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What is the role of increased cytosolic calcium concentration after firing, in neuronal cell bodies?

What is the role of increased cytosolic calcium concentration after firing, in neuronal cell bodies?


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I've come across several studies in which scientists were investigating various questions related to neural activity by focusing on neuronal cell bodies using Calcium imaging. As this article suggests for example, cytosolic Ca concentration in the soma will significantly increase if the neuron fires. I understand the process behind this. But I'd like to find out if this somatic Ca increase has significant downstream effects inside the soma.

Could anyone shed some light on such known processes/pathways?

I've thought of one possibility based on an article that showed Calcium would upregulate mitochondrial enzymes, such as PDH and OGDC, responsible for TCA reactions. Considering that there's a coupling (source) between cytosolic and mitochondrial Ca concentrations, this could lead to a very quick increase in metabolic activity that covers up for the ATP loss related to neuron firing. However, I'm a bit skeptical about this idea. I'd think that the lack of ATP and low NADH/NAD ratio should speed up the TCA enough already and Ca might be more critical somewhere else instead.


LSXL7C LaunchPad Week 2

(1) opening of voltage-gated calcium channels
(2) change in membrane potential of the postsynaptic neuron
(3) fusion of secretory vesicles with the plasma membrane of axon terminal
(4) binding of neurotransmitters to receptors on the postsynaptic neuron
(5) increase in cytosolic calcium levels in the presynaptic neuron
(6) arrival of action potential at the axon terminal of the presynaptic neuron

a. Ion channels bind the ligand and open.
2. Na⁺ is pumped out of the cell, and the membrane potential is restored.
3. Acetylcholinesterase breaks down acetylcholine.
4. Na⁺ enters the postsynaptic cell, and the membrane potential changes.
5. Acetylcholine is released in the synapse.
6. Na⁺ ion channels close.

An excitatory postsynaptic potential (EPSP) of sufficient strength to reach threshold occurred at point 1 on the figure.

The voltage differences shown in the figure are measured across the plasma membrane.

Resting potential is generated mainly by the outward movement of K⁺ ions from inside the cell.

All voltage changes along the plasma membrane of the axon have the characteristics shape shown on the graph.


Intracellular Calcium Dysregulation: Implications for Alzheimer’s Disease

Alzheimer’s Disease (AD) is a neurodegenerative disorder characterized by progressive neuronal loss. AD is associated with aberrant processing of the amyloid precursor protein, which leads to the deposition of amyloid-β plaques within the brain. Together with plaques deposition, the hyperphosphorylation of the microtubules associated protein tau and the formation of intraneuronal neurofibrillary tangles are a typical neuropathological feature in AD brains. Cellular dysfunctions involving specific subcellular compartments, such as mitochondria and endoplasmic reticulum (ER), are emerging as crucial players in the pathogenesis of AD, as well as increased oxidative stress and dysregulation of calcium homeostasis. Specifically, dysregulation of intracellular calcium homeostasis has been suggested as a common proximal cause of neural dysfunction in AD. Aberrant calcium signaling has been considered a phenomenon mainly related to the dysfunction of intracellular calcium stores, which can occur in both neuronal and nonneuronal cells. This review reports the most recent findings on cellular mechanisms involved in the pathogenesis of AD, with main focus on the control of calcium homeostasis at both cytosolic and mitochondrial level.

1. Introduction

Alzheimer’s Disease (AD) is the most common type of dementia affecting millions of people. According to Alzheimer’s Disease International (ADI), as of 2015 people suffering from dementia worldwide accounted for estimated 46.8 million. Approximately 70% of these cases were attributed to AD. This amount will increase to an estimated 74.7 million in 2030 and 131.5 million in 2050, with a parallel rise of healthcare costs. As a matter of fact, global costs of dementia have increased from US$ 604 billion in 2010 to US$ 818 billion in 2015, for a 35.4% increase. The incidence rate for AD grows exponentially with age, with the main onset time observed in people aged over 60, in particular between the age of 70 and 80 [1, 2]. AD has also a sex-related incidence, making women 1.5–3 times more vulnerable than men [3]. It has been widely assumed that the higher risk observed in females is related to the loss of the neuroprotective effect of sex steroid hormones during menopause, resulting in estrogen deficiency in the brain [4–6].

AD is a progressive neurodegenerative disorder leading to severe cognitive, memory, and behavioral impairment [7]. The majority of cases is idiopathic however a rare variant of AD, known as Familial Alzheimer’s Disease (FAD), accounts for a small percentage (1–5%) [2, 8] of all cases. FAD features an autosomal dominant heritability and an early disease onset (<65 years old) [7, 9]. Three genetic mutations have been identified as being responsible for FAD. They involve genes for amyloid precursor protein (APP) on chromosome 21 [10], presenilin 1 (PS1) on chromosome 14 [11], and presenilin 2 (PS2) on chromosome 1 [12]. Both forms of AD share two main pathological hallmarks: the abnormal extracellular accrual and deposition of amyloid-β (Aβ) peptides and the intracellular accumulation of neurofibrillary tangles (NFTs). Aβ peptides are cleaved products of APP obtained via sequential proteolysis by two membrane-bound endoproteases, aspartyl β-secretase and presenilin-dependent secretase (γ-secretase) [13, 14]. APP can also be cleaved by α-secretase to produce nontoxic fragments, which are thought to antagonize Aβ peptides generation [15]. Aβ is a protein consisting of 39–43 amino acids, and it mainly exists in two isoforms: soluble A

5–10%) [15, 16]. In particular, due to a greater tendency to aggregate than A , A seems to be the main pathological isoform [17]. Interestingly, it has been described that soluble Aβ globular oligomers can form along a new aggregation pathway independent of Aβ fibril formation. These globular Aβ oligomers have been found in the brain of patients affected by AD and APP transgenic mice, and they bind specifically to neurons and affect synaptic plasticity, as demonstrated by Barghorn and coworkers [18]. The disturbance afforded by soluble Aβ oligomers has also been supported by evidence showing that they can bind to glutamate receptors (both ionotropic and metabotropic), thereby impairing glutamatergic neurotransmission [19, 20]. It is interesting to underline, however, that APP products and very low concentrations of soluble Aβ can be involved in important physiological processes, such as synapse activity and behavior [21, 22].

As for NFTs, it has been found that their major constituent is the protein tau. Tau is the predominant microtubule-associated protein found in mammalian brain [28]. During early stages of development tau is highly phosphorylated however phosphorylation decreases with brain aging [29, 30], leading to an unphosphorylated form that binds to microtubules, thereby making them more stable. In AD, tau is aberrantly misfolded and abnormally hyperphosphorylated [7, 13]. Several factors might be involved in tau hyperphosphorylation, including Aβ-mediated caspases activation, Aβ-mediated oxidative stress, chronic oxidative stress, and reduced insulin-like growth factor 1-mediated oxidative stress [31]. Over the course of AD, hyperphosphorylation contributes to the loss of tau physiological functions and it prepares this protein to form neurotoxic aggregates. It has been shown that, in this pathological form, tau can also ectopically enter the somatodendritic compartment where, in conjunction with Aβ oligomers, it promotes excitotoxicity. Additionally, tau phosphorylation can modulate DNA integrity and global changes in transcriptional events [32].

Aβ plaques and NFTs, often referred to as “positive features” [13], occur in specific regions rather than diffusely throughout the brain: in particular hippocampus and cortex are mainly affected [8, 13]. In addition, negative features of AD have also been described, including typical losses of neurons, neuropil, and synaptic elements, that mostly parallel NFTs formation. However, a causative relationship between NFTs and neuronal loss still remains to be clarified [33–40]. Growing evidence supports the involvement of neuroinflammation in AD [41], focusing on its critical role within brain regions where Aβ plaques are mainly distributed. Aβ-deposition renders cells more likely to develop inflammatory responses that involve the production of neuronal and glial cytokines belonging to the Tumor Necrosis Factor-α (TNF-α) superfamily [42]. Interestingly, it has been shown that neutralization of the Tumor Necrosis Factor Related Apoptosis Inducing Ligand (TRAIL) protects human neurons from Aβ-induced toxicity [43]. In this context, in vitro experiments conducted using the differentiated human neuroblastoma cell line SH-SY5Y demonstrated that the nonsteroidal anti-inflammatory derivative CHF5074 abrogates neurotoxic effects of both A

and TRAIL [44], suggesting a potential role of this drug as neuroprotective agent.

AD patients show symptoms that can be divided into two main categories: cognitive and psychiatric. Cognitive symptoms include loss of long term memory, aphasia, apraxia, and agnosia, while psychiatric symptoms include personality changes, depression, and hallucinations (Alzheimer’s Foundation of America, Last Update: January 29, 2016 [8]). AD is a complex multifactorial disorder, neuronal death is a subtle phenomenon, and it is difficult to identify a single cause. The idea that energy/mitochondrial dysfunction and oxidative stress may have a central role in the pathogenesis of AD is widely supported by literature [45–49]. Research on the pathogenesis of AD has recently stressed the role of mitochondria, based on the finding that mutation in APP and tau may directly affect mitochondrial function and dynamics [8], and now it is accepted that the impairment of mitochondrial function may affect other crucial cell signaling pathways, as in calcium signaling. A central role for calcium dysregulation in the pathogenesis of AD has been extensively suggested [7, 50]. This review attempts to clarify connections between mitochondrial pathways impairment and the pathogenesis of AD, drawing attention to the calcium homeostasis deregulation as a potential consequence of mitochondrial function disturbance and to the proteins mainly involved in this process, such as the sodium-calcium exchanger (NCX).

