Ca²⁺-dependent endoplasmic reticulum stress correlation with astrogliosis involves upregulation of KCa3.1 and inhibition of AKT/mTOR signaling

Frozen tissues and paraffin-embedded brain slices from the hippocampi of postmortem human samples of control and AD patients were obtained from the Netherlands Brain Bank (Netherlands Institute for Neuroscience, Amsterdam, Netherlands). The Netherlands Brain Bank obtained written informed consent for brain autopsy specimens for research purposes after death. The Braak stage, neuropathological and clinical diagnoses, age and gender distributions, as well as diagnostic groupings are presented in Table 1. The frozen tissues were used to isolate proteins, and expression of proteins was evaluated by Western blotting as described below. Immunofluorescence was performed on 8-μm sections using antibodies against glial fibrillary acidic protein (GFAP), KCa3.1, Orai1, and GRP78 antibodies as described below.

Wild-type (WT) C57BL/6 mice (male, 25–30 g) were procured from the Shanghai Laboratory Animal Center (Shanghai, China). KCa3.1^−/− mice were obtained from the Jackson Laboratory (Bar Harbor, ME, USA) and housed as described previously [18]. APP/PS1 transgenic mice (an AD model) were also purchased from the Jackson Laboratory (no. 003378) [15]. KCa3.1^−/− mice were crossed with APP/PS1 mice and the offspring were intercrossed to generate mice with the KCa3.1^−/−/APP/PS1 genotype. Mouse cohorts of the four genotypes (WT, KCa3.1^−/−, KCa3.1^−/−/APP/PS1, and APP/PS1) were generated and used for behavioral analysis. The protocols of all animal experiments were approved by the Animal Experimentation Ethics Committee of Shanghai Jiao Tong University School of Medicine, Shanghai, China (ethics protocol number: A-2015-010).

The modified Morris water maze (MWM) test was performed as previously described [21]. Briefly, the test requires the mice to receive 5 consecutive training days (1 day with the visible platform and 4 days with the hidden platform) in a circular water tank (diameter = 80 cm; height = 50 cm) containing opaque water (22 °C, 25 cm deep). A spatial probe trial was performed on day 6 with no platform present. During the hidden platform training days, the mice were allowed to swim for 60 s to find the hidden platform (1 cm below the water surface). The platform was removed from the pool on day 6 of the spatial probe trial and the mice swam freely for 60 s. Performance of the MWM test was recorded using a video camera and analyzed using a video tracking system (Jiliang Software Technology Co., Ltd., Shanghai, China).

Confocal microscopy was performed as previously described [18]. Briefly, mice were anesthetized with chloral hydrate and perfused with 4% paraformaldehyde. Then, 10% goat serum in 0.01 M phosphate-buffered saline (PBS) was used to block 12-μm sections of the mice brain or 8-μm sections of the human brain for 1 h at room temperature. The brain sections were then incubated with the following primary antibodies: rabbit anti-GFAP (1:1000; Dako, Glostrup, Denmark), mouse anti-GFAP (1:200; Merck Millipore, Burlington, MA, USA), rabbit anti-NeuN (1:100; Merck Millipore, Burlington, MA, USA), rabbit anti-Iba1 (1:500; Wako Pure Chemical Industries, Ltd., Osaka, Japan), mouse anti-KCa3.1 (1:100; Alomone Labs, Ltd., Jerusalem, Israel), rabbit anti-GRP78 (1:100; Cell Signaling Technology, Inc., Beverly, MA, USA), mouse anti-NG2 (Sigma-Aldrich Corporation, St. Louis, MO, USA), and rabbit anti-Orai1 (1:100; Santa Cruz Biotechnology, Inc., Dallas, TX, USA). After incubation at 4 °C overnight, the brain sections were incubated with the respective Alexa Fluor® 488- or 568-conjugated secondary antibodies (1500; Invitrogen Corporation, Carlsbad, CA, USA). Images were collected using a TCS SP8 confocal laser scanning microscope (Leica Microsystems, Wetzlar, Germany) and processed with Leica LASAF Lite imaging software. The same reference position was used for each brain slice for quantification, i.e., between three and five random 0.01-mm² microscopic fields. Quantification was measured from six sections per brain (120-μm intervals) in a blinded manner.

