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Keywords:

  • dendritic spines;
  • ectodomain shedding;
  • nectin;
  • secretase

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

J. Neurochem. (2012) 120, 741–751.

Abstract

Synaptic remodeling has been postulated as a mechanism underlying synaptic plasticity and cell adhesion molecules are thought to contribute to this process. We examined the role of nectin-1 ectodomain shedding on synaptogenesis in cultured rat hippocampal neurons. Nectins are Ca2+-independent immunoglobulin-like adhesion molecules, involved in cell-cell adherens junctions. Herein, we show that the processing of nectin-1 occurs by multiple endoproteolytic steps both in vivo and in vitro. We identified regions containing two distinct cleavage sites within the ectodomain of nectin-1. By alanine scanning mutagenesis, two point mutations that disrupt nectin-1 ectodomain cleavage events were identified. Expression of these mutants significantly alters the density of dendritic spines. These findings suggest that ectodomain shedding of nectin-1 regulates dendritic spine density and related synaptic functions.

Abbreviations used:
CAM

cell adhesion molecule

CTF

COOH-terminal fragment

DIV

days in vitro

GFP

green fluorescence protein

Ig

immunoglobulin

MPP3

membrane palmitoylated protein 3

PAT

puncta adherentia junction

PBS

phosphate-buffered saline

SAP97

synapse-associated protein 97

SDS

sodium dodecyl sulfate

Synapses are specialized intercellular junctions that are formed when a pre-synaptic terminal contacts a post-synaptic neuron. These pre- and post-synaptic connections are mediated by numerous cell adhesion molecules (CAMs). Several CAMs undergo proteolytic shedding of their extracellular NH2-terminal domains and a subsequent intramembranous cleavage event mediated by presenilin-dependent gamma-secretase (Kim et al. 2002; Marambaud et al. 2003; Maretzky et al. 2005a,b; Uemura et al. 2006). This process may be enhanced with neuronal activity (Tian et al. 2007; Conant et al. 2010; Kim et al. 2010). Ectodomain shedding and presenilin-dependent gamma-secretase cleavage of synaptic CAMs would comprise a rapid and elegant means by which neurons might remodel spine structure in response to synaptic transmission. This change ultimately leads to long-term changes in synaptic function, which are required for higher order processes in the brain such as learning and memory (Shiosaka 2004; Zhang et al. 2005; Lee et al. 2008).

Nectin is a Ca2+-independent, immunoglobulin-like adhesion molecule involved in various cell–cell adherens junctions (Takai and Nakanishi 2003; Takai et al. 2003). Nectins are composed of four members – nectin-1, 2, 3 and 4. In the CNS, nectin-1 and 3 localize at the pre- and post-synaptic sides of puncta adherentia junctions (PAJs) formed in the CA3 pyramidal region of adult mouse hippocampus (Mizoguchi et al. 2002). At the synapse, nectin co-localizes with afadin, an F-actin binding protein (Mizoguchi et al. 2002; Lim et al. 2008). The addition of nectin-1 or nectin-3 inhibitors to cultured rat hippocampal neurons alters the cellular distribution of synaptophysin and postsynaptic density 95 (PSD-95) and decreases the size, but increases the number, of synapses (Mizoguchi et al. 2002). Mutations in the nectin-1 gene cause cleft lip/palate-ectodermal dysplasia and, in some cases, mental retardation consistent with a role in the development of ectodermally derived tissues (Suzuki et al. 2000; Sozen et al. 2001). While nectin-1 −/− and nectin-3 −/− mice show no dramatic organismal phenotypes (Inagaki et al. 2005), both mutant mice exhibit an abnormal mossy fiber trajectory and a reduction in the number of synaptic PAJs between the mossy fiber terminals and the dendrites of the CA3 pyramidal cells (Honda et al. 2006). Hippocampal neurons derived from nectin-1-null mice exhibit a reduction in spine head width and an increase in spine length (Togashi et al. 2006). Nectin-1 is initially expressed at excitatory and inhibitory synapses but is progressively lost at inhibitory synapses during their maturation (Lim et al. 2008). These data suggest that nectin plays an important role in synaptogenesis.

Nectin-1 undergoes ectodomain shedding by alpha-secretase and subsequent proteolytic processing by gamma-secretase (Kim et al. 2002, 2010, 2011; Tanaka et al. 2002). Ectodomain shedding and intramembrane cleavage of nectin-1 occur in both pre-synaptic and post-synaptic compartments in constitutive and regulated manners (Kim et al. 2010, 2011). The activity-dependent cleavage of nectin-1 mediated by alpha-secretase requires an influx of Ca2+ through NMDA receptors and an activation of calmodulin in mature cortical neurons (Kim et al. 2010). ADAM10 is the major alpha-sheddase responsible for nectin-1 ectodomain cleavage in neurons and in the brain (Kim et al. 2010).

Currently, it is not well understood how ectodomain shedding of synaptic adhesion molecules modulates synapse formation and synaptic plasticity. In this study, we examined the roles of nectin-1 shedding in synaptogenesis in cultured hippocampal neurons. Through a series of truncation mutants and alanine scanning point mutants, we identified constructs that were refractory to ectodomain shedding. By expressing these point mutants, we dissected the role of nectin-1 shedding in synaptogenesis. The results of our mutational analyses indicate that shedding of nectin-1 plays a role in the regulation of the density of dendritic spines. These observations support the critical role of nectin-1 in the formation of synapses and suggest that it functions as a regulator of synaptic plasticity through its modulation of synaptic connections.

