (1106–1124) [53], to interfere with the CaMKII-MyoVa interaction (Fig. 5C, D). The efficiency of Tat-MyoVa in blocking the CaMKII-MyoVa interaction was confirmed by co-IP experiments on hippocampal homogenates with antibodies to MyoVa (Fig. S6A, B). Control experiments were also performed to see that Tat-MyoVa (4 μmol/L) has no influence on CaMKII activities (Fig. S6C) or NMDA EPSCs (Fig. S6D, E) under basal conditions.

Tat-MyoVa inhibition in the CaMKII-MyoVa association was accompanied by a drop in the MyoVa-GluN1 association (PMA, 1.61 ± 0.09, P <0.01; PMA + Tat-MyoVa, 1.01 ± 0.07, P >0.05; PMA + Tat-Scr, 1.47 ± 0.08, P <0.05; n = 4; Fig. 5E, F), suggesting that the CaMKII-MyoVa association affects the interaction between NMDARs and MyoVa. This supports the existence of a complex composed of CaMKII, MyoVa, and NMDARs. This CaMKII-MyoVa-NMDAR complex places CaMKII in a strategic position to activate MyoVa for cargo binding and subsequently enhance MyoVa-NMDAR interaction, which in turn facilitates PMA-induced NMDAR transport. In line with this notion, interfering with CaMKII-MyoVa association using Tat-MyoVa suppressed the PMA-induced enhancement in postsynaptic NMDAR expression (GluN2A: PMA, 1.30 ± 0.07, P <0.05; Tat-MyoVa-PMA, 0.93 ± 0.03; P >0.05; GluN1: PMA, 1.39 ± 0.05; P <0.01; Tat-MyoVa-PMA, 0.98 ± 0.03, P >0.05; n = 5; Fig. 5G, H), as revealed by Western blotting assays on TIF preparations that roughly represent the postsynaptic compartment [31, 33, 34]. Furthermore, both PMA- and TBS-elicited LTP of NMDA EPSCs (Fig. 5I, N) and NMDA fEPSPs (Fig. S6F, K), which have been shown to be mediated by postsynaptic NMDAR transport, were also abolished by Tat-MyoVa (4 μmol/L). As a control, peptides with scrambled sequences failed to have any significant effects on MyoVa-NMDAR associations or the LTP of NMDA EPSCs (PMA: control, 1.75 ± 0.11, n = 6; Tat-MyoVa, 1.08 ± 0.08, n = 9; P <0.01; Tat-Scr, 1.76 ± 0.11, n = 8, P >0.05; TBS: control, 1.59 ± 0.15, n = 6; Tat-MyoVa, 1.01 ± 0.06, n = 9, P <0.01; Tat-Scr, 1.57 ± 0.09, n = 6, P >0.05; Fig. 5I, N).

Together, our results suggest that CaMKII regulates NMDAR transport through its direct interaction with MyoVa.

MyoVa Traffics NMDARs by Interacting with Rab11/FIP3

To attach to its different cargoes, the myosin uses organelle-specific receptors that often comprise Rab GTPases that are inserted in the organelle membrane by geranylgeranyl moieties [22, 55]. In particular, members of the Rab family of small GTPases have emerged as potential mediators of vesicle transport by members of the myosin V family [20, 56]. Moreover, recent studies have revealed that the small GTPase Rab11 associates with NMDARs and is required for NMDAR surface transport [57,58,59]. These findings raise the possibility that MyoVa traffics NMDARs by interacting with Rab11. To test this hypothesis, we first corroborated the binding of MyoVa to Rab11. We carried out GST pull-downs targeting different domains of the MyoVa C terminus and found that the MyoVa globular tail interacted with Rab11 from both cultured hippocampal neurons and hippocampal slices, whereas the medial tail of MyoVa did not (Fig. 6A).