Mice were anesthetized with chloral hydrate and perfused with saline. Mice or human brain tissues and cells were lysed on ice with radioimmunoprecipitation assay buffer (50 mM Tris (pH 7.4), 150 mM NaCl, 1% Triton X-100; 1% sodium deoxycholate, 0.1% sodium dodecyl sulfate, sodium orthovanadate, sodium fluoride, ethylenediaminetetraacetic acid, leupeptin) containing 1% phenylmethanesulfonyl fluoride. The supernatants were collected after centrifuging at 13,500 rpm for 5 min at 4 °C. Equal concentrations of proteins were separated by 10% (w/v) sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and then transferred to a polyvinylidene difluoride membrane, which was blocked for 1 h at room temperature (RT) in 5% milk in Tris-buffered saline with 0.05% Tween 20. The membrane was incubated with the following primary antibodies overnight at 4 °C: mouse anti-KCa3.1 (1:100; Alomone Labs, Ltd.), mouse anti-STIM1 (1:500; Santa Cruz Biotechnology, Inc.), rabbit anti-Orai1 (1:1000; Santa Cruz Biotechnology, Inc.), rabbit anti-GRP78, rabbit anti-mTOR, rabbit anti-phospho-mTOR (Ser2448), rabbit anti-phospho-PERK (Thr980), rabbit anti-phospho-eIF2α, rabbit anti-phospho-Akt (Thr308), rabbit anti-Akt, rabbit anti-phospho-4E-BP1, rabbit anti-phospho-p70 S6 Kinase, mouse anti-CHOP, rabbit anti-iNOS, rabbit anti-COX2 (1:1000; Cell Signaling Technology), and mouse anti-β-actin (1:1000; Beyotime Institute of Biotechnology, Haimen, China). The membranes were subsequently washed and incubated with anti-rabbit or anti-mouse HRP for 1 h at RT, and developed using BeyoECL solution (P0018A; Beyotime Institute of Biotechnology). Image Studio Lite Ver 5.2 software (LI-COR Biosciences, Lincoln, NE, USA) was used to analyze the proteins.

ELISA was performed using the kit for TNF-α and IL-1β (Rapidbio Labs, Langka Trade Co. Ltd., Shanghai, China). The procedures were conducted according to manufacturer’s protocols.

The oligomeric Aβ was prepared as described previously (Yi et al. 2016b). Briefly, monomeric peptide Aβ was dissolved in 1,1,1,3,3,3-hexafluoro 2-propanol (Sigma-Aldrich Corporation, St. Louis, MO, USA) at 1 mg/mL. The dried monomeric peptide Aβ was resolved in ddH₂O at a concentration of 100 mM and incubated at 4 °C for 24 h to induce aggregation.

Primary cortical astrocyte cultures derived from newborn (P0–P2) WT or KCa3.1^−/− C57BL/6 mouse brains were prepared from mixed glial cultures (10–14 days in vitro) as described previously [22]. Briefly, the cerebral cortices were dissociated into single cell suspensions. Dissociated single cells were grown in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum at 37 °C under an atmosphere of 5% CO₂. When the astrocytes grew to confluence (10–14 days later), the flasks were shaken overnight (200 rpm, 37 °C) to deplete microglia and oligodendrocytes. The purified astrocytes were then plated onto 6-well or 96-well plates in serum containing medium. Astrocytes were incubated with serum-free DMEM for 24 h after reaching confluence again, and then were treated with Aβ for different time periods before harvest. The culture medium (CM) was changed to neurobasal medium (NB) with B27 supplement (Invitrogen Corporation) after confluent astrocytes were serum-free for 24 h. The NB/B27-based WT and KCa3.1^−/− astrocytes were then treated with 5 μM Aβ oligomer. The CM from the NB/B27-based cells was collected and used immediately. The primary cultured cortical neurons were isolated from neonatal (P0–P2) C57BL/6 mice and cultured as described previously (Yi et al. 2016b).

