Dense core vesicles (DCVs) and synaptic vesicles are specialised secretory vesicles in neurons and neuroendocrine cells, and abnormal release of their cargo is associated with various pathophysiologies. Endoplasmic reticulum (ER) stress and inter-organellar communication are also associated with disease biology. To investigate the functional status of regulated exocytosis arising from the crosstalk of a stressed ER and DCVs, ER stress was modelled in PC12 neuroendocrine cells using thapsigargin. DCV exocytosis was severely compromised in ER-stressed PC12 cells and was reversed to varying magnitudes by ER stress attenuators. Experiments with tunicamycin, an independent ER stressor, yielded similar results. Concurrently, ER stress also caused impaired DCV exocytosis in insulin-secreting INS-1 cells. Molecular analysis revealed blunted SNAP25 expression, potentially attributed to augmented levels of ATF4, an inhibitor of CREB that binds to the CREB-binding site. The effects of loss of function of ATF4 in ER-stressed cells substantiated this attribution. Our studies revealed severe defects in DCV exocytosis in ER-stressed cells for the first time, mediated by reduced levels of key exocytotic and granulogenic switches regulated via the eIF2α (EIF2A)–ATF4 axis.

Disruption of protein homeostasis and vesicle trafficking is associated with pathological conditions such as Alzheimer's disease (Ashraf et al., 2014; Barranco et al., 2021), Parkinson's disease (Zou et al., 2021) and amyotrophic lateral sclerosis (Cicardi et al., 2021; Gagliardi et al., 2021), to name a few (Sweeney et al., 2017; Wang et al., 2014). The endoplasmic reticulum (ER) is the principal site of protein synthesis, folding and sorting, and becomes compromised under stress. An ER-stressed cell uses the unfolded protein response (UPR) as a survival strategy mediated by three pathways: IRE1-α, PERK and ATF6. They initiate signalling cascades to regulate transcriptional and translational alterations, enhancing protein folding and cell survival against ER stress (Shen et al., 2004). ER stress leads to IRE1-α activation, causing changes in spliced XBP1 levels. XBP1 regulates the activity of several genes, which ultimately upregulates the expression of chaperones to counteract the ER stress (or the UPR). Similarly, ATF6 activation contributes to the UPR and determines cell fate to microautophagy. In contrast, PERK activation attenuates global translation; upon sustained stressed conditions, it induces apoptosis through the PERK–ATF4–CHOP pathway (Hetz, 2012). Dense core vesicles (DCVs) are specialised subcellular organelles in neurons and neuroendocrine cells in which peptides, hormones and neurotransmitters are stored. They undergo regulated secretion when an appropriate physiological stimulus is provided. DCVs comprise two major structural compartments, the matrix and the membrane. The matrix includes chromogranin and/or secretogranin family proteins, out of which chromogranin A (CGA or CHGA), chromogranin B (CGB) and secretogranin II (SCGII or SCG2) are the most important as they participate in the loading of neuropeptides into secretory vesicles. Their functionality also lies in regulating the matrix constituent and its exocytosis (Huttner et al., 1991). The release of cargo to the extracellular space is highly regulated and coupled with membrane depolarisation and Ca2+ influx, ultimately leading to the docking and release of the vesicle (Barclay et al., 2005; Neher and Sakaba, 2008). Neuroendocrine cells also have synaptic-like vesicles (SLVs) similar to neurons, packed with neurotransmitters, which help in intercellular communication (Shi et al., 1998). The release of hormones, neurotransmitters, peptides and other molecules of physiological relevance is tightly regulated by vesicle (v-)SNAREs (VAMP1) and target (t-)SNAREs (SNAP25).

Neuropeptides and neurotransmitters are DCV cargoes regulating diverse functions such as sleep, behaviour, cardiovascular regulation, body weight and blood glucose levels, and their abnormal levels are associated with various diseases, including neurological disorders (Beal and Martin, 1986; Ganten et al., 1991; McCall, 1990; Teleanu et al., 2022; Yang et al., 2021). Likewise, a stressed ER causes disturbances in cellular homeostasis by altering functions such as protein folding and is associated with several conditions such as Huntington's disease and diabetes (Yoshida, 2007). It is noteworthy that misfolded proteins, such as mutated α-synuclein, are associated with disrupted fusion pore formation during exocytosis (Burré et al., 2010; Sulzer and Edwards, 2019), neurodegeneration (Bennett, 2005) and ER stress (Colla et al., 2012). This unfolds potential common crosstalk between ER stress and DCV trafficking. Although DCV biogenesis and release have been studied for decades, their alterations in response to ER stress and subsequent effects have not yet been explored. A growing body of evidence suggests that ER stress can potentially impact regulated secretion, which has been implicated in defective insulin biogenesis and subsequent insulin release (Yong et al., 2016, 2021). The molecular underpinnings beyond this are also not known. Besides, whether this is true across other cell lineage-specific regulated secretory vesicles, such as DCVs storing monoamines and neuropeptides, and SLVs storing neurotransmitters, is not clearly understood. Notably, insulin granules differ from DCVs in several aspects related to signature proteins and exocytotic machinery. Likewise, synaptic vesicles differ in release kinetics and composition (Wegrzyn et al., 2010; Brunner et al., 2007; Lang et al., 1997; Shin et al., 2002; Gondré-Lewis et al., 2012). This study was conceived based on these points with the hope of gaining clear insights into the impact of ER stress on regulated exocytosis with some understanding of the molecular underpinnings.

To study the association of DCV and SLV exocytosis with ER stress, we chose PC12 cells, a well-known cellular model widely used to study the phenomenon of exocytosis and pharmacologically induced ER stress by thapsigargin (Tg) treatment as described previously (Lindner et al., 2020; Westerink and Ewing, 2008). Following this, we evaluated the functional status of DCVs, which suggested strongly compromised DCV exocytosis in ER-stressed cells. To test the specificity of the contribution of ER stress in impaired DCV exocytosis, we investigated the functional status of regulated exocytosis in ER-stressed cells supplemented with docosahexaenoic acid (DHA), an ER stress attenuator (Begum et al., 2014). To our surprise, impaired regulated secretion was restored to near physiological levels in Tg- and DHA-co-treated cells. These findings were similar using a different ER stress attenuator, sodium 4-phenylbutyrate (PBA), albeit to a different magnitude (Movie 11). Transmission electron microscopy (TEM) revealed that ER stress did not change the size and number but altered the spatial localisation of DCVs. Molecular analysis revealed ‘expression switching,’ i.e. altered abundances of key granulogenic proteins and exocytotic signature candidates (CGA, SCGII and SNAP25) were associated with an impediment in exocytosis via the eIF2α (EIF2A)–ATF4 signalling cascade. This was further accompanied by enhanced localisation of CGA to the lysosomes. Intriguingly, SLVs also displayed impaired exocytosis, which did not completely reverse to normal following DHA treatment.

Furthermore, similar experiments using tunicamycin, an independent ER stressor, caused similar effects on impaired exocytosis of regulated secretion. Finally, we also tested the functional status of DCVs in Tg-mediated ER-stressed insulin-secreting (INS-1) cells, which also suggested exocytotic defects of DCVs. In summary, our studies for the first time provide evidence that ER stress has detrimental effects on DCV exocytosis, plausibly resulting from cumulative effects of expression switching of SNAP25 and CGA, potentially modulated by eIF2α–ATF4 axis.

Thapsigargin causes ER stress in PC12 cells without inducing apoptosis

Thapsigargin (Tg) has been used as a pharmacological ER stress inducer in the PC12 cell line (Szegezdi et al., 2008). We carried out a cell viability assay for various doses of Tg and a specific dose of 10 nM that had minimum cell death but was able to activate the ER stress pathways was chosen for all subsequent experiments (Fig. 1A,B) (Brodnanova et al., 2021). At this specific dose, 80% of the cells were viable without enhanced expression of the pro-apoptotic marker caspase 3 (Casp3) (Fig. 1C) but with the activation of the PERK–eIF2α arm of the ER stress pathway, as evident by the phosphorylation of eIF2α (p-eIF2α; Fig. 1D) (Novoa et al., 2003). However, the other two arms of the UPR (indicated by xbp1 and atf6 expression) were unaltered (Fig. S1A,B) in Tg-treated cells throughout various time points starting from 6 h (Shen et al., 2004). Further, BIP (or HSPA5) levels also remained unaltered (Fig. S1C,D).

Fig. 1.

Thapsigargin causes ER stress in PC12 cells. (A) Treatment protocol: 24 h after plating, cells were treated with 10 nM thapsigargin (Tg) under reduced serum conditions, and experiments were performed after 24 h. (B) Cell viability assay for different Tg doses showed around 80% cell viability at 10 nM dosage (one-way ANOVA with Tukey's multiple comparison test, N=6). (C) Transcript levels of caspase 3 (Casp3) in control and Tg-treated cells (Mann–Whitney test, N=4). (D) Representative immunoblot showing p-eIF2α and total eIF2α levels (left) and their corresponding quantification (right) in control and Tg-treated cells. β-tubulin was used as a loading control (two-tailed unpaired Student's t-test, N=5). N denotes the number of biological replicates. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; ****P<0.0001.

Fig. 1.