2. Calcium and AD

Calcium can be considered a ubiquitous intracellular messenger within cells acting as a regulator in multiple physiological functions. As a divalent cation, calcium can bind to several proteins, receptors, and ion channels. All of these properties are of great importance within neurons, where continuous firing of action potentials leads to calcium cycling, and it implies an influx through the calcium channels at the plasma membrane level, intracellular buffering, and an efflux through the calcium plasma membrane transporters. This cycling involves several subcellular compartments and proteins. In particular, two organelles play a major role in calcium buffering, namely, endoplasmic reticulum (ER) and mitochondria, whereas ATPase calcium pump and NCX are the two main systems involved in calcium efflux through the plasma membrane (Figure 1). Perturbation in such delicate balance may have deleterious consequences for cells and in particular for neurons, leading to necrosis and/or apoptosis and subsequently to stroke and neurodegeneration.

2.1. Intracellular Calcium Homeostasis

There is a large body of evidence documenting a connection between calcium homeostasis disruption and the development of neurodegenerative diseases such as Alzheimer’s [50]. The involvement of calcium in the pathogenesis of AD has been suggested long time ago by Khachaturian [51], and since then many efforts have been made to clarify this hypothesis [7, 52–56]. Despite the significant progresses made in explaining this theory, several aspects are to be defined. For instance, growing in vitro evidence suggests that neuroprotection could be mediated by the restoration of calcium homeostasis. Different calcium channel blockers have been reported to be effective in preventing long- and short-term memory impairment induced by A (the shortest Aβ fragment processed in vivo by brain proteases, retaining the toxicity of the full-length peptide [57]) and in decreasing Aβ production, inflammation, and oxidative stress. For example, Rani et al. described the effect of a calcium channel blocker clinically used in angina, in a mouse model of dementia. Interestingly, Morris water maze test, plus maze test and different biochemical analysis, demonstrated the restoration of normal learning and memory functions. Moreover, SCR-1693 (a nonselective calcium channel blocker) has been described to attenuate A -induced death in SH-SY5Y cells and to regulate Aβ-induced signal cascade in neurons [58–60]. However, the use of calcium channel blockers to mitigate AD outcomes is still much debated. For example, at least three clinical studies emphasized that elderly people, taking calcium channel blockers as antihypertensive drugs, were significantly more likely to experience cognitive decline than those using other agents [61–63].

At cellular level, it is well documented that abnormal amyloid metabolism induces an upregulation of neuronal calcium signaling, firstly resulting in a decline of memory and then leading to apoptosis [7, 50, 51, 64, 65]. An interesting connection between Aβ, calcium, and AD has been postulated by Arispe and coworkers [66], who suggested that Aβ oligomers can form calcium-permeable channels in membranes. It seems that energy deficits can promote this association, consistently with the observation that neurons with low cytosolic ATP levels showed a pronounced vulnerability to Aβ-induced toxicity [67]. In line with these reports, studies conducted in animal models (i.e., transgenic mice) highlighted an increase in calcium resting levels in the spines and dendrites of pyramidal cortical neurons [68, 69], supporting the hypothesis that calcium-permeable channels can form in the neuronal plasma membrane close to the Aβ plaques, thanks to the high concentration of Aβ oligomers found in these areas [67]. Tau protein is also able to form ion channels in planar lipid bilayer, with lack of ion selectivity and multiple channels conductance, thus contributing to lower membrane potential, dysregulate calcium, depolarize mitochondria, or deplete energy stores [70]. Within neurons, the increase in intracellular calcium levels stimulated by Aβ does not seem to be necessarily sustained by extracellular calcium influx. By using the human neuroblastoma SH-SY5Y cell line, Jensen and coworkers [71] interestingly described that the increase in intracellular calcium levels elicited by the A fragment can occur in the absence of extracellular calcium. Such observation supports the role of calcium release from the ER [72] to the generation of these signals. In addition, they demonstrated that this phenomenon relies only partially on inositol 1,4,5-trisphosphate (IP3) signaling, based on the fact that they observed the calcium mobilizing effect of A when the fragment was applied to permeabilized cells deficient in IP3 receptors (IP3R). Notably, this effect could underpin an additional direct effect of A upon the ER and a mechanism for induction of toxicity by intracellular A [71]. As a matter of fact, ryanodine receptors (RyR) can also contribute to the Aβ-induced calcium release from ER, as described by Ferreiro and coworkers [73, 74]. Exposing rat primary cortical neurons to A or to A peptides, the authors observed an increase in cytosolic calcium levels that was counteracted by either xestospongin C or dantrolene, pharmacological inhibitors of IP3R and RyR, respectively. Once calcium has been mobilized, it can initiate a cascade of events promoting free radicals generation, cytochrome c release from mitochondria, and activation of caspases, culminating in apoptotic cell death [73, 74]. It is worth mentioning that the balance between intracellular calcium levels and ER content involves not only IP3R and RyR, but also the activity of sarcoendoplasmic reticulum calcium ATPase (SERCA), which transports calcium ions from the cytoplasm into the ER (Figure 1). In this regard, Ferreiro and coworkers performed a comparative study by using the selective SERCA blocker thapsigargin [74]. They demonstrated that thapsigargin induced the loss of intracellular calcium homeostasis and the activation of caspase-3, leading to apoptotic cell death, as observed after incubation with A or A peptides. These findings lend support to the hypothesis that intracellular calcium deregulation induced by ER stress may be critical in the neurodegenerative processes triggered by Aβ peptide. Furthermore, the role of SERCA has been also investigated in the context of the FAD. Specifically, it has been proposed that SERCA activity is physiologically regulated by the interaction with presenilin [75], the membrane intrinsic protein that localizes predominantly to the ER membrane, which is responsible for the generation of the Aβ fragment. The finding that the modulation of SERCA activity would alter Aβ production may entail a possible role of the SERCA in the pathogenesis of AD [76].

The alteration of the glutamatergic system may be another important factor causing calcium imbalance in AD. Once released at glutamatergic synapses, glutamate is cleared from the extracellular space by the activity of the high affinity sodium-dependent glutamate transporters (Excitatory Amino Acid Transporters, EAATs) [77], which represent the most prominent system involved in terminating the excitatory signal, recycling the transmitter, and regulating extracellular levels of glutamate. As a result of overproduction and/or impaired clearance from synapses, glutamate may become excitotoxic. In this case, a prolonged exposure to glutamate induces an excessive activation of glutamate receptors, which is associated with a massive calcium influx through the receptor’s associated ion channel. The resulting calcium overload is particularly neurotoxic, leading to the activation of several degradation pathways which can have deleterious consequences on the cell fate [78–80]. Marked changes in functional elements of the glutamatergic synapses, such as glutamatergic receptors and transporters, have been described in AD. In 1996, Masliah and coworkers observed a deficit in glutamate transport activity in AD brains, likely occurring at neuronal level [81]. In line with this report, more recent findings suggested that soluble Aβ oligomers can disrupt neuronal glutamate uptake and promote long-term synaptic depression (LTD), a form of synaptic plasticity. In particular, the elegant study by Li and coworkers [82] showed that soluble Aβ oligomers from several sources, including human brain extracts, facilitated electrically evoked LTD in the mouse hippocampal CA1 region, involving both metabotropic and ionotropic glutamate receptors, and high extracellular glutamate levels. Accordingly, neuronal synaptic glutamate uptake was significantly decreased by Aβ. It is interesting to note that Aβ-facilitated LTD was mimicked by the action of the glutamate reuptake inhibitor DL-threo-beta-benzyloxyaspartate (TBOA), confirming that Aβ oligomers ability to perturb synaptic plasticity may rely upon glutamate recycling alteration at the synaptic level. In this regard, a dramatic reduction in the expression of two members of the EAAT family, EAAT1 and EAAT2, has been described at both gene and protein levels in hippocampus and gyrus frontalis medialis of AD patients [83]. Interestingly, in the same regions, glutamate receptors of the kainate type were significantly upregulated, further supporting the hypothesis that excitotoxic mechanisms can have a role in the pathogenesis of AD [79]. Such upregulation was accompanied by downregulation of the other ionotropic glutamate receptors, namely, N-methyl-D-aspartate (NMDA) and α-amino-3-hydroxyl-5-methyl-4-isoxazole-propionate (AMPA) receptors. Considering that both NMDA and AMPA receptors are known to mediate long-term potentiation [84, 85], the fundamental molecular mechanism of learning, memory, and cognition, their impairment may be considered a causative factor of the reduced cognitive functions observed in AD patients [83].

Although the observed alterations in intracellular calcium homeostasis in neurons significantly contribute to the pathogenesis of AD, more recent findings suggest that calcium dysregulation occurring in other cell types that support neuronal activity may contribute to degenerative processes [86]. In this regard, Fonseca and colleagues have recently demonstrated that Aβ may imbalance calcium homeostasis in brain endothelial cells with an increase in oxidative stress [87]. Using rat brain microvascular endothelial cells, they showed that the exposure to a toxic dose of Aβ alters ER ability to buffer calcium, and it enhances the mitochondrial and cytosolic response to ATP-stimulated ER calcium release. Although these responses are compensated after a longer exposure to Aβ, the early increase in oxidant levels and the concomitant decrease of antioxidant defenses induce deleterious effects on endothelial cells that undergo apoptosis, contributing to the cerebrovascular impairment observed in AD [87].

