WT C57BL/6J mice were used for all experiments. Animal procedures were performed in accordance with the guidelines of the Canadian Council for Animal Care and the Montreal General Hospital Facility Animal Care Committee.

Recombinant SPARC protein was purchased from R&D Systems (Mouse; Cat no. 942-SP-050). In all experiments, a concentration of 0.5 μg/ml SPARC was used. This concentration was determined by titration of different SPARC concentrations and analysis of its effect on surface GluA1 levels (Supplementary Figure 1).

Dissociated mouse hippocampal neurons were grown suspended above astrocyte feeder cultures using a method modified from Kaech and Banker, 2006 published previously (described in detail in Jones et al., 2012). An astrocyte feeder layer was prepared by plating mouse astrocytes at 20,000 cells/cm² onto 12-well dishes coated with poly-D-lysine (0.1 mg/ml) (Sigma) in Minimal Essential Medium containing Earle’s salts and L-glutamine supplemented with 10% horse serum, 0.6% glucose, and 1% penicillin/streptomycin (Invitrogen). Media was replaced with Neurobasal-A medium supplemented with 2% B27, 1 mM GlutaMax, and 1% penicillin/streptomycin (Invitrogen) 24 h before neuron dissection. After 5 days of growth of astrocyte cultures, hippocampal neurons were isolated from P0 pups. Hippocampi were dissociated by treatment with papain (0.1% papain, 0.02% BSA in Neurobasal-A medium, 15 min at 37°C) followed by trituration with a fire-polished glass pipette in Neurobasal-A media containing trypsin inhibitor (1%, Sigma) and BSA (1%). Neurons were plated onto poly-L-lysine-coated coverslips (0.1 mg/ml Sigma) at a density of 20,000 cells/cm² for a period of 3 h before transfer to dishes containing the astrocyte feeder layers. Coverslips were suspended above the feeder layer on wax dots adhered to the bottom of the culture dish well.

We used a protocol described previously (Stellwagen et al., 2005; Jones et al., 2011) to visualize surface GluA1 and GluA2-containing AMPARs under non-permeabilizing conditions. Briefly, 14-day-old cultures were washed with ice-cold PBS then fixed with 4% paraformaldehyde/0.1 M phosphate buffer for 10 min on ice. The coverslips were blocked in 5% BSA and incubated with an antibody against the N terminus of GluA1 (Millipore clone RH95 1:30) and GluA2 (Millipore clone 6C4 1:200) for 1 h. To analyze surface GluA1 and GluA2 levels, blinded images were thresholded using ImageJ (NIH) to exclude background noise and values were held constant across each experiment. For each neuron, we measured a 50 μm region of three dendrites (at least one body distance away). The average pixel intensity was recorded for each dendrite (>70 measured per condition for each experiment) and used to compare surface GluA1 and GluA2 levels across conditions.

To examine the surface expression of AMPAR subunits GluA1 and GluA2, neurons were gently washed twice with ice-cold PBS (containing Ca²⁺ and Mg²⁺) and then incubated with Sulfo-NHS-SS-Biotin/PBS (0.2 mg/ml) for 30 min at 4°C. Cells were then washed twice with 100 mM glycine/PBS to quench the biotin reaction, followed by lysis on ice with PBS/0.1% Triton X-100/0.1% SDS supplemented with protease inhibitors and sodium orthovanadate. Protein levels were assessed by BCA assay and normalized with lysis buffer to ensure equal input into the immunoprecipitation. An aliquot of lysate was kept for analysis of total receptor levels and the remainder was incubated with streptavidin beads (Sigma) for 2 h at 4°C on a rotating platform. Biotinylated proteins were eluted with 3X sample buffer, resolved by SDS-PAGE and analyzed by immunoblotting with GluA1 (Millipore clone RH95; 1:9000) and GluA2 (Millipore clone 6C4; 1:2000) antibodies. Immunoblotting with a GAPDH antibody demonstrated that only cell surface proteins were isolated with this method. Densitometric quantification was carried using ImageJ (NIH). Cell surface receptor values were adjusted relative to total receptor levels (GluA1/GluA2).