Thapsigargin causes ER stress in PC12 cells. (A) Treatment protocol: 24 h after plating, cells were treated with 10 nM thapsigargin (Tg) under reduced serum conditions, and experiments were performed after 24 h. (B) Cell viability assay for different Tg doses showed around 80% cell viability at 10 nM dosage (one-way ANOVA with Tukey's multiple comparison test, N=6). (C) Transcript levels of caspase 3 (Casp3) in control and Tg-treated cells (Mann–Whitney test, N=4). (D) Representative immunoblot showing p-eIF2α and total eIF2α levels (left) and their corresponding quantification (right) in control and Tg-treated cells. β-tubulin was used as a loading control (two-tailed unpaired Student's t-test, N=5). N denotes the number of biological replicates. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; ****P<0.0001.

ER stress inhibits exocytosis of DCVs

Cell lines stably expressing NPY–mApple (fluorophore tagged to neuropeptide Y, a soluble DCV cargo) were generated and the increase in NPY–mApple fluorescence intensities upon treatment with an external stimulus was quantified using a plate reader as a measure of regulated secretion. ER-stressed cells displayed blunted response to stimulus-coupled secretion of NPY–mApple, suggesting an impediment in cargo release from DCVs (Fig. 2A,B).

Fig. 2.

ER stress causes an impediment to regulated secretion in DCVs. (A) Schematic showing transfection and treatment protocol for imaging and plate reader assay experiments. (B) Quantitative measurement of NPY–mApple fluorescence intensity in control and Tg-treated PC12 cells transfected with NPY–mApple showing the percentage of secretion with respect to total NPY–mApple levels, measured using a plate reader (one-way ANOVA with Tukey's multiple comparison test, N=4). Bas, basal; Stim, stimulated. (C) Representative images of NPY–pHTomato (red)-expressing PC12 cells showing basal and KCl-stimulated conditions. Scale bars: 10 µm. (D) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg). (E) Quantitative analysis of the maximum change in NPY–pHTomato puncta fluorescence relative to basal fluorescence (Δf/f0) in control and Tg-treated PC12 cells (two-tailed unpaired Student's t-test, control, n=10; Tg, n=16). (F,G) Numbers of responding (Mann–Whitney test) (F) and non-responding (two-tailed unpaired Student's t-test) (G) NPY–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=10; Tg, n=16) (N=3). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; ****P<0.0001.

Fig. 2.

ER stress causes an impediment to regulated secretion in DCVs. (A) Schematic showing transfection and treatment protocol for imaging and plate reader assay experiments. (B) Quantitative measurement of NPY–mApple fluorescence intensity in control and Tg-treated PC12 cells transfected with NPY–mApple showing the percentage of secretion with respect to total NPY–mApple levels, measured using a plate reader (one-way ANOVA with Tukey's multiple comparison test, N=4). Bas, basal; Stim, stimulated. (C) Representative images of NPY–pHTomato (red)-expressing PC12 cells showing basal and KCl-stimulated conditions. Scale bars: 10 µm. (D) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg). (E) Quantitative analysis of the maximum change in NPY–pHTomato puncta fluorescence relative to basal fluorescence (Δf/f0) in control and Tg-treated PC12 cells (two-tailed unpaired Student's t-test, control, n=10; Tg, n=16). (F,G) Numbers of responding (Mann–Whitney test) (F) and non-responding (two-tailed unpaired Student's t-test) (G) NPY–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=10; Tg, n=16) (N=3). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; ****P<0.0001.

To further assess the possible defect in DCV exocytosis following Tg treatment, we transfected PC12 cells with NPY–pHTomato (a pH-sensitive DCV marker) (Gandasi et al., 2015; Hummer et al., 2017). We then observed the change in NPY–pHTomato puncta fluorescence in response to 100 mM KCl stimulation and individual exocytotic events were monitored by live-cell imaging using a spinning-disk confocal microscope. 100 mM KCl acts as a secretagogue and is known to induce depolarisation of the plasma membrane, leading to the fusion and exocytosis of DCVs. We observed increased NPY–pHTomato fluorescence upon 100 mM KCl stimulation in control cells. However, we saw distinct non-responsive dynamics in ER-stressed cells (Fig. 2C,D, Movie 1). Next, we quantified the change in NPY–pHTomato puncta fluorescence (Δf) relative to basal fluorescence (f0) and plotted maximum Δf/f0 to measure DCV exocytosis. We found a significant reduction in DCV exocytosis in Tg-treated cells compared to that in control cells (Fig. 2E). These findings were in line with the NPY–mApple plate reader-based secretion experiments. Furthermore, we assessed the global response of NPY–pHTomato by analysing individual exocytotic events. We observed an overall decrease in responsiveness as seen by increased non-responsive NPY–pHTomato puncta in ER-stressed cells (Fig. 2F,G) (Hummer et al., 2017). We also investigated the functional status of DCV exocytosis in ER-stressed INS-1 cells by NPY–pHluorin imaging and observed similarly compromised DCV exocytosis (Fig. S2A–E; Movie 9) (Ji et al., 2017). To rule out the observed cellular effect as a consequence of Tg, not ER stress, we tested another ER stressor with a different mechanism of action, tunicamycin (Tm) (Guha et al., 2017). Tm also induced ER stress without driving the cell towards apoptosis at a 0.05 µg/ml dose (Fig. S3A,B); hence, this dose was chosen for treatment. Tm induced the phosphorylation of eIF2α (Fig. S3C) without altering CHOP and XBP1 levels (Fig. S3D–F) and impeded the secretion of DCVs, similar to the effects of Tg, and DCV secretion was subsequently restored to normal levels upon DHA treatment (Fig. S3H–L; Movie 10). Our findings show an impediment in DCV exocytosis in multiple cellular models of ER stress using multiple ER stressors.

ER stress causes defects in regulated exocytosis in SLVs

Neuroendocrine cells also have SLVs apart from DCVs. Extending our observation to SLVs, we investigated the functional status of regulated secretion in SLVs. PC12 cells were transfected with synaptophysin–pHTomato. Synaptophysin (SYP) is a membrane glycoprotein that localises to synaptic vesicles in neurons and to SLVs in neuroendocrine cells and hence serves as an important marker for studying the dynamics of such vesicles (Kwon and Chapman, 2011). Individual exocytotic events were tracked by live-cell imaging using a spinning-disk confocal microscope under a 63× oil objective. Tg-treated cells showed exocytotic defects and non-responsive dynamics of SLVs upon 100 mM KCl stimulation compared to control cells, which responded with a fluorescence burst upon stimulation with the same secretagogue (Fig. 3A–C; Movie 2). We calculated the maximum Δf/f0 as a measure of synaptic vesicle exocytosis, thus reflecting the change in fluorescence between the basal and stimulated conditions. Stressed cells showed reduced to almost no response upon stimulation with KCl secretagogue compared to control cells (Fig. 3D,E) and an increase in the number of non-responsive synaptophysin–pHTomato puncta (Fig. 3F). Hence, a stress-induced impediment to regulated secretion is replicated in SLVs, similar to in DCVs.

Fig. 3.

ER stress causes an impediment to regulated secretion in SLVs. (A) Schematic showing transfection and treatment protocol for imaging experiments. (B) Representative images of synaptophysin–pHTomato (red)-expressing PC12 cells showing basal and KCl-stimulated conditions. Scale bars: 10 µm. (C) Representative time traces of mean synaptophysin–pHTomato fluorescence intensities showing the average of all puncta per cell (green, control; red, Tg-treated cells). (D) Quantitative analysis of the maximum Δf/f0 in control and Tg-treated PC12 cells (Mann–Whitney test; control, n=7; Tg, n=11; N=3). (E,F) Numbers of responding (Mann–Whitney test) (E) and non-responding (two-tailed unpaired Student's t-test) (F) synaptophysin–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=7; Tg, n=11; N=3). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; **P<0.01; ***P<0.001.

Fig. 3.

ER stress causes an impediment to regulated secretion in SLVs. (A) Schematic showing transfection and treatment protocol for imaging experiments. (B) Representative images of synaptophysin–pHTomato (red)-expressing PC12 cells showing basal and KCl-stimulated conditions. Scale bars: 10 µm. (C) Representative time traces of mean synaptophysin–pHTomato fluorescence intensities showing the average of all puncta per cell (green, control; red, Tg-treated cells). (D) Quantitative analysis of the maximum Δf/f0 in control and Tg-treated PC12 cells (Mann–Whitney test; control, n=7; Tg, n=11; N=3). (E,F) Numbers of responding (Mann–Whitney test) (E) and non-responding (two-tailed unpaired Student's t-test) (F) synaptophysin–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=7; Tg, n=11; N=3). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; **P<0.01; ***P<0.001.

DHA reverses the defective DCV exocytosis phenotype by reducing ER stress

DHA, one of the most abundant omega-3 fatty acids in the brain, has been previously shown to attenuate ER stress in several in vitro studies (Begum et al., 2012, 2013, 2014; Dyall, 2015). In line with previous studies, we saw a reduced activation of ER stress as indexed by restoring the phosphorylated (p-)eIF2α/total eIF2α ratio to near control levels (Fig. 4A,B). This gave us the first line of data implying the reversal of the ER stress phenotype in Tg-treated cells upon co-treatment with DHA.

Fig. 4.