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Animals

C57/BL6 pregnant mice and Sprague–Dawley pregnant rats were purchased from Charles River Laboratories (Wilmington, MA, USA). All animals were humanely killed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and the institutional guidelines established by the Institutional Animal Care and Use Committee at Georgetown University Medical Center (Assurance A3282-01 and protocol 10-073).

Cell lines, antibodies and reagents

HEK293 and Neuro-2A cells were purchased from American Type Culture Collection. All cell lines were maintained in Dulbecco’s modified Eagle’s medium supplied with 10% of fetal bovine serum and antibiotics (Invitrogen, Carlsbad, CA, USA). Rabbit anti-nectin-1 was prepared as described (Lim et al. 2008). This antibody was created using a synthetic peptide that corresponded to amino acids 500–513 of the C-terminal of the human nectin-1 (Fig. 1a). Mouse anti-flag (m2), anti-v5, anti-synaptophysin, and rabbit anti-actin antibodies were purchased from Sigma (St. Louis, MO, USA). Mouse anti-N-cadherin and anti-afadin antibodies were from BD Transduction Laboratory (San Diego, CA, USA). CK6 antibody was from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Texas-Red-conjugated phalloidin was purchased from Invitrogen. Gamma-secretase inhibitor X was purchased from Calbiochem (San Diego, CA, USA). All restriction endonucleases were purchased from New England Biolabs (Ipswich, MA, USA).

image

Figure 1.  Nectin-1 ectodomain shedding is developmentally regulated in mouse brains and mediated by several proteases. (a) Schematic representation of human nectin-1. Nectin-1 is a type I transmembrane protein composed of an extracellular domain, a single transmembrane domain, and a cytoplasmic domain. The sequences for the antibody binding site is shown in bold. (b) 50 μg of RIPA cortical extracts harvested from E13.5, E15.5 E17.5, P1, P6, P10, and P60 mice were separated on 4–12% Tris–Glycine gels and analyzed by immunoblotting with nectin-1, afadin, N-cadherin, and actin antibodies. (c) Relative ratios of full-length nectin-1α, 30 and 34 kDa CTFs. The intensities of full-length and CTF bands were measured by densitometric analysis using the blots with the least saturated pixels. Average densities of samples were calculated relative to the control (the highest intensity) from three independent western blots, and ratios of relative densities were determined. (d) Rat cortical neurons at 7 DIV were transduced with an adenovirus vector expressing a COOH-terminal flag-tagged nectin-1 for 24 h at an multiplicity of infection (MOI) of 500. Neurons were treated without (1ane 2), with 1 μM (lanes 3 and 4), or with 2 μM γ-secretase inhibitor X (lanes 5 and 6) for 24 h. These cells were harvested in reducing sample buffer and analyzed on a 4–12% Tris–Glycine gel. The immunoblot was probed with m2 monoclonal antibody. The western blot exhibits six distinct molecular weight bands at 90, 64, 37, 34, 30, and 24 kDa. The three molecular bands at 37, 34 and 30 kDa were increased in the presence of the γ-secretase inhibitor X while the 24 kDa band was abolished.

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Generation of constructs and recombinant adenovirus

Human nectin-1 (a gift from Dr Patricia G Spear, Northwestern University) was flag tagged at the C-terminus and inserted into the pVAX vector (Invitrogen). Nectin-1-flag was subcloned into the pShuttle-CMV vector or pAD-Track-CMV (a kind gift from Dr. Vogelstein, Johns Hopkins University School of Medicine, Baltimore MD) using these two primers (Primer 1: 5′-GTGGTCGACATGGGGCTTGCGGGCGCCGCT-3′; Primer 2: 5′-TTTGCG GCCGCCTACTTATCGTCGTCATCCTTGTAATCCACGTACCACTCCTTCTTGGA-3′). To generate V5-nectin1-flag, a set of two adjacent non-overlapping oligonucleotides containing the V5 epitope were generated. Using nectin-1 pVAX as a template, inverse PCR mutagenesis was performed to generate V5-nectin1-flag. This construct was used for a series of external truncation mutants. The external truncation mutant was generated by inverse PCR mutagenesis using two oligonucleotides, which encoded the desired truncation (Chen and Courey, 1999). The PCR product was treated with DpnI (New England BioLabs) to digest methylated parental plasmids and subsequently introduced into competent cells through transformation. Point mutants were made using the Quick change site-directed mutagenesis kit (Stratagene, La Jolla, CA, USA). Sequence fidelity of all constructs and PCR inserts was verified by sequencing. Recombinant adenoviruses were generated as previously described (He et al. 1998; Kim et al. 2010).

