The Laboratory Animal Care and Use Committee of Shenzhen University approved all animal related experimental protocols. Male Sprague–Dawley rats (purchased from the Experimental Animal Center Southern Medical University, Guangzhou, Guangdong, China) weighing 300 g to 400 g were anesthetized with isoflurane (4% for induction, 1.75% for maintenance) in N₂O:O₂ (70%:30%) during surgical procedures and the body temperature was maintained through a heated pad. A 4–0 silicone-coated monofilament nylon suture was introduced into the right intra-carotid artery to occlude the opening of the MCA as we previously described (Liu et al., 2009). MCAO was lasted for 3 h, and the animals were then deeply anesthetized with isoflurane and euthanized by decapitation. Successful MCAO was confirmed by 2,3,5-triphenyltetrazolium chloride (TTC, Sigma-Aldrich, St. Louis, MI, USA) staining of the 2-mm-thick brain coronal section 6 mm away from the tip of the front lobe as we previously described (Liu et al., 2008).

Rat primary cortical neurons were cultured using a method as we described previously (Liu et al., 2007). Briefly, cerebral cortices were removed from the embryos of Sprague–Dawley pregnant rats at 15–18 days gestation (Shouthern Medical University Experimental Animal Center). After removing the meninges, the cortical tissue was minced and incubated with 0.05% trypsin for 30 min at 37°C with gentle trituration. After digestion, the neurons were achieved and suspended in neurobasal medium containing 2% B27 supplement and 0.5 mM L-Glutamine. Before seeding, culture vessels including 96-well plates, 1.2 cm glass slides or 6 cm dishes were coated with poly-L-lysine (PLL; 50 μg/mL, Sigma-Aldrich) at room temperature overnight. Neurons were maintained at 37°C in a humidified 5% CO₂ incubator and half of the culture medium was changed every 3 days. The neurons were subjected to experiments 8 days after seeding.

PC12 cells, a rat PC12 cell line, were obtained from the Cell Resource Center of the Institute of Basic Medical Sciences, Peking Union Medical College/Chinese Academy of Medical Sciences (Beijing, China). PC12 cells were grown as a monolayer in dulbecco’s modified eagle medium (DMEM) supplemented with 10% horse serum and 5% fetal bovine serum (FBS), 100 U/ml penicillin and 100 μg/ml streptomycin at 37°C in a humidified incubator gassed with 5% CO₂ and 95% room air. For neuronal differentiation, PC12 cells were seeded in PLL pre-coated plates and allowed to adhere for 24 h. Following adherence, the culture medium was replaced with nerve growth factor (NGF, 50 ng/ml; New England Biolabs, MA, USA) containing medium. The NGF-containing medium was replaced every other day. Seven days after the supplement of NGF, the NGF-induced differentiation of PC12 cells were determined using immunofluorescence staining with antibodies against the neuron-specific marker microtubule associated protein 2 (MAP2; Supplementary Figure S1).

Recombinant adenoviral constructs, including Ad-control (control construct), Ad-Nrdp1 (overexpressing Nrdp1), Ad-si-control (expressing non-targeting control siRNA) and Ad-si-Nrdp1 (expressing Nrdp1 siRNA) were generated as described previously (Zhang et al., 2011a). Eight days after plating, cells were infected with Ad-control, Ad-Nrdp1, Ad-si-control or Ad-si-Nrdp1 for 24 h before OGD treatment.

To mimic ischemic condition in vitro, primary neurons or PC12 cells were exposed to OGD as described previously (Liu et al., 2012). In brief, neurons or PC12 cells were subjected to OGD by replacing the normal growth medium with glucose free medium (DMEM without glucose) pre-equilibrated with 95% N₂ and 5% CO₂. The cells were then incubated in a humidified airtight chamber (Biospherix Ltd., Lacona, NY, USA) for 1 h, 3 h, or 6 h. Control cultures were incubated with normal DMEM medium without FBS at 37°C in 5% CO₂/95% air. OGD was terminated by removing cells from the hypoxic chamber and the cells were collected separately for further measurement.

