TRPM7 is critical for short-term synaptic depression by regulating synaptic vesicle endocytosis

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Version of Record published: October 14, 2021 (This version)
Accepted Manuscript published: September 27, 2021 (Go to version)
Accepted: September 10, 2021
Preprint posted: January 25, 2021 (Go to version)
Received: January 20, 2021

1. Of interest
Glucagon-like peptide-1 receptor activation stimulates PKA-mediated phosphorylation of Raptor and this contributes to the weight loss effect of liraglutide

Thao DV Le, Dianxin Liu ... Julio E Ayala

Research Article Dec 1, 2023
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Abstract

Transient receptor potential melastatin 7 (TRPM7) contributes to a variety of physiological and pathological processes in many tissues and cells. With a widespread distribution in the nervous system, TRPM7 is involved in animal behaviors and neuronal death induced by ischemia. However, the physiological role of TRPM7 in central nervous system (CNS) neuron remains unclear. Here, we identify endocytic defects in neuroendocrine cells and neurons from TRPM7 knockout (KO) mice, indicating a role of TRPM7 in synaptic vesicle endocytosis. Our experiments further pinpoint the importance of TRPM7 as an ion channel in synaptic vesicle endocytosis. Ca²⁺ imaging detects a defect in presynaptic Ca²⁺ dynamics in TRPM7 KO neuron, suggesting an importance of Ca²⁺ influx via TRPM7 in synaptic vesicle endocytosis. Moreover, the short-term depression is enhanced in both excitatory and inhibitory synaptic transmissions from TRPM7 KO mice. Taken together, our data suggests that Ca²⁺ influx via TRPM7 may be critical for short-term plasticity of synaptic strength by regulating synaptic vesicle endocytosis in neurons.

Introduction

Transient receptor potential melastatin 7 (TRPM7), a ubiquitously expressed member of transient receptor potential (TRP) superfamily (Nadler et al., 2001; Ramsey et al., 2006; Venkatachalam and Montell, 2007), is a cation channel fused with an unique alpha-kinase domain on its C-terminus (Nadler et al., 2001). TRPM7 is able to transport divalent cations, particularly Mg²⁺ and Ca²⁺ (Monteilh-Zoller et al., 2003). TRPM7 is involved in physiological and pathological processes, including embryonic development, organogenesis, and organism survival (Jin et al., 2008; Ryazanova et al., 2010). The cellular functions of TRPM7 have been extensively studied in non-neuronal cells (Abumaria et al., 2018). The kinase domain of TRPM7 is believed to be important for cell growth and proliferation through either eEF2-kinase activation (Perraud et al., 2011) or chromatin remodeling (Krapivinsky et al., 2014). On the other hand, it has been suggested that the ion channel region of TRPM7 may be critical for cell survival by regulating Mg²⁺ homeostasis within cells, such as DT40 cell lines and embryonic stem cells (Ryazanova et al., 2010; Schmitz et al., 2003).

TRPM7 is widely distributed in the nervous system (Grube et al., 2003; Krapivinsky et al., 2006), and it may regulate animal behaviors, such as learning and memory in mouse and rat (Liu et al., 2018) and escape response in zebrafish (Low et al., 2011). A TRPM7 mutation with a defective ion channel property has been identified in human patients with Guamanian amyotrophic lateral sclerosis and Parkinson’s disease (Hermosura et al., 2005), indicating a link between TRPM7 and neurodegenerative diseases. For the cellular physiology of neurons, TRPM7’s kinase domain is reported to be important for normal synaptic density and long-term synaptic plasticity in excitatory neurons from the central nervous system (CNS) (Liu et al., 2018). The ion channel region of TRPM7 may regulate acetylcholine release in sympathetic neurons from the peripheral nervous system (PNS) (Krapivinsky et al., 2006; Montell, 2006), while the role of TRPM7 as an ion channel in CNS neurons has been focused on pathological neuronal death. TRPM7 may mediate neuronal Ca²⁺ overload, resulting in neuronal death under prolonged oxygen/glucose deprivation in neuronal culture (Aarts et al., 2003). Meanwhile, knocking down TRPM7 prevents neuronal death induced by either anoxia (Aarts et al., 2003) or ischemia (Sun et al., 2009). Furthermore, pharmacological blockage of TRPM7 reduces brain damage during neonatal hypoxic injury (Chen et al., 2015). TRPM7 as an ion channel is therefore considered as a potential molecular target for drug design to prevent neuronal death (Bae and Sun, 2013). Yet, the physiological role of the TRPM7’s ion channel region in CNS neurons remains unexplored.

A critical function of TRPM7 in membrane trafficking has been highlighted from non-neuronal cells (Abiria et al., 2017). Along this line, TRPM7 has been implicated in presynaptic vesicle recycling in both neuroendocrine cells (Brauchi et al., 2008) and PNS neurons (Krapivinsky et al., 2006). In PNS sympathetic neurons (Krapivinsky et al., 2006), it is reported that TRPM7 may regulate release of positively charged acetylcholine during vesicle fusion through a proposed ion compensation mechanism (Krapivinsky et al., 2006; Montell, 2006). However, it remains largely unknown what the physiological function of presynaptic TRPM7 is in CNS neurons. By using a combination of biophysical, molecular biology, electrophysiological, and live-cell imaging methods, the present study has identified that Ca²⁺ influx via TRPM7 may be critical for short-term changes of synaptic strength by regulating synaptic vesicle endocytosis in CNS neurons.

Results

Endocytic kinetics is slower in TRPM7 KO chromaffin cells

To understand potential roles of TRPM7 in exocytosis and endocytosis, TRPM7 knockout (KO) chromaffin cells were harvested from conditional KO newborn pups containing both Trpm7^fl/fl (Jin et al., 2008) and Th-Cre (a Cre specific to catecholaminergic cells; Savitt et al., 2005) genes. The depletion of TRPM7 expression in medulla of adrenal glands was further confirmed by RT-PCR (Figure 1—figure supplement 1A). Single vesicle endocytosis was monitored in neuroendocrine chromaffin cells using cell-attached capacitance recordings with a millisecond time resolution, and representative endocytic events from wild-type (WT) or TRPM7 KO cells were shown in Figure 1A. Data from cell-attached capacitance recordings (Figure 1A) showed that, while the number of endocytic events (Figure 1B), the capacitance size of endocytic vesicles (Figure 1C), and the fission-pore conductance (Gp) (Figure 1D) were indistinguishable between WT and KO cells, the fission-pore duration (Figure 1A and E; Figure 1—source data 1) was significantly increased in TRPM7 KO cells, suggesting a critical role of TRPM7 in regulating endocytic kinetics. This finding was further confirmed with data re-analysis, which has a shorter duration cutoff and thus includes additional endocytic events with fission-pore durations in the range of 5–15 ms (Figure 1—figure supplement 2, Figure 1—figure supplement 2—source data 1), indicating that the increase in the fission-pore duration may be independent of durations of endocytic events in TRPM7 KO cells.

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Transient receptor potential melastatin 7 (TRPM7) is important for endocytic fission during endocytosis in chromaffin cells.

(A) Representative endocytic events, as membrane conductance (Re), membrane capacitance (Im), and the fission-pore conductance (Gp), in wild-type (WT) and TRPM7 knockoout (KO) cells. (**B–E**) The number of endocytic events (WT: n = 83 cells; KO: n = 110 cells) (B), the capacitance Cv of endocytic vesicles (C), and the fission-pore Gp (D) were statistically comparable between WT and KO cells; the log-transformed fission-pore duration (E) was significantly increased in KO cells (WT: n = 48 events; KO: n = 50 events). (F) Representative membrane capacitance (Im) and patch membrane current (I_patch) traces at +40 mV in WT and KO cells. Dashed lines in cyan represent linear fitting lines before or after capacitance drop induced by endocytosis. (G) The current-voltage relationship indicating that the endocytosis-associated current drift is reduced at positive membrane potentials in TRPM7 KO cells. The x-axis represents voltages across the patch membrane. n is 12, 9, 13, 12, 13, 8, and 11 for membrane voltages at –40, –30, –20, 0, +20, +30, and +40 mV in WT cells, and 12, 7, 8, 10, 11, 13, and 11 for membrane voltages at –40, –30, –20, 0, +20, +30, and +40 mV in KO cells, respectively. *p < 0.05 and **p < 0.01, unpaired two-tailed Student’s t-test.

Figure 1—source data 1 TRPM7 and single vesicle endocytosis in chromaffin cells.: https://cdn.elifesciences.org/articles/66709/elife-66709-fig1-data1-v2.xlsx
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Since localizations of TRPM7 to synaptic vesicles has been implied from previous studies (Brauchi et al., 2008; Krapivinsky et al., 2006), to examine any potential TRPM7 openings during endocytosis, we performed double (whole-cell/cell-attached) patch recordings, in which the cell-attached patch pipette was utilized to monitor both endocytic events and the ionic current across the patch membrane. With the cell constantly clamped at −60 mV by the whole-cell patch pipette, the patch of membrane captured by the cell-attached pipette was held at different potentials by varying the voltage in the cell-attached pipette. Figure 1F showed at +40 mV an ionic current drift (I_patch) associated with endocytosis in WT cells, and the I-V relationships of the endocytosis-associated ionic current drift in WT cells (Figure 1G, Figure 1—source data 1) seemed to display the outwardly rectifying characteristics of whole-cell TRPM7 currents as reported in the literature (Nadler et al., 2001).

It is important to note that the ionic current drift (Figure 1F) is synchronized with the capacitance decrease, due to single vesicle endocytosis (Yao et al., 2012), which indicates that ion channels contributing to the current drift may be located on the endocytic vesicle. The upward drift in the membrane current at positive potentials may be due to a loss of an outward ionic current on the endocytic vesicle (Figure 1F–G), while the downward drift at negative potentials may be attributed to a loss of an inward ionic current upon the separation of the endocytic vesicle from the plasma membrane (Figure 1G). The endocytosis-associated ionic current drift cannot be attributed to a loss of non-specific leak channels on the endocytic vesicle, for the consequence would be predicted to have a linear current-voltage relationship at both negative and positive potentials. On the contrary, the endocytosis-associated ionic current drift we observed displays a non-linear current-voltage relationship (Figure 1G). It is of note that TRPM7-mediated currents have been missed in previous whole-cell recordings in chromaffin cells, which were typically performed, either at its resting membrane potential of around –80 mV (Smith and Neher, 1997), or in the presence of 0.5–1 mM Mg²⁺ in the intracellular solution (Smith and Neher, 1997; Wu et al., 2021) that substantially inhibits TRPM7 currents (Nadler et al., 2001).

Quantifications showed that ionic current drifts at positive membrane potentials were substantially reduced in KO cells as compared to WT cells (Figure 1F–G), suggesting TRPM7 openings during endocytosis. Meanwhile, the ionic current drift seemed to be associated with single vesicle endocytosis regardless of their capacitance size (Figure 1—figure supplement 3, Figure 1—figure supplement 3—source data 1). Interestingly, there was no such ionic current drift associated with single vesicle exocytosis in WT cells (Figure 1—figure supplement 4A), suggesting openings of TRPM7 channels specifically coincide with endocytosis but not exocytosis. Consistently, the biophysical calculation identified an extra conductance associated with endocytosis rather than exocytosis (Figure 1—figure supplement 4B, C, Figure 1—figure supplement 4—source data 1).

Exocytosis remains unaltered in TRPM7 KO chromaffin cells

To determine whether TRPM7 is involved in exocytosis, catecholamine release from single large dense-core vesicles (LDCVs) was analyzed using carbon fiber amperometry. Secretion induced by 90 mM KCl application for 5 s was recorded by a carbon fiber electrode placed in a direct contact with cells (Figure 2A). The frequency of amperometrical spikes within the first 15 s of stimulation was indistinguishable between WT and KO cells (Figure 2B). Furthermore, no noticeable differences were observed between WT and KO cells for a variety of spike parameters such as peak amplitude (Figure 2C), halfwidth (Figure 2D), the time constant of falling phase (Figure 2E), and quantal size (Figure 2F; Figure 2—source data 1). Amperometry can also provide detailed millisecond information about the initial release through a transient fusion pore (Chow et al., 1992), the foot signal, preceding the subsequent spike of catecholamine release (Figure 2G). Our data showed no difference in foot duration between WT and TRPM7 KO cells (Figure 2H–I; Figure 2—source data 1). Therefore, our amperometrical results indicate that TRPM7 may be dispensable for catecholamine release from LDCVs, which is confirmed by our whole-cell capacitance recordings by detecting exocytosis induced by membrane depolarizations (Figure 2—figure supplement 1, Figure 2—figure supplement 1—source data 1). Our results are consistent with a previous study showing that TRPM7 knockdown or expression of a dominant negative TRPM7 mutant does not affect LDCVs secretion in PC12 cells (Brauchi et al., 2008).

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No change in large dense-core vesicles (LDCVs) exocytosis in transient receptor potential melastatin 7 (TRPM7) knockout (KO) chromaffin cells detected by carbon fiber amperometry.

(A) Amperometrical detection of catecholamine release from wild-type (WT) (upper) and KO (lower) chromaffin cells stimulated with 90 mM KCl for 5 s. Each amperometrical spike represents release from an individual LDCV. (**B–F**) No differences between WT (n = 28) and TRPM7 KO (n = 31) cells in a variety of amperometric parameters, such as spike frequency (B), height (C), halfwidth (D), the exponential time constant of falling phase (E), and quantal size (F). (G) Diagram of the parameters analyzed in amperometric spikes. (**H–I**) Analysis of foot duration. Normalized cumulative distributions of all events in all cells (WT: n = 482; KO: n = 642) (H) and average of median values obtained from individual WT (n = 28) and KO (n = 31) cells (I). Unpaired two-tailed Student’s t-test.