Organotypic hippocampal slices were prepared as described previously (Stoppini et al., 1991; Murai et al., 2003; Zhou et al., 2007). Briefly, 300 μm slices from postnatal day 6–7 mouse pups were made with an automatic tissue chopper and transferred onto semiporous tissue culture inserts (0.4 μm pore size; Millipore) containing media [50% minimum essential medium, 25% horse serum, 25% HBSS, 6.5 mg/ml D-glucose, and 0.5% penicillin/streptomycin (Invitrogen)]. Medium was replaced three times per week. For immunofluorescence (IF) labeling slices were fixed in 4% paraformaldehyde/0.1M phosphate buffer for 30 min, washed with Tris-buffer saline (TBS) and blocked for 1 h at room temperature in 10% donkey serum (Jackson Immunoresearch Laboratories) in TBS containing 0.2% Triton X-100 (TBS-T) and then incubated with a goat anti-SPARC antibody (R&D Systems AF942 1:300) overnight. The next day, slices were washed four times with TBS-T before incubation with donkey anti-goat Alexa-568 (1:1000 Molecular Probes) for 1 h at room temperature. Slices were then washed four times with TBS-T and imaged with confocal microscopy. For synaptosomal preparations, slices were lysed in Syn-PER Synaptic Protein Extraction Reagent Thermo Scientific #87793, characterized in Palavicini et al., 2013; Wang et al., 2016 and in Cook et al., 2014. The synaptosomal fraction was extracted as per supplier’s directions. Briefly, slices were scraped into 200 μl of Syn-PER containing protease inhibitors and dounced using a glass homogenizer. The homogenate was centrifuged at 1200 g for 10 min at 4°C to remove cell debris. The supernatant (‘crude’ fraction) was then centrifuged for 20 min at 15,000 g at 4°C. The resulting supernatant was removed, and the pellet (‘synaptic’ fraction) was resuspended in Syn-PER reagent. The protein concentration of each fraction was determined by BCA assay. The crude and synaptic fractions were diluted in 3X sample buffer, resolved by SDS-PAGE, and analyzed by immunoblotting with the following synaptic antibodies: mouse anti-GluA1 (Millipore clone RH95; 1:9000), mouse anti-GluA2 (Millipore clone 6C4; 1:2000), mouse anti-vGlut1 (Neuromab clone N28/9; 1:7000), mouse anti-PSD-95 (Neuromab clone K28/43; 1:300,000), mouse anti-GluN1 (BD Pharmingen; 1:3000) and mouse anti-GAPDH (loading control; Millipore clone 6C5 1:100,000). For other immunoblot analysis, slices were lysed on ice in Triton lysis buffer (20 mM Tris pH 7.4, 137 mM NaCl, 25 mM beta-glycerophosphate, 2mM EDTA, 1% Triton X-100, 10% glycerol, and 0.1% SDS supplemented with protease inhibitors and sodium orthovanadate). Lysates were centrifuged at 13,000 rpm for 10 min at 4°C to pellet cell debris. Supernatants were diluted with 3X sample buffer, resolved by SDS-PAGE, and analyzed by immunoblotting with antibodies anti-SPARC (R&D Systems AF942; 1:3000) and GAPDH as a control for protein levels.

Following synaptosomal purification, we carried out co-immunoprecipitation experiments for GluA1 and GluA2 using a previously published protocol (Kang et al., 2012; Cook et al., 2014). The immunoprecipitated AMPAR complexes were diluted in 3X sample buffer, resolved by SDS-PAGE and analyzed by immunoblotting using GluA1 and GluA2 antibodies (as above). The unbound supernatant fraction from the co-immunoprecipitation was blotted for GAPDH to ensure equal protein loading.

Miniature excitatory postsynaptic current (mEPSCs) were recorded by whole-cell patch recordings made on CA1 pyramidal cells from mouse organotypic hippocampal slices after 13–19 DIV. Slice cultures were maintained at 32°C in a carbogenated (5% CO₂/95% O₂) interface chamber under artificial cerebrospinal fluid (ACSF) perfusion. Recordings were made with an Axopatch 200B (Molecular Devices) using low-resistance pipettes (2–5 MΩ) containing 140 mM K-gluconate, 5 mM NaCl, 2 mM MgCl₂, 0.1 mM CaCl₂, 1.1 mM EGTA, 7 mM Na₂-phosphocreatine, 10 mM HEPES, 4 mM Mg-ATP, and 0.4 mM Na₃-GTP. Cells were held at -65 mV with 1 μM TTX and 50 μM picrotoxin. Membrane currents were monitored in voltage-clamp mode using pClamp software (Molecular Devices). Series resistance was compensated and checked before and after every recording period; cells containing >20% resistance change were excluded from the analysis. mEPSCs were collected over a 5–10 min period from control or 48h SPARC-treated hippocampal slices (0.5 μg/ml), with or without Philanthotoxin 433 (5 μM). Custom written software (Courtesy Pablo Mendez, University of Geneva, Medical School, Switzerland) was used for analyzing mEPSC events. Briefly, individual events were detected with a threshold-shape process. Detection criteria based on threshold was adjusted to ignore slow membrane fluctuations and electric noise. Events smaller than -4 pA were discarded. To obtain the cumulative mEPSC amplitude, all event amplitudes were collected for each cell recording. An average amplitude was then calculated by cumulating the first 100 events from all experiments. The mean mEPSC frequency was obtained by dividing the number of event by the duration of a given recording. An average frequency was then calculated across experiments. All data are presented as mean ± SEM, and comparisons were made using ANOVA with post hoc Holm–Sidak test as specified. Differences were considered to be statistically significant ^∗∗∗p < 0.001.