DHA rescues the phenotype by reducing ER stress in DCVs. (A) Treatment protocol: after plating, cells were serum-starved for 2 h, followed by pre-incubating the cells with 10 µM DHA. After 24 h, 10 nM Tg was given to the cells to induce ER stress in the Tg+DHA-co-treated and Tg-alone cells. (B) Representative immunoblots (left) and quantification (right) of protein levels of p-eIF2α and total eIF2α. β-tubulin and EF2 served as a loading control [two-tailed unpaired Student's t-test, control, Tg, Tg+DHA, N=6 (top); control, DHA only, N=3 (bottom)]. (C) Quantitative measurement of NPY–mApple fluorescence intensities in control, Tg-treated, Tg+DHA-co-treated and only-DHA-treated PC12 cells transfected with NPY–mApple, showing the percentage of secretion (one-way ANOVA with Tukey's multiple comparison test, N=3). Bas, basal; Stim, stimulated. (D) Quantification of total basal NPY–mApple fluorescence intensities (cell lysates and supernatants) (two-tailed unpaired Student's t-test, N=3). A.U., arbitrary units. (E) Coomassie staining showing total protein contents of constitutive secretion. (F) Immunoblots of the constitutive secretion marker HSP90. N=3. (G) Representative NPY–pHTomato (red) images in PC12 cells showing basal and KCl-stimulated conditions in control, Tg- and Tg+DHA-co-treated cells. Scale bars: 10 µm. (H) Quantitative analysis of the maximum Δf/f0 in control, Tg-treated, and Tg+DHA-co-treated PC12 cells (Mann–Whitney test; control, n=12; Tg, n=15; Tg+DHA, n=13; N=3). (I) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, Tg+DHA-co-treated cells). (J,K) Numbers of responding (J) and non-responding (K) NPY–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (Mann–Whitney test; control, n=12; Tg, n=15; Tg+DHA, n=13; N=3). (L) Representative NPY–pHTomato (red) fluorescence images in PC12 cells showing basal and KCl-stimulated conditions in control, Tg-treated and DHA only cells. Scale bars: 10 µm. (M) Quantitative analysis of the maximum Δf/f0 in control and DHA-only treated PC12 cells (Mann–Whitney test; control, n=10; Tg, n=8; DHA n=8; N=3). (N) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, DHA-only treated cells). N denotes the number of biological replicates and n is the number of cells taken for quantification. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

Fig. 4.

DHA rescues the phenotype by reducing ER stress in DCVs. (A) Treatment protocol: after plating, cells were serum-starved for 2 h, followed by pre-incubating the cells with 10 µM DHA. After 24 h, 10 nM Tg was given to the cells to induce ER stress in the Tg+DHA-co-treated and Tg-alone cells. (B) Representative immunoblots (left) and quantification (right) of protein levels of p-eIF2α and total eIF2α. β-tubulin and EF2 served as a loading control [two-tailed unpaired Student's t-test, control, Tg, Tg+DHA, N=6 (top); control, DHA only, N=3 (bottom)]. (C) Quantitative measurement of NPY–mApple fluorescence intensities in control, Tg-treated, Tg+DHA-co-treated and only-DHA-treated PC12 cells transfected with NPY–mApple, showing the percentage of secretion (one-way ANOVA with Tukey's multiple comparison test, N=3). Bas, basal; Stim, stimulated. (D) Quantification of total basal NPY–mApple fluorescence intensities (cell lysates and supernatants) (two-tailed unpaired Student's t-test, N=3). A.U., arbitrary units. (E) Coomassie staining showing total protein contents of constitutive secretion. (F) Immunoblots of the constitutive secretion marker HSP90. N=3. (G) Representative NPY–pHTomato (red) images in PC12 cells showing basal and KCl-stimulated conditions in control, Tg- and Tg+DHA-co-treated cells. Scale bars: 10 µm. (H) Quantitative analysis of the maximum Δf/f0 in control, Tg-treated, and Tg+DHA-co-treated PC12 cells (Mann–Whitney test; control, n=12; Tg, n=15; Tg+DHA, n=13; N=3). (I) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, Tg+DHA-co-treated cells). (J,K) Numbers of responding (J) and non-responding (K) NPY–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (Mann–Whitney test; control, n=12; Tg, n=15; Tg+DHA, n=13; N=3). (L) Representative NPY–pHTomato (red) fluorescence images in PC12 cells showing basal and KCl-stimulated conditions in control, Tg-treated and DHA only cells. Scale bars: 10 µm. (M) Quantitative analysis of the maximum Δf/f0 in control and DHA-only treated PC12 cells (Mann–Whitney test; control, n=10; Tg, n=8; DHA n=8; N=3). (N) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, DHA-only treated cells). N denotes the number of biological replicates and n is the number of cells taken for quantification. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

We studied the pathway connected to ER stress inhibition and DCV exocytosis and biogenesis using DHA. Interestingly, DHA restored the impaired regulated exocytosis of DCV in Tg-treated cells to near control levels, as seen from the NPY–mApple fluorescence-based plate reader assay (Fig. 4C,D). We then checked for the signature abundance of the constitutively secreted proteome by staining the gel containing cell lysates with Coomassie Blue (Fig. 4E) and also by immunoblotting for the specific constitutive secretory marker HSP90 (with an antibody specific to HSP90AA1) at basal conditions without stimulation (Fig. 4F) (Guo et al., 2017; Hummer et al., 2017). This ruled out impediments in the constitutive secretory pathway.

Additionally, we also observed an overall shift in the dynamics of DCV exocytosis as that of control cells upon Tg+DHA co-treatment (Fig. 4G–I; Movies 3 and 4) and in DHA-only treated cells (Fig. 4L–N). We also observed that the relative responsiveness of the cells to stimulus-coupled secretion was restored, as evident from the increased numbers of responsive NPY–pHTomato puncta upon Tg+DHA co-treatment (Fig. 4J,K). Hence, DHA treatment as an attenuator of ER stress causes a change in DCV dynamics by the reversal of the impaired regulated secretion phenotype as seen in the Tg-treated cells towards normal conditions. These experiments convey the specificity of the ER stress-associated perturbed regulated exocytosis, besides flagging the therapeutic potential of DHA in conditions where ER stress causes compromised regulated exocytosis. Additionally, we tested this with another stress inhibitor, PBA, and observed a similar reversal in impairment, although it was to a lesser magnitude (Fig. S4A–D).

Tg treatment does not change the size and number but alters the spatial localisation of DCVs

In light of the severe disruption in DCV exocytosis, we wanted to estimate the size and distribution of the vesicles. TEM experiments followed by morphometry analyses suggested non-significant differences in the DCV size and number (Fig. 5A–D,F) across all treatments. However, DCVs were significantly closer to the plasma membrane in the control cells than in the ER-stressed counterparts (Fig. 5E) and this was mildly reversed upon DHA treatment.

Fig. 5.

Transmission electron microscopy of PC12 cells. (A) Representative transmission electron micrograph showing electron-dense vesicles. (B–D) Quantification of the number of DCVs by cell surface area (in μm2) (B), dense core (DC) diameter) (C) and DCV diameter (D) (two-tailed unpaired Student's t-test; control, n=5; Tg, n=3; Tg+DHA, n=5; N=2) showing no significant difference. Data are shown as mean±s.e.m. (E) Quantification of DCV distance from the nearest plasma membrane, showing significant difference in DCV localisation from the membrane (Mann–Whitney test; control, n=5; Tg, n=3; Tg+DHA, n=5; N=2). Box plots show the 25–75th percentiles, whiskers show the range, and the median is marked with a line. (F) Histogram of DCV diameter distribution. N denotes the number of biological replicates and n is the number of cells taken for quantification. ns, statistically non-significant; **P<0.01.

Fig. 5.

Transmission electron microscopy of PC12 cells. (A) Representative transmission electron micrograph showing electron-dense vesicles. (B–D) Quantification of the number of DCVs by cell surface area (in μm2) (B), dense core (DC) diameter) (C) and DCV diameter (D) (two-tailed unpaired Student's t-test; control, n=5; Tg, n=3; Tg+DHA, n=5; N=2) showing no significant difference. Data are shown as mean±s.e.m. (E) Quantification of DCV distance from the nearest plasma membrane, showing significant difference in DCV localisation from the membrane (Mann–Whitney test; control, n=5; Tg, n=3; Tg+DHA, n=5; N=2). Box plots show the 25–75th percentiles, whiskers show the range, and the median is marked with a line. (F) Histogram of DCV diameter distribution. N denotes the number of biological replicates and n is the number of cells taken for quantification. ns, statistically non-significant; **P<0.01.

DHA reverses the impediment to regulated secretion in SLVs

In line with our previous observation, we checked the effect of Tg+DHA co-treatment on the exocytosis of SLVs in response to 100 mM KCl in ER-stressed cells. Although there was a subtle restoration of impaired regulated secretion of SLVs in Tg+DHA-co-treated ER-stressed cells, the magnitude of restoration was not on the order of DCV exocytosis (Fig. 6A–C; Movies 5 and 6). We also observed that most of the synaptophysin–pHTomato puncta showed non-responsive behaviour in Tg-treated cells (Fig. 6D,E), which was rescued to near control levels upon co-treatment with DHA. Treatment with DHA alone did not have any effect on synaptophysin-pHTomato exocytosis (Fig. 6F–H). We report that treatment with DHA, as an inhibitor of ER stress, also causes a mild reversion in SLV dynamics towards normal or wild-type conditions, along with the reversal of the impaired regulated secretion phenotype as seen in the ER-stressed cells.