Figure 2—source data 1 No change in exocytosis in KO chromaffin cells.: https://cdn.elifesciences.org/articles/66709/elife-66709-fig2-data1-v2.xlsx
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Synaptic vesicle endocytosis is impaired in neurons from TRPM7 KO animals

To examine potential roles of TRPM7 in synaptic transmission, we cultured TRPM7 KO neurons from conditional KO newborn pups containing both Trpm7^fl/fl (Jin et al., 2008) and Nestin-Cre (a brain specific Cre; Tronche et al., 1999) genes, and the depletion of TRPM7 in cortical neurons was confirmed by RT-PCR (Figure 1—figure supplement 1B). Next, to directly examine synaptic vesicle endocytosis, we performed live-cell imaging of neurons with synaptophysin-pHluorin (sypHy) (Fernández-Alfonso and Ryan, 2004; Granseth et al., 2006; Soykan et al., 2017; Voglmaier et al., 2006) expression driven by the human synapsin promoter (Nathanson et al., 2009; Yaguchi et al., 2013). There was an obvious increase in the time constant of decay in sypHy fluorescent signals in TRPM7 KO neurons at either room temperature (Figure 3A and B, Figure 3—source data 1) or physiological temperature of 34°C (Figure 3—figure supplement 1, Figure 3—figure supplement 1—source data 1), indicative of an endocytic defect of synaptic vesicles. A similar endocytic defect was observed in neurons expressing vGlut1-pHluorin, which revealed a longer time constant for fluorescence decay in KO neurons as well (Figure 3C–D, Figure 3—source data 1). This endocytic defect cannot be the consequence from any changes in vesicle reacidification, since the reacidification rate of newly formed endocytic vesicles, as demonstrated in Figure 3—figure supplement 2, was comparable between WT and KO neurons measured with sypHy (Figure 3E–F, Figure 3—source data 1). Moreover, the endocytic defects we have observed in TRPM7 KO neurons are unlikely due to any potential alterations in endocytic protein levels, as there was no change in expression levels of several key endocytic proteins we tested, such as dynamin 1, AP2, clathrin, and Syt1, in both chromaffin cells (Figure 3—figure supplement 3, Figure 3—figure supplement 3—source data 1) and presynaptic terminals in neurons (Figure 3—figure supplement 4, Figure 3—figure supplement 4—source data 1) from TRPM7 KO animals. Collectively, our data indicates that TRPM7 may be important for synaptic vesicle endocytosis in neurons.

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Synaptic vesicle endocytosis is impaired in transient receptor potential melastatin 7 (TRPM7) knockout (KO) neurons using pHluorin-based optical imaging assays.

(A) Normalized fluorescence changes of synaptophysin-pHluorin (sypHy) signals in wild-type (WT) (n = 139) and KO (n = 127) neurons at 2 mM [Ca²⁺]_e. (B) Bar graph comparing the endocytic time constant from sypHy experiments in A. (C) Normalized fluorescence changes of vGlut1-pHluorin signals in WT (n = 127) and KO (n = 105) neurons at 2 mM [Ca²⁺]_e. (D) Bar graph comparing the endocytic time constant from vGlut1-pHluorin experiments in C. (E) Normalized sypHy fluorescence traces (WT: n = 121; KO: n = 101) with two periods of perfusion with pH 5.5 MES buffer as indicated in the figure (black bar). Newly endocytosed vesicles ‘trapped’ during acidic buffer perfusion starting 5 s after 100 action potentials (APs) at 20 Hz. (F) Bar graph for the reacidification rate of endocytic vesicles, which was obtained by exponential fittings of fluorescence decay, indicates no change in reacidification in KO neurons. n is the total number of boutons pooled from three animals. ***p < 0.001, unpaired two-tailed Student’s t-test.

Figure 3—source data 1 Defects in synaptic vesicle endocytosis in KO neurons.: https://cdn.elifesciences.org/articles/66709/elife-66709-fig3-data1-v2.xlsx
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Conversely, paired whole-cell recordings in pyramidal neurons of cortical culture revealed no change in the amplitude of inhibitory postsynaptic currents (IPSCs) evoked by single presynaptic stimulation with 1 ms depolarization to +30 mV in KO neurons (Figure 3—figure supplement 5A,B, Figure 3—figure supplement 5—source data 1). There was also no obvious change in the paired pulse ratio (Figure 3—figure supplement 5C, Figure 3—figure supplement 3—source data 1), indicating no alteration in release probability in KO neurons. Consistently, vesicular release probability measured in vGlut1-pHluorin experiments was not altered in TRPM7 KO neurons (Figure 3—figure supplement 6, Figure 3—figure supplement 6—source data 1 ). These results suggest that TRPM7 may not be critical for exocytosis of synaptic vesicles in inhibitory synapses with electrically neutral GABA or excitatory synapses with negatively charged glutamate as the predominant neurotransmitters, although it is aware that a previous study implies that TRPM7 may have a post-fusion role by supplying counterions during release of positively charged acetylcholine in PNS sympathetic neurons (Krapivinsky et al., 2006; Montell, 2006).

WT TRPM7 (TRPM7^WT), but not a non-conducting TRPM7 mutant (TRPM7^LCF), rescues endocytic defects in TRPM7 KO chromaffin cells and neurons

To further explore whether the ion channel region of TRPM7 is relevant to TRPM7-dependent synaptic vesicle endocytosis, we generated a non-conducting TRPM7 mutant with a loss of ion channel functions (TRPM7^LCF), by mutating highly conserved amino acids located in the sixth putative transmembrane domain (amino acids 1090–1092, NLL mutated to FAP) (Krapivinsky et al., 2006). As reported previously (Krapivinsky et al., 2006), when compared to TRPM7^WT, there was an almost complete suppression of TRPM7^LCF currents in HEK 293 cells with lentiviral infection (Figure 4A–B, Figure 4—source data 1 and Figure 4—figure supplement 1). Using cell-attached recordings, we examined kinetics of single endocytic events in KO chromaffin cells expressing either TRPM7^WT or TRPM7^LCF (Figure 4C), with similar TRPM7^WT or TRPM7^LCF expression levels as confirmed by immunostaining (Figure 4—figure supplement 2, Figure 4—figure supplement 2—source data 1). While the number of endocytic events (Figure 4D), the capacitance size of endocytic vesicles (Figure 4E), and the fission-pore Gp (Figure 4F) were comparable between these two groups, the fission-pore duration was reduced in KO cells expressing TRPM7^WT as compared to TRPM7^LCF (Figure 4G; Figure 4—source data 1). Consistently, the time constant of sypHy signal decay was shorter in KO neurons expressing TRPM7^WT than TRPM7^LCF from live-cell imaging experiments (Figure 4H–I; Figure 4—source data 1). These findings thus conclude that TRPM7’s ion channel region may be important for synaptic vesicle endocytosis in neurons. Alternatively, since TRPM7’s kinase activity could be potentially altered in the TRPM7^LCF mutant, it could be possible that the endocytic defect we observed in KO neurons might reflect roles of TRPM7’s C-terminal kinase in synaptic vesicle endocytosis. However, this possibility is unlikely, since annexin A1 (Dorovkov et al., 2011), the so far identified substrate of TRPM7 kinase important for endocytosis (Futter and White, 2007), is largely expressed in ependymal and glial cells rather than neurons in the brain (Solito et al., 2008).

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Transient receptor potential melastatin 7 (TRPM7) as an ion channel is critical for endocytosis in both chromaffin cells and neurons.

(A) Averaged whole-cell TRPM7 currents, recorded from HEK 293 cells transduced with lentivirus encoding TRPM7^WT or TRPM7^LCF, in response to voltage ramps from –100 to +100 mV. (B) Quantifications showing TRPM7^LCF currents (n = 16 cells) at +100 mV were almost completely blocked as compared to TRPM7^WT currents (n = 13 cells). (C) Representative endocytic events, as membrane conductance (Re), capacitance (Im), and the fission-pore Gp traces in KO cells expressing either TRPM7^WT (left) or TRPM7^LCF (right). (**D–G**) The number of endocytic events (TRPM7^WT: n = 141 cells; TRPM7^LCF: n = 128 cells) (D), the capacitance Cv of endocytic vesicles (E), and the fission-pore Gp (F) were indistinguishable between these two groups; the log-transformed fission-pore duration (G) was significantly increased in TRPM7^LCF-expressing KO cells (TRPM7^WT: n = 44 events; TRPM7^LCF: n = 53 events). (**H–I**) Normalized fluorescence changes of synaptophysin-pHluorin (sypHy) signals (H) and bar graph (I), comparing the endocytic time constant from sypHy experiments in KO neurons expressing TRPM7^WT (n = 69) or TRPM7^LCF (n = 87) neurons, indicate that endocytic kinetics is slower in TRPM7^LCF-expressing KO neurons. n is the total number of boutons pooled from six animals. *p < 0.05 and ***p < 0.001, unpaired two-tailed Student’s t-test.

Figure 4—source data 1 Lentivirus-mediated TRPM7 versions in HEK 293 cells.: https://cdn.elifesciences.org/articles/66709/elife-66709-fig4-data1-v2.xlsx
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The presynaptic Ca²⁺ increase upon a train of stimulations is reduced in TRPM7 KO neurons

Our sypHy experiments showed that there was no difference in the endocytic time constant of fluorescent signal decay between WT and KO neurons perfused with 0 mM [Ca²⁺]_e right after the cessation of stimulations (Figure 5A–B, Figure 5—source data 1), suggesting a role of TRPM7 in Ca²⁺-dependent rather than Ca²⁺-independent synaptic vesicle endocytosis. Therefore, it is speculated that Ca²⁺ influx via TRPM7 may be critical for synaptic vesicle endocytosis. To test this idea, we compared presynaptic Ca²⁺ signaling between WT and TRPM7 KO neurons by using synaptophysin-GCaMP6f (SyGCaMP6f) fusion protein as a presynaptic Ca²⁺ reporter (de Juan-Sanz et al., 2017). While neurons responded to electrical stimuli with a robust increase in SyGCaMP6f fluorescence (Figure 5C–D), the increase in the SyGCaMP6f signal was substantially reduced in response to a stimulation of 100 action potentials (APs) at 20 Hz in TRPM7 KO neurons (Figure 5C–D, Figure 5—source data 1). The reduction in Ca²⁺ rise in response to stimulation train is unlikely due to any changes in Ca²⁺ influx via voltage-gated Ca²⁺ channels (VGCCs), since the peak of SyGCaMP6f signals induced by single AP (Brockhaus et al., 2019; Kim and Ryan, 2013) was indistinguishable between WT and KO neurons (Figure 5E–F, Figure 5—source data 1). Meanwhile, the resting Ca²⁺ concentration remained unaltered in both chromaffin cells (Figure 5—figure supplement 1, Figure 5—figure supplement 1—source data 1) and presynaptic terminals in neurons from TRPM7 KO animals (Figure 5—figure supplement 2, Figure 5—figure supplement 2—source data 1), and the time constant of Ca²⁺ signal decay right after the cessation of stimulations was similar between WT and TRPM7 KO neurons (Figure 5—figure supplement 3, Figure 5—figure supplement 3—source data 1).

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Transient receptor potential melastatin 7 (TRPM7) is important for presynaptic Ca²⁺ increase upon a train of stimulations.

(A) Normalized fluorescence changes of synaptophysin-pHluorin (sypHy) signal in wild-type (WT) (2 mM: n = 131 buttons; 0 mM: n = 68 buttons) and TRPM7 knockout (KO) (2 mM: n = 116 buttons; 0 mM: n = 67 buttons) neurons at 2 and 0 mM [Ca²⁺]_e after the cessation of stimuli. (B) Bar graph comparing the endocytic time constant from sypHy experiments in A. (C) Sample traces showing normalized changes in SyGCaMP6f fluorescence stimulated with 100 action potentials (APs) at 20 Hz from an individual coverslip of WT (top) (25 regions of interest [ROIs]) or TRPM7 KO (bottom) (25 ROIs) neurons at 2 mM [Ca²⁺]_e. Gray lines are changes for individual ROIs with the black or red line as the averaged response. (D) Bar graph comparing peak values of Ca²⁺ transients from SyGCaMP6f experiments in (C) displays a significant decrease of Ca²⁺ signals in TRPM7 KO neurons (WT: n = 11 coverslips; KO: n = 10 coverslips). (E) Sample traces showing normalized changes in SyGCaMP6f fluorescence induced by single AP from an individual coverslip of WT (top) (36 ROIs) or TRPM7 KO (36 ROIs) neurons at 2 mM [Ca²⁺]_e. Gray lines are changes of individual ROIs with black or red line as the averaged response. (F) Bar graph comparing peak values of Ca²⁺ transients as shown in (E) revealed no change in Ca²⁺ signals induced by a single AP (WT: n = 11 coverslips; KO: n = 10 coverslips). **p < 0.01, unpaired two-tailed student’s t-test; ***p < 0.001, Newman of one-way ANOVA in (B).

Figure 5—source data 1 Decreased presynaptic Ca2+ signals during stimulation train.: https://cdn.elifesciences.org/articles/66709/elife-66709-fig5-data1-v2.xlsx
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Short-term depression of synaptic transmission is enhanced in TRPM7 KO neurons

Our data presented so far indicates that Ca²⁺ influx via TRPM7 may be critical for synaptic vesicle endocytosis. Since synaptic vesicle endocytosis is an essential presynaptic factor for short-term synaptic plasticity (Regehr, 2012; Zucker and Regehr, 2002), we next examined whether short-term synaptic depression is altered in TRPM7 KO neurons.

For inhibitory synaptic transmission, evoked IPSCs in pyramidal neurons were recorded by stimulating presynaptic inhibitory neurons with 250 stimuli at 10 Hz through a presynaptic whole-cell pipette (Figure 6A). With the amplitude of evoked IPSCs normalized to the first response, KO neurons displayed a more profound short-term synaptic depression (Figure 6A–B) and a 56% reduction of the steady-state IPSCs (insert in Figure 6B, Figure 6—source data 1), suggesting an important role of TRPM7 in the short-term synaptic depression of inhibitory synaptic transmission.

Figure 6

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Transient receptor potential melastatin 7 (TRPM7) is critical for presynaptic short-term depression in both inhibitory and excitatory synaptic transmissions.

(**A–B**) Representative traces of inhibitory postsynaptic currents (IPSCs) evoked by 250 stimulations at 10 Hz in wild-type (WT) (top in A) and TRPM7 knockout (KO) (bottom in A) neurons (A) and graph of averaged normalized IPSCs changes (B). Bar graph of the steady-state IPSCs calculated as the average of last 10 evoked responses in WT and TRPM7 KO neurons are shown in insert in B. Number of recordings is 10 for WT and 8 for KO. (C) Representative images showing fluorescence levels of the same dendritic arbor expression iGluSnFR at rest (left) or upon stimulations (right). Scale bar, 5 µm. (**D–E**) Representative traces, from WT (top in D) and TRPM7 KO (bottom in D) neurons (D), and normalized graphs (E) of changes in iGluSnFR fluorescence (ΔF/ΔFmax) evoked by 50 stimulations at 10 Hz. Bar graph of the steady state of iGluSnFR ΔF/ΔFmax calculated as the average of last five evoked responses in WT and TRPM7 KO neurons are shown in insert in (E). Number of coverslips is 21 for WT and 20 for KO. **p < 0.01 and ***p < 0.001, unpaired two-tailed Student’s t-test.