Surgery leading to focal cerebral ischemia was conducted as described previously (Zarruk et al., 2012) and a variant of a model described earlier (Chen et al., 1986; Liu et al., 1989). In brief, animals were put under anesthesia with isoflurane in O₂ (0.5 L/min) during the whole procedure. During surgery, body temperature was maintained at 37.0 ± 0.5°C using a homeothermic system with a rectal probe (Harvard apparatus). A small craniotomy was made over the trunk of the left middle cerebral artery (MCA) and above the rhinal fissure. The permanent middle cerebral artery occlusion (MCAO) was done by ligature of the trunk of the MCA just before its bifurcation between the frontal and parietal branches with a 9-0 suture. Complete interruption of blood flow was confirmed under the operating microscope. Additionally, the left common carotid artery was then occluded. Mice in which the MCA was exposed but not occluded served as sham-operated controls (sham). After surgery, animals were returned to their cages with free access to water and food. No spontaneous mortality occurred after MCAO. For IF labeling, mice were perfused with 4% paraformaldehyde and 14 μm thick coronal sections were prepared by a cryostat (Leica, CM3050 S) and processed attached to slides. After 1 h incubation in blocking solution (10% donkey serum in TBS-T), sections were incubated overnight at 4°C with the following primary antibodies: SPARC (1:300), GFAP (1:500) and IBA-1 (1:500). Slides were washed three times with TBS and primary antibodies were revealed with 2 h incubation with Alexa Fluor secondary antibodies (1:300, 10% donkey serum, TBS-T) and confocal microscopy. To assess which cell type upregulated SPARC post-MCAO, we used ImageJ (NIH) to count the total number of microglia (IBA+) and astrocytes (GFAP+) and calculate the proportion of each cell type expressing SPARC.

For immunoblotting, the cortex of the ipsilateral MCAO and sham control C57BL/6J mice was dissected at different time points (n = 3 mice per group) following intracardiac perfusion with PBS. Total protein was extracted with 1% Nonidet P-40 (Sigma), 1% sodium deoxycholate (BDH Chemicals), 2% SDS, 0.15 M sodium phosphate (pH 7.2), 2 mM EDTA, containing a mixture of protease inhibitors (Roche Diagnostics) as described previously (Zarruk et al., 2015). Naive animals served as controls. Samples (25 μg) were separated by SDS-PAGE gels 4-12% (Novex, life Technologies), transferred to polyvinylidene fluoride membrane (Millipore, Billerica, MA, United States).

Oxygen-glucose deprivation followed by normoxic reoxyge-nation was used to simulate ischemic damage in vitro. Subsequent cell death was assessed by measuring cellular uptake of PI. To induce OGD we used a protocol modified from Lushnikova et al. (2004), Kawano et al. (2006) and Montero et al. (2007). A humidified modular incubator chamber (Billups-Rothenberg Inc.) and glucose-free and serum-free medium (Neurobasal A media without glucose 0050128DJ, Invitrogen), supplemented with 1 mM Sodium Pyruvate) were pre-warmed at 37°C for 30 min. The chamber was then flooded by 95% N₂/5% CO₂ gas for 4–6 min (to reduce the O₂ gas level to zero), then sealed and incubated at 37°C for 15 min to deoxygenate the media. The semi-porous inserts containing the hippocampal slices (11–14 DIV) were transferred into dishes containing 1 ml of the warm deoxygenated media and washed with 0.5 ml of the media, (which was placed on top of the slices and then aspirated) before being transferred to the chamber where air was again replaced by 95% N2/5% CO₂ for 4–6 min. The chamber was then sealed and placed in an incubator at 37°C for 45 min. Following OGD, the cultures were returned to fresh media and normoxic conditions for 24, 48, or 72 h (as indicated in the figure legend) prior to analysis. Untreated slice cultures were used as control. For experiments involving PI (Invitrogen), we added 2 μM into the slice growth media following OGD. Prior to fixation, slices were washed twice with PBS to remove excess PI and then fixed in 4% paraformaldehyde/0.1 M phosphate buffer for 30 min. Slices were permeabilised with PBS/0.2% Triton-X100 and incubated with TO-PRO-3 iodide (Invitrogen) (1:10,000) in PBS to label all cell nuclei (‘total’). The percentage of apoptotic nuclei was determined by counting the number of PI-positive nuclei relative to total number of nuclei (TO-PRO-3) using ImageJ (NIH). For these experiments, imaging was performed blindly and at least 10 images were taken per condition for each experiment.