The MyoVa-Rab11 association hints that MyoVa may use Rab11 as the adaptor protein for NMDAR transport. Thus, we next examined the possible alteration in the association of MyoVa and NMDAR with Rab11 during NMDAR transport. We used an anti-Rab11 antibody to precipitate the protein complex and to examine how the association between these molecules would change during PMA-induced NMDAR transport. We found that upon PMA treatment (0.5 μmol/L) the amounts of MyoVa and NMDAR subunits (GluN1 and GluN2A) that interacted with Rab11 (MyoVa-Rab11, GluN1-Rab11, and GluN2A-Rab11) were significantly increased (MyoVa, 1.33 ± 0.08, P <0.01; GluN1, 1.50 ± 0.09, P <0.01; GluN2A, 1.39 ± 0.08, P <0.01; n = 7; Fig. 6B, C). The finding of increased MyoVa-Rab11 association and GluN1-Rab11 association was further confirmed by standard co-IP experiments with antibodies to MyoVa (Rab11, 1.31 ± 0.05, n = 7, P <0.01; Fig. 6B, C) and GluN1 (1.53 ± 0.11, n = 4, P <0.05; Fig. S7A, B), respectively.

One recent study has characterized multiple novel Rab-MyoVa interactions and revealed that MyoVa interacts with multiple Rab GTPases of the endosomal and secretory pathways [60]. By performing a series of individual mutations on residues in MyoVa, the study further showed that the Q1750A mutant in humans selectively abolishes the MyoVa interaction with Rab11A, without any effect on the MyoVa interaction with other Rab GTPases. Therefore, we next determined whether disturbing MyoVa-Rab11 with the corresponding Q1725A mutation in rats (homologous to 1750 in humans) would affect the surface expression of NMDARs in cultured hippocampal neurons (Fig. S8A). In contrast to the dramatic restoration of surface NMDAR expression by MyoVa rescue (Fig. 3D, G), the expression of shRNA-insensitive MyoVa with the Q1725A mutation failed to reverse the jeopardized surface expression of NMDARs, determined by both surface biotinylation (GluN1, 1.02 ± 0.05; GluN2A, 1.06 ± 0.07; n = 7; P >0.05; Fig. 6D, E) and immunofluorescence assays (Q1725A, n = 19 neurons, 0.96 ± 0.02; Q1725A-PMA, n = 19 neurons, 0.98 ± 0.05; P >0.05; Fig. 6F, G). As a control, NMDAR surface expression without PMA stimulation was unaffected in MyoVa KD and Q1725A mutant neurons (Fig. S8B, C). In line with these results, in hippocampal slices with MyoVa KD (Fig. S8D, E) the blocked LTP of NMDA fEPSPs elicited by either PMA or TBS was not restored by MyoVa expression with the Q1725A mutation (PMA: control, 1.52 ± 0.11, n = 8; Q1725A, 1.00 ± 0.03, n = 10, P <0.01; TBS: control, 1.53 ± 0.09, n = 9; Q1725A, 0.93 ± 0.02, n = 8, P <0.01; Fig. 6H, M). Collectively, these results demonstrate that the direct interaction between MyoVa and Rab11 is crucial for NMDAR transport to the membrane.

The ability of MyoVa to regulate NMDAR transport could thus rely on its interaction with the GTPase Rab11 and its effector Rab11-family interacting protein (Rab11-FIP, see Fig. 7A) [21, 61, 62]. Although numerous downstream effectors have been identified [55, 63], the molecular processes through which Rab11 cooperatively regulates NMDARs remain unknown. We next investigated the role of Rab11-FIPs in this process. Among the five key effectors by which Rab11 regulates endosomal trafficking, FIP3 attracted our attention, as it can interact with traffic and cytoskeleton regulators [61, 64, 65] and couples to Rab11 to mediate endosomal trafficking [66,67,68]. We thus investigated whether MyoVa binds to FIP3. Co-IP with anti-FIP3 antibody revealed that the interaction of FIP3 with both MyoVa and NMDARs increased during PMA-induced NMDAR transport, suggesting that MyoVa forms a complex with FIP3 to facilitate NMDAR transport (MyoVa, 1.33 ± 0.06, P <0.01; GluN1, 1.44 ± 0.06, P <0.01; GluN2A, 1.45 ± 0.05, P <0.01; Rab11, 1.38 ± 0.08, P <0.01; n = 9; Fig. 7B, C).

If FIP3 associates with MyoVa for NMDAR transport, then the association between FIP3 and NMDARs should also increase. To test this speculation, we carried out standard co-IP experiments with antibodies against GluN1 to assess the possible alteration in the binding of FIP3 and NMDARs (Fig. 7D, E). Our results revealed an enhanced interaction between FIP3 and NMDARs upon PMA treatment (1.37 ± 0.12, P <0.05, n = 8; Fig. 7E). In contrast, we did not detect any association between GluN1 and FIP2 (Fig. S9A).