Astrocytes are also emerging as active players in AD [88], as highlighted in a recent paper by Dal Prà and coworkers [89]. They suggested an interesting issue concerning Aβ interaction with the Calcium Sensing Receptor (CaSR) [90]. The CaSR is a member of the largest family of cell surface receptors, the G protein-coupled receptors involved in calcium homeostasis. CaSRs expression is ubiquitous within the brain [91], where they are involved in several physiological processes, including synaptic plasticity and neurotransmission [92]. They showed that, in astrocytes, CaSR-Aβ interaction induces a downregulation of CaSR, leading the neighboring neurons to oversecrete de novo synthesized Aβ as well as nitric oxide (NO) and the toxic peroxynitrite (ONOO − ) [90, 93]. Recently, they have shown that the interaction occurring between Aβ and CaSR in human astrocytes may activate a signaling able to stimulate de novo production and secretion of vascular endothelial growth factor (VEGF) [89], whose excessive production can have toxic effects on neurons, astrocytes, and brain–blood barrier [94–97].

In general, the available literature suggests that the prolonged intracellular calcium elevation occurring within brain cells may be a crucial early event in AD pathogenesis, even though the mechanisms have not been fully explained.

In terms of proteins contributing to the calcium homeostasis in the brain, particular attention should be focused on NCX. NCX is a transporter that can move sodium across the membrane in exchange for calcium, operating in either calcium-efflux/sodium-influx mode (forward mode) or calcium-influx/sodium-efflux mode (reverse mode) depending upon the electrochemical ion gradients [24]. Three NCX isoforms have been described, namely, NCX1, NCX2, and NCX3, whose pattern of expression is tissue-specific [98]. Recent reports demonstrated the main role of NCX1 in controlling energy metabolism in several cells types, including neurons and astrocytes [99, 100]. In detail, our group recently reported a functional interaction between NCX1 and the sodium-dependent Excitatory Amino Acid Carrier 1 (EAAC1), at both plasma membrane and mitochondrial level in neuronal, glial, and cardiac models [99, 100]. Notably, we found that NCX1 reverse activity is necessary to restore transmembrane sodium gradient after glutamate entry into the cytoplasm, supporting glutamate utilization as a metabolic substrate that, in turn, enhances ATP production.

The role of NCX isoforms in the pathogenesis of AD is still under investigation. In 1991 Colvin and coworkers [101], measuring NCX activity in cerebral plasma membrane vesicles purified from human postmortem brain tissues of normal, AD, and non-AD origin dementia, identified a transporter altered kinetic in the vesicles of AD patients. The surviving neurons showed an increased NCX activity, leading authors to speculate that this phenomenon could help the surviving neurons to overtake the neurodegenerative process of AD, reinforcing the idea that the increase in intracellular calcium levels can play a major role in the pathogenesis of AD entailing the death of nonsurviving neurons. The hypothesis of an altered activity of NCX in AD patients represents an attractive mechanism that could, at least partially, be accountable for the calcium dysregulation observed in neurodegenerative processes accompanying the pathology [102]. The impairment of NCX activity can be related to the main features of AD. For instance, aggregated Aβ could interact with the hydrophobic surface of NCX, leading to an altered activity of the transporter [103] however, it cannot be excluded that the observed interaction of Aβ oligomers with the plasma membrane could be per se responsible for the alteration of NCX transport properties [103]. The pioneering study of Colvin has inspired further studies that explained the specific role of different NCX isoforms in AD in this regard, the study by Sokolow and coworkers offered a better understanding of the actual role of NCXs [104]. The analysis of NCX1, NCX2, and NCX3 expression in AD parietal cortex disclosed a specific pattern of expression within nerve terminals. In particular, NCX1 is the main isoform expressed in nerve terminals of cognitively normal patients, while NCX2 and NCX3 seem to be modulated in the parietal cortex in a late AD stage, as NCX2 expression is increased in positive terminals, while NCX3 expression is reduced [104]. Interestingly, the three isoforms colocalize with Aβ, supporting the hypothesis that the NCX activity modulation can be connected to a direct interaction with Aβ furthermore, in all synaptic terminals containing Aβ, NCX1-3 expression is upregulated [104]. It could be possible that the altered expression of NCX isoforms represents the neurons attempt to counterbalance the Aβ-induced alteration in calcium homeostasis. But, the different pattern observed in NCX isoforms expression can underpin a specific role for each isoform within the neurodegenerative process accompanying AD. In this regard, a specific alteration has been demonstrated for NCX3 isoform, leading to inactivation. NCX proteins can be inactivated by specific calpain 1 operated cleavage, and this can produce an increase of intracellular calcium levels contributing to the neurodegenerative calcium overload [105, 106]. In AD, the overproduction of Aβ increases calpain-mediated cleavage of NCX3, resulting in a decreased NCX3 activity [107]. Interestingly, the localization of NCX3 in dendrites and astrocytes processes contacting excitatory synapses [108] suggests the major role of NCX3 in regulating calcium current during synaptic activity, which is crucial for normal learning and memory. Therefore, reduced NCX3 activity can strongly contribute to the altered calcium levels associated with neuronal dysfunctions in AD [107].

3. Mitochondria and AD

Mitochondria are essential organelles for both cell survival and death, as they produce the largest part of cellular energy in the form of ATP and they play an active role in apoptosis induction [109, 110]. Mitochondria take part in cellular calcium signaling and act as highly localized buffers, thereby acting in the regulation of cytosolic calcium transient [111–113] (Figure 1). A crucial role in neurodegenerative disorders has been suggested for mitochondria, and AD patients have shown evidence of impaired mitochondrial function [114]. Reddy and coworkers demonstrated the upregulation of genes related to mitochondrial energy metabolism and apoptosis in an AD transgenic mouse model overexpressing a mutant form of APP at different stages of AD progression [115]. Mutant APP and soluble Aβ may enter mitochondria, which generate reactive oxygen species leading to oxidative damage, thereby affecting mitochondrial function. That is why the upregulation of mitochondrial genes could be a compensatory response to mitochondrial dysfunction induced by mutant APP or Aβ [115, 116].

In healthy neurons synaptic activity can be influenced by mitochondrial dynamics, such as fission and fusion events [117]. A number of studies demonstrate that essential proteins for fission and fusion are altered when APP is overexpressed [118, 119]. It has been shown that dynamin-like protein 1 (DLP1) and optic atrophy (OPA1) protein are significantly decreased, whereas levels of fission 1 (Fis1) are significantly increased in cell lines overexpressing APP [119] this leads to mitochondrial fragmentation and abnormal distribution, which contribute to mitochondrial and neuronal dysfunction [119]. These findings were confirmed by Gan and coworkers [120] that observed significant changes in mitochondria morphology and function in cytoplasmic hybrid (cybrid) neurons, where platelet mitochondria from AD and non-AD human subjects were incorporated into mitochondrial DNA-depleted neuronal cells. They found an impairment of fission/fusion proteins expression and function that was reverted by antioxidant treatment. Interestingly, they showed that oxidative stress negatively affects the extracellular-signal-regulated kinases (ERK) transduction pathway, which alters the expression levels of mitochondrial fission/fusion protein in AD cybrids [120].