Brain tissue preparation

Mouse brain tissues were harvested at E13.5, E15.5, E17.5, P1, P6 and P60. To obtain sufficient amount of embryonic brain samples, we combined 2–4 brains per blot. We used 6–12 embryos for three western blot analyses. For postnatal brain samples, we used three brains for each time point. Tissue samples were homogenized with a dounce pestle in modified RIPA buffer with protease inhibitors (50 mM Tris–HCl pH 7.4, 1% NP-40, 0.25% sodium deoxycholate, 150 mM NaCl, 1 mM EDTA). After 20 min on ice, lysates were cleared by centrifugation at 10 200 g for 10 min. The detergent-insoluble pellet was solubilized by boiling in UREA/sodium dodecyl sulfate (SDS) sample buffer. Protein concentration was determined using the BCA assay (Bio-Rad, Hercules, CA, USA).

Primary hippocampal culture

For the immunocytochemistry study, neurons were prepared from E18 Sprague Dawley rat embryos as previously described (Goslin and Banker 1991; Lim et al. 2000, 2008). For experiments requiring higher cell densities, neurons were plated on polylysine-coated (1 mg/mL) 24-well tissue culture plates at a density of 1.3 × 105 cells/well in the absence of an astrocytes feeder layer. Hippocampal neurons were maintained in neurobasal medium (Gibco, Grand Island, NY, USA) containing 2% B27 supplement (Gibco) and 500 μM L-glutamine (Sigma), and the medium was changed every 3–4 days. These neurons were also treated with 5 μM cytosine arabinoside to inhibit the proliferation of non-neuronal cells. Immunostaining indicates that less than 5% of cells were non-neuronal cells at 6 days in vitro (DIV) in this culture system.

Transient transfection of HEK293 cells

Cells were transfected using LipoD293TM (SignaGen, Ijamsville, MD, USA) according to manufacturer’s protocol. For each experiment, 24 h after plating, cells were transfected with a freshly prepared DNA-LipoD293TM complex that contained the DNA of interest at a final concentration of 20 mg/mL. After 1 h incubation at 37°C, the media were exchanged with fresh Dulbecco’s modified Eagle’s medium. Transfected cells were lysed in reducing sample buffer 48 h after transfection.

Viral transduction and gamma-secretase treatment in hippocampal neurons

Recombinant adenovirus at an multiplicity of infection of 200 was applied to the middle of the culture and gently rocked. Cultures were placed into 37°C incubators and shaken every 15 min for 1 h. Cultures were washed with warm phosphate-buffered saline (PBS) and replaced with the previously conditioned media. One day after infection, transduced neurons were treated with 1 and 2 μM γ-secretase inhibitor X for 24 h.

Immunoblot and statistical data analysis

Cell homogenates for primary neurons were prepared from high-density hippocampal cultures at 21 or 28 DIV by directly solubilizing them in 100 μL reducing SDS sample buffer. Cell homogenates for HEK293 cells were prepared by directly solubilizing transfected HEK293 cells in 500 μL reducing SDS sample buffer. Samples were boiled for 10 min. Equal amounts of mouse brain tissue or equivalent volumes of cell lysates were separated on 12% Tris–Glycine gels, blotted onto nitrocellulose membranes (Bio-Rad, Hercules, CA, USA) and probed with antibodies as indicated in the figure legends. Chemiluminescence signals were captured on BioMAX (Kodak, Rochester, NY, USA) or by a cooled-CCD device, BioChemi System (UVP Inc., Upland, CA, USA). Densitometry was preformed with VisionWorksLS (UVP BioImaging Systema, Cambridge, UK). Average densities of samples were calculated relative to the controls from three independent Western blots, and ratios of relative densities were determined. Y-error bars were determined by calculating the standard deviations. p-values were calculated by utilizing T-tests to determine if the ratios of densities were significantly different. p-values of less than 0.05 are typically considered significant. Western blots captured on film were digitized by scanning on a UMAX Super Vista S-12 scanner.

Immunocytochemistry and microscopy analysis

Neurons were fixed in 4% paraformaldehyde, 4% sucrose in PBS for 30 min at 24°C. The neurons were rinsed and permeabilized in PBS containing 0.05% Triton X-100 for 30 min. Non-specific binding sites were blocked by incubation for 30 min at 24°C in Blotto-T (4% non-fat dry milk powder in 20 mM Tris, pH 7.5, 150 mM NaCl, 0.05% Triton X-100). Cells were incubated for 1 to 2 h at 24°C with anti-M2 antibodies. HEK293 cells were grown on poly-l-lysine coated glass coverslips. Cells were washed three times in ice-cold PBS (10 mM phosphate buffer, pH 7.4, 150 mM NaCl) and then fixed for 30 minutes at 4°C in a freshly prepared 3% paraformaldehyde solution in PBS. For cell surface labeling, HEK293 cells were then incubated for 1 or 2 h with 1 μg/mL V5 antibody prior to permeabilization. Excess V5 antibody was removed by washing three times with PBS. The cells were then permeabilized in 0.05% Triton X-100 for 30 min, and rinsed in PBS, containing 0.05% Triton X-100. Non-specific binding sites were blocked by incubation for 30 min at 24°C in Blotto-T. Cells were immunostained for 1 to 2 h at 24°C with nectin-1 cytoplasmic specific antibody. Immunostaining was visualized by incubation with appropriate secondary antibodies at 1 : 2000 dilutions. Cells were then washed three times in PBS-T. The coverslips were mounted in mowiol on microscope slides. Fluorescent images were captured using Scion image 1.60 software on an Axioskop camera (Zeiss) with a 100×, 1.4 N. A. lens. Images were prepared for presentation using Adobe Photoshop software. To quantitate the immunocytochemical data, pyramidal neurons were chosen randomly for image acquisition (10–12 cells each from three separate experiments). The numbers of dendritic spines were counted on 50–100 μm dendritic lengths. Spines were defined as protrusions from the dendritic stalk and contained a rounded head region. Only spines with defined edges were included in the analysis. The lengths of individual spines were measured from the tip of the spine head to the interface with the dendritic stalk. The widths of spines were taken as a diameter of the spine head perpendicular to the length of the spines. Dendritic spine length and head size were measured using Scanalytic’s IPLab software (Fairfax, VA, USA).