Cells were stained with primary antibody against microtubule-associated protein 2 (MAP2) and Alexa Fluor® 568-conjugated secondary antibody. A Cellomics Kinetic Scan reader (Thermo Fisher Scientific, Waltham, MA, USA) was used to scan the MAP2-positive cells. Extended Neurite Outgrowth software (Thermo Fisher Scientific) was used to analyze the data.

The purified astrocytes were plated in the wells of 96-well plates containing DMEM supplemented with 10% fetal bovine serum. After once again reaching confluence, the cells were incubated with the fluorescent calcium indicator Fluo-4AM (1.6 μM; Beyotime Institute of Biotechnology) for 25 min and then washed three times with 0.01 M PBS. Afterward, 2 μM ethylene glycol-bis (β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid was used to chelate the calcium in the DMEM and then 1 μM thapsigargin (Tg) was used to induce [Ca²⁺]_i release. After the [Ca²⁺]_i concentration was stable, 2 mM CaCl₂ solution was added to the DMEM to induce Ca²⁺ influx. Fluorescence signals were recorded and analyzed using a FlexStation 3 Multi-Mode Microplate Reader (Molecular Devices, Sunnyvale, CA, USA).

All data are presented as the mean ± standard error of the mean. Statistical analyses were performed using Prism software (GraphPad Software, Inc., La Jolla, CA, USA). Data were tested for Gaussian distribution with the Kolmogorov–Smirnov normality test and then analyzed by one-way analysis of variance (ANOVA) and the Dunnett’s post-hoc test. Data were analyzed with an unpaired, two-tailed Student’s t test when comparing between two groups or the non-parametric Mann–Whitney test was applied. Statistical significance was set at p < 0.05.

ER stress induced by increased protein misfolding and upregulation of the UPR has been observed in AD patients and in an AD mouse model [23, 24]. Disruption of protein folding in the ER triggers the UPR via activation of three ER pathways: PERK, ATF6, and IRE1 [9, 10]. The GRP78 dissociates from PERK, ATF6, and IRE1 during ER stress, and then initiates proapoptotic signaling via activation of CHOP. GRP78 and CHOP protein levels were increased in both the brains of AD patients (p < 0.05, Fig. 1a, b) and the hippocampi of APP/PS1 mice (p < 0.05, Fig. 1c, d), as compared to control tissues. The Aβ oligomer was reported to disrupt Ca²⁺ homeostasis in the ER of astrocytes, which leads to ER stress-induced RA [25]. The addition of Aβ to primary cultured astrocytes significantly upregulated the protein levels of GRP78 and CHOP at 12 h, as compared with control cells, indicating that Aβ induces the activation of the UPR in astrocytes (p < 0.05, Fig. 1e, f). Overall, these data obtained from AD patients and APP/PS1 AD mouse models demonstrate a potential mechanism of ER stress activation in AD.

For evaluation of SOCE, Tg (1 μM), an ER Ca²⁺ ATPase pump blocker, was used to deplete ER Ca²⁺ stores. Without extracellular Ca²⁺ (0 Ca²⁺), Tg-induced upregulation of [Ca²⁺]_i (first peak) showed no significant changes in response to treatment with 5 μM Aβ at 12 h, as compared to the control astrocytes (p > 0.05, Fig. 1g, h). However, SOCE, initiated by external Ca²⁺ (2 mM), was increased at 12 h (p < 0.05, Fig. 1g, h) and 24 h (p < 0.01, Fig. 1g, h) in response to treatment with 5 μM Aβ, as compared to control cells. Colocalization experiments were conducted using antibodies against the ER stress marker GRP78 and the RA marker GFAP. Double-labeled staining showed that GRP78 and GFAP colocalized in the AD brain tissues, but not the controls (Fig. 1i).