In the developing brain, SPARC is highly expressed in CNS astrocytes and microglia, where it has been shown to regulate synapse development and maturation (Jones et al., 2011; Kucukdereli et al., 2011). Conversely, in the mature CNS, the expression of SPARC is reduced and largely confined to expression in microglia, Bergmann glia in the cerebellum and Muller glia in the retina (Vincent et al., 2008). However, there are several studies that show that following increased neuronal activity (Jones et al., 2011) or upon injury or disease, SPARC levels are upregulated (Ikemoto et al., 2000; Liu X. et al., 2005; Ozbas-Gerceker et al., 2006; Au et al., 2007; Baumann et al., 2009; Lloyd-Burton et al., 2013) to modulate neuronal behavior. Given our previous work showing that SPARC can regulate the strength of synapses during development, we decided to investigate the effect of increasing SPARC on surface AMPARs in hippocampal neuron cultures grown with a feeder layer of astrocytes. We assessed the surface levels of AMPAR subunits GluA1 and GluA2 by immunofluorescence using non-permeabilizing conditions and found that surface GluA1 but not GluA2 was significantly increased upon administration of exogeneous recombinant SPARC (48 h, 0.5 μg/ml; Figures 1A,B). This result was confirmed by performing surface biotinylation experiments (Figures 1C,D), which showed a specific increase in surface GluA1 without a change in overall levels in neurons. Next, we examined the effect of SPARC in organotypic hippocampal slices where the three-dimensional architecture of synapses and their interaction with astrocytes is preserved (Haber et al., 2006). In contrast to the dissociated neuron cultures, we found that SPARC treatment (48 h, 0.5 μg/ml) not only increased synaptic GluA1 but also total GluA1 levels. Similar to the dissociated cultures, surface and total GluA2 levels were not significantly changed. We did observe a trend for an increase in total and synaptic GluA2 levels following SPARC treatment. However, performing additional experiments did not identify a significant difference with GluA2. Overall GluN1 and PSD-95 levels were also not affected, however, we observed a small but significant increase in synaptic GluN1 levels (Figures 2A,B). Therefore, increasing SPARC levels induces an increase in synaptic and surface GluA1-containing AMPARs in hippocampal slices and dissociated neurons.

In the hippocampus, both GluA1-containing homomeric (GluA1-GluA1) and heteromeric (GluA1–GluA2) complexes exist, with the majority being composed of both GluA1 and GluA2 proteins (∼80%; Lu et al., 2009). Given that SPARC increased the levels of synaptic GluA1, we sought to investigate whether there was a change in the levels of GluA1-GluA1 containing AMPARs. To test this, we co-immunoprecipitated GluA1 subunits from synaptosomes (Kang et al., 2012; Cook et al., 2014) and examined the levels of associated GluA1 and GluA2 in control and SPARC-treated hippocampal slices (Figure 2C). We found that there was an increase in the amount of GluA1–GluA1 AMPAR interactions in SPARC treated slices compared to control, whereas GluA1–GluA2 interactions remained unaffected, suggesting that SPARC promotes interactions among GluA1 subunits in synaptic fractions.

We next investigated whether the increase in GluA1 induced by SPARC affected synaptic function. To do this, we recorded AMPAR-mediated mEPSCs (miniature EPSCs) from CA1 neurons in control and SPARC-treated hippocampal slices. We found that mEPSCs in SPARC-treated slices were significantly greater in amplitude whereas their frequency was unchanged (Figures 3A,B and Supplementary Figures 2A,B) indicating that SPARC promotes an increase in synaptic strength. To determine whether the increase in mEPSC amplitude was due to homomeric GluA1-containing AMPAR, we treated slices with Philanthotoxin 433 (Phx-433), a polyamine-containing toxin which specifically inhibits calcium permeable AMPARs (Washburn and Dingledine, 1996), the majority of which are assumed to be GluA2-lacking, homomeric GluA1 AMPARs (Rozov et al., 2012; Mattison et al., 2014). In SPARC-treated slices, Phx-433 reduced mEPSC amplitude to baseline levels, suggesting that the increase in synaptic strength induced by SPARC was a result of additional GluA1-containing AMPARs at the synapse (Figure 3B). Taken together, these results suggest that SPARC induces a selective increase in functional synaptic GluA1-containing AMPARs.