Figure 6—source data 1 Enhanced short-term depression in KO neurons.: https://cdn.elifesciences.org/articles/66709/elife-66709-fig6-data1-v2.xlsx
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For excitatory synaptic transmission, high-speed optical imaging of evoked glutamate release using iGluSnFR (Marvin et al., 2018) was utilized to directly monitor the presynaptic short-term plasticity (Figure 6C). Neurons on coverslips with iGluSnFR expression were challenged with 50 stimuli at 10 Hz to induce short-term depression of evoked glutamate release, fluorescent healthy dendrite arbors were focused and imaged at 50 Hz (Figure 6C–D). Similar to that observed in the synaptic plasticity measured with evoked IPSCs (Figure 6A–B), normalized changes of the iGluSnFR fluorescence during the train also revealed an enhanced short-term depression in TRPM7 KO neurons (Figure 6D–E). The normalized steady-state iGluSnFR signals at the end of the train was reduced by 36% in TRPM7 KO neurons (p < 0.001) (insert in Figure 6E, Figure 6—source data 1), implying a significance of TRPM7 in short-term depression of excitatory synaptic transmission.

Taken together, the enhanced presynaptic short-term depression in both inhibitory and excitatory neurons from TRPM7 KO animals (inserts in Figure 6B and E) corroborated each other. Given the well-established significance of synaptic vesicle endocytosis in short-term depression (Henkel and Betz, 1995; Hermosura et al., 2005), our data may suggest that TRPM7 is critical for presynaptic short-term synaptic depression via its regulation in synaptic vesicle endocytosis.

Discussion

TRPM7 may serve as a Ca²⁺ influx pathway for synaptic vesicle endocytosis

With VGCCs as the well-established Ca²⁺ influx channels for synaptic vesicle exocytosis, the identity of Ca²⁺ influx channels for endocytosis remains uncertain. Since VGCCs have been shown to be important for endocytosis (Midorikawa et al., 2014; Perissinotti et al., 2008; Xue et al., 2012), it is implied that Ca²⁺ for endocytosis at the periactive zone may be derived from diffusion of VGCCs-mediated Ca²⁺ influxes at active zones. On the other hand, exocytosis can be triggered, without activations of VGCCs, by black widow spider venom (Ceccarelli and Hurlbut, 1980; Henkel and Betz, 1995), caffeine (Zefirov et al., 2006), or veratridine (Kuromi and Kidokoro, 2002). Interestingly, extracellular Ca²⁺ is still necessary for synaptic vesicle endocytosis in these paradigms, suggesting distinct Ca²⁺ channels independent of VGCCs for endocytosis. Indeed, a La³⁺-sensitive Ca²⁺ channel is indicated to be required for synaptic vesicle endocytosis in Drosophila (Kuromi et al., 2004).

While it has been proposed that a Ca²⁺-permeable Flower channel may be necessary for synaptic vesicle endocytosis at the neuromuscular junction in Drosophila (Yao et al., 2009), the function of Flower as a Ca²⁺ influx channel in synaptic vesicle endocytosis of mammalian systems remains questionable, since the role of this Ca²⁺-permeable protein in synaptic vesicle endocytosis was not confirmed at the calyx of Held (Xue et al., 2012). Additionally, a recent study showed that the protein itself, but not the ion channel property of Flower, is important for synaptic vesicle endocytosis elicited by moderate stimulations in rat hippocampal neurons (Yao et al., 2017). Furthermore, the time course of Ca²⁺ influx via Flower protein, in the order of 10–60 min (Yao et al., 2009), may be too slow for synaptic vesicle endocytosis.

Our results showed that Ca²⁺ increase during stimulation train is reduced in TRPM7 KO neurons (Figure 5C–D), indicating an importance of TRPM7-mediated Ca²⁺ influx in maintaining Ca²⁺ levels during stimulations. Alternatively, this reduction could be due to slower VGCCs depression and/or slower Ca²⁺ extrusion mechanisms during stimulation train. On the other hand, sypHy experiments indicate that post-stimulus Ca²⁺ influx may be critical for the TRPM7-mediated synaptic vesicle endocytosis (Figure 5A–B). Interestingly, Ca²⁺ signals returned to the resting level quickly, with time constant ~0.6 s, right after the cessation of stimulations, which is similar as described in the literature (Akerboom et al., 2012; Brockhaus et al., 2019; Sgobio et al., 2014; Singh et al., 2018), and there was no alteration in the resting Ca²⁺ level in presynaptic terminals in TRPM7 KO neurons (Figure 5—figure supplement 2). Taken together, it is reasonable to consider that TRPM7 may regulate local Ca²⁺ levels in specific presynaptic regions close to the periactive zone, sites for synaptic vesicle endocytosis (Saheki and De Camilli, 2012), although the global Ca²⁺ level as we have detected within presynaptic terminals is quickly recovered to the resting level after stimulations. However, measurements of local Ca²⁺ signals of a specific region within presynaptic terminals are challenging for techniques currently utilized in the field.

Collectively, our data suggest that TRPM7 channels may serve a Ca²⁺ influx pathway for synaptic vesicle endocytosis in neurons. Along this line, it is of note that Ca²⁺ influx via TRPM7 may be essential for endocytosis of toll-like receptor 4 in the non-neuronal microphage (Schappe et al., 2018). A recent study indicates that TRPC5, another TRP superfamily member, may serve as a Ca²⁺ influx pathway for synaptic vesicle endocytosis in hippocampal neurons (Schwarz et al., 2019). Since TRP channels are widely expressed in the brain (Kunert-Keil et al., 2006), it is speculated that members of the TRP superfamily may serve as general Ca²⁺ channels for synaptic vesicle endocytosis in different regions of the nervous system.

Ca²⁺ influx via TRPM7 may regulate the kinetics rather than the number of single endocytic events

The rate of synaptic vesicle endocytosis is determined by the number of vesicles involved and the kinetics of single endocytic events. Our results, showing slower kinetics but no change in the number of single endocytic events in TRPM7 KO chromaffin cells, indicate that Ca²⁺ influx via TRPM7 may regulate the rate of synaptic vesicle endocytosis by regulating the kinetics of single endocytic events but not the number of vesicles involved. This result is in line with our previous study, which demonstrates an importance of extracellular Ca²⁺ for the kinetics but not the number of single vesicle endocytosis (Yao et al., 2012). Consistently, extracellular Ca²⁺ may modulate endocytic kinetics of single synaptic vesicles in neurons, although no analysis was performed on whether Ca²⁺ affects the number of synaptic vesicles involved (Leitz and Kavalali, 2011).

TRPM7 may be a vesicular protein in presynaptic terminals

By simultaneously monitoring ionic current and membrane capacitance in neuroendocrine chromaffin cells, we have identified an endocytosis-associated ionic current drift, which is substantially reduced in TRPM7 KO cells (Figure 1F–G). Since the endocytosis-associated ionic current drifts rather than displaying a pattern of single channel openings (Figure 1F), we reason that the endocytosis-associated ionic current drift may be attributed to a loss of multiple opening ion channels with small conductance on the endocytic vesicle upon its separation from the plasma membrane. Consistently, our data shows that the endocytosis-associated current drift is synchronized with capacitance drop, which reflects a separation of the endocytic vesicle from the plasma membrane. In fact, TRPM7 is a channel that can be activated by PI(4,5)P₂ and inactivated upon PI(4,5)P₂ hydrolysis (Runnels et al., 2002). Interestingly, PI(4,5)P₂ is concentrated on the plasma membrane but absent from synaptic vesicles, due to PI(4,5)P₂ hydrolysis by phosphatases such as synaptojanin 1 during uncoating after vesicle pinch-off, a late stage of endocytosis (Cremona and De Camilli, 2001). Therefore, the specific openings of TRPM7 during endocytosis may be attributed to the tight coupling between PI(4,5)P₂ metabolism and synaptic vesicle recycling (Cremona and De Camilli, 2001).

Consistent with our data, previous studies have indicated TRPM7 as a vesicular protein in non-neuronal cells (Abiria et al., 2017), neuroendocrine cells (Brauchi et al., 2008), and neurons (Krapivinsky et al., 2006). Immunostaining in PC12 cells, a cell line of neuroendocrine chromaffin cells, has found that TRPM7 is localized to small synaptic-like vesicles (Brauchi et al., 2008). A biochemical analysis of purified rat brain synaptosomes and synaptic vesicles supports this finding by revealing that TRPM7 is concentrated in those regions but absent from postsynaptic densities (Krapivinsky et al., 2006). Further, the same study suggests that TRPM7 may form a molecular complex with vesicular proteins such as SYN1 and synaptotagmin 1 (Krapivinsky et al., 2006). On the other hand, a systematic survey of synaptic vesicle proteins did not detect TRPM7 as a synaptic vesicle protein (Takamori et al., 2006). However, this survey has missed some well-known synaptic vesicle proteins, such as SVOP, the chloride channels CIC3 and CIC7, and ever the putative vesicular neurotransmitter transporters (VMATs and VACHT). Indeed, as claimed by the authors, their proteomics results in an overall coverage of ~80% of all known vesicle membrane proteins (Takamori et al., 2006). Therefore, it is possible that the systematic survey of synaptic vesicle proteins may have failed to detect TRPM7 on synaptic vesicles.

Potential roles of TRPM7 in short-term synaptic depression

Our results indicate an important role of TRPM7 in short-term synaptic depression in both excitatory and inhibitory neurons (Figure 6). It is notable that the onset of short-term depression may be different from electrophysiological recordings and iGluSnFR-based live-cell imaging experiments. As compared to WT neurons, the depression in TRPM7 KO neurons seemed to be apparent after ~15 stimuli in electrophysiological recordings (Figure 6B), while the depression started to show up earlier after ~5 stimuli from iGluSnFR experiments (Figure 6E). This discrepancy could be due to different sensitivities between these two approaches. Alternatively, it may indicate a slight difference in synaptic vesicle recycling between excitatory neurons from our imaging experiments and inhibitory neurons from electrophysiological experiments. Since it is postulated that clearance of endocytic components from release sites is an important factor for short-term depression (Hua et al., 2013; Neher, 2010), the enhanced short-term depression we have observed in TRPM7 KO neurons may indicate a potential role of TRPM7 in release site clearance, in addition to synaptic vesicle endocytosis as we have proposed. It is also reported that the degree of short-term depression can be modulated by exocytic parameters, such as release probability and recruitment of release-ready vesicles (Neher and Sakaba, 2008). While our data showed no difference in the release probability between WT and TRPM7 KO neurons (Figure 3—figure supplement 6), a reduction in presynaptic Ca²⁺ signals during sustained activity in TRPM7 KO neurons (Figure 5C–D) could slow down recruitment of release-ready vesicles, by reducing activations of vesicle priming and Ca²⁺ sensors in fusion (Chapman, 2008; Pang and Südhof, 2010), which could also lead to an enhanced short-term depression.

The physiological role of TRPM7 in CNS neurons

As an ion channel fused with a C-terminal alpha-kinase (Nadler et al., 2001), TRPM7’s ion channel region may be critical for synaptic vesicle endocytosis (the present study) while its kinase domain may be important for normal synaptic density (Liu et al., 2018) in CNS neurons under physiological conditions. It is reported that TRPM7 may be involved in learning and memory in mouse and rat, as knockdown or conditional KO of TRPM7 impairs spatial learning (Liu et al., 2018). The authors indicate that the role of TRPM7 in learning and memory could reflect its importance in regulating synaptic density and long-term synaptic plasticity in excitatory neurons (Liu et al., 2018). However, this interpretation is conflicting with a previous study showing that TRPM7 knockdown does not change evoked synaptic responses, indicative of no change in synaptic density and LTP (long-term potentiation) (Sun et al., 2009). Interestingly, recent studies suggest that TRPC5, which regulates synaptic vesicle endocytosis through a mechanism (Schwarz et al., 2019) similar to TRPM7 as we have observed in the present study, is critical for both short-term synaptic plasticity in excitatory neurons (Schwarz et al., 2019) and learning and memory of animals (Bröker-Lai et al., 2017). Therefore, the role of TRPM7 in short-term synaptic plasticity in excitatory neurons identified here may presumably account for its functions in learning and memory of animals as reported previously (Liu et al., 2018). Notably, physiological roles of TRPM7 in inhibitory neurons remain largely unknown, although TRPM7 is indeed expressed in both excitatory and inhibitory neurons from single cell transcriptome (Huntley et al., 2020). Our study identifies that TRPM7 is also important for short-term synaptic plasticity in inhibitory neurons. Given the increasing significance of inhibitory neuronal circuits in learning and memory (Barron et al., 2017), the reported roles of TRPM7 in learning and memory (Liu et al., 2018) could, alternatively, be attributed to its modulations of inhibitory synaptic transmission.

Implications

In summary, our study indicates that Ca²⁺ influx via TRPM7 is critical for synaptic vesicle endocytosis and consequently short-term synaptic depression, thus implying that TRPM7 may be critical for filtering and gain control of synaptic transmission, as short-term synaptic plasticity is an important factor of synaptic computation (Abbott and Regehr, 2004).

During electrophysiological recordings, TRPM7 can be fully inhibited by intracellular Mg²⁺ (Nadler et al., 2001), which justifies our choice of whole-cell pipette solutions with no Mg²⁺ for electrophysiological recordings in both chromaffin cells and HEK 293 cells. However, roles of influx of ions, such as Ca²⁺, via TRPM7 in neurons have been documented under both physiological (Krapivinsky et al., 2006) and pathological (Aarts et al., 2003; Sun et al., 2009) conditions, indicating that additional cytoplasmic signaling molecules, which remains unknown so far, may be able to facilitate TRPM7 to bypass tonic inhibitions by intracellular Mg²⁺ within intact neurons. In addition, TRPM7 can be blocked by extracellular Mg²⁺ (Nadler et al., 2001), and this Mg²⁺-dependent TRPM7 blockage may be relieved upon membrane depolarizations (Ramsey et al., 2006), a mechanism similar to postsynaptic NMDA receptors (Hunt and Castillo, 2012). Therefore, it is likely that membrane depolarizations will enhance Ca²⁺ influx via TRPM7 and thus accelerate synaptic vesicle endocytosis. Interestingly, many neurons, such as cortical neurons, frequently generate bursts of action potentials in vivo (Baranyi et al., 1993; Gray and McCormick, 1996), and these bursts can occur at a very high frequency, up to 300 Hz, in awake animals as reported (Gray and Viana Di Prisco, 1997). Since synaptic vesicle endocytosis becomes a limiting factor for synaptic transmission in neurons during high-frequency action potential firings (Kawasaki et al., 2000), it is likely that neurons may require an up-regulation of synaptic vesicle endocytosis to maintain synaptic transmission. Therefore, it is speculated that roles of TRPM7 in synaptic vesicle endocytosis will be enhanced during high-frequency firings, which may couple synaptic vesicle endocytosis to neuronal activities.