After OGD treatment, cell cytotoxicity was determined by the release of lactate dehydrogenase (LDH), a cytoplasmic enzyme released from cells. LDH release into the culture medium was detected using a CytoTox 96^®Non-Radioactive Cytotoxicity Assay Kit (Promega Corporation. Madison, WI, USA). Briefly, 50 μl of each sample medium (i.e., pure culture medium for measuring background LDH release, culture media collected from control or OGD-treated cells for measuring experimental LDH release and lysis buffer-treated cells for measuring maximum LDH release) was collected to assay LDH release. The samples were incubated with reduced form of nicotinamide adenine dinucleotide and pyruvate for 30 min at room temperature and the reaction was terminated by adding Stop Solution. LDH release was assessed by measuring the absorbance of supernatants at 490 nm. Cell death rate was calculated as follows: cell death rate = (experimental LDH release-background LDH release) /(maximum LDH release-background LDH release) × 100%. The results were presented as fold increase of the control cells.

Total RNA was isolated from neurons using Trizol reagents (Invitrogen Life Technologies, Carlsbad, CA, USA). RNA samples (2 μg) were reverse-transcribed to generate first-strand cDNA. After reverse transcription using TaqMan^® Reverse Transcription Kits (Applied Biosystems), reverse-transcribed products were amplified with the 7900HT real-time PCR System (Applied Biosystems) using SYBR^® Green PCR Master Mix (Applied Biosystems, Foster City, CA, USA) under the following conditions: 30 s at 95°C, followed by a total of 40 cycles of two temperature cycles (15 s at 95°C and 1 min at 60°C). Primer sequences were as follows: rat Nrdp1 forward: 5′-ATGGGGTATGATGTAACCCGG-3′ and reverse: 5′-GATGCAGGCGTTGCAGAAG-3′; Rat GAPDH served as endogenous control, and the primers were forward: 5′-CAATGTGTCCGTCGTGGATCT-3′; reverse: 5′-GTCCTCAGTGTAGCCCAAGATG-3′. The Ct value was calculated by the comparative ∆∆C_t method using the SDS Enterprise Database software (Applied Biosystems).

Apoptosis was analyzed by TUNEL assay using Click-iT^® Plus TUNEL Assay (Life Technologies, Inc., Carlsbad, CA, USA) according to manufacturer’s instruction. Briefly, at the end of the indicated treatments, primary rat cerebral cortical neurons grown on coverslips were incubated with TdT reaction mixture for 2 h at 37°C, followed by 30-min incubation with the Alexa Fluor^® 594 dye. Then, the cells were counterstained with DAPI (Sigma-Aldrich) for 20 min and observed under a fluorescence microscope (Leica, Germany). The TUNEL-positive nuclei of six non-overlapping fields per coverslip were counted by a researcher blinded to treatment, and these counts were converted to percentages by comparing the TUNEL-positive counts to the total number of cell nuclei as determined by DAPI counterstaining, that is TUNEL-positive ratio = (number of red nuclei/number of blue nuclei) × 100%.

Protein samples were prepared from cultured neurons or PC12 cells using extraction buffer as described previously (Liu et al., 2012). The protein samples were electrophoresed on SDS-PAGE gels and transferred to a nitrocellulose membrane. After blocking with 5% non-fat milk for 1 h at room temperature, the membranes were incubated with the indicated primary antibodies overnight and then with horseradish peroxidase-conjugated secondary antibody for 1 h. The blots were developed using a chemiluminescent system, and the bands were scanned, and densitometry analysis was performed with Gel-pro 4.5 Analyzer (Media Cybernetics, Silver Spring, MD, USA). The primary antibodies were anti-cleaved-PARP, anti-PARP, anti-Bcl-2, anti-Bax, anti-USP8 and anti-β-actin. Anti-Nrdp1 and anti-HIF-1α antibodies were purchased from BETHYL Laboratories and Novus Biologicals, respectively, and the rest primary antibodies were purchased from Santa Cruz Biotech. Relative protein levels were quantified after normalization to the loading control β-actin.

After the indicated treatments, PC12 cells were lysed in RIPA buffer and centrifuged at 12,000× g for 10 min at 4°C. The supernatant were incubated overnight at 4°C with 4 μg of anti-USP8 (Santa Cruz Biotech, Chicago, IL, USA), followed by precipitation with 50 μl of Dynabeads protein A (Pierce Biotechnology, Rockford, IL, USA) for 10 min at room temperature. The protein A were then washed extensively with binding buffer, resuspended in SDS-PAGE buffer, and boiled for 5 min. Samples of 30 μg total cell lysate were used as an input control. The precipitated complexes were separated on SDS-PAGE gels, and transferred to nitrocellulose membranes, and immunoblotted with anti-Nrdp1 (BETHYL Laboratories, Montgomery, TX, USA), K48-linkage-specific anti-ubiquitin antibody (Abcam, Cambridge, MA, USA) or anti-HIF-1α antibody (Novus Biological, Littleton, CO, USA) to detect the presence of these proteins in the complex. Normal rabbit IgG (Santa Cruz Biotech) was used as a loading control.