Fig. 6.

DHA rescues the phenotype by reducing ER stress in synaptic-like vesicles. (A) Representative images of synaptophysin–pHTomato (red) in PC12 cells showing basal and KCl-stimulated conditions in control, Tg-treated and Tg+DHA-co-treated cells. Scale bars: 10 µm. (B) Quantitative analysis of the maximum Δf/f0 in control, Tg-treated and Tg+DHA-co-treated PC12 cells (two-tailed unpaired Student's t-test; control, Tg, Tg+DHA, n=8, N=3). (C) Representative time traces of mean synaptophysin–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, Tg+DHA-co-treated cells). (D,E) Numbers of responding (Mann–Whitney test) (D) and non-responding (two-tailed unpaired Student's t-test) (E) synaptophysin–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, Tg, Tg+DHA, n=8, N=3). (F) Representative synaptophysin–pHTomato (red) fluorescence images in PC12 cells showing basal and KCl-stimulated conditions in control, Tg and DHA only cells. Scale bars: 10 µm. (G) Quantitative analysis of the maximum Δf/f0 in control, Tg- and DHA-only-treated PC12 cells (Mann–Whitney test; control, n=8; Tg, n=6; DHA n=9; N=3). (H) Representative time traces of mean synaptophysin–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, DHA-only-treated cells). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

Fig. 6.

DHA rescues the phenotype by reducing ER stress in synaptic-like vesicles. (A) Representative images of synaptophysin–pHTomato (red) in PC12 cells showing basal and KCl-stimulated conditions in control, Tg-treated and Tg+DHA-co-treated cells. Scale bars: 10 µm. (B) Quantitative analysis of the maximum Δf/f0 in control, Tg-treated and Tg+DHA-co-treated PC12 cells (two-tailed unpaired Student's t-test; control, Tg, Tg+DHA, n=8, N=3). (C) Representative time traces of mean synaptophysin–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, Tg+DHA-co-treated cells). (D,E) Numbers of responding (Mann–Whitney test) (D) and non-responding (two-tailed unpaired Student's t-test) (E) synaptophysin–pHTomato puncta/cell, considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, Tg, Tg+DHA, n=8, N=3). (F) Representative synaptophysin–pHTomato (red) fluorescence images in PC12 cells showing basal and KCl-stimulated conditions in control, Tg and DHA only cells. Scale bars: 10 µm. (G) Quantitative analysis of the maximum Δf/f0 in control, Tg- and DHA-only-treated PC12 cells (Mann–Whitney test; control, n=8; Tg, n=6; DHA n=9; N=3). (H) Representative time traces of mean synaptophysin–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; blue, DHA-only-treated cells). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

ER stress alters the abundance of signature genes associated with exocytosis in an ATF4-dependent manner

We conducted experiments to understand the molecular basis for impaired exocytosis by pharmacological ER stress triggered by Tg. Previous reports suggested blunted expression of key exocytotic genes such as Snap25, syntaxin 1A (Stx1a), Vamp2 (Liew et al., 2010; Goda, 1997) and Syt1 (Sørensen et al., 2003), causing failure of exocytosis. Hence, we hypothesised that diminished expression of these genes can cause impaired exocytosis, and estimated the abundances of key SNAREs, tethers and Rab proteins. The results suggested that, except for SNAP25 (Fig. 7A,B), other candidates did not show a significant change in expression (Fig. S1E–J). These findings were consistent with those for another ER stressor, Tm (Fig. S3G). Additionally, when we overexpressed constitutively active SNAP25–GFP in Tg-stressed cells, we noticed a significant rescue of the impaired regulated exocytosis. These findings strengthen the definitive role of SNAP25 in the disruption of regulated secretion in ER-stressed cells (Fig. 7K–P; Movie 7).

Fig. 7.

ER stress alters the abundance of signature gene expression associated with exocytosis in an ATF4-dependent manner. (A) Quantification of the transcript levels of Snap25, encoding a t-SNARE that helps in DCV exocytosis, by qRT-PCR (N=4). (B) Representative blots of SNAP25 (left) and their corresponding quantification. EF2 served as a loading control (N=4). (C) Quantification of Atf4 transcript levels by qRT-PCR (N=3). (D) Representative blots of ATF4 (left) and their corresponding quantification (right). GAPDH served as a loading control (N=3). (E) Schematic representation of the transcriptional switch of CREB during augmented ATF4. (F,G) Representative blots of ATF4 (N=3) (F, left) and SNAP25 (N=5) (G, top) and their corresponding quantification (F, right; G, bottom) upon treatment with ISRIB. EF2 served as a loading control. (H–J) Representative blots of SNAP25 (top) and their corresponding quantification (bottom) in non-targeting (NT) (N=4) (H), shRNA 1 (Sh1) (I) and shRNA 2 (Sh2) (J) Atf4 knockdown (N=3) cells with or without 5 µg/ml doxycycline (Dox) treatment. EF2 and β-tubulin served as loading controls. Two-tailed unpaired Student's t-test was used for statistical analysis in A–D,F–I. (K) Schematic showing the overexpression and imaging protocol of SNAP25–GFP overexpression. (L) Representative fluorescence images of NPY tagged with pHTomato (red) showing basal and KCl-stimulated conditions for control cells, Tg-treated cells and Tg-treated cells ectopically expressing SNAP25. Images beneath show both SNAP25–GFP (green) and NPY–pHTomato (red) signals for a cell ectopically expressing SNAP25. Scale bars: 10 µm. (M) Quantitative analysis of the maximum Δf/f0 in control, Tg and Tg+SNAP25 PC12 cells (Mann–Whitney test; control, n=12; Tg, n=8; Tg+SNAP25, n=12; N=3). (N) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; pink, Tg+SNAP25-treated cells). (O,P) Numbers of responding (O) and non-responding (P) NPY–pHTomato puncta/cell (Mann–Whitney test), considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=12; Tg, n=8; Tg+DHA, n=12; N=3). N denotes the number of biological replicates and n is number of cells. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

Fig. 7.

ER stress alters the abundance of signature gene expression associated with exocytosis in an ATF4-dependent manner. (A) Quantification of the transcript levels of Snap25, encoding a t-SNARE that helps in DCV exocytosis, by qRT-PCR (N=4). (B) Representative blots of SNAP25 (left) and their corresponding quantification. EF2 served as a loading control (N=4). (C) Quantification of Atf4 transcript levels by qRT-PCR (N=3). (D) Representative blots of ATF4 (left) and their corresponding quantification (right). GAPDH served as a loading control (N=3). (E) Schematic representation of the transcriptional switch of CREB during augmented ATF4. (F,G) Representative blots of ATF4 (N=3) (F, left) and SNAP25 (N=5) (G, top) and their corresponding quantification (F, right; G, bottom) upon treatment with ISRIB. EF2 served as a loading control. (H–J) Representative blots of SNAP25 (top) and their corresponding quantification (bottom) in non-targeting (NT) (N=4) (H), shRNA 1 (Sh1) (I) and shRNA 2 (Sh2) (J) Atf4 knockdown (N=3) cells with or without 5 µg/ml doxycycline (Dox) treatment. EF2 and β-tubulin served as loading controls. Two-tailed unpaired Student's t-test was used for statistical analysis in A–D,F–I. (K) Schematic showing the overexpression and imaging protocol of SNAP25–GFP overexpression. (L) Representative fluorescence images of NPY tagged with pHTomato (red) showing basal and KCl-stimulated conditions for control cells, Tg-treated cells and Tg-treated cells ectopically expressing SNAP25. Images beneath show both SNAP25–GFP (green) and NPY–pHTomato (red) signals for a cell ectopically expressing SNAP25. Scale bars: 10 µm. (M) Quantitative analysis of the maximum Δf/f0 in control, Tg and Tg+SNAP25 PC12 cells (Mann–Whitney test; control, n=12; Tg, n=8; Tg+SNAP25, n=12; N=3). (N) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; pink, Tg+SNAP25-treated cells). (O,P) Numbers of responding (O) and non-responding (P) NPY–pHTomato puncta/cell (Mann–Whitney test), considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=12; Tg, n=8; Tg+DHA, n=12; N=3). N denotes the number of biological replicates and n is number of cells. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

Our investigation focused on the hypothesis that elevated ATF4 (Fig. 7C–J), a prominent downstream effector of the activated eIF2α pathway (Lu et al., 2004), might contribute to downregulating SNAP25 expression by acting as a repressor. To assess this hypothesis, we employed two distinct approaches: the use of ISRIB, a well-established inhibitor of ATF4 (Fig. 7F) (Yasuda et al., 2021; Bugallo et al., 2020), and the knockdown of Atf4 through short hairpin RNA (shRNA) (Fig. S6B–D). SNAP25 levels did not change in ER-stressed cells treated with ISRIB (Fig. 7G) or upon Atf4 knockdown (Fig. 7H–J). Remarkably, the effective reversal of the blunted stimulus-coupled secretion phenotype upon ATF4 inhibition and/or depletion strongly supports the conclusion that the diminished levels of SNAP25 are indeed due to the negative modulatory effects of ATF4 on SNAP25 expression in ER-stressed cells (Fig. S6E–J).