Recent studies have revealed a tight link between neurodegenerative diseases, particularly Parkinson’s disease, and proteins that participate in synaptic vesicle endocytosis, such as auxilin (Song et al., 2017), synaptojanin 1 (Cao et al., 2017), and endophilin (Trempe et al., 2009). Our evidence for a role of TRPM7 in synaptic vesicle endocytosis suggests that the consequence on synaptic vesicle endocytosis attributed to TRPM7 dysfunctions could be a potential molecular mechanism for TRPM7-related neurodegenerative diseases, such as Guamanian amyotrophic lateral sclerosis and Parkinson’s disease (Hermosura et al., 2005).

Materials and methods

Key resources table

Reagent type (species) or resource	Designation	Source or reference	Identifiers	Additional information
Strain, strain background (Mus musculus)	TRPM7^fl/fl	Jackson LaboratoryStock No: 018784	RRID:IMSR_JAX:018784
Strain, strain background (Mus musculus)	Nestin-Cre	Jackson LaboratoryStock No: 003771	RRID:IMSR_JAX:003771
Strain, strain background (Mus musculus)	TH-Cre	Jackson LaboratoryStock No: 008601	RRID:IMSR_JAX:008601
Cell line (human)	HEK 293FT	Thermo Fisher Scientific	Catalog number: R70007
Chemical compound, drug	CNQX	Tocris Bioscience	Cat# 0190
Chemical compound, drug	AP5	Tocris Bioscience	Cat# 0106
Chemical compound, drug	TTX	Tocris Bioscience	Cat# 1078
Chemical compound, drug	Fura-2 AM	Thermo Fisher Scientific	Cat# F1221
Antibody	Anti-FLAG(Mouse monoclone M2)	Sigma-Aldrich	Cat# F1804RRID:AB_262044	ICC (1:1000)
Antibody	Anti-TH(Rabbit polyclonal)	Abcam	Cat# ab6211RRID:AB_2240393	ICC (1:2000)
Antibody	Anti-synaptophysin (Rabbit polyclonal)	Aviva Systems Biology	Cat# ARP45435_P050 RRID: AB_2048301	ICC (1:1000)
Antibody	Anti-synaptophysin (Mouse monoclonal)	Sysy	Cat# 101011RRID: AB_887824	ICC(1:1000)
Antibody	Anti-AP2(Mouse monoclonal)	abcam	Cat# ab2730RRID: AB_303255	ICC(1:200)
Antibody	Aanti-Syt1(Mouse monoclonal)	Sysy	Cat# 105011RRID: AB_887832	ICC(1:1000)
Antibody	Anti-clathrin(Mouse monoclonal)	Pierce	Cat# MA1-065RRID: AB_2083179	ICC(1:500)
Antibody	Anti-dynamin(Rabbit polyclonal)	Abcam	Cat# ab3456RRID: AB_303818	ICC (1:1000)
Antibody	Anti-mouse 555(Donkey polyclonal)	Thermo Fisher Scientific	Cat# A-31570RRID:AB_2536180	ICC(1:1000)
Antibody	Anti-rabbit 488(Donkey polyclonal)	Thermo Fisher Scientific	Cat# A-21206 RRID:AB_2535792	ICC(1:1000)
Antibody	Anti-mouse 647(Donkey polyclonal)	Thermo Fisher Scientific	Cat# A32787 RRID:AB_2762830	ICC(1:1000)
Antibody	Anti-rabbit 555(Goat polyclonal)	Thermo Fisher Scientific	Cat# A32732 RRID:AB_2633281	ICC(1:1000)
Antibody	Anti-mouse 488(Goat polyclonal)	Thermo Fisher Scientific	Cat# A32723 RRID:AB_2633275	ICC(1:1000)
Antibody	Anti-rabbit 647(Goat polyclonal)	Thermo Fisher Scientific	Cat# A32733TR RRID:AB_2866492	ICC(1:1000)
Recombinant DNA reagent	pCDH-EF1-MCS-T2A-copGFP	System Biosciences	Cat# CD526A-1	A lentiviral vector
Recombinant DNA reagent	CMV::SypHy 4 A	Dr Leon LagnadoPubMed 16982422	Cat# 24478RRID:Addgene_24478	SypHy was subcloned into a pCDH lentiviral vector containing a synapsin 1 promoter in lab
Recombinant DNA reagent	pCDH-SYN1-sypHy	This paper	N/A	Plasmid was packaged into lentivirus, which was used to transduce primary neurons
Recombinant DNA reagent	pCDH-EF1-sypHy-T2A-TRPM7^WT	This paper	N/A	Plasmid was used to transfect TRPM7 KO neurons
Recombinant DNA reagent	pCDH-EF1-sypHy-T2A-TRPM7^LCF	This paper	N/A	Plasmid was used to transfect TRPM7 KO neurons
Recombinant DNA reagent	pCDH-EF1- TRPM7^WT (Flag tag on N-term)	This paper	N/A	Plasmid was packaged into lentivirus, which was then used to infect TRPM7 KO chromaffin cells
Recombinant DNA reagent	pCDH-EF1-TRPM7^LCF (Flag tag on N-term)	This paper	N/A	Plasmid was packaged into lentivirus, which was then used to infect TRPM7 KO chromaffin cells
Recombinant DNA reagent	TRPM7 (Flag tag on N-term)	Dr AM ScharenbergPubMed11385574	Cat# 45482, AddgeneRRID:Addgene_45482	DNA was subcloned into a pCDH lentiviral vector containing a EF1 promoter in lab
Recombinant DNA reagent	vGlut1-pHluorin	Dr V HauckePMID:21808019	N/A	DNA was subcloned into a pCDH lentiviral vector containing a synapsin one promoter in lab
Recombinant DNA reagent	CaMKIIα-iGluSnFR	Dr ER ChapmanPMID:32515733	N/A	S72A variant was used in this paper; DNA was subcloned into a pCDH lentiviral vector containing a synapsin 1 promoter in lab
Recombinant DNA reagent	Syn-GCaMP6f	Dr TA Ryan lab PMID:28162809	N/A	DNA was subcloned into a pCDH lentiviral vector containing a synapsin 1 promoter in lab
Software, algorithm	Igor Pro 8	WaveMetrics	RRID:SCR_000325
Software, algorithm	Excel	Microsoft	RRID:SCR_016137
Software, algorithm	Micro-Manager	ImageJ	RRID:SCR_000415
software, algorithm	FIJI	Fiji.sc	RRID:SCR_002285
Software, algorithm	Adobe Illustrator CS2	Adobe	RRID:SCR_010279
Software, algorithm	Prism	GraphPad	RRID:SCR_002798
Software, algorithm	NIS-Elements	Nikon	RRID:SCR_014329

Generation of TRPM7 conditional KO mice and mouse breeding

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All animal experimental studies were approved by the Animal Care and Use Committee of the University of Illinois at Chicago and conformed to the guidelines of the National Institutes of Health (animal protocol number 19–189). All mating cages were exposed to a 12 hr light/dark cycle with food and water provided ad libitum.

Trpm7^fl/fl (018784, a kind gift of Dr Clapham lab [Jin et al., 2008]), Nestin-Cre (003771, a kind gift from Dr Schütz lab [Tronche et al., 1999]), and Th-Cre (008601, a kind gift from Dr Dawson lab [Savitt et al., 2005]) mice were obtained from Jackson lab. TRPM7^fl/fl mice were crossed with Th-Cre or Nestin-Cre line to produce hemizygous Th-Cre or Nestin-Cre mice, which were heterozygous for the loxP targeted (floxed) Trpm7 gene (Th-Cre/Trpm7^fl/+ or Nestin-Cre/Trpm7^fl/+), respectively. Male Th-Cre/Trpm7^fl/+mice were then backcrossed to female Trpm7^fl/fl mice to generate mice with TRPM7 KO catecholaminergic cells including chromaffin cells (Th-Cre/Trpm^fl/fl). Similarly, Nestin-Cre/Trpm7^fl/+ mice were backcrossed to Trpm7^fl/fl mice to generate brain-specific TRPM7 KO mice (Nestin-Cre/Trpm7^fl/fl). Controls were the littermates carrying a single floxed allele of Trpm7 (Trpm7^fl/+).

Cre transgenes were genotyped using tail biopsy DNA samples with PCR primers 5'-GCGGTCTGGCAGTAAAAACTATC-3' and 5'-GTGAAACAGCATTGCTGTCACTT-3' to detect a 100 bp transgenic fragment. The TRPM7 floxed and WT alleles were detected using the primer set: forward, 5′- TTTCTCCAATTAGCCCTGTAGA-3′; reverse, 5′-CTTGCCATTTTACCCAAATC-3′, the products were 300 bp for the floxed allele and 193 bp for the WT allele. RT-PCR of adrenal medulla and cerebral cortex mRNA were used to confirm the excision of loxP-flanked (floxed) sequences, indicated by a 454 bp cDNA fragment of wild-type TRPM7, and a 225 bp fragment from Cre recombinase-excised, as previously described (Jin et al., 2008).

Cultures of chromaffin cells, neurons, and HEK 293 cells

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Chromaffin cells in culture, prepared from adrenal glands of newborn pups with either sex as previously described (Gong et al., 2005; Yao et al., 2013; Yao et al., 2012), were maintained at 37°C in 5% CO₂ and used within 4 days for electrophysiology or immunostaining. Lentiviral infections with TRPM7^WT or TRPM7^LCF in TRPM7 KO cells were carried out at days in vitro (DIV) 0, and electrophysiological recordings or immunostaining staining was performed 72–96 hr after infection.

Neuronal culture from cortex of newborn pups with either sex was prepared as described previously (Gong and De Camilli, 2008). Neurons in culture were maintained at 37°C in 5% CO₂ and used at DIV 14–16 for electrophysiology. Neurons at DIV 5 were transfected with vGlut1-pHluorin, TRPM7^WT-sypHy, or TRPM7^LCF-sypHy plasmids using Lipofectamine LTX (Thermo Fisher Scientific) or infected with sypHy, SyGCaMP6f, or iGluSnFR lentivirus, and imaging experiments were carried out on neurons at DIV 16–20.

HEK 293 cells were cultured in DMEM supplemented with 10% FBS, 0.1 mM MEM non-essential amino acids, 6 mM L-glutamine, 1 mM MEM sodium pyruvate, 1% Pen-Strep, and 500 µg/ml Geneticin antibiotic. Cells, which typically reached 80% confluence every 3 days, were treated with trypsin-EDTA and passaged with a 1:10 ratio for the general maintenance. Cells at low density were seeded onto 24-well culture plates and infected with lentivirus carrying FLAG-tagged TRPM7^WT or TRPM7^LCF, and monoclonal cell lines were then generated by limiting dilution. After confirming the monoclonal cell lines expressing either TRPM7^WT or TRPM7^LCF by FLAG immunostaining, the monoclonal cell lines were maintained at 37°C in a 5% CO₂ incubator for 48–72 hr before being trypsinized and seeded to PDL-coated coverslip for whole-cell patch clamping recordings.

Cell-attached capacitance recordings and analyses of exocytic fusion-pore and endocytic fission-pore

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Cell-attached capacitance recordings of membrane capacitance and conductance were performed as described previously (Yao et al., 2013; Yao et al., 2012). Fire polished pipettes had a typical resistance of ~2 M in the bath solution. The bath solution contained 130 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 10 mM HEPES-NaOH, and 10 mM glucose; the pH was adjusted to 7.3 with NaOH. The solution in the cell-attached pipette contained 40 mM NaCl, 100 mM TEACl, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, and 10 mM HEPES-NaOH; the pH was adjusted to 7.3 with NaOH. As mentioned previously (Yao et al., 2013; Yao et al., 2012), capacitance steps and fission-pore durations were considered as reliably detected for step sizes > 0.2 fF, and smaller capacitance steps are not included in the analysis. The number of endocytic events per patch detected in the cell-attached recordings is counted as the total number of downward capacitance steps within the first 5 min of recordings (Yao et al., 2013; Yao et al., 2012).

The fusion-pore conductance (Gp) for exocytic events or the fission-pore Gp for endocytic events can be calculated from both Re and Im traces as Gp_(Im+Re) = $\frac{{R e}^{2} + {I m}^{2}}{R e}$ or from Im trace alone as Gp_(Im) = $\frac{ω ∙ C v}{\sqrt{(\frac{ω ∙ C v}{I m}) - 1}}$ , as described (Gong et al., 2007; Yao et al., 2013; Yao et al., 2012). Analysis of fission-pore kinetics were restricted to fission-pores with durations > 15 ms, since shorter events were distorted by the lock-in amplifier low-pass filter (set to 1 ms, 24 dB) (Yao et al., 2013; Yao et al., 2012).

Double (whole-cell/cell-attached) patch recordings in chromaffin cells

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Double (cell-attached/whole-cell) patch recordings were performed to analyze endocytosis-associated current drift, in which the cell-attached patch pipette was utilized to simultaneously monitor endocytic events and the ionic current across the patch membrane. With the cell constantly clamped at –60 mV by the whole-cell patch pipette via an EPC-10 amplifier (HEKA Elektronik, Germany), the patch membrane was held at different voltages by varying the voltage in the cell-attached pipette. The solution in the whole-cell pipette contained 20 mM CsCl, 90 mM KCl, 20 TEACl, 10 mM HEPES-KOH, 2 mM EGTA, 2 mM Na₂ATP, and 0.3 Na₃GTP; the pH was adjusted to 7.3 with KOH with the osmolarity of 290 mmol/kg. The basal Ca²⁺ level in the whole-cell pipette solution was buffered by 1.1 mM CaCl₂ and 2 mM EGTA to give a final free Ca²⁺ level of ~150 nM. All the settings for cell-attached recordings and the solution in the cell-attached pipette were identical to descriptions in Cell-attached capacitance recordings and analyses of exocytic fusion-pore and endocytic fission-pore. Ionic currents from the cell-attached pipette were low-pass filtered at 500 Hz. As illustrated in Figure 1F, the endocytosis-associated current drift was calculated as the difference between linear fittings of a 20–100 ms duration in the I_patch trace before and after the capacitance drop due to endocytosis.