To confirm this result, we further applied an immunofluorescence assay to cultured hippocampal neurons (Fig. 7F, G). As shown in Fig. 7F, FIP3 displayed a partial colocalization with NMDAR subunit GluN1 under basal conditions. PMA treatment (0.5 μmol/L) substantially enhanced the colocalization value (control, n = 20 neurons, 0.62 ± 0.01; PMA, n = 26 neurons, 0.71 ± 0.01, P <0.01; Fig. 7G), indicating that FIP3 increases its association with NMDARs during NMDAR transport. Consistent with this, FIP3 KD decreased the co-localization of MyoVa and NMDARs, as shown by a significant reduction in the Pearson coefficient for MyoVa/GluN1 in FIP3 KD neurons (Fig. S9B, C), further demonstrating that FIP3 is involved in NMDAR transport.

We next investigated the functional role of FIP3 in NMDAR transport by assessing the effect of FIP3 KD on surface biotinylation assays and the LTP of NMDA fEPSPs, which has been shown to be mediated by postsynaptic NMDAR transport [7, 12, 32, 69]. As shown in Fig. S9D and E, PMA-induced enhancement of surface NMDAR expression was absent in FIP3 KD neurons. In hippocampal slices prepared from animals injected with AAV targeting FIP3, both the LTP of NMDA fEPSPs induced by PMA (0.5 μmol/L; control: 1.69 ± 0.07, n = 7; FIP3 KD: 1.01 ± 0.01, n = 8, P <0.01; scrambled: 1.79 ± 0.09, n = 7, P >0.05; Fig. 7I, K) and TBS were absent (control: 1.69 ± 0.06, n = 7; FIP3 KD: 0.98 ± 0.02, n = 7, P <0.01; scrambled: 1.75 ± 0.08, n = 7, P >0.05; Fig. 7L, N). As a control, the LTP of NMDA fEPSPs was intact in slices injected with scrambled shRNAs. To assess whether the impaired NMDAR LTP is due to altered basal synaptic transmission, we measured NMDAR mEPSCs in FIP3 KD neurons and found no changes in either mEPSC amplitude or frequency (Fig. S9F–H).

Together, our results suggest that MyoVa traffics NMDARs by interacting with Rab11/FIP3.

FIP3 KD Impairs Hippocampal Memory

NMDAR transport has been reported to be important for memory processes [1, 7, 12, 59, 70,71,72]. Taking into consideration the crucial role of MyoVa in NMDAR transport, we hypothesized that suppressing the MyoVa adaptor protein FIP3 would selectively disturb the coupling between NMDARs and MyoVa. As a result, memory formation could be affected. To test this hypothesis, we stereotactically injected AAV containing shRNAs targeting FIP3 in the dorsal hippocampal CA1 region (Fig. 8A) of 8-week-old rats. The efficiency of FIP3 KD was then validated (KD: FIP3, 0.33 ± 0.05, n = 5, P <0.01; Figs 8B and S9I, J). The animals were allowed to recover for two weeks after the AAV injection, and then were trained and tested on the following days. FIP3 KD rats displayed normal locomotion and motivation as indicated by the absence of any differences in distance traveled, mean speed, and mobile time in open-field analysis (Fig. S9K–N). We then conducted various memory tasks that are classically associated with hippocampal functions to evaluate the memory processes of FIP3 KD rats [13, 36,37,38, 59, 73, 74].

We first assessed associative memory by contextual and trace auditory fear conditioning tasks (Fig. S9O, P). Upon re-exposure to the training context 24 h later after conditioning, the conditioned rats with FIP3 KD displayed significantly less context-evoked freezing behavior (control, n = 14 rats, 62.46% ± 3.52%; FIP3 KD, n = 9 rats, 32.33% ± 5.13%, P <0.01; scrambled, n = 8 rats, 62.23% ± 5.56%; P >0.05; Fig. 8C). Moreover, deficits in conditioned freezing behavior during the trace interval and the inter-trial intervals were also detected in the trace fear conditioning task (control: n = 14 rats, tone, 58.82% ± 6.40%; trace 68.73% ± 6.40%; ITI, 56.60% ± 5.45%; FIP3 KD: n = 8 rats, tone, 32.76% ± 7.21%; trace, 38.96% ± 7.94%; ITI, 33.39% ± 7.72%; P <0.05; scrambled: n = 8 rats, tone, 59.90% ± 8.03%; trace, 71.62% ± 6.31%; ITI, 65.72% ± 6.51%; P >0.05; Fig. 8D), suggesting that FIP3 deficiency caused the deficits in fear memory processes.