ER stress lowers the abundance of key granulogenic signatures

Given the well-documented evidence on the granulogenic potential of chromogranins, specifically SCGII and CGA (Kim et al., 2001; Lin et al., 2022), we wanted to test whether there were any significant changes in the relative abundance of chromogranins in ER-stressed cells. Interestingly, CGA levels were significantly decreased at the mRNA and protein levels, which remained unchanged in Tg+DHA-co-treated cells (Fig. 8A,B) (Mahata and Corti, 2019). A similar trend was also observed at the transcript and protein levels for SCGII (Fig. S5A,B). Furthermore, overexpression of CGA in stressed cells effectively reversed the secretory impairment. However, the reversal was comparatively mild compared to that of SNAP25 (Fig. 8G–L; Movie 8).

Fig. 8.

Change in DCV matrix proteins. (A) Transcript levels of Cga in control, Tg-treated and Tg+DHA-co-treated cells by qRT-PCR (Mann–Whitney test, N=4). (B) Representative blots of CGA (left) and quantification of the protein levels of CGA (right). GAPDH served as a loading control (N=6). (C) Representative fluorescence images showing DAPI (blue) and CGA (green) immunostaining in PC12 cells transfected with mCherry-Lysosomes-20 (expressing LAMP1–mCherry, red). Scale bar: 10 µm (2048×2048 pixels). (D) Quantification of CGA fluorescence intensity levels in control, Tg-treated and Tg+DHA-co-treated cells (images quantified were captured in 512×512 pixels; n=5; N=3). (E) Quantification showing Mander's correlation coefficient between CGA and LAMP1 (fraction of CGA overlapping LAMP1; n=6, N=3). (F) Representative blot of PDI (left) and quantification of PDI expression (right) by western blot analysis. GAPDH served as a loading control (N=4). Two-tailed unpaired Student's t-test was used in B,D–F. (G) Schematic showing the overexpression and imaging protocol of CGA–GFP. (H) Representative NPY–pHTomato (red) fluorescence images in PC12 cells showing basal and KCl-stimulated conditions (top) and chosen cells expressing both NPY–pHTomato and CGA–GFP (bottom). Scale bars: 10 µm. (I) Quantitative analysis of the maximum Δf/f0 in control, Tg and Tg+CGA cells (Mann–Whitney test; control, n=11; Tg, n=11; Tg+CGA, n=8; N=3). (J) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; orange, Tg+CGA-co-treated cells). (K,L) Numbers of responding (K) and non-responding (L) NPY–pHTomato puncta/cell (Mann–Whitney test), considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=11; Tg, n=11; Tg+DHA, n=8; N=3). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

Fig. 8.

Change in DCV matrix proteins. (A) Transcript levels of Cga in control, Tg-treated and Tg+DHA-co-treated cells by qRT-PCR (Mann–Whitney test, N=4). (B) Representative blots of CGA (left) and quantification of the protein levels of CGA (right). GAPDH served as a loading control (N=6). (C) Representative fluorescence images showing DAPI (blue) and CGA (green) immunostaining in PC12 cells transfected with mCherry-Lysosomes-20 (expressing LAMP1–mCherry, red). Scale bar: 10 µm (2048×2048 pixels). (D) Quantification of CGA fluorescence intensity levels in control, Tg-treated and Tg+DHA-co-treated cells (images quantified were captured in 512×512 pixels; n=5; N=3). (E) Quantification showing Mander's correlation coefficient between CGA and LAMP1 (fraction of CGA overlapping LAMP1; n=6, N=3). (F) Representative blot of PDI (left) and quantification of PDI expression (right) by western blot analysis. GAPDH served as a loading control (N=4). Two-tailed unpaired Student's t-test was used in B,D–F. (G) Schematic showing the overexpression and imaging protocol of CGA–GFP. (H) Representative NPY–pHTomato (red) fluorescence images in PC12 cells showing basal and KCl-stimulated conditions (top) and chosen cells expressing both NPY–pHTomato and CGA–GFP (bottom). Scale bars: 10 µm. (I) Quantitative analysis of the maximum Δf/f0 in control, Tg and Tg+CGA cells (Mann–Whitney test; control, n=11; Tg, n=11; Tg+CGA, n=8; N=3). (J) Representative time traces of mean NPY–pHTomato fluorescence intensities showing an average of all puncta per cell (green, control; red, Tg-treated cells; orange, Tg+CGA-co-treated cells). (K,L) Numbers of responding (K) and non-responding (L) NPY–pHTomato puncta/cell (Mann–Whitney test), considering a minimum Δf/f0 of 0.3 upon KCl stimulation as a response (control, n=11; Tg, n=11; Tg+DHA, n=8; N=3). N denotes the number of biological replicates and n is the number of cells taken for quantification. Data are shown as mean±s.e.m. ns, statistically non-significant; *P<0.05; **P<0.01; ***P<0.001; ****P<0.0001.

Concomitantly, there was enhanced colocalisation of CGA with the lysosomal marker LAMP1 in LAMP1–mCherry-positive Tg-treated PC12 cells, which remained unchanged upon co-treatment with DHA (Fig. 8C–E). These findings were substantiated by the augmented levels of protein disulfide isomerase (PDI or P4HB), an ER-resident chaperone known for its role in rerouting of misfolded proteins to the lysosomes (Fig. 8F) (Cha-Molstad et al., 2015; Wang et al., 2022; Wilkinson and Gilbert, 2004). In summary, these experiments suggested that major granins, known for their granulogenic functions, were depleted both transcriptionally and translationally, which was further accompanied by enhanced lysosomal localisation, leading to a severe impediment in regulated DCV exocytosis in ER-stressed conditions.

Previous studies have shown that a disruption in the release of components of DCVs (Barranco et al., 2021) and SLVs (Zou et al., 2021) is tightly related to almost all neurodegenerative diseases. Regulated exocytosis displayed by DCVs and SLVs is a highly orchestrated major cell physiological pathway. DCVs and SLVs contain neuropeptides, monoamines and neurotransmitters essential for cell survival and growth (Birinci et al., 2020; Salio et al., 2006; Speese et al., 2007; Zhang et al., 2011), and the compromised release of their contents creates a significant challenge to cellular physiology and ultimately leads to apoptosis, a consequence of which is neurodegeneration (Ulrich et al., 2002). The basic pathways related to the biogenesis of these vesicles are well understood. However, specific areas, such as crosstalk with other organelles and their behaviour during certain physiologically challenging conditions, such as metabolic stress, ER stress or autophagy, are unexplored research avenues. We have pharmacologically modelled ER stress in professional secretory cells, such as PC12 and INS-1 cells, without activating cell death and multiple ER stress pathways. We refer to this as ‘mild ER stress’.

Interestingly, ER stress caused severe impairment in DCV exocytosis in these two separate secretory cell lines. Similar defects were also observed in SLV exocytosis, whereas no defects were observed in the constitutive secretory pathway. To check the specificity of the ER stress pathway for its contribution to the impediment of regulated secretion, we rescued the cells from ER stress by using a well-known ER stress attenuator – an omega-3 fatty acid, DHA – which also happens to be a neuroprotective compound (Begum et al., 2014, 2013, 2012; Dyall, 2015). DHA co-treatment reversed eIF2α phosphorylation and strikingly negated the DCV exocytotic defects. Treatment with multiple ER stressors and attenuators in two independent cell lines substantiated our findings of impaired regulated secretion in ER-stressed cells. Interestingly, stimulus-driven Ca2+ extracellular influx via voltage-dependent ion channels, the major source for Ca2+-dependent DCV exocytosis, remains unchanged in Tg-treated cells compared to in control cells (Zorec, 1996; Preissler et al., 2020) (Fig. S1K–M).

To understand the molecular underpinning of ER stress-associated impaired exocytosis of DCVs, we quantified the expression of exocytotic genes, which are fundamentally responsible for secretory vesicle exocytosis. Expression switches of such regulatory genes are critical for active functional exocytosis. SNAP25 is an important membrane-associated t-SNARE required for the fusion of specialised secretory vesicles with the plasma membrane (Arora et al., 2017; Tsuboi and Fukuda, 2005; Walter et al., 2010). Intriguingly, impaired exocytosis in ER-stressed cells was correlated with reduced expression of SNAP25 levels, whereas the levels of several SNAREs, tethers and Rabs involved in exocytosis remain unchanged. Ectopic SNAP25 rescue experiments in Tg-stressed cells effectively reversed the phenotype and facilitated DCV exocytosis upon stimulation, suggesting the crucial role of SNAP25 in ER stress-mediated blunted exocytosis. The decrease in SNAP25 levels appears to be associated with the activity of ATF4, a downstream effector molecule in the PERK–eIF2α pathway. ATF4, also known as CREB2, has been proposed to act as a suppressor of CREB-dependent genes, suggesting a competitive inhibitory role on CREB (Smith et al., 2020; Liew et al., 2010; Oliveira et al., 2021). ATF4 ablation in ER-stressed cells by pharmacological agents such as ISRIB or shRNA restored the SNAP25 levels towards normalcy. Corroborative, functional assays also revealed a reversal of exocytosis defects. These findings highlight the significance of the eIF2α–ATF4 axis in regulating SNAP25, a crucial molecular switch that governs the exocytosis of DCVs (Fig. S6).