Electron microscopy

Ultrastructural preparation was carried out by the URMC Electron Microscopy Core (Rochester, NY, USA). Cultured hippocampal neurons were grown on glass coverslips, fixed for 30 min in 0.1 M phosphate buffered 4% paraformaldehyde, pH 7.4, rinsed in 0.1 M sodium phosphate buffer, cryoprotected in 10%, 20% then 30% sucrose, freeze/thawed on dry ice three times, and rinsed in phosphate buffer. The cells were incubated overnight at 4°C in a blocking solution (0.8% bovine serum albumin, 0.1% cold water fish gelatin, 5.0% normal goat serum and 0.05% Triton X-100) diluted in 0.1 M sodium phosphate buffer. T Cells were incubated with polyclonal rabbit primary antibody to nectin-1 for 2 days at 4°C with gentle agitation at a working dilution of 1 : 50 in the same blocking solution as stated, except with normal goat serum at 1.0%. The cells were rinsed with 0.1 M phosphate buffer 10 times at 4°C with gentle agitation and incubated overnight at 4°C with anti-rabbit biotinylated secondary antibody (Vector Laboratories, Burlingame, CA, USA) diluted 1 : 100 in the same blocking solution as the primary antibody. The cells were rinsed with 0.1 M phosphate buffer 10 times at 4°C with gentle agitation and incubated for 2 h at 24°C in extrAvidin (Sigma) diluted 1 : 100 in phosphate buffer only. The cells were rinsed at 24°C 10 times in 0.1 M phosphate buffer, three times in 0.1 M Tris–HCl buffer pH 7.4 with gentle agitation, pre-soaked in a 0.15% diaminobenzidine–HCl (Sigma) diluted in Tris–HCl for 20 min and activated with the addition of 0.03% hydrogen peroxide. The cells were developed in the working solution of diaminobenzidine–HCl for 7 min and rinsed four times in Tris–HCl buffer, two times in 0.1 M sodium cacodylate (Electron Microscopy Sciences) buffer and fixed overnight at 4°C in 2.5% glutaraldehyde diluted in sodium cacodylate buffer. The cells were post-fixed with a cacodylate buffered 1.0% osmium tetroxide solution for 20 min, dehydrated in a graded series of ethanol, infiltrated in Spurr epoxy resin overnight, embedded, polymerized overnight at 70°C and, using a “pop-off” technique (de Mesy Jensen and di Sant’Agnese 1992), the labeled cells were removed from the glass coverslips using liquid nitrogen. The ‘pop-off’ blocks containing labeled cells were inverted and examined under a light microscope to determine the area to be trimmed and thin sectioned for electron microscopy. Sections of the labeled ‘popped-off’ cells were thin sectioned at 80 nm with a diamond knife, placed onto formvar coated 100 mesh nickel grids, and stained with aqueous uranyl acetate and lead citrate. The grids were examined with a Hitachi 7100 Electron Microscope and digital images were captured using a Megaview III digital camera (Soft Imaging System, Lakewood, CA, USA).

Results

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Ectodomain cleavage of nectin-1 occurs in vivo and in vitro and is regulated by multiple sheddases

As nectins are present in synapses in vivo and in vitro (Mizoguchi et al. 2002; Lim et al. 2008) and localize to PAJs, which are adhesive sites between pre- and post-synaptic membranes (Honda et al. 2006), we surmised that shedding of nectins may be a regulated event influencing synaptic adhesion under conditions of synaptic plasticity. To investigate to what extent nectin-1 ectodomain shedding occurred during developmental synaptogenesis, we examined levels of full-length nectin-1 and COOH-terminal fragments (CTFs) in embryonic and postnatal mouse brains. We performed Western blotting analysis of mouse cortices sampled at E13.5, E15.5, E17.5, P1, P6, P10 and P60 using a polyclonal nectin-1 antibody directed against the COOH-terminal of nectin-1 (Fig. 1a), and commercial antibodies against afadin and beta-actin. Consistent with earlier studies (Schmitt et al. 1977; Lazarini et al. 1991), actin protein levels appeared to decline evenly throughout development (Fig. 1b). Levels of full-length nectin-1 uniformly increased during mouse brain embryonic and postnatal development (Fig. 1b and c). However, levels of the 30 kDa nectin-1 CTF displayed a bimodal distribution peaking at E13.5 and P10 (Fig. 1b and c). Interestingly, two additional species of 34 and 37 kDa also appear around E13.5 and P1 respectively, and their levels increase in a time dependent manner, suggesting these two additional CTFs are also derived from the activity of two unidentified secretases expressed in different developmental stages. These two bands are more prominent in P60 mice, while the levels of 30 kDa species decreased, suggesting that ADAM10 is competing with two unidentified secretases for nectin-1 cleavage events, or that ADAM10 may undergo a shift in substrate preference. However, we were not able to detect a 24-kDa fragment, which is a γ secretase product (Kim et al. 2002, 2010). This is possibly caused by instability or low abundance of the 24 kDa fragment. Interestingly, nectin-1 and afadin were detected in the early developmental stages, suggesting the important role of their association in the development of the CNS in addition to the formation of synapses (Fig. 1b).