Although it was common to focus primarily on Aβ, recently there has been an increasing interest on the role of the hyperphosphorylated form of tau. Hyperphosphorylation can decrease tau binding to microtubules, thereby affecting their stability and axonal transport of organelles, including mitochondria [8, 31]. Recent studies have begun to explore the effect of this altered protein on mitochondrial dynamics. Interesting findings come from the experiments performed by Schulz and coworkers [121] in SH-SY5Y wild-type (wt) and overexpressing P301L mutant tau. They demonstrated that P301L overexpression results in a substantial complex I deficit accompanied by decreased ATP levels and increased vulnerability to oxidative stress. Interestingly, those events were paralleled by pronounced changes in mitochondrial morphology and decreased fusion/fission rates, observed as reduced expression of several fission and fusion proteins such as OPA-1 or DLP- 1 [121]. An imbalance in fission/fusion proteins has also been shown by Manczak and Reddy [122] who demonstrated a physical link between phosphorylated tau and DLP1. The authors concluded that the interaction between phosphorylated tau, DLP1, and Aβ can cause an excessive mitochondrial fragmentation and both mitochondrial and synaptic deficiencies, leading to neuronal damage and cognitive decline [122]. Regardless of its connection with fission/fusion events, the synergistic action of Aβ and tau has been further investigated in a recent study by Quintanilla and colleagues who demonstrated that, in aging neuronal cultures, phosphorylated tau potentiates Aβ-induced mitochondrial dysfunction by affecting mitochondrial membrane potential and increasing oxidative stress [123]. In a previous study, the same group demonstrated that also a truncated form of tau, cleaved at Asp421 by caspases [124], significantly increases oxidative stress response in cortical neurons treated with sublethal concentrations of Aβ [125]. Moreover, interesting results in this field have been obtained by using triple transgenic mice. This model has been obtained by cross-breeding tau transgenic pR5 mice, characterized by tangle formation, and double-transgenic APP152 mice developing Aβ plaques. Only triple transgenic mice, combining both pathologies, at early age (8 months old) showed a reduction of the mitochondrial membrane potential, while at the age of 12 months they showed the strongest defects on oxidative phosphorylation, synthesis of ATP, and reactive oxygen species formation, emphasizing synergistic and age-associated effects of Aβ and tau in perishing mitochondria [126]. Globally, these findings clearly demonstrate that mitochondrial function can be seriously impaired by Aβ and that hyperphosphorylation of tau can enhance the Aβ-induced mitochondrial neuronal damage. Notably, mitochondria are also involved in the maintenance of cellular activities through the contact they establish with ER [127, 128]. Mitochondria-associated ER membranes (MAMs) are intracellular lipid rafts regulating calcium homeostasis and several metabolic pathways, such as glucose, phospholipids, and cholesterol metabolism [127, 129]. The physical interaction between these organelles has been extensively studied, and several MAMs-associated proteins have been identified. A recent research has shown that the contact sites between mitochondria and ER are enriched in PS1 and PS2 [130], components of the γ-secretase complex which processes APP to produce Aβ [131]. A large body of evidence indicates PS1 and PS2 mutations as being responsible for the Aβ overproduction by γ-secretase activity leading to FAD [132, 133]. Recently, it has been shown that mutations in PS1, PS2, and APP can upregulate MAMs function and produce a significant increase in ER-mitochondrial connectivity, suggesting that presenilins can negatively regulate this phenomenon [134]. However, the same upregulation in MAMs function and ER-mitochondrial communication has been found in fibroblasts from patients with sporadic AD (SAD), in which there are no mutations in PS1, PS2, and APP structure [134]. This interesting finding suggests that the upregulated function of MAMs, as a common feature in both FAD and SAD, may represent a pathogenic initiator of AD [134]. A recent study by Schreiner and colleagues [135] supports this hypothesis. In this work, the authors determined the production of Aβ in subcellular fractions isolated from mouse brain. They found that a large amount of Aβ was produced at mitochondria-ER contact sites. They postulated that the enhanced Aβ production may perturb mitochondria and mitochondria-ER contact site functions, leading to neurodegeneration and, therefore, to AD [135]. As a matter of fact, the MAMs structure has been postulated to modulate calcium signals and synaptic and integrative activities at neuronal level [127, 136]. In this regard, it has been suggested that MAMs may host important physiological functions related to neuronal integrity, as they have been reported to be uniformly distributed throughout hippocampal neurons and at synaptic level [127]. In particular, two main proteins have been identified as being crucial for MAMs activity and, consequently, for neuronal integrity: phosphofurin acidic cluster sorting protein-2 (PACS-2) and σ1 receptor (σ1R) [127]. These proteins contribute to maintaining MAMs homeostasis. Specifically, PACS-2 is a multifunctional sorting protein controlling ER-mitochondria communication and apoptosis [137], whereas σ1R promotes calcium transport into mitochondria from the ER by interacting with the IP3R [138]. Their knockdown results in neurodegeneration, and this highlights the importance of these proteins in the maintenance of neuronal integrity [127]. Furthermore, exposure to Aβ results in the increase of MAMs-associated proteins expression and of the amount of contact points between ER and mitochondria in different AD models (namely, APP transgenic mice, primary neurons, and AD brain) [127]. In turn, the alteration in MAMs-associated proteins expression can affect calcium homeostasis, which has been considered an underlying and integral component of AD pathology [7, 50, 67]. This issue is further discussed in the following section.

3.1. Role of Mitochondria in Intracellular Calcium Balance

Intracellular calcium dysregulation is a central event in neurodegeneration it involves plasma membrane transporters and also intracellular organelles, such as mitochondria, thereby creating an endless futile cycle that can have several consequences on neuronal survival [67]. The excess of intracellular calcium is taken up by mitochondrial calcium uniporter (MCU) that, through the large electrochemical gradient across the inner mitochondrial membrane, drives calcium from the cytosol to the mitochondrial matrix [67] (Figure 1). Calcium is then released back into the cytosol through the activity of mitochondrial NCX (mNCX, Figures 1 and 2(a)) [67]. However, mNCX may reverse its mode of operation (Figure 2(b)) from a calcium efflux system to an influx pathway allowing the access of calcium ions into the mitochondrial matrix [26]. Although the molecular identity of mNCX has been extensively researched and strongly debated, our group has provided data showing that plasma membrane NCX (plmNCX) isoforms can contribute to mNCXs. Exploring the subcellular distribution of NCX in the central nervous system by western blot and in situ electron microscopy immunocytochemistry in rat neocortex and hippocampus, we observed a large population of neuronal and astrocytic mitochondria expressing NCX1–3 [26, 27] (Figures 3 and 4). Thus, these mitochondrial calcium transporters manage intracellular changes of this “versatile” ion, impacting several cell functions, including cell metabolism. As a matter of fact, the activity of several intramitochondrial dehydrogenases is enhanced by increased mitochondrial calcium levels, thereby stimulating ATP synthesis [139, 140]. The brain is one of the most metabolically active organs in the body. The brain’s high energy requirements are mainly due to maintenance and restoration of ion gradients dissipated by signaling processes such as postsynaptic and action potentials, as well as uptake and recycling of neurotransmitters. In AD, the impairment in energy production is one of the factors greatly contributing to the vulnerability of neuronal cells [141]. One of the main works demonstrating the cooperative action of tau and Aβ shows, through a proteomic analysis, that one-third of the deregulated proteins in different AD mouse models is made up of mitochondrial proteins involved in oxidative phosphorylation [126]. Hence, it is tempting to speculate that modulation of mitochondrial calcium transporter activity toward the increase in ATP production could have beneficial effects on neuronal survival during the neurodegenerative processes that characterize AD. In this context, it has been suggested that a partial inhibition of mNCX would lead to an increase of the mitochondrial matrix calcium concentration to a higher physiological steady-state level that could stimulate calcium-sensitive dehydrogenase activity and the rate of ATP synthesis [67, 139, 140]. Therefore, calcium may play a dual role within cells: on the one hand it can help vulnerable neurons increase the rate of ATP synthesis on the other hand it can be harmful and activate cell death through the induction of the apoptotic pathways [142]. Thus, there must be a critical point representing the boundary between cytoprotective and cytotoxic effect due to the increase in mitochondrial calcium concentration [67]. An increased rate of ATP synthesis can be achieved stimulating the cell in several ways. Recently, our group found that both plmNCX and mNCX can act synergically to sustain the increase in ATP synthesis promoted by glutamate [99, 100]. As reported above, this metabolic response results from a physical and functional interaction between NCX (particularly NCX1) and EAATs, with particular reference to EAAC1, occurring at both plasma membrane and mitochondrial level [99, 100]. The fact that some substrates, such as glutamate, can modulate ATP synthesis may have several implications for AD too, and this can reverse the traditional view of a predominantly harmful effect of this amino acid, towards a benefic role that is able to rescue vulnerable neurons from death. At present, the role of mNCX in AD is still largely unexplored [26]. In an interesting paper, Thiffault and Bennett [143] reported indirect evidence of an involvement of the exchanger in AD. In particular, they showed that cells, lacking endogenous mitochondria and repopulated with mitochondria from AD patients, virtually lack the spontaneous fluctuations in mitochondrial membrane potential (

), also called “ flickering,” which is normally induced by cyclosporine. It is worth noting that mNCX blockade with CGP-37157 suppresses flickering in control cells, thus recreating a condition similar to the one observed in AD. The role of mNCX in AD is also supported by the work of Chin and colleagues [144], who observed that Aβ potentiates the increase in cytosolic calcium concentration evoked by nicotine in dissociated rat basal forebrain neurons in a CGP-37157-sensitive way.


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DISCUSSION

Overall, we demonstrated that GCaMP-based imaging allows fission yeast to serve as a powerful model organism for the study of calcium transients during various cellular processes. Using this method, we discovered cytokinetic calcium spikes in this unicellular model organism for the first time. These calcium transients are similar to those first uncovered in animal embryos. Calcium likely plays a critical role in promoting ring constriction and daughter cell integrity through these transients.

GCaMP-based calcium imaging of fission yeast

To our knowledge, our study is the first to examine the feasibility of GCaMP-based calcium imaging in fission yeast cells. We are fortunate that fission yeast is quite tolerant of the expression of this calcium indicator. It does not perturb many cellular processes that we examined, even when it is expressed constitutively. More importantly, GCaMP is also highly sensitive as a probe for time-lapse microscopy, allowing us to capture calcium transients at a relatively low sampling rate. Compared with synthetic calcium probes, GCaMP exhibited two key advantages in fission yeast. First, unlike synthetic probes such as Calcium Green (Chang and Meng, 1995), GCaMP can be maintained at a constant intracellular concentration through the homogenous expression of this reporter. Combined with quantitative microscopy, this largely eliminated the heterogeneity of fluorescence intensities among cells. In contrast, the intracellular concentration of the synthetic probes can be highly variable among cells. This advantage of GCaMP was exemplified by the identification of outliers with high calcium concentration among a large number of cells. The other advantage of GCaMP is its more uniform cellular distribution, as compared with the synthetic probes. This allows detection of calcium transients throughout the cytoplasm as well as the nucleus.

When combined with yeast genetics, GCaMP can be a very versatile tool for studying calcium signaling. As an example, we constructed both GCaMP and GCaMP-mCherry, both of which have potential to be employed in the future studies. We employed GCaMP, not GCaMP-mCherry, throughout this study, primarily to preserve the spectrum for imaging the fluorescence protein markers of cytokinesis. Nevertheless, the tandem reporter can be used for ratiometric imaging of intracellular calcium, potentially providing a higher signal-to-noise ratio. Overall, application of GCaMP will make fission yeast an attractive model organism for studying calcium transients and homeostasis in nonexcitable cells.