Whole-cell capacitance recordings in chromaffin Cells

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Whole-cell patch recordings, with pipettes of 2–4 MΩ resistance, on chromaffin cells were performed using an EPC-10 amplifier together with PULSE software (HEKA Electronics, Lambrecht/Pfalz, Germany). The external bathing solution contained 140 mM NaCl, 2.8 mM KCl, 10 mM CaCl₂, 1 mM MgCl₂, 10 mM HEPES, and 5 mM glucose (pH 7.25). The pipette solution contained 135 mM Cs-Glu, 2.8 mM KCl, 1 mM MgCl₂, 10 mM HEEPES, 2 mM MgATP, and 0.3 Na₃GTP (pH 7.25). Basal Ca²⁺ concentration in the pipette solution was buffered by a combination of 2 mM CaCl₂ and 5 mM EGTA to give a final free Ca²⁺ concentration of ~200 nM. Capacitance measurements were performed using the software lock-in module of PULSE. A 1 kHz, 40 mV peak-to-peak sinusoid stimulus was applied on a DC potential of –90 mV. The resulting current was processed by using the Lindau–Neher technique (Lindau and Neher, 1988) to give estimates of the equivalent circuit parameters (Cm, Gm, and Gs). The reversal potential of the measured DC current was assumed to be 0 mV. ΔCm values were analyzed with Igor software (RRID: SCR_000325, WaveMetrics, Lake Oswego, OR). Exocytosis was estimated by using four 100 ms depolarizations to 0 mV (Rettig and Neher, 2002). Cm was averaged over a 50 ms pre-depolarization segment to give a baseline value. Our previous study has suggested that the transient gating capacitance artifact was negligible 40 ms after the end of a depolarization (Gong et al., 2005), thus, Cm was averaged over the 40–100 ms interval after depolarizations, and the baseline Cm value was subtracted to obtain the amount of exocytosis. Evoked Ca²⁺ currents were measured under conditions in which most of the K⁺ currents were blocked by intracellular Cs⁺. Tetrodotoxin was not used to block Na⁺ conductance in these experiments because it has been shown to prolong non-secretory capacitance transients in chromaffin cells. Instead, the first 10 ms of evoked inward current was excluded for estimations of Ca²⁺ influx. No correction for leak currents was made and cells with a leak of >150 pS were discarded.

Paired whole-cell recordings of evoked IPSCs in neurons

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Paired whole-cell patch-clamp recordings were performed on DIV 14–16 neurons, continuously perfused with an extracellular solution containing 130 mM NaCl, 3 mM KCl, 10 mM HEPES-NaOH, 2 mM CaCl₂, 1 mM MgCl₂, and 10 mM glucose, pH adjusted to 7.3 with NaOH. Evoked IPSCs were isolated by adding amino-5-phosphonopentanoate (AP5, Tocris) at 50 μM and 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX, Tocris) at 20 μM in the bath solution to block excitatory synaptic transmission. The pipette solution for presynaptic neurons contained 125 mM KCl, 10 mM HEPES, 2 mM EGTA-KOH, 1 mM CaCl₂, 1 mM MgCl₂, 2 mM Na₂ATP, 0.4 mM Na₃GTP, and 5 mM phosphocreatine disodium salt, the pH adjusted to 7.3 with KOH. For postsynaptic neurons, the pipette solution contained 125 mM CsCl, 10 mM HEPES, 2 mM EGTA-CsOH, 1 mM CaCl₂, 1 mM MgCl₂, 2 mM Na₂ATP, 0.4 mM Na₃GTP, 5 mM QX-314, and 5 mM phosphocreatine disodium salt, pH adjusted to 7.3 with CsOH.

IPSCs were recorded at –70 mV on pyramidal neurons using dual whole-cell configurations by evoking a nearby inhibitory neuron with a 1 ms depolarization from –70 to +30 mV. Recordings were acquired with an EPC-10 patch-clamp amplifier for the presynaptic voltage clamping and an EPC-7 plus patch-clamp amplifier for postsynaptic recordings. Data were filtered at 1 kHz and digitized at 5 kHz. Patch pipettes had a typical resistance of 2–3 MΩ. Criteria for inclusion required that Rs remained below 10 MΩ and did not increase by more than 20% during recordings.

Analysis was carried out with Igor software, using custom-written analysis procedures. Peak amplitudes of IPSCs were extracted by subtracting the extrapolated residual current caused by previous stimulation. The amplitude of IPSCs during train stimulations was normalized by dividing the peak of each response to the peak amplitude of the first response.

Whole-cell recordings of TRPM7 currents in HEK 293 cells

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TRPM7 currents were monitored in HEK 293 cells using the whole-cell configuration as previously reported (Aarts et al., 2003). The bath solution contained 130 mM NaCl, 5 mM KCl, 1 mM CaCl₂, 1 mM MgCl₂, 10 mM HEPES-NaOH, and 10 mM glucose, 0.1 mM CdCl₂, 1 µM TTX; the pH was adjusted to 7.3 with NaOH, and the osmolarity was ~310 mmol/kg. The solution in the whole-cell pipette contained 120 mM Cs-MeSO₃, 8 mM NaCl, 2 mM CaCl₂, 10 mM EGTA-CsOH, and 10 mM HEPES; the pH was adjusted to 7.3 with CsOH, and the osmolarity was adjusted to 290 mmol/kg. The voltage ramp protocol holds cells at 0 mV, steps to –100 mV for 40 ms, and then ramps to +100 mV over 500 ms, holding at +100 mV for 40 ms before stepping back to 0 mV, and this protocol is repeated every 5 s during recording. Series resistance was monitored by applying a voltage step of 10 mV. Current signals were filtered at 3 kHz and digitized at 5 kHz. Only recordings with a pipette-membrane seal resistance >2 GΩ were included. Current traces and time courses were analyzed with customized macro for Igor software.

Carbon fiber amperometry

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Chromaffin cells were incubated in the bath solution containing 140 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 10 mM HEPES-NaOH, and 10 mM glucose, pH 7.3 with NaOH. The stimulation solution contained 45 mM NaCl, 90 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 10 mM HEPES-NaOH, and 10 mM glucose. Conventional carbon fiber amperometry for catecholamine detection used 5 µm carbon fibers (ALA Scientific, Farmingdale, NY) as described previously (Gong et al., 2007; Gong et al., 2005; Yao et al., 2012). The freshly cut tip of the carbon fiber electrode was positioned closely against the cell surface to minimize the diffusion distance from release sites, and cells were stimulated with 90 mM KCl solution for 5 s by a pressurized perfusion system (VC3, ALA Scientific) placed ~40 µm away from the cell. The amperometric current, generated by catecholamine oxidation at the exposed tip of the carbon fiber electrode due to stimulation, was measured using an EPC-7 plus amplifier at a holding potential of +700 mV. Amperometric signals were low-pass filtered at 3 kHz and digitized at 5 kHz. Amperometric recordings were collected and then analyzed with a customized macro for Igor software to extract spike information according to the criteria of Chow et al., 1992. The following criteria were set in single-spike analysis: (i) only spikes > 10 pA were considered; (ii) the maximum number of spikes analyzed per cell was set to 100; (iii) foot duration was delimited by the baseline and spike onset as shown previously (Chow et al., 1992), and (iv) spikes with a foot duration of <0.5 ms were excluded for the assays. Data are presented as mean ± SEM of the cell medians. Thus, the numbers used for statistical tests are the number of cells. The number of amperometric spikes was counted as the total number of spikes with an amplitude >10 pA within 15 s after stimulation.

pHluorin-, SyGCaMP6f- and iGluSnFR-based live-cell imaging in neurons

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Neurons on coverslips were continuously perfused at a flow rate of ~1 ml/min with bath solution (130 mM NaCl, 2.8 mM KCl, 2 mM CaCl₂, 1 mM MgCl₂, 10 mM glucose, 10 mM HEPES [pH 7.4]; ~ 310 mOsm). All imaging experiments were performed on an Olympus IX51 microscope with a 60× oil immersion objective at room temperature (25°C ± 1°C), with a GFP-3035B filter cube (472/30 nm excitation, 520/35 nm emission, 495 nm dichroic mirror) (Semrock, Rochester, NY) as the filter set. To elicit responses, APs were delivered by electric field stimulation at 10 V/cm with 1 ms duration via two parallel platinum wires embedded within the perfusion chamber with 7 mm spacing; 20 μM CNQX (Tocris Bioscience) and 50 μM D,L-2-amino-5-phosphonovaleric acid (AP5, Tocris Bioscience) were included in the bath solution to prevent recurrent activities.

For train stimulation experiments using sypHy, vGlut1-pHluorin, or SyGCaMP6f, a 120 W mercury vapor short Arc lamp (X-cite, 120PC, Exfo) was utilized as the light source for excitation. Images were captured by a Photometrics HQ2 coolSNAP CCD camera controlled by μManager software (micro-manager.org) at a 1–2 s interval with 100 or 200 ms exposure time at 25% illumination. To measure reacidification rate of newly formed endocytic vesicles using sypHy as described previously (Atluri and Ryan, 2006), neurons on coverslip were rapidly superfused with a valve controlled pressurized perfusion system (ALA Scientific, Farmingdale, NY). The perfusing buffer was switched between the standard pH 7.3 bath solution described above and pH 5.5 buffer with the HEPES substituted with equi-molar MES. The electrical stimulation was thus time-locked to frame acquisition and buffer changes.

For single AP stimulation using SyGCaMP6f and train stimulation using iGluSnFR, Lambda 421 Optical Beam Combining System (Sutter instrument, Novato, CA) was used as the light source for illumination. To achieve high quantum efficiency, we used the Prime BSI Scientific CMOS (sCMOS) camera (Teledyne Photometrics). With 20 ms exposure time, the illumination via Lambda 421 and data acquisition via camera were controlled by Igor software to achieve a sampling rate of 50 Hz. The light intensity of illumination was set at 10% to minimize photobleaching.

Fura-2 AM-based Ca²⁺ imaging in chromaffin cells and neurons

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Chromaffin cells or neurons in culture were loaded with freshly thawed, cell permeant Fura-2 AM (Thermo Fisher Scientific, Waltham, MA) at the final concentration of 5 µM for 30 min at room temperature (Jiang et al., 2019). Cells or neurons were then gently rinsed three times with the regular extracellular solution of chromaffin cells or neurons to remove the excess dye and recovered for 20 min in fresh bath solution before experiments.

Chromaffin cells or neurons loaded with Fura-2 AM were imaged with a Prime 95B sCMOS camera mounted onto a Nikon Eclipse Ti2 inverted microscope using a 40×/1.30 oil-immersion objective with 1.5× external magnification. Fluorescence emission images (525 nm) of chromaffin cells or neurons were acquired for 10 s continuously in response to alternating 340 and 380 nm excitation light delivered by a Retra Light Engine with 50 ms exposure. Both diodes were co-illuminated and transmitted to the same optical path through a dichroic beam combiner. Presynaptic terminals in neurons were identified by lentivirus-mediated expressions of vGlut1-pHluorin. Imaging acquisition was performed using NIS-elements software (RRIS: SCR_014329).

Immunofluorescence and confocal imaging of chromaffin cells and neurons

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Chromaffin cells or neurons in culture were fixed with 4% paraformaldehyde for 10 min at room temperature. After permeabilization in blocking PBS buffer containing 3% BSA, 5% normal donkey serum, and 0.1% Triton X-100 for 15 min, the fixed cells or neurons were incubated with primary antibodies overnight at 4°C, followed by fluorescent dye-conjugated secondary antibodies at 1:1000 dilution for 1 hr at room temperature.

Confocal fluorescence images of chromaffin cells or neurons were captured using an Olympus Fluoview FV10i laser-scanning confocal microscope with a 60× oil-immersion objective (NA 1.4) and 473, 559, and 635 nm lasers. The pinhole was set at 50 μm for imaging capturing, and the image size was 1024 × 1024. For comparisons, the settings of the laser intensity and sensitivity of each channel are identical during image capture between WT and TRPM7 KO cells for each individual protein we tested. For different protein we tested, however, the laser intensity and sensitivity of each channel were adjusted differently to minimize photobleaching and saturation of fluorophores.

Image analysis

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Images were analyzed in ImageJ (RRID: SCR_000415, http://rsb.info.nih.gov/ij) using custom-written plugins (http://rsb.info.nih.gov/ij/plugins/time-series.html) and Igor software using custom-written procedures. Photobleaching was less than 2% for all images included within analysis and thus was not corrected. To analyze images from sypHy, vGlut1-pHluorin, or SyGCaMP6f experiments, regions of interest (ROIs) of identical size (4 × 4 pixels) were placed in the center of individual synaptic boutons reacting to stimuli, and fluorescence changes were tracked throughout the image stack. In general, 10–20 ROIs were analyzed from each coverslip for sypHy or vGlut1-pHluorin experiments, and 20–40 ROIs for all SyGCaMP6f experiments. For iGluSnFR analysis, ROIs were created as previously reported (Vevea and Chapman, 2020). Briefly, ROIs (>10 pixels), defined by a series of image subtractions and thresholding (Vevea and Chapman, 2020), were used to measure fluorescence changes of image stacks.

Fluorescent intensities of individual ROIs prior to stimuli were averaged as baseline (F0). The fluorescent changes (ΔF) were normalized to the peak of fluorescent increase (ΔFmax) as ΔF/ΔFmax for sypHy, vGlut1-pHluorin, or iGluSnFR experiments and as ∆F/F0 for SyGCaMP6f experiments. Normalized ΔF of individual ROIs were taken as independent replicates (n) for sypHy and vGlut1-pHluorin experiments with the average of normalized ΔF of ROIs for each coverslip as the independent replicates (n) for SyGCaMP6f and iGluSnFR experiments.

For analysis of Fura-2 AM data, each individual chromaffin cell was defined as a round ROI, while line ROIs with two-pixel thickness were drawn across presynaptic terminals identified by vGlut1-pHluorin in neurons. The fluorescence emission intensity ratio corresponding to 340 nm/380 nm excitation was obtained for chromaffin cells and presynaptic terminals in neurons.

For quantifications of expression levels of endocytic proteins from confocal images, maximum intensity Z-series stacks were created in the image analysis program ImageJ (imagej.nih.gov/ij). The whole cell was selected as ROI for chromaffin cells and ROI of presynaptic terminals, identified by synaptophysin immunostaining, were defined as a 4 × 4 pixel area in neurons. The fluorescent intensity of ROI was subtracted by fluorescent intensity of a background ROI with the same size.