Next, we addressed whether ectodomain shedding of nectin-1 occurs in cultured rat cortical neurons. Technical difficulties preclude detection of endogenous CTFs, particularly the 30 and 24 kDa, forms as a result of low levels in neurons. To circumvent this, we over-expressed a C-terminal flag-tagged nectin-1 in neurons using an adenovirus vector. The immunoblot of cell lysates exhibited six distinct bands consisting of protein species of 100, 64, 37, 34, 30, and 24 kDa (Fig. 1d). The presence of 37, 34, and 30 kDa species indicates that these primary neurons process nectin-1 in a similar manner in vivo and that there are several sites that undergo processing. Interestingly, when transduced neurons were treated with 1 and 2 μM γ-secretase inhibitor X, the 24 kDa band was reduced and the three other bands (37, 34, and 30 kDa) were markedly increased, indicating that the 24 kDa is a product of γ-secretase and that the three CTFs are putative substrates for γ-secretase (Fig. 1d; long exposure). These data indicate that processing of nectin-1 occurs by multiple endoproteolytic steps, one of which is catalyzed by γ-secretase.

External truncation mutants reveal the minimal domains containing nectin-1 ectodomain cleavage sites

To identify the minimal ectodomain region harboring cleavage sites, we generated a panel of progressively truncated mutants (Fig. 2). Five external domain truncation mutants, deleted between amino acids 28–349, were engineered. Nectin-1 is a type I membrane glycoprotein, which consists of three immunoglobulin (Ig)-like domains, followed by a transmembrane domain and a small cytoplasmic tail (Takai et al. 2003). In these constructs, the first and second Ig-like loops were completely deleted, whereas the third Ig-like loop was progressively shortened. The first Ig-like domain participates in cis-dimer (Fabre et al. 2002; Krummenacher et al. 2002; Takai et al. 2003) and trans-dimer formation (Miyahara et al. 2000; Reymond et al. 2001; Sakisaka et al. 2001; Krummenacher et al. 2002; Momose et al. 2002). The second Ig-like domain is necessary for cis-dimer formation (Momose et al. 2002). However, the function of the third Ig-like domain is unknown. To facilitate the analysis, a V5 epitope tag was appended after the signal sequence and a flag tag was added to the C-terminus (Fig. 2).

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Figure 2.  Schematic diagram of the nectin-1 ectodomain deletion mutants. A panel of progressive external truncation mutants was generated by inverse PCR mutagenesis. V5 and flag tags were inserted at the N-terminus right after the signal sequence and C-terminus respectively.

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First, we analyzed whether each fusion construct was expressed on the cell surface since shedding occurs at this location. HEK293 cells were transiently transfected with wild type, epitope tagged, full-length nectin-1, and truncation mutants. Cell surface expression was detected with a V5 antibody in the absence of detergent permeabilization and total protein expression was examined with cytoplasmic specific nectin-1 antibody after cell membrane permeabilization. Double-tagged full-length nectin-1 was detected on the cell surface (Fig. 3a). This indicated that V5 and flag epitope tags did not disrupt the cell surface expression of nectin-1. Surface expression of four truncation mutants, Δ28–250, Δ28–300, Δ28–312, and Δ28–324 were well detected with anti-V5 antibody (Fig. 3a). However, the surface expression of the truncation mutant of Δ28–349 was not detected although the immunostaining with cytoplasmic specific antibody exhibited staining (Fig. 3a). We speculate that because this construct has a very short extracellular domain, the V5 epitope tag may be buried and inaccessible to surface applied antibody.

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Figure 3.  Characterization of truncation mutants in cell lines. (a) HEK293 cells were transfected with nectin-1 and mutants. Cells were fixed 48 h after transfection. Cell surface expression (red, first column panels) was examined with CK6 for wild type and anti-V5 antibody for truncated mutants in the absence of detergent permeabilization, then total protein expression (Green, second column panels) was examined with cytoplasmic specific nectin-1 antibody after cell membrane permeabilization with 0.1% Triton X-100 extraction. The cell nucleus was visualized with DAPI (blue, third column panels). Merged images are shown on the right panel. Scale bar = 10 μm. (b) HEK293 cells were transfected with each mutant. After 24 h, cells were lysed in reducing sample buffer and analyzed on a 4–20% Tris–Glycine gradient gel. Samples were transferred to nitrocellulose, and the blot was probed with V5 epitope antibody to distinguish truncation mutants from CTFs. The results shown are representative of three independent experiments. (c) The membrane shown in (b) was stripped and reprobed with a cyto-specific nectin-1 antibody. The smallest molecular weight fragments recognized by v5 antibody are indicated by * to distinguish CTFs from truncation mutants.