The accumulation of Aβ oligomers and amyloid plaques in the brain is associated with neuroinflammation and abnormal gliosis, including reactive astrocytes [3, 26]. We previously reported that KCa3.1 gene deletion attenuated ER stress in RA via the c-Jun/JNK and MAPK/ERK signaling pathways in ischemic mice [12]. Given the key role of Aβ-induced astrocytes ER stress due to SOCE-induced Ca²⁺ overload, we then measured the expression levels of KCa3.1 and SOCE channels following stimulation by prolonged Aβ treatment. Although the exact SOCE molecular components in astrocytes are unknown, KCa3.1 and Orai1 have been reported as components of SOCE channels in some cell types [8, 27, 28]. Western blot analysis was performed to evaluate changes in the expression profiles of KCa3.1, Orai1, and STIM1, which revealed upregulation of the KCa3.1 and Orai1 channels in response to stimulation with 5 μM Aβ for 24 h, as compared to the control group, whereas there was no obvious change in the expression of STIM1 (a regulator of KCa3.1 and Orai1) (p > 0.05, Fig. 2a, b). Colocalization experiments were conducted using specific KCa3.1 and Orai1 antibodies. Double-labeled staining showed that the colocalization of KCa3.1 and Orai1 was increased in primary astrocytes in response to stimulation with 5 μM Aβ for 24 h, as compared to the control cells (Fig. 2c).

We then further studied the potential relevance of KCa3.1 and Orai1 in the hippocampi of control and AD patients. The results showed that the expression levels of the KCa3.1 and Orai1 proteins, but not STIM1, were upregulated in the hippocampi of AD patients, as compared to the control tissues (p < 0.05, Fig. 2d, e). Colocalization experiments were conducted by immunostaining the brain tissues of AD patients and controls with specific KCa3.1 and Orai1 antibodies. Double-labeled staining showed that the colocalization of KCa3.1 and Orai1 was increased in the brain tissues of AD patients, as compared to the controls (Fig. 2f). In the present study, colocalization between KCa3.1 and astrocytes was detected in the brains of AD patients and 15-month-old APP/PS1 mice. Age-matched control humans and WT littermates were used as controls. In AD patients (Fig. 2g) and APP/PS1 mice (Fig. 2h), colocalization of KCa3.1 and GFAP⁺ astrocytes was increased, as compared to the controls.

The findings of the above experiments suggest that KCa3.1 plays a key role during the process of SOCE and in maintaining Ca²⁺ homeostasis in the ER. Pharmacological blockade of KCa3.1 was then conducted to further confirm the critical role of KCa3.1 in Ca²⁺ homeostasis and stress in the ER. The results showed that blockade of KCa3.1 with 1 μM TRAM-34 attenuated Aβ-induced SOCE-induced Ca²⁺ overload, as compared with Aβ-treated cells (p < 0.05, Fig. 3a, b). Pharmacological blockade of KCa3.1 with 1 μM TRAM-34 decreased the upregulation of GRP78 and CHOP, which was induced by treatment with 5 μM Aβ for 24 h, indicating that blockade of KCa3.1 could inhibit prolonged UPR activation (p < 0.05, Fig. 3c, d). KCa3.1 gene deletion (KCa3.1^−/−) from astrocytes was stimulated with 5 μM Aβ or 1 μM Tg for 24 h, as compared to WT control cells. KCa3.1 gene deletion also decreased Aβ or Tg-induced upregulation of GRP78 (p < 0.05, Fig. 3e–h). As a critical transducer of the UPR, phosphorylation of PERK (p-PERK) was increased after stimulation with 5 μM Aβ or 1 μM Tg, but decreased in KCa3.1^−/− astrocytes (p < 0.05, Fig. 3e–h). Similarly, prolonged UPR has been shown to activate eukaryotic initiation factor 2α (eIF2α), an important downstream signaling target, which was also increased in Aβ or Tg-treated WT astrocytes, but was restored to normal in KCa3.1^−/− cells (p < 0.05, Fig. 3e–h).