Besides regulating exocytotic switches, the ER-stressed cells also exhibited reduced granin levels, specifically for CGA and SCGII, which remained lower upon Tg and DHA co-treatment and which remains to be further explored. There is a scope for alteration of vesicle composition for DCVs given that reduced granin levels in ER-stressed cells can alter the biochemical signature of the DCVs, as granins are known to mobilise, accumulate and concentrate constituents, such as Ca2+ and catecholamines. These events are key as only mature DCVs are competent to undergo exocytosis. The chromogranin family mainly includes CGA, CGB and SCGII, which account for more than 90% of the DCV matrix proteins and are responsible for the accumulation and concentration of neuropeptides and Ca2+ (Kim et al., 2001; Mosley et al., 2007). Recent studies have also pointed out that biophysical events such as liquid–liquid phase separations of granins are also linked to granulogenic functions (Parchure et al., 2022).

It is noteworthy that CREB, a master regulator of several neurotropic genes, has been demonstrated to regulate SNAP25 and SCGII expression positively and CGA in a bidirectional manner (both positively and negatively) (Fig. S6A) (Wang et al., 2019; Cramer et al., 2008; Huttunen et al., 2002; Höcker et al., 1998; Mahapatra et al., 2003; Cibelli et al., 1996; Satoh et al., 2009). Besides, our bioinformatics analysis revealed the existence of a CREB-binding site, and pharmacological inhibition using 666-15, a highly specific CREB inhibitor (Xie et al., 2019; Li et al., 2016), caused depletion of SNAP25. Thus, we are tempted to speculate that reduced expression of CGA and SCGII can be attributed to augmented ATF4 levels, which is well known to bind the CREB-binding site, inhibiting CREB binding to its target, like a repressor. Moreover, expression switching of granulogenic genes was also accompanied by the enhanced colocalisation of CGA with a lysosomal marker. This finding can be potentially attributed to CGA being rerouted towards lysosomes by PDI, an ER-resident chaperone that routes some of the proteins to the autophagy–lysosomal pathway besides assisting in proper folding (Cha-Molstad et al., 2015; Wang et al., 2022).

Despite reduced expression of key granins, DHA treatment was able to reverse the altered regulated secretory phenotype. To understand this unexpected finding, we performed TEM and morphometric analysis to quantify DCV number, dense core size (DCVs comprise the dense core and a membrane surrounding the core) and DCV diameter, which were not significantly different. However, DCVs were more distantly located from the plasma membrane in Tg-stressed cells than in control cells, an observation reverted with the ER stress attenuator DHA (although this was not statistically significant). Altered granin and DCV localisation away from the plasma membrane reduces the propensity of their release by decreasing the chances of vesicle docking and release. We propose that these can be linked to the well-known and documented action of SNAP25, a key component of the STX1–SNAP25 acceptor complex (de Wit et al., 2009). Notably, SNAP25 levels were depleted in ER-stressed cells and were restored upon DHA treatment. Its overexpression rescued the impairment in secretion. However, SYT1 and STX1 levels remain unchanged.

Our study identified a novel hindrance in releasing specialised secretory vesicles (DCVs and SLVs) during ER stress, demonstrating distinct impacts on various cell types. To our knowledge, this study marks the inaugural exploration into the intricate interplay between a stressed ER and specialised secretory vesicles, shedding light on a previously uncharted realm of inter-organellar crosstalk. Our research unveils a ‘double-edged sword’ scenario, wherein compromised exocytosis of specialised secretory vesicles (DCVs) and ER stress can synergistically intensify, leading to detrimental health and physiological effects. This cumulative disturbance of ER stress and the regulated secretory pathway poses a significant risk, potentially worsening conditions associated with neurodegeneration, diabetes, and other systemic and organic disorders. A notable discovery is the ability of DHA, a well-established ER stress attenuator, to reverse the compromised exocytosis of DCVs (Fig. S7). This finding holds immense pharmacological promise, suggesting a potential avenue for correcting impaired DCV exocytosis in conditions linked to ER stress, offering hope for therapeutic interventions in diseases characterised by such dysregulation. For instance, a recent study demonstrated the translational benefits of ATF4 inhibition in memory and synaptic plasticity (Oliveira et al., 2021) in an Alzheimer's disease model. Our findings have substantial translational relevance, providing valuable insights into the intricate relationship between ER stress and regulated secretion. This knowledge opens avenues for targeted therapeutic approaches, particularly in conditions in which ER stress and neuroprotective functions of neuromodulators and/or neurotransmitters intersect, presenting opportunities for mitigating the impact of these pathophysiological conditions (de Diego et al., 2020; Ozcan and Tabas, 2012; Marciniak et al., 2022). Although these are very interesting findings, we acknowledge certain limitations. For instance, the induced ER stress is unnatural and limited to cell culture experiments. Additionally, how ER stress attenuators mechanistically restore impaired exocytosis remains to be further investigated. In the future, it would also be interesting to look at the crosstalk of ER stress and investigate its association with the functional status of regulated exocytosis in animal models under natural pathophysiological conditions such as diabetes, obesity and neurodegenerative diseases, such as Huntington's disease and Alzheimer's disease, given the known regulatory roles of ER stress and DCV cargo in these diseases (Cortès-Saladelafont et al., 2016; Thomas-Reetz and De Camilli, 1994).

Cell culture and treatment

Rat PC12 cells originally generated by Greene and Tischler (1976) were a gift from Nitish Mahapatra, Indian Institute of Technology Madras. Early passage cells (<8) were used for all experiments. The cells were treated routinely with mycoplasma removal reagent (MP Biomedicals, 30-50044). For routine maintenance, the cells were grown in T25 flasks (Nunc, USA) in Dulbecco's modified Eagle medium (DMEM) GlutaMAX (Gibco, 12100046), supplemented with 10% horse serum (Gibco, 16050114), 5% heat-inactivated fetal bovine serum (FBS; Sigma-Aldrich, F7524), 25 mM HEPES (Sigma-Aldrich, H3784) and 1% antibiotic-antimycotic (Gibco, 15240062) in 5% CO2 at 37°C in a humidified incubator (Sahu et al., 2012). The insulin-secreting cell line INS-1 (a gift from Prof. Claes B. Wollheim, University Medical Center, Geneva, Switzerland) was cultured in RPMI-1640 (Sigma-Aldrich), supplemented with 10% FBS, 2 mM L-glutamine (Gibco GlutaMAX supplement, 35050), 25 mM HEPES, 1 mM sodium pyruvate (Gibco, 11360-070) and 50 μM β-mercaptoethanol (Sigma-Aldrich, M3148) in a humidified atmosphere (5% CO2, 37°C) according to the previously described method (Asfari et al., 1992). All treatments were performed in reduced serum media and 10 nM Tg (Sigma-Aldrich, T9033) or 0.05 μg/ml Tm (Cayman Chemical, 11445) was used for 24 h to induce ER stress in PC12 or INS-1 cells. For rescue experiments, 10 µM DHA (Sigma-Aldrich, D2534) was used, and cells were primed with DHA for 24 h before the induction of ER stress (Begum et al., 2014). 1 mM PBA (Cayman Chemical, 11323) was administered as co-treatment with 10 nM Tg. Cells grown in reduced serum media were used as the control. For ATF4 inhibition, 30 nM ISRIB (Cayman Chemical, 16258) was used along with Tg. Similarly, 5 µM of the CREB inhibitor 666-15 (Cayman Chemical, 30780) was used.

Cell viability assay

The MTT assay assessed cytotoxicity to optimise the treatment dose for Tg and Tm (van Meerloo et al., 2011). 104 PC12 cells/well plated in a 48-well plate were allowed to settle for 24 h before starting Tg treatment. The medium was then replaced with reduced serum medium for 2 h, after which 6, 10 or 30 nM of Tg or 0.05, 0.1, 0.2 or 0.5 µg/ml of Tm was administered to the cells for 24 h. Next, cells were incubated in 0.5 mg/ml of MTT (Sigma-Aldrich, 475989) for 4 h. Formazan crystals formed were dissolved via gentle trituration in DMSO (Sigma-Aldrich, 67-68-5). Optical density was recorded at 570 nm with a microplate reader (Tecan Spark multimode reader). The IC50 value was defined as the concentration of Tg or Tm exhibiting 50% cell viability.

