Serotonin regulates glucose-stimulated insulin secretion from pancreatic β cells during pregnancy

In preparation for the metabolic demands of pregnancy, β cells in the maternal pancreatic islets increase both in number and in glucose-stimulated insulin secretion (GSIS) per cell. Mechanisms have been proposed for the increased β cell mass, but not for the increased GSIS. Because serotonin production increases dramatically during pregnancy, we tested whether flux through the ionotropic 5-HT3 receptor (Htr3) affects GSIS during pregnancy. Pregnant Htr3a^−/− mice exhibited impaired glucose tolerance despite normally increased β cell mass, and their islets lacked the increase in GSIS seen in islets from pregnant wild-type mice. Electrophysiological studies showed that activation of Htr3 decreased the resting membrane potential in β cells, which increased Ca²⁺ uptake and insulin exocytosis in response to glucose. Thus, our data indicate that serotonin, acting in a paracrine/autocrine manner through Htr3, lowers the β cell threshold for glucose and plays an essential role in the increased GSIS of pregnancy.Pregnancy places unique demands on the metabolism of the mother. As the pregnancy progresses and the nutrient requirements of the fetus increase, rising levels of placental hormones reduce maternal insulin sensitivity, thereby maintaining the maternal/fetal gradient of glucose and the flow of nutrients to the fetus. The mother balances the resulting increase in insulin demand with structural and functional changes in the islets that generate increased and hyperdynamic insulin secretion. β cell numbers increase, the threshold for glucose decreases, and glucose-stimulated insulin secretion (GSIS) increases (1–3). Failure to reach this balance of insulin demand with insulin production results in gestational diabetes (4).However, the changes in the maternal islets are not simply a response to increased insulin demand, as they precede the development of insulin resistance. Instead, these changes correlate more closely with levels of circulating maternal lactogens (prolactin and placental lactogen) that signal through the prolactin receptor on the β cell (5–9). Downstream of the prolactin receptor, multiple pathway components have been identified that contribute to the maternal increase in β cell mass (10–16), but not the changes in GSIS.In response to the lactogen signaling during pregnancy, levels of both isoforms of tryptophan hydroxylase, the rate-limiting enzyme in the synthesis of serotonin (5-hydroxytryptamine; 5-HT), rise dramatically in the islet (13, 17, 18). Islet serotonin acts in an autocrine/paracrine manner through the Gα_q-coupled serotonin receptor 5-HT2b receptor (Htr2b) to increase β cell proliferation and mass at midgestation and through Gα_i-coupled 5-HT1d receptor (Htr1d) to reduce β cell mass at the end of gestation (13). These dynamic changes in 5-HT receptor (Htr) expression can explain the shifts in β cell proliferation during pregnancy.In addition to Htr2b and Htr1d, β cells also express Htr3a and Htr3b (13). Unlike the 12 other Htr genes in the mouse genome, which encode G-protein coupled serotonin receptors, Htr3a and Htr3b encode subunits of the serotonin-gated cation channel Htr3 (19, 20). Five identical Htr3a subunits or a mixture of Htr3a and Htr3b make up a functional Htr3 channel (21). The channel is predominantly Na⁺- and K⁺-selective, and its opening in response to serotonin actives an inward current and depolarizes the cell membrane (22, 23). Glucose also depolarizes β cells: Rising ATP from glucose catabolism depolarizes the cell by closing ATP-sensitive K⁺ channels, which causes Ca²⁺ to enter the cell through voltage-gated Ca²⁺ channels and trigger insulin granule exocytosis (24).Therefore, we tested the possibility that Htr3 may regulate β cell insulin secretion during pregnancy. We found that lactogen-induced serotonin in the pregnant islet acts through Htr3 to depolarize β cells, thereby lowering the threshold for glucose and enhancing GSIS during pregnancy.     