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To define the minimal region that may contain the cleavage sites within the ectodomain of nectin-1, proteins from the truncated mutants were analyzed by Western blotting. Each construct was expressed in HEK293 cells and harvested 24 hr post-transfection. In cells transfected with wild-type nectin-1, 30, 34, and 37 kDa CTFs were detected (Fig. 3b). We named the 30 kDa species α-CTF, and the 34 kDa species CTF-1. In Δ28–250 transfected cells, all three CTFs were detected, but in Δ28–300 transfected cells, we could verify the presence of the 30 and 34 kDa CTFs but not for the 37 kDa band because of interference of the truncated mutant (Fig. 3a and b). CTF-1 was not detected in Δ28–312 and Δ28–324, indicating that CTF-1 cleavage site may be located between amino acids 300 to 324 (Fig. 3c). The α-CTF was detected in Δ28–324, but we could not verify the presence of α-CTF in Δ28–349 because of interference of the truncated mutant (Fig. 3c). Therefore, we concluded that the α cleavage site is also located between amino acid 324–351. All these external deletion mutants produced a 24-kDa fragment, which is a γ-product that we named γ-CTF. This form is readily detected in longer exposure films (Fig. 3c; bottom blot) and, as predicted, γ-secretase cleaves its substrate within the transmembrane domain. The external truncation mutant analysis reveals that the ectodomain shedding is not dependent on cis- and trans-dimerization since the first and second Ig-like domains were deleted in these constructs. These loops are not necessary for nectin-1 processing.

Point mutants identify critical amino acids important for ectodomain nectin-1 processing

To identify the residues that are necessary for cleavages of alpha-CTF and CTF-1 cleavage, we utilized site-directed alanine scanning mutagenesis. We mutated each residue between amino acid 301–351 to alanine, with the exception of residues 308, 315 and 346. Residues 308 and 315 were mutated to leucine, and residue 346 was mutated to isoleucine. Each of these mutant polypeptides was expressed in HEK293 cells and cell lysates were collected for western blot analysis. We found 10 amino acids that affect nectin-1 processing (Fig 4a). Mutation of amino acid residues 310 and 311 substantially reduced both 30 and 34 kDa CTFs (Fig. 4b and d). Mutation of amino acid residues 303, 313 and 323 caused an accumulation of 50–60 kDa intermediates but still produced 30 and 34 kDa CTFs at low levels (Fig. 4b and d). Interestingly, mutation I303A shifted the full-length nectin-1 to a higher molecular weight. This mutation may affect post-translational modification. Mutation of amino acid residues 335, 336, 339, 342, and 343 selectively inhibited or substantially reduced the production of CTF-1 but not α-CTF, suggesting that these residues play an important role in interactions with an unidentified protease. Interestingly mutation of amino acid residue 336 exhibited another band just above the α-CTF (Fig. 4b; indicated by a red dot in the bottom blot), suggesting that this residue may have changed the cleavage site for an unidentified protease. These point mutational analyses also indicate that the CTF-1 cleavage event is not required for the α-cleavage event. The mutation of residue 329 specifically reduced high molecular full-length nectin-1 presumably because of altering N-glycosylation (Fig. 4c). However this mutation did not affect the formation of CTF-1 and α-CTFs (Fig. 4b and d). The remainder of the mutants produced CTFs identical to that of wild-type nectin-1 (Figure S1). We also analyzed these mutants in neuro-2A neuronal cells and obtained similar results (Figure S2), suggesting that nectin-1 is processed by similar or identical sheddases in various cells. These data indicate that ten residues may play an important role in CTF-1 and/or α-cleavages, perhaps in positioning the substrate sites for important positioning for ADAM10 or unidentified cleavage enzyme(s).

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Figure 4.  Analysis of nectin-1 point mutations that affect nectin-1 processing. HEK293 cells were transfected with nectin-1 and mutants in triplicates. The cells were collected in reducing sample buffer 24 h after transfection and analyzed by western blotting. The blots were probed with anti-nectin-1 antibody. Chemiluminescence signals were detected by a cooled-CCD device and densitometry was performed on CTF-1 and α-CTFs. Protein levels for each mutant were normalized to wild-type nectin-1. The results shown are representative of three independent experiments. (a) The schematic diagram of nectin-1. The mutated residues are indicated by circles, and those mutations that affect nectin-1 processing are indicated by red color. (b) Western blotting of point mutations that alter nectin-1 processing. The doublet of mutation of amino acid residue 336 is indicated by an asterisk. The results shown are representative of six independent experiments. (c) Western blotting of wildtype and N329A. The molecular weight of mature nectin-1 was substantially reduced by mutation of amino acid residue 329. (d) Densitometer analysis of α-CTFs and CTF-1 of mutants is shown. The graph shows an average of three independent experiments. *< 0.05.

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Expression of point mutants alters dendritic spine density

We addressed how these cleavage refractory mutants affect the synaptogenesis in rat hippocampal neurons, in which nectin-1 localizes to both pre- and post-synaptic compartments shown by an electron microscopy study (Figure S3) and preferentially to excitatory synapses in mature neurons (Lim et al. 2008). We over-expressed point mutants using Ca2+ phosphate mediated transfection at early neuronal development stage (3–4 DIV) and examined synaptogenesis at 21 DIV.