Next, we assessed the recognition memory of FIP3 KD rats using the NOR task (Fig. 8E). It has been shown that the dorsal hippocampus plays an important role during NOR memory formation, especially when spatial or contextual information is relevant [75, 76]. We found that both control and FIP3 KD rats spent a similar amount of time exploring two identical objects during training sessions (control, n = 10 rats, 49.58% ± 1.43%; FIP3 KD, n = 12 rats, 49.34% ± 2.87%; scrambled, n = 10 rats, 51.09% ± 2.21%; P >0.05; Fig. 8F), indicating normal locomotor abilities and levels of curiosity in FIP3 KD rats to explore the objects. In retention tests, one of the objects used in the training session was replaced with a novel object, and animals were allowed to explore the vicinity for 10 min. In 1-day retention sessions, both control and scrambled KD rats displayed a significant preference for the novel object, whereas FIP3 KD rats exhibited lower preference for the novel object (control, n = 10 rats, 65.62% ± 1.67%; FIP3 KD, n = 12 rats, 52.87% ± 3.92%, P <0.05; scrambled, n = 10 rats, 67.65% ± 3.70%, P >0.05; Fig. 8G).

We also applied the NPP test, which is similar to the NOR task and is known to rely on the hippocampus, to measure the recognition memory of FIP3 KD rats (Fig. 8H). In the encoding phase, FIP3 KD rats showed performance comparable to groups of control/scrambled rats and spent similar amounts of time exploring the two identical objects (control, n = 13 rats, 50.66% ± 2.88%; FIP3 KD, n = 14 rats, 49.44% ± 2.48%; scrambled, n = 12 rats, 51.70% ± 2.90%; P >0.05; Fig. 8I). One of the objects was then moved to a novel place and rats were returned 24 h later for another round of exploration, during which the time spent investigating each object was measured. In the test phase, the FIP3 KD rats failed to show a preference for the moved object (FIP3 KD, n = 14 rats, 50.74% ± 3.01%, P <0.01; Fig. 8J). By contrast, control and scrambled rats showed a significant preference to explore the moved object (control, n = 13 rats, 68.02% ± 3.12%; scrambled, n = 12 rats, 64.38% ± 3.92%, P >0.05).

Finally, we assessed a temporal order memory task in FIP3 KD rats (Fig. 8K), a task that has been shown to depend on hippocampal CA1 function [77]. In the test phase, control and scrambled rats preferred the object explored earlier to that explored later (control, n = 11 rats, 68.31% ± 3.20%; scrambled, n = 11 rats, 68.07% ± 3.43%, P >0.05), whereas the FIP3 KD rats showed a comparable preference, indicating impaired performance of FIP3 KD rats in this task (FIP3 KD, n = 15 rats, 52.14% ± 3.45%, P <0.01; Fig. 8L).

Altogether, these results suggest that FIP3 deficiency in the CA1 region, which likely impedes NMDAR transport, causes deficits in hippocampus-associated memories.

Discussion

MyoVa and Its Interaction with NMDAR is Essential for NMDAR Transport

Dendritic transport of NMDARs depends on motor proteins that belong to the kinesin superfamily (KIFs). Specifically, KIF3B and KIF17 are respectively required for the transport of GluN2A and GluN2B subunits [10, 13, 18, 78,79,80]. Before entering the dendritic spine, NMDARs need to be released from KIF and then transferred to myosin proteins that are associated with F-actin. Corresponding to the “track switch” from microtubule to actin, a specific type of myosin protein is required for NMDAR transport in the dendritic spine. Studies of three subtypes of class V myosins (Va, Vb, and Vc) have revealed dominant expression of MyoVa and MyoVb in neuronal tissue [81, 82]. The structures of MyoVa and MyoVb in mammals exhibit substantial similarity [20, 22]. They both include a head motor domain to bind actin filaments, hydrolyzing ATP and generating force, a neck domain with six calmodulin-binding IQ motifs, and a tail domain with a proximal coiled-coil sequence and a distal globular region. We here demonstrate that activity-dependent NMDARs in response to external stimuli (PKC activation by PMA or electrical inputs by TBS), which underlies the LTP of NMDAR-mediated synaptic responses [6, 28, 44], require MyoVa and MyoVa-associated adaptor proteins. This does not contradict the absence of change in spontaneous or miniature NMDAR-mediated currents [83] or in NMDAR surface expression [84], as all these results were obtained in Flailer mutant mice under baseline conditions.