DNA constructs

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GFP in the lentiviral vector (pCDH-EF1-MCS-T2A-GFP) was removed and replaced with an artificial DNA fragment containing Pme I and Apa I restriction sites using sense oligo: 5’-CCGGAATGTTTAAACGGGCCCG and antisense oligo: 5’-TCGACGGGCCCGTTTAAACATT-3’. To engineer pCDH-EF1-sypHy, sypHy (#24478 of Addgene, a kind gift from Dr L Lagnado [Granseth et al., 2006]) was inserted into MCS by adding an Xba I site to the 5’ end (primer: 5’-CTTTTCTAGAATGGACGTGGTGAATCA-3’) and a Not I site to the 3’ end (primer: 5’-CAGCGGCCGCCATCTGATTGGAGAAGGAG-3’). The EF1 promoter was then replaced with a human synapsin 1 (SYN1) promoter to produce pCDH-SYN1-sypHy, which was packaged into lentivirus and used for neuronal infections.

SyGCaMP6, a kind gift from Dr Timothy A Ryan’s lab, was generated by adding GCaMP6f to the C-terminus of the mouse sequence of synaptophysin as previously reported (Kim and Ryan, 2013). SyGCaMP6f was first cloned by the standard PCR protocol using Phusion DNA polymerase (forward primer: 5’-ACTCGGATCCCCTCTAGAATGGACGTGGTGAATCAGCTG-3’; reverse primer: 5’- GATTGATATCTCACTTCGCTGTCATCATTTG-3’). The PCR fragment was then enzyme-digested and ligated between the BamHI and EcoRV sites to obtain pCDH-SYN1- SyGCaMP6f by replacing sypHy in the pCDH-SYN1-sypHy vector.

Modified iGluSnFR (Marvin et al., 2018), with the final four amino acid removed and a Golgi export sequence and ER exit motif included, is a kind gift from Dr Edwin Chapman’s lab (Bradberry et al., 2020). iGluSnFR was subcloned and inserted between the BamH1 and Not1 restriction sites in the lentiviral vector (pCDH-CaMKIIα -MCS, a construct we created in lab) to obtain pCDH-CaMKIIα-iGluSnFR (forward primer: 5’- ATAAGGATCCATGGAGACAGACACACTCCTGCTATG-3’, and reverse primer: 5’- CGATGCGGCCGCTTAGAGGGCAACTTCATTTTCATAGC-3’).

To express TRPM7 in KO chromaffin cells with lentivirus, TRPM7 (#45482 of Addgene, a kind gift of Dr A Scharenberg’s lab [Nadler et al., 2001]) was subcloned into a lentiviral vector via Pme I and Apa I restriction sites to generate pCDH-EF1-TRPM7^WT. To engineer TRPM7 mutant with a loss of channel function (TRPM7^LCF, amino acid 1090–1092, NLL > FAP) (Krapivinsky et al., 2006), site-directed mutagenesis was performed using Pfu hotstart DNA polymerase (forward primer: 5’-CAGTATATCATTATGGTTTTCGCTCCTATCGCATTTTTCAATAAT-3’, and reverse primer: 5’-ATTATTGAAAAATGCGATAGGAGCGAAAACCATAAT GATATACTG-3’). Meanwhile, to express TRPM7 in KO neurons, TRPM7^WT or TRPM7^LCF was digested out with Pme I and Apa I restriction sites and ligated into pCDH-EF1-sypHy with the artificial DNA fragment to produce pCDH-EF1-sypHy-T2A-TRPM7.

Lentiviral productions

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Low-passage HEK 293 cells (Thermo Fisher Scientific) were co-transfected with 10 μg lentiviral vector (pCDH-SYN1-sypHy, pCDH-SYN1-SyGCaMP6f, pCDH-CaMKIIα-iGluSnFR, pCDH-EF1-TRPM7^WT, or pCDH-EF1-TRPM7^LCF) along with 7 μg packaging vector psPAX2 and 3 μg envelope vector pMD2.VSVG using the polyethylenimine-mediated transfection method. The supernatant containing lentiviral particles was collected and filtered through 0.45 μm filter to remove cell debris at 48 and 72 hr. Lentiviral particles, concentrated by PEG-it precipitation kits from System Bioscience (Palo Alto, CA), were re-suspended in cold PBS and stored at –80°C.

There is a logarithmic inverse relationship between packaging size and lentiviral titer, that is, the titer drops by one log for every 2 kb of insert (Kumar et al., 2001). Since Trpm7 is a relatively large gene of >5.5 kb, to increase the packaging efficiency, Flag was tagged to the 5’-end of Trpm7 variants for lentivirus, which was utilized to infect KO chromaffin cells. Our experiments showed that the majority of chromaffin cells were infected by anti-FLAG immunostainings (94% ± 1.4% from three independent experiments).

Statistical analysis

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Data were tested for normal distribution and, if necessary, log-transformed for the fission-pore duration in Figures 1E and 4G, and Figure 1—figure supplement 2B to fulfill the criteria of normal distributions for Student’s t-tests. All statistical analyses were performed with Prism (RRID: SCR_002798, GraphPad, San Diego, CA).

No statistical method was used to predetermine sample size, but our samples sizes are similar to those reported in previous studies (Fernández-Alfonso and Ryan, 2004; Granseth et al., 2006; Soykan et al., 2017; Yao et al., 2013; Yao et al., 2012). Data was expressed as mean ± SEM, animals or cells were randomly assigned into control or experimental groups. Statistical analysis was performed with unpaired two-tailed Student’s t-test except paired two-tailed Student’s t-test in Figure 1—figure supplement 4B and Newman of one-way ANOVA in Figure 5B and Figure 4—figure supplement 2B.

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files.

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Decision letter

Nils Brose

Reviewing Editor; Max Planck Institute of Experimental Medicine, Germany
Lu Chen

Senior Editor; Stanford University, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Acceptance summary:

Your paper describes an interesting new role of the ion channel TRPM7 in the regulated release of neurotransmitters. The corresponding data provide important new insights into the mechanisms by which synaptic transmitter release is fine-tuned.

Decision letter after peer review:

Thank you for submitting your article "TRPM7 is critical for short-term synaptic depression by regulating synaptic vesicle endocytosis" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Lu Chen as the Senior Editor. The reviewers have opted to remain anonymous.

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

Essential revisions:

Essential – Not Requiring Additional Experimental Work

1. The TRPM7 KO strategy is based on the deletion of exon 17. The initial description of the KO (Jin et al., 2008) does not clearly show whether there is a possibility of the generation of a truncated variant of the channel or one with a deletion within the sequence upstream of the TMDs. Can this been excluded?

2. For all data in the manuscript, the authors need to show data points, not just bar graphs with mean and SE.

3. The data in Figure 1E showing longer fission pore durations in the KO are interesting (and similarly, the data in Figure 4H on the rescue), but in the legend it becomes clear that only 42 events in each group were considered, which yielded p<0.05. Strikingly, though, the authors measured from no less than 83 and 110 cells with approximately 6 events per cell (Figure 1B), so overall, <10% of the events were included in panel E. Based on the Methods, the authors only considered events with step sizes >0.2 fF and durations >15 ms, citing a low-pass filter set at 1 ms (24 dB). The reviewers understand that the method has limitations (and the authors are clearly competent in the field), but it is a concern to be concluding based on such a small selection of events. With a filter setting at 1 ms, could the authors attempt to visualize the entire distribution of events down to 5 ms, to inspect if the difference between WT and KO is found only for the longest events, or independently of the event duration?

4. In the example of the current flowing through the patch of a WT but not KO cell in Figure 1F, the capacitance change appears to be >10 times in size compared to the example shown in Figure 1A. If this is not a simple typo, an explanation is required as to whether this finding even applies to the smaller events.

5. The authors aimed to study the physiological function of presynaptic TRPM7, and they conclude that TRPM7 might act as a Ca²⁺-influx pathway. However, the pipette-solutions for whole-cell experiments (in chromaffin cells and HEK-cells, where TRPM7-related currents were apparently detected directly, in Figure 1F and Figure 4A) did not contain Mg²⁺ (in chromaffin cells Na-ATP and Na-GTP were used; in HEK-cells no nucleotides or magnesium were included), and TRPM7 is described as a ion channel that is fully inhibited by 2 mM intracellular Mg-ATP (Nadler et al., 2001). Therefore, it is not straight forward to see how Ca²⁺ would permeate the channel under physiological conditions. The authors have data (e.g. CGaMP6) consistent with Ca²⁺-influx, and they show that a mutation (LCF) blocking permeation also blocks function in chromaffin cells, but this is not conclusive evidence that permeation is involved. The authors need to discuss the negative regulation of TRPM7 by Mg-ATP carefully (it seems to not even be mentioned), and whether physiologically relevant conditions are likely to arise where this block would be relieved. They should also discuss their choice of pipette solutions. Finally they should discuss whether their data might be consistent with other scenarios than Ca²⁺-influx through TRPM7.

6. It is unclear why the KO-induced changes in vesicle fusion do not show-up as a slower rise in the SypHy measurements shown in Figure 3. This should be briefly discussed.

7. The explanatory connection that the authors make between the TRPM7-KO effects on calcium signals, endocytosis, and short-term plasticity seems forced. The KO reduces presynaptic calcium signals (likely also with endocytosis during stimulation trains) and increases synaptic depression already very early in the train. There are more parsimonious explanations for the short-term plasticity change: Reduced bulk calcium levels could cause reduced calcium-dependent activation of the vesicle priming machinery or even reduced activation of calcium sensors in fusion (Synaptotagmins). This aspect should be considered in the discussion part of the paper. It may well be that the short-term plasticity defect in the TRPM7 KO neurons is not directly related to the endocytosis defect. Further, it seems hard to imagine that a selective slow-down of endocytosis would become manifest as an appreciable change in exocytosis already after only 10 APs – unless one considers 'site clearing' as proposed by Neher and Sakaba.

8. It is unclear why the 'endocytosis-associated conductance' the authors describe was missed in the many studies employing whole-cell recordings in chromaffin cells. Do the authors assume that detection failed in these cases due to the outward-rectifying character and/or the fact that typical measurements are done at -80 mV? This issue should be discussed.

9. The time scale of changes in endocytosis kinetics in TRPM7 KO chromaffin granules is of no known physiological impact. This should be discussed.

Essential – Requiring Additional Experiments

1. A basic analysis of the morphology of TRPM7 KO chromaffin cells and neurons is needed to complement the present data. Further, it should be tested if the TRPM7 KO affects the expression or localization of proteins that are critically involved in presynapse function, especially in endocytosis and endocytotic fission (e.g. Dynamins).

2. It should be properly verified that WT and mutant versions of TRPM7 are expressed and targeted in the same fashion. Figure S5 does not support the case that the mutant protein is correctly trafficked. It is unclear what the anti-FLAG staining corresponds to in the KO cells infected with a virus with an empty vector. This staining appears to be vesicular, which is also present in the TRPM7-WT infected cells, but not in the LCF infection. A higher resolution approach is required to localize the FLAG staining (confocal might be sufficient). If TRPM7 is no longer targeted correctly when it has the conduction mutation, one cannot conclude that the current through the channel is what is important.

3. Given the ability of the authors to perform very high-end electrophysiological analyses of transmitter secretion from chromaffin cells, the use of KCl^-depolarization to assess TRPM7-KO-induced changes in exocytosis from chromaffin cells appears 'crude'. This type of stimulation is massive and somewhat unphysiological. The key question that remains open is whether exocytosis triggered by more physiological stimuli is affected by the KO (e.g. as assessed by short depolarisation stimuli combined with capacitance measurements). This should be tested in a small KO-vs.WT analysis.

4. Figure 3E-F show reacidification in WT and KO neurons. The description of this experiment is insufficient. The rationale of the experiment is a bit diffuse and it is unclear how the quantification (i.e. on what part of the trace). Moreover, the trace in panel E seems to be inconsistent with the quantification in panel F. In panel E, the reduction in DeltaF in the presence of the second low-pH pulse is steeper in the WT than in the KO. Can this be explained? In essence, these data in their present state are not sufficient to draw unequivocal conclusions regarding reacidification. The authors should include controls with bafilomycin, to understand whether the slopes they measure are actually reporting reacidification.

5. Reacidification kinetics are not resolved, yet are reported as a single parameter of an exponential fit. Furthermore, when the acid was removed, the KO cells should have had higher fluorescence, since it is decaying more slowly (e.g. difference WT vs. KO at the 80 s time point). The acid quenching experiment showed no difference. Given that the authors are not resolving an exponential decay of the acid-quenched surface signal, it would be more appropriate to simply report what fraction of the signal early on after the end of the stimulus – where there is a real difference in the amount of the signal that has decayed (e.g. at the 60 s time point) – can be quenched by acid perfusion. If it is the same fraction for WT and KO, the differences are not explained by the build-up of an alkaline pool, which is really what the authors want to assess here.

6. It is important to know if the phenomena described in the present study occur under physiological conditions (i.e. 37{degree sign}C, ~1.25 mM Ca). The temperature in particular is important as cooling can substantially distort the relevance of different molecular processes. The reviewers do not expect a repetition of all experiments at 37{degree sign}C. The easiest might be an experiment on pHluorin assessment of vesicle recycling in neurons, showing that at 37{degree sign}C there is a measurable impact on the pHluorin recovery kinetics due to the absence or presence of external Ca²⁺.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "TRPM7 is critical for short-term synaptic depression by regulating synaptic vesicle endocytosis" for further consideration by eLife. Your revised article has been reviewed by 3 peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Lu Chen as the Senior Editor.

Your manuscript has been very substantially improved. However, the reviewers have identified three issues that need to be resolved before the paper can be published in eLife:

1. The new confocal images added to Figure 3 (supplement 3, panels A-D; supplement 4, panels A, C, E, G) and to Figure 4 (supplement 2, panel C) are of very poor quality (very low resolution, blurred, pixelated) and do not look like confocal images (rather like ones that have been improperly digitally compressed). In fact, the new panel C in Figure 4, supplement 2 appears to be of even less resolution than the original images in panel A. In essence, the problematic panels indicated above are not of publication quality and need to be redone.

2. The description of confocal microscopy in the methods part is insufficient and requires more detail (i.e. numerical aperture and type of objective, light source, pinhole setting, etc.).

3. Regarding Figure 5, supplement 1, it seems that the 340 nm images in panel A are saturated. If so, it would be impossible to do a ratio analysis as shown in panel B. This issue needs to be explained and resolved.

https://doi.org/10.7554/eLife.66709.sa1

Author response

Essential revisions:

Essential – Not Requiring Additional Experimental Work

1. The TRPM7 KO strategy is based on the deletion of exon 17. The initial description of the KO (Jin et al., 2008) does not clearly show whether there is a possibility of the generation of a truncated variant of the channel or one with a deletion within the sequence upstream of the TMDs. Can this been excluded?