Immunoblotting

After treatment, PC12 cells were lysed using 125 µl of RIPA buffer [10 mM Tris-HCl (Sisco Research Laboratory, 37969), 150 mM NaCl (MP Biomedicals, 194848), 1 mM NaVO4 (Sigma-Aldrich, S-6508), 30 mM Na4P2O7 (Sigma-Aldrich, 011K0307), 50 mM NaF (Sigma-Aldrich, 7681-49-4), 1% Nonidet P-40 (Sigma-Aldrich), 0.1% SDS (Affymetrix USB Products, 151-21-3), 1 mM PMSF (Sigma-Aldrich, 52K0052), 1% Triton X-100 (Sigma-Aldrich, 9002-93-1), 0.5% C24H39NaO4 (Sigma-Aldrich, 145224-92-6) and dissolved protease inhibitor cocktail (Roche Diagnostics, 11714900) in water, pH 7.4] and the protein contents of the cell lysate were quantified using a BCA Protein Assay Kit (Takara, T9300A) following the manufacturer's protocol (Song et al., 2021). The samples containing 10 µg protein were separated using a 10% SDS–polyacrylamide gel and transferred onto nitrocellulose membranes. After blocking with 3% bovine serum albumin (BSA; Sigma-Aldrich, A4503), the membranes were incubated with primary antibodies against eIF2α (Cell Signaling Technology, 9722s, 1:5000), p-eIF2α (Cell Signaling Technology, 9721s, 1:2000), PDI (Cell Signaling Technology, 3501P, 1:3000), SCGII (Proteintech, 20357-1-AP, 1:1000), CGA (Proteintech, 10529-1-AP, 1:3000), CREB (Cell Signaling Technology, 9197s, 1:4000), p-CREB (Cell Signaling Technology, 9191s, 1:1000), SNAP25 (Synaptic Systems, 111011, 1:5000), ATF4 (Proteintech, 10835-1-AP, 1:5000), VAMP2 (Synaptic Systems, 104202, 1:2000), STX6 (Synaptic Systems, 110062, 1:5000), SYT1 (Synaptic Systems, 105011, 1:1000), HSP90 (Proteintech, 13171-1-AP, 1:2000), GAPDH (Proteintech, 60004-1-Ig, 1:10,000), EF2 (Santa Cruz Biotechnology, sc-166415, 1:5000) or β-tubulin (Proteintech, 10094-1-AP, 1:5000) overnight at 4°C, followed by incubation with HRP-conjugated goat anti-mouse [Peroxidase AffiniPure goat anti-mouse IgG (H+L), Jackson Immuno Research, 115-035-003, 1:10,000] and goat anti-rabbit [Peroxidase AffiniPure goat anti-rabbit IgG (H+L), Jackson Immuno Research, 111-035-003, 1:10,000] secondary antibodies. The membrane was washed three times for 10 min after primary and secondary antibody incubation to remove excess antibodies. ECL [luminol (Sigma, A8511-5G), p-coumaric acid (Sigma, C9008), hydrogen peroxide (Merck, 107210)] was prepared using the previously described protocol, and bands were visualised using the Uvitec Mini HD9 gel imaging system (Mruk and Cheng, 2011). Densitometry quantifications were done using ImageLab (6.0.1) (Sahu et al., 2019). Uncropped blots are shown in Fig. S8.

Quantitative real-time PCR

Primers for quantitative real-time PCR (qRT-PCR) were ordered from Integrated DNA Technologies for the following genes: Gapdh,Casp3, Cga, ScgII, Snap25, Atf4, Xbp1 (spliced), Xbp1 (unspliced), Bip, Vamp1, Stx1a and Rab27a. RNA extraction of the samples was done using a Nucleospin RNA isolation kit (Macherey–Nagel, 740955.50) and RNA yields were estimated using a spectrophotometer (NanoDrop ND-1000). The corresponding cDNAs of the samples were prepared using the iScript Select cDNA synthesis kit (Bio-Rad, 1708891). Primers for qRT-PCR were used at a final concentration of 0.2 µM, and a final amount of 80 ng cDNA was added. The mRNA expression of target genes was determined using the real-time cycler CFX 96 Maestro (Bio-Rad) following the manufacturer's protocol for iQ SybrGreen (Bio-Rad, 1725121) (Talbot et al., 2004). 2−ΔΔCq was used to calculate the relative expression of the target gene across various treatments, normalised to the housekeeping control Gapdh or 18s rRNA (Schmittgen and Livak, 2008).

The following primers were used: Atf4, forward primer (FP), 5ʹ-CCTGAACAGCGAAGTGTTGG-3ʹ, and reverse primer (FP), 5ʹ-TGGAGAACCCATGAGGTTTCAA-3ʹ; Atf6, FP, 5ʹ-TCGCCTTTTAGTCCGGTTCTT-3ʹ, and RP, 5ʹ-GGCTCCATAGGTCTGACTCC-3ʹ; Casp3, FP, 5ʹ-GGACAGCAGTTACAAAATGGATT-3ʹ, and RP, 5ʹ-CGGCAGGCCTGAATGATGAAG-3ʹ; Snap25, FP, 5ʹ-CGAAGAGAGTAAAGATGCTGGC-3ʹ, and RP, 5ʹ-GTTTTGTTGGAGTCAGCCTTCT-3ʹ; Cga, FP, 5ʹ-CGGCAGCATCCAGTTCTCA-3ʹ, and RP, 5ʹ-AGCCCCTGTCTTTCCATTCA-3ʹ; ScgII, FP, 5ʹ-CCTACTTGAGAAGGAATTTGC-3ʹ, and RP, 5ʹ-ACCAACCCATTTGGTTTCTC-3ʹ; Xbp-1 (spliced), FP, 5ʹ-TCAGACTACGTGCGCCTCT-3ʹ, and RP, 5ʹ-TCAGACTACGTGCGCCTCT-3ʹ; Xbp-1 (unspliced), FP, 5ʹ-CTGAGTCCGCAGCAGGTG-3ʹ, and RP, 5ʹ-CCACATCCGCCGTAAAAGAATG-3ʹ; Gapdh, FP, 5ʹ-CGTATTGGGCGCCTGGTCAC-3ʹ, and RP, 5ʹ-CGGCCTCACCCCATTTGATG-3ʹ; Bip, FP, 5ʹ-GGTACATTTGATCTGACTG-3ʹ, and RP, 5ʹ-CACTTCCATAGAGTTTGCTG-3ʹ; Vamp1, FP, 5ʹ-GGTTTCCATTGTGTCTGTC-3ʹ, and RP, 5ʹ-ATCTGTCACATGCCTTTGGT-3ʹ; Stx1a, FP, 5ʹ-TACAACGCCACTCAGTCAGA-3ʹ, and RP, 5ʹ-GAGTCCATGATGATCCCAGA-3ʹ; Rab27a, FP, 5ʹ-TTCAGGGACGCTATGGGTTT-3ʹ, and RP, 5ʹ-CCGCAGAGCACTATATCTGGG-3ʹ; and 18S, FP, 5ʹ-GTAACCCGTTGAACCCCATT-3ʹ, and RP, 5ʹ-CCATCCAATCGGTAGTAGCG-3ʹ.

Transfections

PC12 and INS-1 cells were transfected following the manufacturer's protocol using Metafectene Pro (Biontex, lot no. RKP205/RK081621) (Grindheim et al., 2014). In brief, plasmid DNA and Metafectene Pro were mixed in DMEM/RPMI medium without any supplements and incubated for 20 min at room temperature to form complexes. These complexes were directly added to the cells. Cells were cultured in G418 (Geneticin, G-418 sulfate; GOLDBIO, G-418-25)-containing medium for 14 days for stable cell line generation. The following plasmids were used: NPY-mApple (Addgene, 83498), NPY-pHTomato (Addgene, 83501), NPY-pHluorin (a kind gift from Sebastian Barg, Uppsala University, Sweden), synaptophysin-pHTomato (a kind gift from Yulong Li laboratory, Department of Molecular and Cellular Physiology, Stanford University School of Medicine, Stanford, CA, USA), mCherry-Lysosomes-20 (Addgene, 55073), His-GFP SNAP25 (Addgene, 170872) and CGA-GFP (a kind gift from Nitish Mahapatra laboratory).

NPY–mApple plate reader assay

Stably selected NPY–mApple PC12 cells were plated at 3×105 cells/well in a six-well plate. Then, cells were washed with ice-cold 1× PBS (Gibco, 21600-069) and incubated in Tyrode's buffer (basal: 150 mM NaCl, 5 mM KCl, 2 mM CaCl2, 10 mM HEPES, pH 7.4; stimulation: 55 mM NaCl, 100 mM KCl, 2 mM CaCl2, 10 mM HEPES, pH 7.4) for 20 min. The supernatant was collected and cells were lysed on ice. Lysed cells were sonicated and spun at 14,000 g for 15 min at 4°C. 10 μl of the cellular lysate was used and the supernatant was diluted at a 1:10 ratio in 1× PBS. 100 µl of the supernatant and diluted cellular lysate was loaded in a dark 96-well plate to eliminate cross-contamination of the fluorescence signal from adjacent wells. NPY–mApple fluorescence signals were then recorded using the Tecan plate reader (excitation wavelength, 568 nm; emission wavelength, 592 nm), subtracting them from the autofluorescence of the plate itself (Wemhöner et al., 2006). All values were expressed as (Zhang and Martin, 2018):

Immunofluorescence and confocal imaging

Approximately 3×104 cells/coverslip were seeded onto poly-L-lysine (Sigma-Aldrich, P2636)-coated coverslips. After treatment, cells were gently rinsed with 1× PBS and fixed with 4% paraformaldehyde (PFA; Thermo Fisher Scientific, 30525-89-4) at room temperature for 20 min, followed by three 5 min washes with 1× PBS to remove excess PFA. Cells were then permeabilised with 0.1% Triton X-100 for 10 min and blocked using 1% BSA for 1 h. Next, they were incubated with goat anti-CGA primary antibody (Santa Cruz Biotechnology, sc-1488, 1:500 in blocking buffer) in a humidified chamber for 1 h, followed by three gentle washes with 1× PBS to remove the excess antibody. Alexa Fluor 488-conjugated donkey anti-Goat (Invitrogen, A11055, 1:2000) secondary antibody was then added to the cells for 45 min in a humidified chamber in the dark. Samples were again rinsed three times with 1× PBS, and coverslips were mounted using Flouroshield with DAPI (Sigma-Aldrich, F6057) to reduce photobleaching (Berlier et al., 2003). Images were captured using a Nikon A1HD25 confocal microscope with 60× oil objective (NA 1.4) at a 512×512 pixel resolution (Representative images were captured at a 2048×2048 pixel resolution).