Evolutional conservation of the cytokinetic calcium transients

Our study provides fresh evidence for the existence of calcium transients in a unicellular organism. Similarly to the earlier studies (Fluck et al., 1991 Chang and Meng, 1995 Noguchi and Mabuchi, 2002), we employed live fluorescence microscopy to determine the temporal correlation between the calcium transients and cytokinesis. With the availability of fluorescence protein markers of cytokinesis, we now determined the temporal regulation of these spikes with more precision. In addition to time-lapse microscopy, we sought out an alternative approach to illustrate this close correlation. We analyzed just snapshots of many GCaMP-expressing cells to reveal the relationship between cell division and intracellular calcium level. Compared with the movies, the snapshots examined all cells regardless of their cell-cycle stage, presenting a less biased view of the potential calcium change in them. Combined with the previous studies of animal embryos, our results strongly support the conclusion that a temporal increase of calcium during cytokinesis may be evolutionally conserved.

The temporal regulation of fission yeast cytokinetic calcium spikes bears strong similarities to that found in the animal embryos. The constriction spikes of fission yeast are comparable to the first “furrowing wave” observed in the animal embryos (Fluck et al., 1991), which initiates just as the cleavage furrow starts to ingress (Chang and Meng, 1995 Noguchi and Mabuchi, 2002). The separation spikes are comparable to the second “zipping wave” of the embryos, which starts just when the two daughter cells separate (Fluck et al., 1991 Noguchi and Mabuchi, 2002). Both the calcium waves in the embryos and the spikes in fission yeast can last for minutes, far longer than other known calcium transients (Jaffe and Creton, 1998). Although no calcium spikes have been identified yet during budding yeast cytokinesis, Carbo et al. (2017) observed higher frequency of calcium bursts during G1 to S phase transition in synchronized cells, which may be similar to the separation calcium spikes in fission yeast. Overall, our study suggests that the regulatory mechanism of calcium during cytokinesis may have been conserved as well.

There are also key differences between the calcium spikes and these embryonic calcium waves. Chief among them is the spatial regulation of these calcium transients. In fission yeast, the calcium spikes propagate globally throughout the cytoplasm and nucleus. In contrast, the calcium waves of animal embryos are restricted to the cleavage furrow (Fluck et al., 1991 Chang and Meng, 1995) or its surrounding region (Noguchi and Mabuchi, 2002). Although this difference may be due to a difference in sensitivity of the calcium probes, we propose that this distinction is likely due to the relatively small size of fission yeast cells, measuring only 4 µm wide and 14 µm long, compared with embryos of hundreds of micrometers. So far, we have found no evidence that the constriction spike of fission yeast triggers contractile ring constriction, even though there is evidence that the furrowing waves trigger the ring constriction (Fluck et al., 1991 Miller et al., 1993 Chang and Meng, 1995). Last, the asymmetry of the separation spikes is also unique to fission yeast. It may be linked to the asymmetric turgor pressure in the two daughter cells during separation.

Mechanism of the calcium spikes and their potential roles

Although the regulatory mechanism for these cytokinetic calcium spikes remains to be explored, our data suggest that they likely draw calcium from both the influx and internal release. Inhibition of the influx through depleting extracellular calcium inhibited the spikes significantly but not completely. Under this condition, the cytokinetic calcium spikes still occurred, albeit with greatly diminished amplitude. This points to ER- or vacuole-stored calcium as the other likely source for the spikes. Supporting this hypothesis is our observation that EGTA exhibited a stronger inhibitory effect on cytokinesis, compared with the calcium-free EMM medium. EGTA may have chelated the intracellular calcium slowly during long-term incubation. This is consistent with the frequent lysis of pmr1Δ mutant during the cell separation in the calcium-free medium. In comparison, the embryonic calcium waves draw only from the internal store (Miller et al., 1993 Chang and Meng, 1995). It remains unclear what are the ion channels mediating the cytokinetic calcium transients. Fluck et al. (1991) first proposed the potential role of tension-sensing calcium channels, which can be activated during cytokinesis. This remains a feasible mechanism, considering increased membrane tension on the cleavage furrow. On the other hand, release of internal calcium could be through the store-operated calcium release (SOCE) channels (Chan et al., 2015, 2016). Nevertheless, none of these cytokinetic ion channels have been identified.

Although our study clearly showed that calcium is important for fission yeast cytokinesis, it leaves unanswered whether this is exclusively through the calcium spikes. This is in part due to the technical difficulty of separating the function of the spikes from that of the intracellular calcium level. Our method of depleting extracellular calcium led to both inhibition of calcium spikes and a slight drop of intracellular calcium level. Although we favor the model that these spikes can regulate cytokinesis, our current results cannot rule out the possibility that a threshold level of intracellular calcium is sufficient for cytokinesis. From comparing the inhibition of cytokinesis by 1 and 2 mM EGTA, it is likely that both the spikes and the intracellular calcium level contribute to cytokinesis. Nevertheless, both the constriction and separation spikes likely activate the calcium signaling pathways during cytokinesis. The constriction spikes may increase the activity of type II myosin in the ring through the phosphorylation of myosin regulatory light chain (Fluck et al., 1991 Chang and Meng, 1995). Alternatively, the transient increase of calcium could stimulate the septum biosynthesis through activating the calcium-dependent calcineurin (Yoshida et al., 1994 Cadou et al., 2013 Martin-Garcia et al., 2018). The separation spikes could result from a drop in turgor pressure in the daughter cells following the separation, similar to those triggered by hypoosmotic stress. Alternatively, these spikes may modulate the turgor pressure throughout the cell separation (Proctor et al., 2012 Abenza et al., 2015). Further studies will be required to determine the downstream targets of cytokinetic calcium spikes.


Conclusion

Our findings indicate that nerve injury associated with hyperalgesia depresses resting [Ca 2+ ] c in sensory neurons. This change is most evident in nonnociceptive axotomized neurons of L5 after SNL, which supports a central role of this neuronal group in the genesis of neuropathic pain.9However, these observations do not eliminate the possibly critical contributions of functional changes in calcium signaling of neurons that remain in continuity to their receptive fields, since we also identified decreased resting [Ca 2+ ] c in large L4 neurons after SNL.


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3. Results

Brain-derived neurotrophic factor (BDNF), signaling through its plasma membrane tyrosine kinase receptor TrkB, activates several intracellular signaling cascades (Segal and Greenberg, 1996) which may directly elicit Ca 2+ release from intracellular stores via IP3 production, and/or indirectly modulate Ca 2+ influx through voltage-gated Ca 2+ channels or NMDA receptors via enhanced protein phosphorylation of specific subunits. Our proposed model of BDNF modulation of neuronal Ca 2+ signaling is illustrated in Fig. 1 . To begin addressing the modulation of dendritic Ca 2+ signaling in hippocampal neurons by BDNF, we focused on Ca 2+ influx into proximal apical dendrites and somatic regions during short trains of back-propagating action potentials (APs) evoked by depolarizing current injection.

Hypothesized regulation of the different routes of Ca 2+ influx and/or release from stores by NTs, and its effects on intracellular Ca 2+ levels. (A) CA1 pyramidal neuron at rest (left). Intracellular calcium concentration is represented following the standard convention utilized to display data from imaging experiments using Ca 2+ -sensitive fluorescent dyes (blue to red, low to high Ca 2+ levels, respectively). The diagram to the right is an expanded view of an excitatory synapse on a dendritic spine framed by a box in the apical dendrite at left. (B) Pre-synaptic release of excitatory neurotransmitter from afferent fibers depolarizes the post-synaptic membrane, eliciting Ca 2+ elevations by influx through voltage-dependent Ca 2+ channels and NMDA receptors. Activation of metabotropic glutamate receptors also contributes to Ca 2+ rises by release from IP3-sensitive intracellular stores. Ca 2+ -induced Ca 2+ release from ryanodine stores further amplifies dendritic Ca 2+ elevations. (C) Our model proposes that, in addition to evoke Ca 2+ changes by itself, BDNF enhances dendritic Ca 2+ elevations through other routes evoked by synaptic activity. BDNF binding induces autophosphorylation of TrkB receptors, and subsequent activation of the PLC-γ/IP3 signaling pathway, leading to Ca 2+ release from intracellular stores. Activated TrkB receptors also trigger tyrosine phosphorylation of voltage-dependent Ca 2+ channels and NMDA receptors, enhancing Ca 2+ influx into spines and dendrites.

Whole-cell intracellular recordings were performed under current-clamp from visually identified CA1 pyramidal neurons from 9 to 11 days in vitro (div.) hippocampal slice cultures using patch pipettes containing the fluorescent Ca 2+ indicator bis-fura-2 (250 µM in K + -gluconate intracellular solution). Simultaneous fluorescence digital imaging of dye-filled neurons was restricted to proximal apical dendrites (up to 100 µm from the soma). No significant differences (P > 0.05) were found between CA1 pyramidal neurons from serum-free controls (n = 5) and slices treated with BDNF (250 ng/ml) for 2 to 4 div. (n = 5) with respect to resting membrane potentials (� ± 1 mV versus � ± 2 mV, control versus BDNF), input resistances (103 ± 17 MΩ versus 87 ± 7 MΩ, control versus BDNF), or resting intracellular Ca 2+ concentrations (357/380 ratio = 0.15 ± .02 versus 0.14 ± 0.03, control versus BDNF).