We appreciate reviewer’s comment. As described in supplementary text in the paper by Jin et al. (Jin et al., 2008), the Cre-mediated deletion of exon 17, which encodes the protein sequence preceding the first transmembrane segment of TRPM7, results in a frame shift that disrupts the translation of the downstream sequence, thus generating TRPM7 with only N-terminal lacking all 7 transmembrane domains and the c-terminal tail. Therefore, the possibility of the generation of a truncated variant of the channel can be excluded.

2. For all data in the manuscript, the authors need to show data points, not just bar graphs with mean and SE.

Following the reviewer’s comment, all data in the figures are presented as mean±SE with all original data points except for panels Figure 1G and Figure 3—figure supplement 5C. And the original data points for these two panels are provided in the related source data files in the revised manuscript.

3. The data in Figure 1E showing longer fission pore durations in the KO are interesting (and similarly, the data in Figure 4H on the rescue), but in the legend it becomes clear that only 42 events in each group were considered, which yielded p<0.05. Strikingly, though, the authors measured from no less than 83 and 110 cells with approximately 6 events per cell (Figure 1B), so overall, <10% of the events were included in panel E. Based on the Methods, the authors only considered events with step sizes >0.2 fF and durations >15 ms, citing a low-pass filter set at 1 ms (24 dB). The reviewers understand that the method has limitations (and the authors are clearly competent in the field), but it is a concern to be concluding based on such a small selection of events. With a filter setting at 1 ms, could the authors attempt to visualize the entire distribution of events down to 5 ms, to inspect if the difference between WT and KO is found only for the longest events, or independently of the event duration?

To answer the reviewer’s question, we have performed a re-analysis of this data set. Results from this re-analysis showed that the percentage of endocytic events in different duration ranges (<5 ms; between 5 ms and 15 ms; >15 ms) was comparable between WT and KO cells. Consistent with an increase in the fission-pore duration in KO cells with inclusion of >15 ms events as shown in Figure 1E in the initial analysis, our re-analysis also showed an increase in the fission-pore duration in TRPM7 KO cells with additional inclusions of >5 ms but <15ms events, which is presented now in Figure 1—figure supplement 2 in the revised manuscript. It is thus likely that the difference in the fission-pore duration between WT and KO cells is independent of event duration. In addition, comparisons between WT and KO cells in the distribution of endocytic events with their fission-pore duration >5 ms or >15 ms are presented in this Figure 1—figure supplement 2 (>15 ms: panel C; >5 ms: panel D). We thank the reviewer for this comment, which prompt us to re-analyze this set of data, making our conclusion on the increase in the fission-pore duration in TRPM7 KO cells more convincing.

4. In the example of the current flowing through the patch of a WT but not KO cell in Figure 1F, the capacitance change appears to be >10 times in size compared to the example shown in Figure 1A. If this is not a simple typo, an explanation is required as to whether this finding even applies to the smaller events.

We would like to clarify that this is not a typo. It is reasonable from our observations that endocytic events with bigger size are typically associated with bigger ionic current drifts. For demonstration purpose, an endocytic event with relatively big size was, therefore, chosen in Figure 1F. To answer the reviewer’s question of whether the ionic current flowing through the patch applies to smaller events, we have compared ionic current drifts with different capacitance sizes (such as < 1 fF vs > 1 fF). To make sure that each group has enough endocytic events for this comparison, we grouped all endocytic events only at positive membrane potentials by neglecting negative membrane potential due to the outwardly rectifying properties of this current drift as shown in Figure 1G. The ionic current drifts were also divided by the voltage across the patch membrane to minimize the impact from the difference in patch membrane voltages. Our results showed similar normalized amplitudes of this endocytosis-associated current drift between relatively smaller (< 1 fF) and bigger (> 1 fF) endocytic events, suggesting that this finding may apply to all endocytic events regardless of their size. This result is added as Figure 1—figure supplement 3 in the revised manuscript.

5. The authors aimed to study the physiological function of presynaptic TRPM7, and they conclude that TRPM7 might act as a Ca²⁺-influx pathway. However, the pipette-solutions for whole-cell experiments (in chromaffin cells and HEK-cells, where TRPM7-related currents were apparently detected directly, in Figure 1F and Figure 4A) did not contain Mg²⁺ (in chromaffin cells Na-ATP and Na-GTP were used; in HEK-cells no nucleotides or magnesium were included), and TRPM7 is described as a ion channel that is fully inhibited by 2 mM intracellular Mg-ATP (Nadler et al., 2001). Therefore, it is not straight forward to see how Ca²⁺ would permeate the channel under physiological conditions. The authors have data (e.g. CGaMP6) consistent with Ca²⁺-influx, and they show that a mutation (LCF) blocking permeation also blocks function in chromaffin cells, but this is not conclusive evidence that permeation is involved. The authors need to discuss the negative regulation of TRPM7 by Mg-ATP carefully (it seems to not even be mentioned), and whether physiologically relevant conditions are likely to arise where this block would be relieved. They should also discuss their choice of pipette solutions. Finally they should discuss whether their data might be consistent with other scenarios than Ca²⁺-influx through TRPM7.

As pointed out by the reviewer, during electrophysiological recordings, TRPM7 can be fully inhibited by intracellular Mg²⁺ (Nadler et al., 2001), which justifies our choice of whole-cell pipette solutions with no Mg²⁺ for electrophysiological recordings in both chromaffin cells and HEK 293 cells. However, roles of influx of ions, including Ca²⁺, via TRPM7 in neurons has been documented under both physiological (Krapivinsky et al., 2006) and pathological (Aarts et al., 2003; Sun et al., 2009) conditions, indicating that additional cytoplasmic signaling molecules, which remains unknown so far, may be able to facilitate TRPM7 to bypass the tonic inhibitions from intracellular Mg²⁺ within intact neurons. In addition, TRPM7may be blocked by extracellular Mg²⁺ (Nadler et al., 2001), and this Mg²⁺-dependent blockage of TRPM7 can be relieved upon membrane depolarizations (Kerschbaum et al., 2003; Li et al., 2007), a mechanism similar to postsynaptic NMDA receptors (Hunt and Castillo, 2012). Therefore, it is likely that membrane depolarizations will enhance Ca²⁺ influx via TRPM7 and thus accelerate synaptic vesicle endocytosis. Interestingly, many neurons, such as cortical neurons, frequently generate bursts of action potentials in vivo (Baranyi et al., 1993; Gray and McCormick, 1996), and these bursts can occur at a very high frequency, up to 300 Hz, in awake animals as reported (Gray and Viana Di Prisco, 1997). Since synaptic vesicle endocytosis becomes a limiting factor for synaptic transmission in neurons during high frequency action potential firings (Kawasaki et al., 2000), it is likely that neurons may require an up-regulation of synaptic vesicle endocytosis to maintain synaptic transmission. Therefore, it is speculated that roles of TRPM7 in synaptic vesicle endocytosis will be enhanced during high frequency firings, which may couple synaptic vesicle endocytosis to neuronal activities. This discussion has been added to “implications” of Discussion section in the revised manuscript.

Following the reviewer’s comment regarding whether our data might be consistent with other scenarios that Ca²⁺ influx through TRPM7, we have also discussed the possibility of TRPM7’s kinase domain in synaptic vesicle endocytosis: While our data, by showing the non-conducting TRPM7^LCF mutant fails to rescue endocytic defects in both TRPM7 KO chromaffin cells and neurons, indicates TRPM7 as an ion channel in synaptic vesicle endocytosis. Alternatively, it could be possible that this might reflect roles of TRPM7’s C-terminal kinase in synaptic vesicle endocytosis, since TRPM7’s kinase activity could be potentially altered in the non-conducting TRPM7 mutant. However, this possibility is less likely, since annexin A1 (Dorovkov et al., 2011), the so far identified substrate of TRPM7 kinase important for vesicle endocytosis (Futter and White, 2007), is largely expressed in ependymal and glial cells rather than neurons in the brain (Solito et al., 2008).

This discussion has been added in the revised manuscript. We appreciate the reviewer’s comments, which promote us to discuss the physiological relevance of Mg²⁺-mediated inhibition on TRPM7 in synaptic vesicle endocytosis.

6. It is unclear why the KO-induced changes in vesicle fusion do not show-up as a slower rise in the SypHy measurements shown in Figure 3. This should be briefly discussed.

As presented in Figure 6A-B in inhibitory synapses and in Figure 6C-E in excitatory neurons, there is an enhanced short-term synaptic depression upon repetitive stimulations in TRPM7 KO neurons. However, as pointed out by the reviewer, this KO-induced enhanced synaptic depression does not show up as a slower increase in the SypHy measurement as shown in Figure 3A and C, which, we believe, may be due to a couple of reasons: (1) in order to estimate synaptic vesicle endocytosis, the traces presented in Figure 3A and C is normalized to the peak of fluorescent signals upon sustained stimulation, which can cover up any potential changes in pHluorin signal increase during stimulations in KO neurons; and (2) as reported previously, there is substantial amount of synaptic vesicle endocytosis during sustained stimulations and the fluorescent signal increase during stimulations is not a good representation of synaptic vesicle exocytosis but rather represents the net balance of exocytosis an endocytosis (Sankaranarayanan and Ryan, 2001).

7. The explanatory connection that the authors make between the TRPM7-KO effects on calcium signals, endocytosis, and short-term plasticity seems forced. The KO reduces presynaptic calcium signals (likely also with endocytosis during stimulation trains) and increases synaptic depression already very early in the train. There are more parsimonious explanations for the short-term plasticity change: Reduced bulk calcium levels could cause reduced calcium-dependent activation of the vesicle priming machinery or even reduced activation of calcium sensors in fusion (Synaptotagmins). This aspect should be considered in the discussion part of the paper. It may well be that the short-term plasticity defect in the TRPM7 KO neurons is not directly related to the endocytosis defect. Further, it seems hard to imagine that a selective slow-down of endocytosis would become manifest as an appreciable change in exocytosis already after only 10 APs – unless one considers 'site clearing' as proposed by Neher and Sakaba.

We agree with the reviewer that an enhanced short-term depression in KO neurons may possibly be due to alterations in exocytosis and release site clearance, in addition to synaptic vesicle endocytosis as we have proposed. As pointed by the reviewer, since it is postulated that clearance of endocytic components from release sites is an important factor for shortterm depression (Hua et al., 2013; Neher, 2010), the enhanced short-term depression we have observed in TRPM7 KO neurons may indicate a potential role of TRPM7 in release site clearance, in addition to synaptic vesicle endocytosis are we have proposed. It is also shown that the degree of short-term depression can be modulated by exocytic parameters, such as recruitments of release-ready vesicles (Neher and Sakaba, 2008). A reduction in presynaptic Ca²⁺ signals during sustained activity in TRPM7 KO neurons (Figure 5C-D) could slow down recruitments of release-ready vesicles, by reducing activations of vesicle priming and/or Ca²⁺ sensors in fusion (Chapman, 2008; Pang and Sudhof, 2010), which could also lead to an enhanced short-term depression. Following the reviewer’s comments, this has been added as part of “potential roles of TRPM7 in short-term synaptic depression” in the Discussion section in the revised manuscript.

8. It is unclear why the 'endocytosis-associated conductance' the authors describe was missed in the many studies employing whole-cell recordings in chromaffin cells. Do the authors assume that detection failed in these cases due to the outward-rectifying character and/or the fact that typical measurements are done at -80 mV? This issue should be discussed.

We agree with the reviewer on this comment. Previous whole-cell recordings in chromaffin cells are typically conducted at its resting membrane potential of around -80 mV (Smith and Neher, 1997), a condition that may have missed the detection of TRPM7 currents due to its outward-rectifying properties (Nadler et al., 2001). Additionally, most of previous whole-cell recordings in chromaffin cells includes 0.5-1 mM Mg²⁺ in the intracellular solution, which substantially inhibits TRPM7 currents (Nadler et al., 2001). This is another big reason for the missed detection of TRPM7 currents by previous studies in chromaffin cells. This discussion has been added in the revised manuscript.

9. The time scale of changes in endocytosis kinetics in TRPM7 KO chromaffin granules is of no known physiological impact. This should be discussed.

Vesicle fission is a crucial step during endocytosis (Kaksonen and Roux, 2018). The importance of vesicle fission in endocytosis can be reflected by significant roles of the GTPase dynamin, a key molecule essential for vesicle fission (Antonny et al., 2016), in synaptic vesicle endocytosis (Ferguson et al., 2007; Newton et al., 2006). Our previous work using cell-attached capacitance recordings, the same approach as the present study, has shown that inhibition of dynamin GTPase slows down endocytic kinetics of single vesicles in chromaffin cells at the millisecond time scale. Therefore, our data shows an increase in the endocytic kinetics of single vesicle in KO chromaffin cells, indicating a potential role of TRPM7 in vesicle fission during endocytosis. The physiological significance of TRPM7 in vesicle fission during endocytosis is reiterated by a slower rate of synaptic vesicle endocytosis in TRPM7 KO neurons. The endocytic rate of synaptic vesicles is decided by the number of endocytic vesicles and the kinetics of single endocytic events. Our results, showing slower kinetics (Figure 1A and E) but no change in the number of single endocytic events (Figure 1B) in TRPM7 KO chromaffin cells, indicate that TRPM7 may regulate the rate of synaptic vesicle endocytosis by regulating the kinetics of single endocytic events but not the number of endocytic vesicles. Following the reviewer’s comment, this discussion has been added to the Discussion section in the revised manuscript.

Essential – Requiring Additional Experiments

1. A basic analysis of the morphology of TRPM7 KO chromaffin cells and neurons is needed to complement the present data. Further, it should be tested if the TRPM7 KO affects the expression or localization of proteins that are critically involved in presynapse function, especially in endocytosis and endocytotic fission (e.g. Dynamins).

Following the reviewer’s comments, we have analyzed, using immunostaining, expression levels of endocytic proteins, such as dynamin 1, AP2, clathrin and synaptotagmin 1, in both chromaffin cells and presynaptic terminals from TRPM7 KO animals. Chromaffin cells were identified with immunostaining of tyrosine hydroxylase (TH) and presynaptic terminals were detected with a synaptophysin antibody, and the fluorescent signals of individual endocytic proteins were normalized to TH signals in chromaffin cells and synaptophysin signals in presynaptic terminals in neurons to minimize variations from experiments to experiments. This analysis showed no alterations in expression levels of all these 4 endocytic proteins in both chromaffin cells and presynaptic terminals from TRPM7 KO neurons, suggesting that the endocytic defects we have observed is unlikely due to any changes in these key endocytic proteins. These additional data have been added as the Figure 3—figure supplement 3 for chromaffin cells and the Figure 1—figure supplement 4 for presynaptic terminals in neurons in the revised manuscript.