All data were expressed as means ± SEM. Differences between groups were evaluated by either an unpaired Student’s t test or one-way ANOVA followed by Tukey’s post hoc test as indicated in the Figure Legends. P < 0.05 was regarded as statistically significant.

Nrdp1 is found to be widely expressed in the brain and is implicated in ischemic damage to the heart (Zhang et al., 2011b). To determine whether Nrdp1 plays a role in ischemic brain injury, we examined the change of Nrdp1 expression in the cerebral cortex isolated from the rats that were subjected to 3-h MCAO without reperfusion. Nrdp1 mRNA expression was analyzed in isolated hemispheric cortex by real time RT-PCR and found that 3-h MCAO induced a significant increase (~1-fold) of Nrdp1 mRNA expression in ischemic hemispheric cortex compared to non-ischemic cortical tissue (Figure 1A, P < 0.05). Consistent with its mRNA change, Nrdp1 protein levels were also significantly increased in ischemic cerebral cortex (Figure 1B). These results demonstrate that Nrdp1 is upregulated in the ischemic brain cortex. To further demonstrate a role of Nrdp1 in ischemic neuron injury and the underlying mechanisms involved, we chose the widely used in vitro model of ischemia (i.e., OGD) for the rest of this study.

To determine the functional role of cerebral Nrdp1 in response to OGD treatment, we examined the expression of Nrdp1 in OGD-treated primary rat cerebral cortical neurons. The cells were exposed to OGD for 1, 3, or 6 h before analyzing Nrdp1 mRNA and protein levels. Real time RT-PCR analysis showed that Nrdp1 mRNA expression was increased in cerebral cortical neurons after exposing to OGD for 1 h and was further increased at 6-h OGD, while no significant difference was seen between 1-h OGD and 3-h OGD (Figure 2A). Western blot analysis showed that Nrdp1 protein levels were significantly increased in cerebral cortical neurons after exposing to OGD for 3 h and 6 h, but not for 1 h (Figure 2B). To further verify the above findings, we assayed the expression of Nrdp1 in PC12 cells exposed to the same OGD treatment as above. Real time RT-PCR analysis showed that Nrdp1 mRNA expression was increased in PC12 cells after exposing to OGD for 3-h and 6-h OGD, but not for 1 h (Figure 2C). Western blot analysis showed that Nrdp1 protein levels were significantly increased in PC12 cells after exposing to OGD for 6 h, but not for 1 h and 3 h (Figure 2D). These data demonstrate that OGD induces Nrdp1 upregulation in cerebral cortical neurons as well as PC12 cells in a time-dependent manner.

To investigate whether Nrdp1 is implicated in OGD-induced apoptosis in neurons, we transfected cerebral cortical neurons with Ad-control, Ad-Nrdp1 or Ad-si-Nrdp1. As shown in Figure 3, the transfection efficiency reached more than 90% at 24 h after transfection (Figure 3A), and western blot analysis showed that incubating the neurons with Ad-si-Nrdp1 and Ad-Nrdp1 for 48 h significantly reduced (~90% reduction) and increased (~2-fold increase) Nrdp1 protein levels, respectively (Figures 3B,C). We assessed the effect of Ad-si-Nrdp1 and Ad-Nrdp1 on OGD-induced neuronal death by measuring LDH release (indicating late apoptosis and necrosis) and TUNEL staining (indicating apoptosis). The data showed that cell death and apoptosis did not differ across the groups under basal conditions (Figures 3D,E and Supplementary Figure S2). However, after OGD treatment, inhibition of Nrdp1 significantly attenuated neuronal death and apoptosis as compared with the Ad-control, while transfection with the Ad-Nrdp1 greatly enhanced OGD-induced neurons death and apoptosis (Figures 3D,E and Supplementary Figure S2).