Several studies have reported that SPARC is upregulated following challenge to the nervous system from elevated neuronal activity, glutamate exposure, or injury (Ikemoto et al., 2000; Liu Q.S. et al., 2005; Ozbas-Gerceker et al., 2006; Jones et al., 2011) including photothrombotic stroke (Lloyd-Burton et al., 2013). We chose to examine SPARC expression in the well-characterized model of ischemic cortical stroke generated by MCAO (Chen et al., 1986; Liu et al., 1989; Carmichael, 2005). MCAO produces an ischemic infarct ‘core,’ characterized by almost complete loss of neurons, surrounded by a ‘penumbra-like’ peri-infarct region, where neuronal cell death, although severe, is delayed and hence represents an important therapeutic target for potential neuroprotection and recovery (Lo, 2008; Murphy and Corbett, 2009; Ramos-Cabrer et al., 2011). Using IF labeling, we examined SPARC expression in the peri-infarct region compared to the same region in the contralateral cortex, where SPARC was present at a low level, predominantly in resting microglia (Figure 4A, top panel), consistent with previous studies (Vincent et al., 2008). In contrast, following MCAO, SPARC was upregulated in both microglia and reactive astrocytes (Figure 4A, bottom panel and Figure 4B). To quantify changes in SPARC expression following MCAO, we performed immunoblotting on cortical tissue dissected at 48 h, 72 h, or 15 days following MCAO or Sham surgery (Figure 4C). Consistent with the IF analysis, we observed a significant increase in SPARC protein levels at 72 h post MCAO, with a return to baseline levels by 15 days. We did not see any changes in SPARC in the contralateral cortex (data not shown). Thus, SPARC shows a delayed upregulation in astrocytes and microglia in response to MCAO.

To better dissect the role of SPARC after CNS injury, we used an in vitro model of ischemia, OGD in organotypic hippocampal slices where we could more carefully control the experimental conditions. Consistent with previous reports (Lushnikova et al., 2004), OGD caused a loss of hippocampal neurons, particularly in the CA1 subfield [Figure 5A; apoptotic cells labeled using propidium iodide (PI)]. We found that approximately 60% of CA1 were PI-positive at 72 h post-OGD (Figure 5B). To determine whether SPARC expression was regulated by OGD in a similar manner to MCAO, we performed IF and immunoblotting analysis on slices subjected to OGD. We found that SPARC protein was robustly increased at 48 h following OGD (Figures 5C,D).

We next sought to investigate how SPARC might be regulating synaptic proteins during OGD. We prepared synaptosomes from slices subjected to OGD with and without SPARC treatment (SPARC treatment was for 48 h before and after OGD). OGD led to a significant loss of the synaptic proteins GluA1, GluA2, vGlut1, GluN1, PSD-95 (Figure 6), suggesting that OGD induces a loss of synapses, consistent with previous studies (Kovalenko et al., 2006; Nikonenko et al., 2009). Interestingly, pretreatment with SPARC prevented synaptic loss of GluA1 and GluN1 following OGD whereas vGlut1, GluA2 and PSD-95 levels were still reduced (Figure 6B). The rescue of synaptic AMPARs occurred even though overall levels of GluA1 were not significantly elevated, suggesting that SPARC preferentially promotes the recruitment and maintenance of GluA1-containing AMPAR complexes at synapses following OGD challenge rather than regulating overall GluA1 expression.

Given these findings, we investigated whether SPARC could affect the survival of CA1 neurons following OGD. To do this, we compared hippocampal slices subjected to OGD and acutely applied SPARC at time of OGD (acSPARC) or pre-treated with SPARC (chSPARC) for 48 h prior to OGD and following reoxygenation. We found that SPARC had a significant protective effect (∼26%) when slices were pre-treated with SPARC but not when it was acutely applied (Figures 7A,B). Interestingly, the protective effect of SPARC could be blocked by co-application of Phx-433 (Figures 7A,D). Phx-433 treatment alone had no effect on the number of apoptotic cells (Figure 7C). Together, these results suggest that SPARC regulation of Phx-433-sensitive GluA1-containing AMPARs at the synapse are important for improved cell survival and recovery following OGD.