First, we transfected neurons with green fluorescence protein (GFP) alone, then co-expressed nectin-1, T310A, or Y311A with GFP. GFP-expressing neurons exhibit stubby-like spines, no longer than 1–2 μm in length, and some filopodia-like protrusions (Fig. 5a). Flag-tagged nectin-1 expressing neurons exhibited mushroom like spines (Fig. 5b) with slightly bigger spine heads than that of GFP neurons (Fig. 5g). However, over-expression of nectin-1 did not increase the density of dendritic spines (Fig. 5e), but increased the length of spines (Fig. 5f). Neurons expressing T310A or Y311A exhibited a dramatic increase of dendritic protrusions (Fig. 5c and d respectively). There were 3- to 5-fold increases in the density of dendritic spines (Fig. 5e). Both mutants did not affect the length of dendritic spines although they increase the size of spine head (Fig. 5f and g). We also examined three other point mutations including S323A, P339A, and H342A. These mutants increased the number of spines but less significantly than did T310A and Y311A (Fig. 5e). These data demonstrate that the shedding of nectin plays an important role in the density of dendritic spines.

image

Figure 5.  The effects of point mutants in dendritic spine density. Neurons 4 DIV were transfected with vectors expressing GFP alone, and co-expressing nectin-1-flag, T310A or Y311A with eGFP. Cells were fixed 14–15 days after transfection. (a) GFP transfected neurons were stained with synaptophysin (red, middle panel). (b) Neurons coexpressing GFP (green, left panel) and nectin-1 with flag tag (red, middle panel). (c) Neurons coexpressing GFP (green, left panel) and T310A (red, middle panel). (d) Neurons coexpressing GFP (green, left panel) and Y311A (red, middle panel). Right panels are superimposed images of the left and middle panels. Scale bar = 5 μm. (e) The density of dendritic spines per 10 μm is shown. To quantitate the total number of protrusions, 10–12 pyramidal neurons were analyzed from three independent experiments. *< 0.05. (f) The lengths of spines were measured from 200–250 spines for each experiment. The spines were defined as protrusions from the dendritic stalk that contained a rounded head region. Only spines with defined edges were included in the analysis. (g) The widths of spines were measured from the same pools used in panel (f). The widths of spines were taken as a diameter of the spine head perpendicular to the length of the spine. *< 0.05.

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The marked increase of dendritic spines in T310A and Y311A expressing neurons suggested that these T310A and Y311A mutations might prolong the nectin-1 function by blocking its ectodomain shedding. This, in turn, leads to a decrease in the turnover rate of spines or an increase in stability of pre-existing spines. Thus, ectodomain shedding of nectin-1 by sheddases is a regulatory step to turn off nectin function in forming adherens junctions including synapses.

Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

In this study, we showed that the processing of nectin-1 occurs by multiple endoproteolytic events both in vivo (Fig. 1b) and in vitro (Fig. 1d), and that the shedding of nectin-1 is developmentally regulated in mouse brains (Fig. 1b), implying the importance of nectin-1 processing during brain development. Nectin-1 localizes to both post-synaptic and pre-synaptic membranes (Figure S2), and ectodomain shedding and intramembrane cleavage of nectin-1 occurs in both compartments (Kim et al. 2011). Therefore, the proteolytic machinery formed by metalloproteases and gamma-secretase must be present and functioning in both sides of the synapse to cleave nectin-1. Recently, we reported that nectin-1 recruits membrane palmitoylated protein 3 (MPP3) to cell–cell contact sites, mediated by a postsynaptic density-95/discs large/zonula occludens-1 (PDZ)-binding motif at the carboxyl terminus of nectin-1 (Dudak et al. 2011). ADAM10 is one of the major sheddases responsible for ectodomain shedding of nectin-1 (Kim et al. 2010), resulting in the generation of a 30 kDa CTF. ADAM10 interacts with synapse-associated protein 97 (SAP97) (Marcello et al. 2007), which increases the cell surface expression of ADAM10. SAP97 interacts with MPP3 through the L27 domains of MPP3 (Karnak et al. 2002). It is possible that the nectin–MPP3 complex may play a role in the recruitment of the ADAM10-SAP97 complex to cell–cell contact sites, and that this recruitment of ADAM10, in turn, increases ectodomain shedding of nectin-1 and other cell adhesion molecules associated with nectin-based adhesion junctions. However, besides ADAM10, it is still unknown which secretases are responsible for the shedding of nectin-1 and whether these cleavage events are sequential or independent of one another in vivo.

Using molecular approaches, we identified the regions containing the two main cleavage sites in the extracellular domain of nectin-1 (Fig. 3b and c). By alanine scanning mutagenesis, we identified point mutations that disrupt nectin-1 cleavage, substantially reducing alpha-CTF and/or CTF-1 (Fig. 4b). Over-expression of wild type nectin-1 had no effect on the dendritic spine density, whereas the expression of cleavage resistant mutants altered the density of dendritic spines (Fig. 5a). These observations suggest that ectodomain shedding of nectin-1 is a regulatory step to turn off nectin function at synapses. At synapses, nectin-1 shedding mediated by ADAM10 is regulated by the activation of NMDA receptors (Kim et al. 2010) or by chemical long-term potentiation (Kim et al. 2011). These data suggest that secretase-mediated cleavage of synaptic adhesion molecules, such as nectin-1, may allow the activated synapse to undergo either an increase or decrease in spine size or density observed during induction of long-term potentiation and long-term depression.