WT and KCa3.1^−/− astrocytes were used to test whether KCa3.1-mediated astrogliosis could induce dendritic damage. More dendritic damage was observed in the CM from 5 μM Aβ-stimulated WT astrocytes (WT/Aβ-CM) than in those treated with CM from WT astrocytes (WT/CM), as shown by immunofluorescent staining of the dendritic marker MAP2 (Fig. 3i). Incubation with WT/Aβ-CM decreased total neurite length (p < 0.01, Fig. 3j) and number of branch points (p < 0.01, Fig. 3k). Incubation with CM from 5 μM Aβ-stimulated KCa3.1^−/− astrocytes (KCa3.1^−/−/Aβ-CM) attenuated the effect of WT/Aβ-CM by increasing total neurite length and the number of branch points (p < 0.05, Fig. 3j, k).

In the present study, APP/PS1 mice were crossed with KCa3.1^−/− mice to study the role of KCa3.1 channels in RA-mediated neuronal toxicity. Offspring intercrossing generated mice with the WT, KCa3.1^−/−, APP/PS1, and KCa3.1^−/−/APP/PS1 genotypes.

Western blot analysis was conducted to identify changes in the expressions patterns of GRP78 and CHOP in the brains of 15-month-old APP/PS1 and KCa3.1^−/−/APP/PS1 mice, as compared with WT and KCa3.1^−/− controls. Similar to previous results in AD patients (Fig. 1a, b), GRP78 and CHOP levels were significantly increased in APP/PS1 mice, as compared to WT and KCa3.1^−/− mice (p < 0.05, Fig. 4a–c), while the expression levels of GRP78 and CHOP were significantly decreased in KCa3.1^−/−/APP/PS1 mice, as compared to APP/PS1 mice (p < 0.05, Fig. 4a–c).

It was reported that upregulation of GRP78 and phosphorylation of eIF2α in response to Aβ-induced ER stress contributed to RA in the AD mouse model and primary cultured astrocytes [25]. We then investigated whether KCa3.1 gene deletion could prevent changes in GRP78 expression levels in the astrocytes of APP/PS1 mice. To compare the response in astrocytes specifically, immunofluorescence was used to quantify GRP78 levels in GFAP⁺ astrocytes in the hippocampi of WT, KCa3.1^−/−, APP/PS1, and KCa3.1^−/−/APP/PS1 mice. In the hippocampi of APP/PS1 mice, GRP78 expression was increased in GFAP⁺ astrocytes, as compared to that of WT controls (p < 0.01, Fig. 4d, e). Meanwhile, hippocampal GRP78 immunoreactivity levels of KCa3.1^−/−/APP/PS1 mice were significantly decreased, as compared to that of APP/PS1 mice (p < 0.01, Fig. 4d, e).

The foregoing data suggest that KCa3.1 gene deletion prevented activation of the UPR and attenuated astrocytic ER stress in an in vivo AD model. However, the signaling intermediates linking KCa3.1 and astrocytic ER stress in AD remain unknown. Given the reported relationship between the phosphatidylinositol 3-phosphate kinase/Akt pathway and ER stress [29], we tested whether the AKT pathway is involved in KCa3.1-mediated astrocytes ER stress.

Decreased phosphorylation of AKT (p-AKT) (Thr308) was observed in the brain tissues of AD patients (p < 0.01, Fig. 5a, b). In primary cultured astrocytes, Aβ treatment for 1 h significantly decreased p-AKT (Thr308) levels without inducing changes in total AKT (p < 0.01, Fig. 5c, d), while KCa3.1 gene deletion (KCa3.1^−/−) inhibited the decrease in p-AKT (Thr308) after Aβ treatment (p < 0.05, Fig. 5c, d).

We then examined whether KCa3.1 gene deletion would activate the AKT/mTOR pathway in an in vivo AD model. Consistent with the in vitro results (Fig. 5c), p-AKT (Thr308) levels were decreased in the brain tissue of APP/PS1 mice (p < 0.01, Fig. 5e, f). KCa3.1 gene deletion in APP/PS1 mice (KCa3.1^−/−/APP/PS1) attenuated p-AKT (Thr308) suppression (p < 0.05, Fig. 5e, f). Importantly, the phosphorylation of mTOR, a kinase that regulates cell survival, was decreased in the brain tissue of APP/PS1 mice (p < 0.01, Fig. 5e, f). mTOR suppression, in turn, suppressed its downstream proteins, including phosphorylated 4EBP1 (p-4EBP1) and p70 S6 kinase (p-p70 S6 kinase) (Thr389), while KCa3.1 gene deletion in APP/PS1 mice (KCa3.1^−/−/APP/PS1) attenuated the suppression of mTOR phosphorylation (p < 0.05, Fig. 5e, f).