3.1. BDNF does not affect Ca 2+ transients evoked by back-propagating action potentials in CA1 pyramidal neuron dendrites

Figure 2 shows the transient elevations of [Ca 2+ ]i in the proximal apical dendrites of CA1 pyramidal neurons (shown at left) in serum-free control and BDNF-treated slices, evoked by a train of 10 APs at 20 Hz elicited by short (5 ms) depolarizing current injections into their somas via the patch pipette. To prevent excessive network activity in the slice during depolarizing current injections into the cell under study, the AMPA receptor non-competitive antagonist GYKI (20 µM) was routinely added to the extracellular recording solution. Calcium concentration changes within the areas marked by the regions-of-interest (ROIs) in the fluorescent images (380 nm excitation, Fig. 2 , left) were acquired at 20� frames per second simultaneously with recordings of membrane potential in the current-clamp mode, and are plotted as a function of time ( Fig. 2 , traces at right). Rapid Ca 2+ elevations within proximal apical dendrites occur immediately after the firing of the first APs, and reach a maximum level by the end of the 10 AP train. Virtually equal spatio–temporal patterns of transient elevations of Ca 2+ concentration were observed in CA1 neurons from serum-free controls (n = 5) and BDNF-treated slices (n = 5). In addition, no significant differences were observed in the peak dendritic Ca 2+ concentrations evoked by these trains of 10 APs at 20 Hz between neurons from serum-free controls (Δ R = 0.033ଐ.007, n = 5) and BDNF-treated slices (Δ R = 0.025 ± 0.005, n = 5, t-test, P > 0.05). These results suggest that long-term exposure to BDNF does not affect voltage-gated Ca 2+ entry in proximal apical dendrites of CA1 pyramidal neurons in postnatal hippocampal slices.

Voltage-dependent Ca 2+ elevations in CA1 pyramidal neurons during back-propagating APs are not affected by BDNF treatment. No differences in the spatio-temporal pattern or in the peak amplitude of voltage-dependent Ca 2+ transients were observed between serum-free controls and BDNF-treated slices. A train of 10 back-propagating APs (middle right traces) was evoked by short depolarizing current pulses (bottom right traces) delivered by the recording patch pipette in the soma. The square pulses shown in the top right traces represent the TTL signal output of the digital camera during image frame exposures used to synchronize optical and electrophysiological recordings. Top left, fluorescence image (380 nm excitation) of a CA1 pyramidal neuron from a serum-free control slice. Bottom left, CA1 pyramidal neuron from a BDNF-treated slice culture (250 ng/ml for 48 h). Topmost right traces show intracellular Ca 2+ concentration levels calculated from the dotted ROIs shown in the fluorescence images at left after background-subtraction from a ROI over the slice but outside the dye-filled cell. Membrane voltage and current traces (at 10 kHz) were acquired synchronously with Ca 2+ concentration traces (at 10 ms frame interval), and are shown with the same time base.

3.2. BDNF does not affect Ca 2+ transients mediated by L-type Ca 2+ channels during back-propagating AP firing in CA1 pyramidal neuron dendrites and somas

Due to their clustering at the soma and in the base of proximal dendrites, high-threshold L-type voltage-gated Ca 2+ channels are the most likely route of Ca 2+ entry in those regions during back-propagating APs (Ahlijanian et al., 1990 Westenbroek et al., 1990). To test if the proportion of Ca 2+ entry mediated by L-type channels was different between serum-starved and BDNF-treated neurons, we used the L-type channel blocker nifedipine (20 µM) and measured the magnitude of the blockade of Ca 2+ transients during APs trains ( Fig. 3 ). The magnitude of the L-type component of the dendritic and somatic Ca 2+ transients observed here is in good agreement with similar Ca 2+ measurements in CA1 neurons in acute slices (Christie et al., 1995), and in CA3 neurons in slice cultures (Elliott et al., 1995). Nifedipine reduced dendritic Ca 2+ transients during APs trains by 19% (serum-free, n = 5) and 26% (BDNF-treated, n = 5), while somatic Ca 2+ transients were reduced by 22 and 20% in serum-free and BDNF-treated neurons, respectively ( Fig. 4 ). Therefore, there were no significant differences (P > 0.05) between serum-free controls and BDNF-treated neurons in the amount of L-type channel block by nifedipine. Similar results were obtained using another dihydropyridine L-type channel blocker, ni-modipine (not shown). These results strongly suggest that long-term exposure to BDNF does not affect dendritic Ca 2+ entry mediated by L-type Ca 2+ channels during the firing of back-propagating APs in CA1 pyramidal neurons.

Voltage-dependent Ca 2+ elevations in CA1 pyramidal neurons evoked by 10 short depolarizing current pulses are partially sensitive to the L-type Ca 2+ channel blocker nifedipine. The top panel shows a fluorescence image (380 nm excitation) of a representative CA1 pyramidal neuron from a BDNF-treated slice, and the traces below show the effect of the L-type Ca 2+ channel blocker nifedipine on the Ca 2+ elevations during a train of 10 back-propagating APs. The order of the bottom traces is the same as in Fig. 2 (top to bottom, Ca 2+ profile from the ROIs in the fluorescence image, expose camera signal TTL, action potentials in membrane voltage, injected current).

The proportion of nifedipine block is not different between neurons from serum-free controls and BDNF-treated slices. Summary bar graphs of all experiments on the proportion of L-type Ca 2+ channel block of Ca 2+ transients during trains of 10 back-propagating APs. Nifedipine blocked between 19 and 26% of dendritic and somatic Ca 2+ transients, and there were no differences between CA1 pyramidal neurons from serum-free controls and BDNF-treated slices (t-test, P > 0.05).


THE CONTRACTILE MECHANISM

In the intact body, the process of smooth muscle cell contraction is regulated principally by receptor and mechanical (stretch) activation of the contractile proteins myosin and actin. A change in membrane potential, brought on by the firing of action potentials or by activation of stretch-dependent ion channels in the plasma membrane, can also trigger contraction. For contraction to occur, myosin light chain kinase (MLC kinase) must phosphorylate the 20-kDa light chain of myosin, enabling the molecular interaction of myosin with actin. Energy released from ATP by myosin ATPase activity results in the cycling of the myosin cross-bridges with actin for contraction. Thus contractile activity in smooth muscle is determined primarily by the phosphorylation state of the light chain of myosin—a highly regulated process. In some smooth muscle cells, the phosphorylation of the light chain of myosin is maintained at a low level in the absence of external stimuli (i.e., no receptor or mechanical activation). This activity results in what is known as smooth muscle tone and its intensity can be varied.