2. It should be properly verified that WT and mutant versions of TRPM7 are expressed and targeted in the same fashion. Figure S5 does not support the case that the mutant protein is correctly trafficked. It is unclear what the anti-FLAG staining corresponds to in the KO cells infected with a virus with an empty vector. This staining appears to be vesicular, which is also present in the TRPM7-WT infected cells, but not in the LCF infection. A higher resolution approach is required to localize the FLAG staining (confocal might be sufficient). If TRPM7 is no longer targeted correctly when it has the conduction mutation, one cannot conclude that the current through the channel is what is important.

As shown in the original supplementary Figure 5A (now Figure 4—figure supplement 2A in the revised manuscript), since KO cells in the mock group are infected with a lentivirus with an empty vector, the anti-FLAG signals detected in this group is likely non-specific staining. We agree with the reviewer that a higher resolution approach may be better to localize the FLAG staining. Following the reviewer’s comment, we have taken confocal images of KO chromaffin cells infected with either WT or mutant TRPM7, showing similar distributions of these 2 TRPM7 versions within KO cells. These confocal images are now added as panel C in this Figure 4—figure supplement 2 in the revised manuscript.

3. Given the ability of the authors to perform very high-end electrophysiological analyses of transmitter secretion from chromaffin cells, the use of KCl^-depolarization to assess TRPM7-KO-induced changes in exocytosis from chromaffin cells appears 'crude'. This type of stimulation is massive and somewhat unphysiological. The key question that remains open is whether exocytosis triggered by more physiological stimuli is affected by the KO (e.g. as assessed by short depolarisation stimuli combined with capacitance measurements). This should be tested in a small KO-vs.WT analysis.

Following the reviewer’s comment, we have compared exocytosis induced by membrane depolarization between WT and TRPM7 KO chromaffin cells using whole-cell capacitance measurements. Our results showed no difference in exocytosis induced by membrane depolarization, suggesting no change in exocytosis in TRPM7 KO chromaffin cells. This result is consistent with our amperometrical data using KCl^-depolarization as shown in Figure 2, and the data from whole-cell capacitance recordings is now added as the Figure 2—figure supplement 1 in the revised manuscript.

4. Figure 3E-F show reacidification in WT and KO neurons. The description of this experiment is insufficient. The rationale of the experiment is a bit diffuse and it is unclear how the quantification (i.e. on what part of the trace). Moreover, the trace in panel E seems to be inconsistent with the quantification in panel F. In panel E, the reduction in DeltaF in the presence of the second low-pH pulse is steeper in the WT than in the KO. Can this be expllained? In essence, these data in their present state are not sufficient to draw unequivocal conclusions regarding reacidification. The authors should include controls with bafilomycin, to understand whether the slopes they measure are actually reporting reacidification.

As pointed out by the reviewer, our data on vesicle reacidification seems confusing, which, we think, could be due to low signal-to-noise ratio of some of our original data on individual synaptic terminals. Therefore, we have performed additional experiments between WT and TRPM7 KO neurons. We believe the data from these new experiments has excellent signal-to noise ratio for individual synaptic terminals, which is evident from the example shown in Figure 3—figure supplement 2. We believe the high quality of new data set will allow us to draw an unequivocal conclusion on roles of TRPM7 in vesicle reacidification. Results from these new experiments show no difference in the averaged trace (Figure 3E in the revised manuscript) and time constant of vesicle acidification (Figure 3F in the revised manuscript) between WT and TRPM7 KO neurons. The revised manuscript has been updated with results from these new data.

Following the reviewer’s comment, we also performed additional experiments with bafilomycin to verify whether the decay we measured are reporting vesicle reacidification. Our data shows that the decay can be blocked by applications of bafilomycin that inhibits vesicular proton pump, demonstrating that the decay we measured does represent vesicle reacidification. A sample trace of an individual terminal in the presence of bafilomycin is presented in the Figure 3—figure supplement 2.

In response to the reviewer’s concern of “insufficient description on vesicle reacidification”. We have now added, in the legends of this Figure 3—figure supplement 2, the following description on our experiments on vesicle reacidification in much more details:

To measure reacidification rate of newly formed endocytic vesicles using sypHy as described by Atluri and Ryan (J. Neurosci., 2006), neurons infected with lentivirus encoding sypHy were rapidly perfused with a valve controlled pressurized local perfusion system (ALA Scientific, Farmingdale, NY). Surface fluorescence of pHluorin is quenched by the first application of acidic buffer (pH 5.5 buffer with the HEPES substituted with equi-molar MES) for 10s to establish the baseline, and the quenched fluorescence is recovered by washing off the acidic solution. To isolate pHluorin signals from newly endocytosed synaptic vesicles, acidic buffer, indicated by the 2^nd bar in the trace, is applied with a 20s duration 5s after the cessation of stimulation train. Newly endocytosed alkaline pool of pHluorin is detached from the plasma membrane and thus resistant to quenching by the applied acidic buffer, and the decay of this portion of fluorescence to the baseline represents acidification of newly endocytosed vesicles. The decay of fluorescent signals, which can be blocked by bafilomycin A1, thus likely reflects reacidification kinetics of newly endocytic vesicles.

Reacidification rate of newly endocytosed vesicles is estimated by an exponential fit, as indicated by the red dashed line, of this fluorescent signal decay from the first to the last point during this 20s duration of acidic buffer application.

We believe these descriptions, in combined with the traces added in the Figure 3—figure supplement 2, will be sufficient and convincing for our experiments on vesicle reacidification.

5. Reacidification kinetics are not resolved, yet are reported as a single parameter of an exponential fit. Furthermore, when the acid was removed, the KO cells should have had higher fluorescence, since it is decaying more slowly (e.g. difference WT vs. KO at the 80 s time point). The acid quenching experiment showed no difference. Given that the authors are not resolving an exponential decay of the acid-quenched surface signal, it would be more appropriate to simply report what fraction of the signal early on after the end of the stimulus – where there is a real difference in the amount of the signal that has decayed (e.g. at the 60 s time point) – can be quenched by acid perfusion. If it is the same fraction for WT and KO, the differences are not explained by the build-up of an alkaline pool, which is really what the authors want to assess here.

As mentioned in our response to reviewer’s previous comment on vesicle reacidification, the data from our new experiment, which has an excellent signal-to-noise ratio as shown in the Figure 3—figure supplement 2, clearly shows that the reacidification kinetics can be reported as a single exponential fit in our hands, which is consistent with previous reports (Atluri and Ryan, 2006; Egashira et al., 2015). Additionally, our data showed that, upon the removal of acidic buffer at the 80 s time point, presynaptic terminals from KO neurons had a higher fluorescence (WT: 0. 221 ± 0.011; KO: 0.287 ± 0.012; p < 0.01, unpaired two-tailed student’s t-test), as shown in Figure 3E, which is in line with the reviewer’s prediction. This higher fluorescence in KO neurons reflects a build-up of surface signals during applications of acidic buffer after stimulations, thus suggesting an endocytic defect in TRPM7 KO neurons.

Alternatively, the reviewer indicates to report whether there is a difference in the fraction of the signal early on after the end of the stimulus – where there is a real difference in the amount of the signal that has decayed (e.g. at the 60 s time point) – can be quenched by acid perfusion. However, as shown in Figure 3 A, C and E, the endocytic defect in TRPM7 KO neurons builds up gradually from the cessation of stimulations to the end of recordings and there is no obvious difference in fluorescent signals within 8 s after the end of stimulation. Since our acid perfusion is applied 5s right after the end of stimulation, synaptic vesicle endocytosis during this 5s duration, from the end of stimulations to the start of acid perfusion, is relatively small. We think our method may not be sensitive enough to catch a potentially very small difference in synaptic vesicle endocytosis, occurring during the 5s duration (from the end of stimulations to the start of acid perfusion), between WT and TRPM7 KO neurons.

We appreciate the reviewers’ comments on vesicle reacidification, which prompt us to perform additional experiments, thus making our conclusion on vesicle reacidification more convincing.

6. It is important to know if the phenomena described in the present study occur under physiological conditions (i.e. 37{degree sign}C, ~1.25 mM Ca). The temperature in particular is important as cooling can substantially distort the relevance of different molecular processes. The reviewers do not expect a repetition of all experiments at 37{degree sign}C. The easiest might be an experiment on pHluorin assessment of vesicle recycling in neurons, showing that at 37{degree sign}C there is a measurable impact on the pHluorin recovery kinetics due to the absence or presence of external Ca²⁺.

To answer the reviewer’s question, we have performed a side-by-side comparisons of synaptic vesicle endocytosis between WT and TRPM7 KO neurons at 34ºC in the presence of 2 mM extracellular Ca²⁺. Our results showed that synaptic vesicle endocytosis is much slower in TRPM7 KO neurons as compared to WT neurons at physiological temperature (p < 0.001), which is consistent with our data observed at room temperature (Figure 3). This result is now added as Figure 3—figure supplement 1 in the revised manuscript. We thank the reviewer for this comment, which strengths our conclusion on the role of TRPM7 in synaptic vesicle endocytosis.

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[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Your manuscript has been very substantially improved. However, the reviewers have identified three issues that need to be resolved before the paper can be published in eLife:

1. The new confocal images added to Figure 3 (supplement 3, panels A-D; supplement 4, panels A, C, E, G) and to Figure 4 (supplement 2, panel C) are of very poor quality (very low resolution, blurred, pixelated) and do not look like confocal images (rather like ones that have been improperly digitally compressed). In fact, the new panel C in Figure 4, supplement 2 appears to be of even less resolution than the original images in panel A. In essence, the problematic panels indicated above are not of publication quality and need to be redone.

We appreciate reviewer’s comment. We realized that the confocal images mentioned by the reviewers were captured with pinhole of 200 µm, which gave a relatively poor quality of images. To obtain confocal images with better quality, we have re-captured the confocal images with pinhole set to 50 µm. Consequently, all the confocal images related to Figure 3 (supplement 3, panels A-D; supplement 4, panels A, C, E, G) and to Figure 4 (supplement 2, panel C) have been replaced with our recaptured images with much higher quality.

2. The description of confocal microscopy in the methods part is insufficient and requires more detail (i.e. numerical aperture and type of objective, light source, pinhole setting, etc.).

Following the reviewer’s comment, we have added more description on confocal imaging in “Materials and methods”. Regarding methods on confocal imaging, it reads as “Confocal fluorescence images of chromaffin cells or neurons were captured using an Olympus FluoviewFV10i laser-scanning confocal microscope with a 60x oil-immersion objective (NA 1.4) and 473nm, 559nm, and 635nm lasers. The pinhole was set to 50 μm for imaging capturing, and the image size was 1024 x 1024.” now in the revised manuscript.

3. Regarding Figure 5, supplement 1, it seems that the 340 nm images in panel A are saturated. If so, it would be impossible to do a ratio analysis as shown in panel B. This issue needs to be explained and resolved.

We appreciate that the reviewer made a valid point, and we agree that any potential saturation will for sure affect data analysis. However, we believe that none of our images used for analysis were saturated. Our Photometrics Prime 95B sCMOS camera saturates when the intensity has reached 65000 a.u. Author response image 1 shows intensity plots for the raw images that were used for panel A of Figure 5, supplement 1, and the intensity seems far below 65000 a.u of our camera’s saturating limit. In the meantime, we realized that the brightness of those images was adjusted for visualization. To minimize any potential concerns on saturation of displayed images, we have replaced with new images that have reduced brightness but exactly same display settings between WT and KO cells in the revised manuscript.

Author response image 1

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https://doi.org/10.7554/eLife.66709.sa2

Article and author information

Author details

Zhong-Jiao Jiang

Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States

Contribution
Conceptualization, Formal analysis, Investigation, Methodology, Software, Validation, Writing – original draft, Writing – review and editing

Contributed equally with
Wenping Li

Competing interests
No competing interests declared
Wenping Li

Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States

Contribution
Conceptualization, Formal analysis, Investigation, Methodology, Writing – original draft, Writing – review and editing

Contributed equally with
Zhong-Jiao Jiang

Competing interests
No competing interests declared
Li-Hua Yao
1. Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States
2. School of Life Science, Jiangxi Science & Technology Normal University, Nanchang, China
Contribution
Investigation

Competing interests
No competing interests declared
Badeia Saed

Department of Chemistry, University of Illinois at Chicago, Chicago, United States

Contribution
Investigation, Methodology

Competing interests
No competing interests declared
Yan Rao

Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States

Contribution
Formal analysis, Investigation, Writing – review and editing

Competing interests
No competing interests declared
Brian S Grewe

Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States

Contribution
Investigation, Methodology

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0002-8287-6588
Andrea McGinley

Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States

Contribution
Investigation, Methodology

Competing interests
No competing interests declared
Kelly Varga
1. Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States
2. Department of Biological Sciences, University of North Texas at Dallas, Dallas, United States
Contribution
Investigation

Competing interests
No competing interests declared
Simon Alford

Department of Anatomy and Cell Biology, University of Illinois at Chicago, Chicago, United States

Contribution
Investigation

Competing interests
No competing interests declared
Ying S Hu

Department of Chemistry, University of Illinois at Chicago, Chicago, United States

Contribution
Investigation, Methodology, Resources

Competing interests
No competing interests declared
Liang-Wei Gong

Department of Biological Sciences, University of Illinois at Chicago, Chicago, United States

Contribution
Conceptualization, Funding acquisition, Project administration, Supervision, Writing – original draft, Writing – review and editing

For correspondence
lwgong@uic.edu

Competing interests
No competing interests declared

"This ORCID iD identifies the author of this article:" 0000-0002-5868-6039

Funding

NIH Office of the Director (R01NS110533)

Liang-Wei Gong

NSF (IOS-1346826)

Brian S Grewe

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work is supported by NIH (R01NS110533) to L-WG and NSF (IOS-1346826) to BSG. The authors are grateful to Dr V Haucke for vGlut1-pHluorin, Dr TA Ryan for SyGCaMP6f, Dr ER Chapman for iGluSnFR, and Dr AM Scharenberg for TRPM7 plasmid (#45482, Addgene).

Ethics

Senior Editor

Lu Chen, Stanford University, United States

Reviewing Editor

Nils Brose, Max Planck Institute of Experimental Medicine, Germany

Version history

Received: January 20, 2021
Preprint posted: January 25, 2021 (view preprint)
Accepted: September 10, 2021
Accepted Manuscript published: September 27, 2021 (version 1)
Version of Record published: October 14, 2021 (version 2)

Copyright

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.