It was reported that blockade of KCa3.1 significantly reduced neuronal loss and memory deficits in both APP/PS1 mice and SAMP8 mice [11, 15]. Given that neuronal loss and decreased expression of synaptic proteins are likely correlated with the severity of AD [30], we compared expression levels of the neuron marker NeuN in the brain tissues of APP/PS1 mice with those of KCa3.1^−/−/APP/PS1 mutants. Immunostaining analysis revealed that NeuN-positive neuron levels were decreased in the CA1 areas of the hippocampi of APP/PS1 mice, as compared with that in the WT group, while KCa3.1 gene deletion in APP/PS1 mice (KCa3.1^−/−/APP/PS1) attenuated the decrease of NeuN staining (p < 0.05, Fig. 6a, b).

Both reactive astrocytes and microglial activation are predominant pathological features of AD [31]. Immunostaining of the RA marker GFAP was measured to determine whether KCa3.1 gene deletion affects RA in APP/PS1 mice. Immunostaining of GFAP⁺ astrocytes was increased in the brains of APP/PS1 mice, as compared with WT mice (p < 0.05, Fig. 7a, b), while KCa3.1 gene deletion in APP/PS1 mice (KCa3.1^−/−/APP/PS1) attenuated the increase in immunostaining of GFAP⁺ astrocytes (p < 0.05, Fig. 7a, b).

We also investigated whether KCa3.1 gene deletion attenuates microglial activation in APP/PS1 mice by measuring expression levels of the microglia marker Iba1. Our data show that the content of Iba1⁺ microglia was increased in the brains of APP/PS1 mice, as compared with that of WT mice (p < 0.01, Fig. 7c, d). KCa3.1 gene deletion in APP/PS1 mice (KCa3.1^−/−/APP/PS1) attenuated the increase of Iba1⁺ microglia immunostaining (p < 0.01, Fig. 7c, d). Both the RA and microglia activation results suggest that, while brains of APP/PS1 mice show increased glial responses, these responses are attenuated in the brains of KCa3.1^−/−/APP/PS1 mice.

KCa3.1 deficiency in KCa3.1^−/−/APP/PS1 mice attenuated the upregulation of COX-2 (Fig. 7e, f) and iNOS (Fig. 7e, g), compared with APP/PS1 mice. Levels of TNF-α and IL-1β in brain homogenates from WT, KCa3.1^−/−, APP/PS1, and KCa3.1^−/−/APP/PS1 group mice were measured by ELISA experiments (Fig. 7h, i). Levels of both TNF-α and IL-1β released were attenuated in the KCa3.1^−/−/APP/PS1 group compared with the APP/PS1 group. Together, these findings indicate that elimination of KCa3.1 ameliorated the pathological hallmarks of AD in KCa3.1^−/−/APP/PS1 mice, such as neuronal loss, microglial activation, RA, and neuroinflammation.

In this study, the MWM spatial memory test was used to determine whether loss of KCa3.1 had any effect on cognitive deficits in APP/PS1 mice. With the use of a hidden platform during the MWM test, 15-month-old APP/PS1 mice showed memory deficits, as compared with WT mice (p < 0.01, Fig. 8a). Furthermore, the KCa3.1^−/−/APP/PS1 mice showed improved spatial learning and memory, as compared with the APP/PS1 mice (Fig. 8a). In the spatial probe trial without an escape platform, significantly more KCa3.1^−/−/APP/PS1 mice were able to cross the target quadrant, as compared to APP/PS1 mice (p < 0.05, Fig. 8b), and spent more time and swam for greater distances in the target quadrant (p < 0.05, Fig. 8c, d). In conclusion, the MWM test results indicate that APP/PS1 mice present significant spatial memory deficits and that elimination of KCa3.1 channels ameliorates these deficits.