By applying biochemical or electrophysiological assays to primary cultured hippocampal neurons and acute hippocampal slices, in combination with genetic manipulation techniques and behavioral tests, we demonstrated that MyoVa is the myosin protein that implements NMDAR transport to the postsynaptic membrane (for complete schematics see Fig. S10). Given that MyoVa and MyoVb are structurally similar, we also examined the possible role of MyoVb in NMDAR transport. The absence of MyoVb association with NMDARs under both basal conditions and PMA-induced NMDAR transport does not support this possibility. This could be due to the differences in protein sequences between MyoVa and MyoVb. Overall, MyoVb exhibits 62% homology with MyoVa in the full length and 74% in the motor head domain [85]. The sequence homology between MyoVb and MyoVa is in line with their similar properties. However, the distinct sequences also have a significant impact on their respective properties. For example, the sequence of loop 2 in the motor domain of MyoVb differs considerably from that of MyoVa, and this variation has been suggested to affect the actin-binding properties of myosin [85]. Therefore, these distinctions between MyoVa and MyoVb may affect their ability to transport specific types of cargo. In this scenario, MyoVa may possess a specific sequence or structure necessary for NMDAR transport, while MyoVb may lack this particular feature.

As for the MyoVa transport of NMDAR, we provide a set of evidence to support this conclusion. Firstly, under basal conditions, MyoVa interacts with NMDARs through its cargo binding domain. Their association increased during NMDAR transport. Secondly, MyoVa KD suppressed PMA-induced NMDAR transport. Accordingly, the LTP of NMDAR fEPSPs, which is reported to be mediated by postsynaptic NMDAR transport, is absent in MyoVa KD rats. It is worth mentioning that MyoVa KD did not affect NMDAR LTD. MyoVa moves from the minus end to the plus end of actin filaments in dendritic spines. This unidirectional movement limits MyoVa to mainly mediating transport processes towards the cell surface, such as exocytosis. In contrast, NMDAR LTD at CA1 synapses requires the receptor to move away from the synaptic site, which is dependent on actin depolymerization and the lateral diffusion of NMDARs. This difference could at least partially explain the finding that MyoVa KD did not influence NMDAR LTD in the hippocampal CA1 region. Thirdly, consistent with the fact that MyoVa is a Ca²⁺-sensitive motor protein, we revealed that CaMKII, the protein kinase that can be activated by increased Ca²⁺/CaM upon NMDAR activation, regulates NMDAR transport through its direct action on MyoVa, which in turn facilitates the interaction between MyoVa and NMDAR for NMDAR transport. Fourthly, we demonstrated that MyoVa traffics NMDARs by interacting with Rab11/FIP3. Rab11/FIP3 functions as the adaptor protein to couple NMDARs and MyoVa during their transport. We report that FIP3 KD impairs various memory processes that depend on hippocampal function. Together, these findings indicate that both MyoVa and its association with NMDARs are essential for NMDAR transport.