Materials and methods

Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Genetic reagent (D. melanogaster)w1118Bloomington Drosophila Stock Center (BDSC)RRID:BDSC_5905
Genetic reagent (D. melanogaster)D42-Gal4BDSCRRID:BDSC_8816
Genetic reagent (D. melanogaster)TH-Gal4BDSCRRID:BDSC_8848
Genetic reagent (D. melanogaster)repo-Gal4BDSCRRID:BDSC_7415
Genetic reagent (D. melanogaster)UAS-mCD8-RFPBDSCRRID:BDSC_27399
Genetic reagent (D. melanogaster)UAS-CD4-tdTomBDSCRRID:BDSC_35841
Genetic reagent (D. melanogaster)Itpr ka1091/+BDSCRRID:BDSC_30739
Genetic reagent (D. melanogaster)Itpr sv35/+BDSCRRID:BDSC_30740
Genetic reagent (D. melanogaster)20XUAS-ChR2.T159C-HABDSCRRID:BDSC_52258
Genetic reagent (D. melanogaster)UAS-Chr2.sBDSCRRID:BDSC_9681
Genetic reagent (D. melanogaster)RyR 16/+BDSCRRID:BDSC_6812
Genetic reagent (D. melanogaster)UAS-NachBacBDSCRRID:BDSC_9467
Genetic reagent (D. melanogaster)UAS-R-GECO1-IR1,UAS-R-GECO1.L-IR2BDSCRRID:BDSC_52222
Genetic reagent (D. melanogaster)UAS-Cam RNAiBDSCRRID:BDSC_34609
Genetic reagent (D. melanogaster)UAS-CanA-14F RNAiBDSCRRID:BDSC_58249
Genetic reagent (D. melanogaster)UAS-CanB RNAiBDSCRRID:BDSC_27307
Genetic reagent (D. melanogaster)UAS-CaMKI RNAiBDSCRRID:BDSC_35362
Genetic reagent (D. melanogaster)UAS-Pka-C1 RNAiBDSCRRID:BDSC_31599
Genetic reagent (D. melanogaster)UAS-Pkc53E RNAiBDSCRRID:BDSC_55864
Genetic reagent (D. melanogaster)UAS-CalpA RNAiBDSCRRID:BDSC_29455
Genetic reagent (D. melanogaster)UAS-CalpB RNAiBDSCRRID:BDSC_25963
Genetic reagent (D. melanogaster)UAS-NFAT RNAiBDSCRRID:BDSC_51422
Genetic reagent (D. melanogaster)UAS-nej RNAiBDSCRRID:BDSC_37489
Genetic reagent (D. melanogaster)UAS-CrebA RNAiBDSCRRID:BDSC_27648
Genetic reagent (D. melanogaster)UAS-CrebB RNAiBDSCRRID:BDSC_63681
Genetic reagent (D. melanogaster)UAS-Imp α1 RNAiBDSCRRID:BDSC_27523
Genetic reagent (D. melanogaster)UAS-Imp α2 RNAiBDSCRRID:BDSC_27692
Genetic reagent (D. melanogaster)UAS-Imp α3 RNAiBDSCRRID:BDSC_27535
Genetic reagent (D. melanogaster)UAS-Imp β1 RNAiBDSCRRID:BDSC_31242
Genetic reagent (D. melanogaster)UAS-Imp 7 RNAiBDSCRRID:BDSC_33626
Genetic reagent (D. melanogaster)UAS-Imp β11 RNAiBDSCRRID:BDSC_55142
Genetic reagent (D. melanogaster)UAS-Tnpo RNAiBDSCRRID:BDSC_50732
Genetic reagent (D. melanogaster)UAS-Tnpo-SR RNAiBDSCRRID:BDSC_56974
Genetic reagent (D. melanogaster)UAS-Ran RNAiBDSCRRID:BDSC_42482
Genetic reagent (D. melanogaster)UAS-Ntf-2 RNAiBDSCRRID:BDSC_28633
Genetic reagent (D. melanogaster)UAS-LuciferaseBDSCRRID:BDSC_35788
Genetic reagent (D. melanogaster)UAS-Hrb87F RNAiBDSCRRID:BDSC_52937
Genetic reagent (D. melanogaster)UAS-HNPNPC RNAiBDSCRRID:BDSC_42506
Genetic reagent (D. melanogaster)UAS-glo RNAiBDSCRRID:BDSC_33668
Genetic reagent (D. melanogaster)UAS-Syp RNAiBDSCRRID:BDSC_56972
Genetic reagent (D. melanogaster)UAS-poly-PR.PO-100BDSCRRID:BDSC_58698
Genetic reagent (D. melanogaster)UAS-poly-GR.PO-100BDSCRRID:BDSC_58696
Genetic reagent (D. melanogaster)Gmr-Gal4BDSCRRID:BDSC_1104
Genetic reagent (D. melanogaster)UAS-MJD-tr78QBDSCRRID:BDSC_8150
Genetic reagent (D. melanogaster)Df(2R)BSC26BDSCRRID:BDSC_6866
Genetic reagent (D. melanogaster)UAS-SERCA RNAiVienna Drosophila Resource Center (VDRC)VDRC: 107446 RRID:FlyBase_FBst0479267
Genetic reagent (D. melanogaster)UAS-Itpr RNAiVDRCVDRC: 106982 RRID:FlyBase_FBst0478805
Genetic reagent (D. melanogaster)UAS-RyR RNAiVDRCVDRC: 109631 RRID:FlyBase_FBst0481295
Genetic reagent (D. melanogaster)UAS-CaMKII RNAiVDRCVDRC: 100265 RRID:FlyBase_FBst0472139
Genetic reagent (D. melanogaster)UAS-Imp α3 RNAiVDRCVDRC: 106249 RRID:FlyBase_FBst0478074
Genetic reagent (D. melanogaster)UAS-RanGAP RNAiVDRCVDRC: 108264 RRID:FlyBase_FBst0480076
Genetic reagent (D. melanogaster)UAS-Rcc1 RNAiVDRCVDRC: 110321 RRID:FlyBase_FBst0481896
Genetic reagent (D. melanogaster)UAS-Hrb98DE RNAiVDRCVDRC: 29524 RRID:FlyBase_FBst0458009
Genetic reagent (D. melanogaster)UAS-Hrb27C RNAiVDRCVDRC: 101555 RRID:FlyBase_FBst0473428
Genetic reagent (D. melanogaster)UAS-HNRNPU1 RNAiVDRCVDRC:106984 RRID:FlyBase_FBst0478807
Genetic reagent (D. melanogaster)UAS-Sm RNAiVDRCVDRC:108351 RRID:FlyBase_FBst0480162
Genetic reagent (D. melanogaster)UAS-CalpA RNAiVDRCVDRC:101294 RRID:FlyBase_FBst0473167
Genetic reagent (D. melanogaster)40D UASVDRCVDRC ID: 60101
Genetic reagent (D. melanogaster)UAS-TBPH-Flag-HABangalore Fly Resource CenterDrosophila
Protein interaction
Map (DPiM)
Genetic reagent (D. melanogaster)ppk 1a -Gal4Han et al., 2011 Yuh Nung Jan (University of California, San Francisco (UCSF))
Genetic reagent (D. melanogaster)UAS-tdTomato P2A GCaMP5G, attp1Daniels et al., 2014 Barry Ganetzky (University of Wisconsin-Madison)
Genetic reagent (D. melanogaster)UAS-3xMyc-RFP-TDP-43Wang et al., 2011 Brian D. McCabe, (Swiss Federal Institute of Technology (EPFL))
Genetic reagent (D. melanogaster)UAS-TBPHWang et al., 2011 Brian D. McCabe, (Swiss Federal Institute of Technology (EPFL))
Genetic reagent (D. melanogaster)UAS-TDP-43 WTVoigt et al., 2010 Aaron Voigt (University Hospital, RWTH Aachen University)
Genetic reagent (D. melanogaster)UAS-TDP-43 G287SVoigt et al., 2010 Aaron Voigt (University Hospital, RWTH Aachen University)
Genetic reagent (D. melanogaster)UAS-Flag-TDP-43Miguel et al., 2011 Magalie Lecourtois (University of Rouen)
Genetic reagent (D. melanogaster)UAS-Flag-TDP-43-ΔNLSMiguel et al., 2011 Magalie Lecourtois (University of Rouen)
Genetic reagent (D. melanogaster)UAS-2xFlag-Imp α3 (vk00002)This paperSB Lab (DGIST)
Genetic reagent (D. melanogaster)UAS-V5-Imp β1 (vk00002)This paperSB Lab (DGIST)
Genetic reagent (D. melanogaster)UAS-CalpA-2xMyc (vk00002)This paperSB Lab (DGIST)
Genetic reagent (D. melanogaster)UAS-emptyPark et al., 2020SB Lab (DGIST)
AntibodyMouse monoclonal anti-Flag (DYKDDDDK)WakoCat#: 012–22384 RRID:AB_10659717IHC (1:400)
AntibodyRat monoclonal anti-HARocheCat#: 11867423001 RRID:AB_390918IHC (1:200)
AntibodyRabbit anti-TBPHLTK BioLaboratories, Taiwan
(Lin et al., 2011) C.-K. James Shen (Taipei Medical University)
IHC (1:100)
AntibodyRabbit polyclonal anti-TDP-43ProteintechCat#: 10782–2-AP, RRID:AB_615042IHC (1:400)
AntibodyGoat polyclonal anti-mouse Alexa Fluor 647InvitrogenCat#: A21236 RRID:AB_2535805IHC (1:400)
AntibodyGoat polyclonal anti-rat Alexa Fluor 647Jackson Immunoresearch LaboratoriesCat#: 112-605-003 RRID:AB_2338393IHC (1:200)
AntibodyGoat polyclonal anti-rabbit Alexa Fluor 647InvitrogenCat#: A21244 RRID:AB_2535812IHC (1:400)
AntibodyGoat polyclonal anti-HRP Alexa Fluor 488Jackson Immunoresearch LaboratoriesCat#: 123-545-021 RRID:AB_2338965IHC (1:400)
AntibodyGoat polyclonal anti-HRP Cy3Jackson Immunoresearch LaboratoriesCat#: 123-165-021 RRID:AB_2338959IHC (1:400)
Recombinant DNA reagentPlasmid: UAS-2xFlag-Imp α3This paper
Recombinant DNA reagentPlasmid: UAS-V5-Imp β1This paper
Recombinant DNA reagentPlasmid: UAS-CalpA-2xMycThis paper
Chemical compound, drugAll-trans-retinal (ATR) powderSigma-AldrichCat#: R2500 CAS: 116-31-41 mM
Software, algorithmZenZeissRRID:SCR_013672
Software, algorithmImageJNIHRRID:SCR_003070
Software, algorithmImageJ Ratio Plus (plug in)NIHPMID:22051797
Software, algorithmGraphPad PrismGraphPad SoftwareRRID:SCR_002798
Software, algorithmEthoVision XTNoldus Information TechnologyRRID:SCR_000441
Software, algorithmAdobe photoshopAdobeRRID:SCR_014199
OtherFlouro-BoxNeo ScienceFLB-001B

Drosophila melanogaster

Fly stocks used were as follows: w1118, D42-Gal4, TH-Gal4, repo-Gal4, GMR-gal4, UAS-mCD8-RFP, UAS-CD4-tdTom, Itpr ka1091/+ , Itpr sv35 /+ , 20XUAS-ChR2.T159C-HA, UAS-ChR2.S, RyR 16/+ , UAS-NachBac, UAS-R-GECO1-IR1,UAS-R-GECO1.L-IR2, MJDtr-78Q(s), elav-Gal4, UAS-Cam RNAi, UAS-CanA-14F RNAi, UAS-CanB RNAi, UAS-CaMKI RNAi, UAS-Pka-C1 RNAi, UAS-Pkc53E RNAi, UAS-CalpA RNAi (Ch.2), UAS-CalpA RNAi (Ch.3), UAS-CalpB RNAi, UAS-NFAT RNAi, UAS-nej RNAi, UAS-CrebB RNAi, UAS-Imp α1 RNAi, UAS-Imp α2 RNAi, UAS-Imp α3 RNAi, UAS-Imp β1 RNAi, UAS-Imp 7 RNAi, UAS-Imp β11 RNAi, UAS-Tnpo RNAi, UAS-Tnpo-SR RNAi, UAS-Ran RNAi, UAS-Ntf-2 RNAi, UAS-Hrb87F RNAi, UAS-HNPNPC RNAi, UAS-glo RNAi, UAS-Syp RNAi, Df(2R)BSC26, UAS-Luciferase, UAS-poly-PR.PO-100, and UAS-poly-GR.PO-100 were obtained from the Bloomington Drosophila Stock Center (BDSC). UAS-SERCA RNAi, UAS-Itpr RNAi, UAS-RyR RNAi, UAS-CaMKII RNAi, UAS-Imp α3 RNAi, UAS-RanGAP RNAi, UAS-Rcc1 RNAi, UAS-Hrb98DE RNAi, UAS-Hrb27C RNAi, UAS-HNRNPU1 RNAi, UAS-Sm RNAi, UAS-CalpA RNAi and 40D UAS were obtained from the Vienna Drosophila Resource Center (VDRC). UAS-TBPH-Flag-HA was obtained from the Bangalore Fly Resource Center. ppk 1a -Gal4 (Han et al., 2011) was a gift from Yuh Nung Jan (UCSF). UAS-tdTomato P2A GCaMP5G (Daniels et al., 2014) was a gift from Barry Ganetzky (University of Wisconsin-Madison). UAS-3xMyc-RFP-TDP-43 was a gift from Brian D. McCabe (EPFL). UAS-Flag-TDP-43 and UAS-Flag-TDP-43-ΔNLS (Miguel et al., 2011) were gifts from Magalie Lecourtois (University of Rouen). UAS-TDP-43 WT and UAS-TDP-43 G287S were gifts from Aaron Voigt (University Hospital, RWTH Aachen University). UAS-empty (Park et al., 2020) was used as a control in Drosophila eye experiment. All Flies were raised at 27°C and 60% humidity.