Live-cell imaging and analysis

PC12 cells were transfected with the NPY-pHTomato plasmid and stably selected, whereas the synaptophysin-pHTomato, His-GFP SNAP25 and CGA-GFP plasmids were transiently transfected in PC12 cells. Similarly, INS-1 cells were transiently transfected with NPY-pHluorin (Gandasi et al., 2015; Hummer et al., 2017). For imaging, cells were plated on 15 mm coverslips (Assistant, Glaswarenfabrik, 41001115) coated with 1× poly-L-lysine, and the same treatment protocol was followed. On the day of the experiment, cells were kept in 2.5 mM KCl (Merck, 61753305001046) containing Tyrode's buffer (basal), and coverslips were placed in an open imaging chamber (Live Cell Instruments, Republic of Korea) followed by stimulation with 100 mM KCl containing Tyrode's buffer (stimulation). Images were captured using a Zeiss spinning-disk confocal microscope with a Yokogawa CSU-XA1 head at 63× oil (1.4 NA) and Nikon eclipse Ti2 at 100× oil (1.30 NA) at 300 ms for 2 s in Tyrode's buffer (basal), after which stimulation was given. Captured time-lapse movies were analysed using ImageJ software, and after background subtraction, the change in puncta intensity per cell (Δf) after 100 mM KCl stimulation compared to basal fluorescence (f0) was plotted as Δf/f0. The maximum Δf/f0 indicates the maximum change in fluorescence intensity post stimulation.

Constitutive secretion

To check the constitutive release of proteins in basal conditions, secreted fractions from unstimulated cells were analysed by SimplyBlue SafeStain (Invitrogen, LC6060) Coomassie staining (Huang et al., 2001; Hummer et al., 2017). PC12 cells were seeded in 60 mm culture dishes at a density of 106 cells per dish. After 24 h, treatments were given, followed by medium removal and incubation in Tyrode's buffer (basal) for 20 min. The supernatants and cell lysates were collected and resolved using a 10% SDS-polyacrylamide gel. The gel was washed twice with Milli-Q water and then stained with SimplyBlue SafeStain for 1 h at room temperature with gentle shaking. To remove the background, the gel was washed twice with Milli-Q water and later imaged in the Azure c300 gel imaging system, and all the automatically detected bands were quantified using ImageLab (6.0.1).

For the HSP90 secretion experiment, cells were seeded in six-well plates and, after completion of treatment, cells were incubated with Tyrode's basal buffer for 3 h. After that, the supernatant containing basal buffer was collected and resolved in a 10% SDS-polyacrylamide gel, following the standard immunoblotting procedure, and probed for HSP90 (Proteintech, 13171-1-AP, 1:2000). Bands were visualised using the Uvitec Mini HD9 gel imaging system (Mruk and Cheng, 2011).

Lentiviral transduction

Glycerol stocks of two different shRNAs against Atf4 were commercially obtained from Dharmacon. HEK 293T cells were seeded at 60% confluency in each well of a six-well plate. 15 h post seeding, cells were transfected with two shRNA Atf4 knockdown constructs (0380 and 3728 from Dharmacon) and packaging plasmids (Pspax2 and PMD2G, generously provided by Zan Hai Bin laboratory, University Of Minnesota, MN, USA) in DMEM containing 10% FBS without antibiotics. 12 h post transfection, the medium was replaced with complete medium (DMEM containing 10% FBS with antibiotics), and medium collection was carried out for the next 2 days. After collecting the virus-containing medium, it was filtered using a 0.45 µm filter, and the virus was concentrated using PEG3000 (Promega, V3011). The precipitated viral pellet was then concentrated ten times, aliquoted into single-use aliquots, and stored at −80°C for individual use.

PC12 cells were plated at 30–40% confluency in six-well plates. 12 h after plating, cells were transfected using the virus in complete medium without antibiotics, supplemented with 8 μg/ml polybrene (A kind gift from Sam Mathew, Associate professor, Regional Centre for Biotechnology, India) to a final concentration. After 20–24 h of viral transduction, the medium was changed, and cells were selected for 2 weeks in puromycin-containing medium. Subsequently, the cells were induced using 5 µg/ml doxycycline and subjected to fluorescence-activated cell sorting using a BD FACSAria Fusion flow cytometer (BD Biosciences), and the final sorted cells with optimum GFP expression were established as the population of cells for all further experiments.

Electron microscopy

Cells were seeded in 100 mm cell culture plates at 60% confluency and, at the end of the treatment, cells were fixed in 2.5% glutaraldehyde (Sigma-Aldrich, MKBG0637V) and 2% PFA in 0.15 M sodium cacodylate buffer (Sigma-Aldrich, C0250) and post-fixed in 1% osmium tetroxide (Sigma-Aldrich, 05500) in 0.1 M sodium cacodylate buffer for 1 h on ice. The cells were then dehydrated in a graded series of ethanol (Merck, 1.00983.0511) (50–100%) on ice, followed by one wash with 100% ethanol and two washes with propylene oxide (Tokyo Chemical Industry, E0016) (15 min each) and embedded with Araldite-502 (Ted Pella, 18060). Ultrathin (∼50–60 nm) sections were cut on a Leica UCT ultramicrotome and picked up on carbon-coated copper grids. Sections were stained with 10% uranyl acetate (Sisco Research Laboratory, 81405) for 5 min and Sato's lead citrate stain (Electron Microscopy Sciences, 17800) for 1 min. Grids were viewed using a JEOL JEM1400-plus transmission electron microscope (Sahu et al., 2019). Images obtained were analysed using ImageJ software, DCV and dense core diameters were calculated, and total DCV distribution was obtained by measuring the shortest distance from the cell membrane. DCVs comprise the dense core and the surrounding membrane. DCV diameter corresponds to the diameter of the dense core with the outer membrane, whereas dense core diameter is the diameter of only the dense core.

Statistical analysis

Statistical comparisons between two groups were performed with Student's two-tailed unpaired t-tests when data were normally distributed with similar variances. Otherwise, the Mann–Whitney test was performed to draw the significance. For comparison among multiple groups, one-way ANOVA with Tukey's multiple comparison post hoc test was performed. A P-value of 0.05 or less was considered statistically significant. Statistical analysis was performed using GraphPad Prism 8.0. The data are presented as means, and error bars show the standard error of the mean (s.e.m.).

We would like to extend our gratitude to James Edgar and Nick Bright, University of Cambridge, for their suggestion on electron microscopy-related experiments. We would like to thank the Regional Centre for Biotechnology NCR Biotech cluster, Faridabad, India, for assistance related to electron microscopy imaging. We would also like to thank Thomas Puccyadil and Dileep Vasudevan for their critical suggestions on the manuscript. B.S.S. thanks his academic mentors, Profs. Nitish Mahapatra, Scottie Robinson, Alessandro Bartolomucci and Sushil Mahata, for their invaluable support. He also thanks Prof. L. S. Shashidhara (Indian Institute of Science Education and Research Pune/Ashoka University), Dr Meenakshi Munshi (former advisor, Department of Biotechnology) and Prof. Anirban Basu, his career mentors. Lastly, B.S.S. thanks all colleagues at National Brain Research Centre (NBRC) for helping settle down amidst the pandemic and providing emotional and infrastructural support to our new lab. We thank NBRC core funds for providing salary support to this study's authors.

Author contributions

Conceptualization: B.S.S.; Methodology: M.M., C.M., V.G., A.J., S.S., S.D.; Software: M.M, C.M, V.G, A.J, S.S.; Validation: M.M., C.M., V.G., A.J., S.S., B.S.S.; Formal analysis: M.M., C.M., V.G., B.S.S.; Investigation: M.M., C.M., V.G., A.J., S.S., S.D., B.S.S.; Resources: B.S.S.; Data curation: M.M., C.M., V.G., B.S.S.; Writing - original draft: M.M., C.M., B.S.S.; Writing - review & editing: M.M., C.M., V.G., B.S.S.; Visualization: M.M., C.M., V.G., B.S.S.; Supervision: B.S.S.; Project administration: B.S.S.; Funding acquisition: B.S.S.

Funding

This study was supported by funds from the Rising Stars in Neuroscience award (formerly Return Home Fellowship) by the International Brain Research Organization. A Ramalingaswami fellowship grant (BT/RLF/Re-entry/38/2016) from the Department of Biotechnology, Ministry of Science and Technology, India, an early career research grant from International Centre for Genetic Engineering and Biotechnology, Trieste, Italy, and core funds from National Brain Research Centre also funded the B.S.S. laboratory towards this study.

Data availability

All relevant data can be found within the article and its supplementary information.

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Competing interests

The authors declare no competing or financial interests.

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