CaMKII-Dependent Regulation of NMDAR Transport

We reveal in the present study that CaMKII is the key player that regulates NMDAR transport. CaMKII is an effector kinase central to synaptic plasticity, learning, and memory [70, 86,87,88,89,90]. During synaptic and behavioral plasticity, the activation of CaMKII by increased Ca²⁺ influx facilitates the interaction among MyoVa, NMDAR, and CaMKII. Each of the three molecules interacts with the other two and thus tends to form a complex. MyoVa establishes tighter associations with NMDARs for NMDAR transport. This interaction between MyoVa and NMDARs is closely regulated by CaMKII via its direct action on MyoVa, as the increased MyoVa-NMDAR association can be reversed by blocking CaMKII activity with AIP. Furthermore, interfering with the CaMKII-MyoVa interaction with a short peptide (Tat-MyoVa) suppressed PMA-induced enhancement in postsynaptic NMDAR expression, as well as the LTP of NMDA EPSCs mediated by postsynaptic NMDAR transport. These findings are consistent with the widely accepted notion that MyoVa is the Ca²⁺-sensitive motor protein. For example, Ca²⁺ at micromolar concentrations can cause the unfolding of MyoVa [46, 48, 49, 54, 91]. We speculate that the Ca²⁺/CaM-activated CaMKII forms a tight association with MyoVa to facilitate its action on the neighboring phosphorylation site. This phosphorylation in turn triggers a conformational switch of MyoVa that exposes the globular tail domain (GTD) and forms a tight MyoVa-NMDAR association for NMDAR transport. Alternatively, CaMKII could directly phosphorylate GluN2A for NMDAR transport, as suggested by a recent study [92].

Role of Rab11 and FIP3 in NMDAR Transport and Memory Formation

In the present study, we also identified the GTPase Rab11 as the adaptor protein that couples myosin Va with its NMDAR cargo. The ability of MyoVa to regulate NMDAR transport depends on the interaction of its GTD with Rab11 and its effector FIP3. Several lines of evidence support this conclusion. Firstly, whereas MyoVa rescued the effect of MyoVa KD, the expression of MyoVa with the Q1725A mutation that selectively abolished the MyoVa interaction with Rab11 did not. These findings support a requirement for the association of MyoVa with Rab11 for NMDAR transport. Secondly, using a combination of biochemical, immunofluorescent, and electrophysiological assays, we demonstrated that both Rab11 and FIP3 associate with MyoVa and are required for PMA- or TBS-induced transport of NMDARs to the plasma membrane. Third, we report that FIP3 KD impairs various types of memory processes that depend on hippocampal function. Specifically, the four types of hippocampal memory tasks, i.e., fear memory, NOR, NPP, and temporal order memory, were measured. Considering the essential role of NMDAR transport in memory consolidation, the memory deficit in FIP3 KD animals is consistent with the involvement of FIP3 in MyoVa-dependent NMDAR transport. We thus propose that Rab11 and FIP3 function as key adaptor proteins to couple MyoVa and NMDAR for NMDAR transport (Fig. S10).

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Acknowledgments

We thank all members of the Lu lab for their discussion and support. This work was supported by grants from the STI2030-Major Projects (2021ZD0203502) and the National Natural Science Foundation of China (31730107, 31970959, and 31671056).

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Ministry of Education (MOE) Key Laboratory of Developmental Genes and Human Disease, School of Life Science and Technology, Southeast University, Nanjing, 210096, China
Ru Gong, Linwei Qin, Yifei Bao & Wei Lu
Department of Neurosurgery, State Key Laboratory of Medical Neurobiology, MOE Frontiers Center for Brain Science, Huashan Hospital, Institute for Translational Brain Research, Fudan University, Shanghai, 200032, China
Wei Lu
Department of Neurobiology, Nanjing Medical University, Nanjing, 210096, China
Linlin Chen, Ning Wang & Wei Lu
Co-innovation Center of Neuroregeneration, Nantong University, Nantong, 226001, China
Wei Lu

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Gong, R., Qin, L., Chen, L. et al. Myosin Va-dependent Transport of NMDA Receptors in Hippocampal Neurons. Neurosci. Bull. (2024). https://doi.org/10.1007/s12264-023-01174-y

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Received: 18 July 2023
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DOI: https://doi.org/10.1007/s12264-023-01174-y

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Myosin Va-dependent Transport of NMDA Receptors in Hippocampal Neurons

Abstract

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Fear Conditioning

Novel Object Recognition (NOR) Task

Novel Place Preference (NPP) Task

Temporal Order Memory Task

Open-field Tests

Quantification and Statistical Analysis

Results

Increased Association Between MyoVa and NMDARs During NMDAR Transport

MyoVa KD Affects NMDAR Transport

CaMKII Regulates NMDAR Transport via MyoVa

MyoVa Traffics NMDARs by Interacting with Rab11/FIP3

FIP3 KD Impairs Hippocampal Memory

Discussion

MyoVa and Its Interaction with NMDAR is Essential for NMDAR Transport

CaMKII-Dependent Regulation of NMDAR Transport

Role of Rab11 and FIP3 in NMDAR Transport and Memory Formation

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