Generation of transgenic fly lines

UAS-2xFlag-Imp α3 and UAS-CalpA-2xMyc transgenes were generated by using the LD13917 (Flybase ID: FBcl0163088) and LD22862 (Flybase ID: FBcl0178847) clones obtained from the Drosophila Genomics Resource Center (DGRC), respectively. UAS-V5-Imp β1 transgene was synthesized by Genscript (USA). All these transgenes were subcloned into pACU2 vector, and the transgenic fly lines were generated by Bestgene Inc (USA).

Immunohistochemistry

Larvae (120 hr AEL), pupae (18 hr APF), and adult flies (10d adult and 40d adult) were dissected in 1x Phosphate Buffered Saline (PBS) to obtain fillet or brain samples for immunohistochemical analyses. Obtained samples were fixed in 4% Paraformaldehyde for 20 min, washed in 0.3% PBST (Triton-X100 0.3% in PBS), and blocked in blocking buffer (5% Normal donkey serum or normal goat serum in 0.3% PBST) for 45 min at room temperature. Samples were then incubated with the following primary antibodies for overnight at 4°C: mouse anti-Flag (1F6, Wako 1:400 dilution), rat anti-HA (3F10, Roche 1:200 dilution), rabbit anti-TBPH (1:100 dilution) (Lin et al., 2011), and goat anti-HRP Alexa Fluor 488 (Jackson Immunoresearch Laboratories 1:400 dilution) antibodies. The next day, samples were washed for 10 min (repeated three times) in 0.3% PBST and incubated with the following secondary antibodies for 4 hr: goat anti-mouse Alexa Fluor 647 (Invitrogen 1:400 dilution), goat anti-rat Alexa Fluor 647 (Jackson Immunoresearch Laboratories 1:200 dilution), and goat anti-rabbit Alexa Fluor 647 (Invitrogen 1:400 dilution) antibodies. Samples were then rinsed with 0.3% PBST for 10 min (repeated three times) and mounted with 70% glycerol in phosphate buffered saline (PBG) for imaging.

Microscope image acquisition

All images were acquired using LSM 780, 800 (Zeiss) confocal microscope and Zen (Zeiss) software. All images of samples after immunohistochemistry experiments were taken at 200x and 400x magnifications using 20x and 40x objective lens, respectively. Images of the C4da sensory neurons were obtained from the abdominal segments A5-A6, where anterior is to the left and dorsal is up. Retinal images were obtained using Leica SP5. One-day-old adult fly eyes (left eyes only) were taken at 160x magnification immediately upon dissection.

The ratiometric calcium imaging and analysis

For ratiometric calcium imaging of C4da neurons in larval (120 hr AEL) and pupal (18 hr APF) stages, genetically encoded calcium indicator (GCaMP) and red fluorescent proteins, tdTomato (tdTom), were co-expressed in C4da neurons using P2A system (Daniels et al., 2014). The pseudo-colored images reflecting relative GCaMP level to tdTom level were generated by ImageJ Ratio plus. The mean pixel intensity of GCaMP/tdTom ratio was then measured using ImageJ to determine relative calcium levels, as previously described (Kardash et al., 2011).

Optogenetic stimulation

Larvae expressing channelrhodopsin in C4da neurons were used for optogenetic experiments (behavioral assay and calcium imaging). Larvae were raised under constant darkness at 27°C and 60% humidity on standard media containing 1 mM ATR (Sigma-Aldrich) and collected at 5 days AEL. ATR inhibits the closed state of cyclic nucleotide–gated channels, including channelrhodopsin: ATR is needed to keep the channels open upon stimulation with blue light. Optogenetic stimulation (470 nm) was achieved by Flouro-Box (FLB-001B). Illumination duration and frequency (denoted conditions in Figure 2G and Figure 2—figure supplement 1H) were controlled manually. Both control and experimental groups were placed in the box for illumination, but control groups were prevented from light exposure by covering the vials with aluminum foil. Optogenetic stimulation was performed in a room temperature of 25°C. The validity of optogenetic stimulation and concomitant calcium uptake of C4da neurons was confirmed by both live imaging of calcium indicator RGECO1 and monitoring nociceptive rolling behavior (Kaneko et al., 2017).

Fluorescence recovery after photobleaching (FRAP) experiment and analysis

FRAP experiment was performed by using Zeiss confocal microscope (LSM800). The control and experimental larvae were fed food without and with ATR, respectively. The optogenetic stimulation protocol from Figure 2—figure supplement 1H was applied to the larvae prior to FRAP. Nuclear area was selected as a region of interest (ROI), and cytoplasmic and extracellular areas of the cell were selected as reference and background, respectively, for normalization. Three pre-bleach images were obtained, and then photobleaching of the ROI was performed with 100% power of DPSS laser (561 nm laser) for three iterations. Images for fluorescence recovery were taken every 2 s during 5 min. For quantitative analysis, first, the mean intensity values of background were subtracted from those of ROI in obtained images. Then, these subtracted values were normalized to the fluorescence intensity of the reference. The normalized values were plotted for comparison, and best-fit curve was applied to the graph by using non-linear regression.

Quantitative analysis of dendrites

All images of dendrites were subjected to skeletonization using ImageJ for subsequent analyses of dendritic length and number of branch points. Sholl analysis protocol was adapted from a previous study (Yadav et al., 2019).

Larval motility assay and analysis

The wandering larvae (120 hr AEL) expressing denoted transgenes in motor neurons using the D42-Gal4 driver were used for motility assays. Prior to the assay, individual larva was gently washed in 1x PBS, then briefly placed in a 90 mm Petri dish containing 25 ml of 3% agar. The larva was then placed in another identical Petri dish inside a dark box equipped with indirect lighting. For each genotype, the time for larvae (n ≥ 23 per genotype) to reach the edge of the Petri dish or for up to 60 s was recorded. Recording only started after larva’s first sign of forward movement. These steps were applied to all genotypes tested.

To analyze the motility assay, EthoVision XT (13 version Noldus Information Technology) video tracking system was used (Noldus et al., 2001). Residence probability until reaching the edge of the dish (up to 60 s) was shown as heat map (the color represented the amount of resident time Red: large, Blue: small). Total distance travelled by larvae during 10 s was analyzed.

Western blot

Fly head samples were prepared in a lysis buffer solution (50 mM Tris-buffered saline (Tris-HCL) pH 7.5, 150 mM NaCl, 1% Triton X100) with protease inhibitor cocktail (Thermo Scientific, #87786 1:100). Samples were centrifuged for 15 min at 13,300 rpm and the supernatants were collected into new tubes. Subsequently, their protein amount was measured using Bradford protein assay. Quantified proteins were mixed with a solution containing 9:1 ratio of Laemmli buffer (Bio-Rad, #161–0747) to 2-mercaptoethanol (BIOSESANG, #60-24-2) and were boiled at 95°C for 5 min. Samples were then loaded onto Mini-PROTEAN TGX Stain-Free, 4–15% gel (#BR456-8083, Bio-Rad). Protein transfer to the membrane (PVDF) was followed by an incubation in 5% skim milk diluted in 1% TBST (blocking buffer) for 1 hr at RT. Then they were incubated with primary antibodies overnight at 4°C. The follow primary antibodies were used: Rabbit anti-TBPH (from Dr. C.-K. James Shen) (1:5000), Rat anti-elav (DSHB 7E8A10) (1:200,000). After washing, the membranes were then incubated for 1 hr at RT with the corresponding secondary antibodies: Goat anti-rat IgG-HRP (Santa Cruz, sc-2006) (1:100,000) and Goat anti-rabbit IgG HRP (Santa Cruz, sc-3837) (1:5000). Finally, after washing five times in 1% TBST at RT, the membranes were incubated with ECL solution prior to detection using ChemiDocTM XRS+.

Quantification and statistical analysis

To calculate the cytoplasm-to-nucleus (Cyt/Nuc) ratios of immunostained proteins (TBPH-Flag-HA, endogenous TBPH, 3xMyc-RFP-TDP-43, and 2xFlag-Imp α3), the mean pixel intensities of them in the nucleus and cytoplasm were measured using ImageJ (NIH) and Adobe photoshop (Adobe).

Statistical analysis was performed using GraphPad Prism (GraphPad Software), with Student’s t-test and one-way ANOVA followed by Tukey’s post hoc analysis. In all figures, N.S., *, **, ***, and **** represent p>0.05, p<0.05, p<0.01, p<1.0×10 −3 , and p<1.0×10 −4 respectively. Error bars are standard errors of the mean (SEM).


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