Interestingly, three residues (T310, Y311, and S323) that most robustly affect nectin-1 processing contain hydroxyl groups. Simple removal of the hydroxyl group by alanine mutation suggests a critical role of the hydroxyl groups themselves, perhaps through participation in hydrogen bonds. These residues could play a critical role in intramolecular hydrogen bonding in positioning the substrate for proteolytic cleavage of nectin-1. However, it is not clear whether these mutations reduce interactions with secretases or simply reduce the ability of secretases to cut. Thus, further investigation is required. Nevertheless, our studies indicate the role of nectin-1 shedding in dendritic spine morphogenesis and perhaps in related synaptic functions.

It is known that soluble extracellular domains of CAMs play roles in various physiological functions (Herreman et al. 1999; Dihne et al. 2003; Kalus et al. 2003; Tian et al. 2007; Conant et al. 2010). Therefore, the soluble fragments of nectin-1 generated by multiple sheddases may also participate in various biological functions, since different soluble fragments interact with different ligands to generate distinct signals. Ectodomain shedding of nectin-1 generates at least two soluble ectodomains derived from either α- or CTF1-cleavage events. These two soluble forms may have different physiological functions based on the observation of APP processing. APP undergoes independent α- or β-secretase cleavage events, releasing two soluble forms of APP: APPsα or APPsβ respectively. APPsα exhibits neurotrophic and neuroprotective properties (Furukawa et al. 1996; Meziane et al. 1998; Stein et al. 2004), whereas APPsβ seems to have a proapoptotic function (Nikolaev et al. 2009). The nectin-soluble ectodomains may act as a signaling protein where they bind to their receptors and mediate signals in an autocrine or paracrine fashion. In addition to the signaling function, these nectin-soluble ectodomains could act as regulators in the postulated cell-cell interaction by binding to either full-length nectin-1 or -3 through homo- or hetero-trans-dimerization. It has been shown that a fusion protein composed of the ectodomain of nectin-1 and the Fc portion of IgG trans-interacts with cellular nectin-1 and nectin-3, and induces filopodia and lamellipodia by activating Rap1, Cdc42, and Rac small G-proteins through the activation of c-Src in Madin-Darby canine kidney (MDCK) and fibroblast cells (Kawakatsu et al. 2002, 2005; Honda et al. 2003a,b). Therefore, binding of nectin ectodomains to full-length nectins may generate similar signaling pathways, resulting in local junctional remodeling. Interestingly, nectin-1 has three isoforms: α, β, and γ. The β form is missing 59 amino acid residues including a conserved postsynaptic density-95/discs large/zonula occludens-1 (PDZ)-binding motif of four amino acid residues at the carboxyl terminus, whereas the γ form is the secreted protein, which lacks the transmembrane region and the entire C-terminus. However, the biological role of the γ form is completely unknown. The presence of nectin-1γ in nature highly suggests that receptor(s) may exist for the soluble nectins and that the association between receptor(s) and soluble nectins may play an important role in cellular functions. It will be interesting to determine what physiological roles the shed ectodomains of nectins and nectin-1γ play and whether they exhibit different roles in vivo. Future studies will be necessary to determine whether extracellular domains of nectin-1 play a role in synaptic plasticity.

In conclusion, nectin-1 ectodomain shedding is regulated by multiple sheddases in vitro and in vivo, and ectodomain shedding of nectin-1 is a regulatory step to turn off nectin function at synapses. In turn, this process modulates the maintenance of dendritic spine densities in rat hippocampal cultures.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

We thank Dr Katherine Conant for comments on the manuscript. This work was supported by NIA RO1 AH027233 to HJF.

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  6. Acknowledgements
  7. References
  8. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Figure S1. Analysis of nectin-1 point mutations from amino acid residues 301 to 351. HEK 293 cells were transfected with each mutant. Twenty-four hrs after transfection, cells were lysed in reducing sample buffer and analyzed on 12% SDSPAGE. Samples were transferred to nitrocellulose, and the blots were probed with antinectin-1 cyto-specific antibody. All experiments were repeated five times.

Figure S2. Analysis of nectin-1 point mutants in neuro-2A cell lines. Neuro-2A cells were transfected with nectin-1 point mutations that alter nectin-1 processing in HEK 293 cells. The cells were collected in reducing sample buffer 24 hrs after transfection and analyzed by Western blotting. The blot was probed with antinectin-1 cyto-specific antibody.

Figure S3. Nectin-1 localizes to both pre- and postsynaptic sites. A. Neurons at 28 DIV were examined at the ultrastructural level by electron microscopy. Synapses were identified by the presence of thickened presynaptic and postsynaptic specializations with intervening dense material indicated by arrows. Nectin-1 immunostaining was observed both in pre- and postsynaptic sites using antibody against the intracellular domain of nectin-1. Over 30 synapses were examined and the immunoreactivity of nectin-1 was observed in all synapses. B. Nectin-1 immunostaining was abrogated by incubation of the antibody with its cognate antigenic peptide.

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