To further verify a role of Nrdp1 in OGD-induced apoptosis in neurons, we assessed the effect of Nrdp1 on several key apoptosis-associated signal proteins including cleaved-PARP and Bax/Bcl-2. As shown in Figures 3F,G, 6-h OGD induced a significant increase in cleaved PARP levels (PARP activation) and a greater ratio of Bax/Bcl-2 in cerebral cortical neurons, and transfection with Ad-si-Nrdp1 abolished these changes. Accordingly, overexpression of Nrdp1 augmented OGD-induced increases in cleaved PARP and Bax/Bcl-2 ratio. As expected, Ad-si-Nrdp1 or Ad-Nrdp1 alone did not affect these apoptosis-associated signal proteins (Figures 3F,G). Taken together, these results clearly indicate that Nrdp1 plays an important role in ischemia-induced apoptosis in cerebral cortical neurons.

HIF-1α acts as an intracellular sensor for hypoxia and promotes the cells to adapt to hypoxic/ischemic conditions (Zis et al., 2015), thus we hypothesized that Nrdp1 might interact with or suppress HIF-1α to promote neuronal cell death under OGD conditions. To test this possibility, we transfected primary cortical neurons and PC12 cells with Ad-control, Ad-Nrdp1, Ad-si-control or Ad-si-Nrdp1 before exposing to OGD for 6 h. HIF-1α protein levels were analyzed by western blot. As shown in Figures 4A,B, 6-h OGD induced a significant increase in the accumulation of HIF-1α protein in both primary neurons and PC12 cells transfected with Ad-si-control, and of note, this increase was further augmented or attenuated when Nrdp1 was knocked down by Ad-si-Nrdp1 or overexpressed by Ad-Nrdp1, respectively (Figures 4C,D). These data suggest that Nrdp1 may act as a negative regulator for HIF-1α expression in neurons under OGD conditions.

The fact that Nrdp1’s substrate USP8 can protect HIF-1α from pVHL-mediated degradation (Troilo et al., 2014) led us to hypothesize that Nrdp1 may act on USP8 to regulate HIF-1α expression under OGD condition. To test this, we investigated the change of USP8 in OGD-treated neurons and the impact of Ad-Nrdp1 and Ad-si-Nrdp1 on USP8 expression. As shown in Figures 4A,B, 6-h OGD induced a significant decrease in the level of USP8 proteins in the cells transfected with Ad-si-control. Importantly, this reduction was partially reversed when Nrdp1 was knocked down by Ad-si-Nrdp1, and was exacerbated when Nrdp1 was overexpressed by Ad-Nrdp1 (Figures 4C,D). Collectively, these results suggest that Nrdp1 may contribute to OGD-induced neuronal cell death via suppressing HIF-1α and USP8 expression.

Lastly, to ambiguously demonstrate the interactions between Nrdp1 and USP8 and between USP8 and HIF-1α in OGD-treated neurons, we performed co-immunoprecipitation experiments. Since Nrdp1 targets USP8 for ubiquitylation, we speculated that overexpression of Nrdp1 could enhance protein ubiquitylation and USP8 degradation in PC12 cells. To test this, we pulled down ubiquitylated species from PC12 cell extracts, and then detected the levels of protein ubiquitilytion in the presence of proteasome inhibitor MG132 as well as the protein levels of Nrdp1 and USP8. As shown in Figure 5A, overexpression of Nrdp1 by Ad-Nrdp1 significantly increased the whole levels of protein ubiquitylation in comparison to Ad-green fluorescent protein (GFP) control, and this change was accompanied by decreased USP8 protein levels.

To demonstrate a direct interaction between USP8 and HIF-1α in OGD-treated PC12 cells, we performed co-immunoprecipitation assays and found that HIF-1α was precipitated by antibody against USP8, but not by control rabbit IgG (Figure 5B). Moreover, under OGD conditions, overexpression of Nrdp1 by Ad-Nrdp1 reduced co-immunoprecipitation between USP8 and HIF-1α, while knockdown of Nrdp1 by Ad-si-Nrdp1 enhanced the interaction between these two proteins (Figure 5C). These data indicate that under ischemic conditions, Nrdp1 upregulation may hinder the stabilization of HIF-1α in neurons via promoting ubiquitin-mediated degradation of USP8, thus attenuating cellular adaptive response to hypoxia/ischemia.