From the Cover: Structural differences in amyloid-β fibrils from brains of nondemented elderly individuals and Alzheimer's disease patients

Although amyloid plaques composed of fibrillar amyloid-β (Aβ) assemblies are a diagnostic hallmark of Alzheimer''s disease (AD), quantities of amyloid similar to those in AD patients are observed in brain tissue of some nondemented elderly individuals. The relationship between amyloid deposition and neurodegeneration in AD has, therefore, been unclear. Here, we use solid-state NMR to investigate whether molecular structures of Aβ fibrils from brain tissue of nondemented elderly individuals with high amyloid loads differ from structures of Aβ fibrils from AD tissue. Two-dimensional solid-state NMR spectra of isotopically labeled Aβ fibrils, prepared by seeded growth from frontal lobe tissue extracts, are similar in the two cases but with statistically significant differences in intensity distributions of cross-peak signals. Differences in solid-state NMR data are greater for 42-residue amyloid-β (Aβ42) fibrils than for 40-residue amyloid-β (Aβ40) fibrils. These data suggest that similar sets of fibril polymorphs develop in nondemented elderly individuals and AD patients but with different relative populations on average.

Amyloid plaques in brain tissue, containing fibrils formed by amyloid-β (Aβ) peptides, are one of the diagnostic pathological signatures of Alzheimer''s disease (AD). Clear genetic and biomarker evidence indicates that Aβ is key to AD pathogenesis (1). However, Aβ is present as a diverse population of multimeric assemblies, ranging from soluble oligomers to insoluble fibrils and plaques, and may lead to neurodegeneration by a number of possible mechanisms (2–7).One argument against a direct neurotoxic role for Aβ plaques and fibrils in AD is the fact that plaques are not uncommon in the brains of nondemented elderly people, as shown both by traditional neuropathological studies (8, 9) and by positron emission tomography (10–13). On average, the quantity of amyloid is greater in AD patients (10) and (at least in some studies) increases with decreasing cognitive ability (12, 14, 15) or increasing rate of cognitive decline (16). However, a high amyloid load does not necessarily imply a high degree of neurodegeneration and cognitive impairment (11, 13, 17).A possible counterargument comes from studies of the molecular structures of Aβ fibrils, which show that Aβ peptides form multiple distinct fibril structures, called fibril polymorphs (18–20). Polymorphism has been demonstrated for fibrils formed by both 40-residue amyloid-β (Aβ40) (19, 21–24) and 42-residue amyloid-β (Aβ42) (22, 25–29) peptides, the two main Aβ isoforms. Among people with similar total amyloid loads, variations in neurodegeneration and cognitive impairment may conceivably arise from variations in the relative populations of different fibril polymorphs. As a hypothetical example, if polymorph A was neurotoxic but polymorph B was not, then people whose Aβ peptides happened to form polymorph A would develop AD, while people whose Aβ peptides happened to form polymorph B would remain cognitively normal. In practice, brains may contain a population of different propagating and/or neurotoxic Aβ species, akin to prion quasispecies or “clouds,” and the relative proportions of these and their dynamic interplay may affect clinical phenotype and rates of progression (30).Well-established connections between molecular structural polymorphism and variations in other neurodegenerative diseases lend credence to the hypothesis that Aβ fibril polymorphism plays a role in variations in the characteristics of AD. Distinct strains of prions causing the transmissible spongiform encephalopathies have been shown to involve different molecular structural states of the mammalian prion protein PrP (30–32). Distinct tauopathies involve different polymorphs of tau protein fibrils (33–37). In the case of synucleopathies, α-synuclein has been shown to be capable of forming polymorphic fibrils (38–40) with distinct biological effects (41–43).Experimental support for connections between Aβ polymorphism and variations in characteristics of AD comes from polymorph-dependent fibril toxicities in neuronal cell cultures (19), differences in neuropathology induced in transgenic mice by injection of amyloid-containing extracts from different sources (44–46), differences in conformation and stability with respect to chemical denaturation of Aβ assemblies prepared from brain tissue of rapidly or slowly progressing AD patients (47), and differences in fluorescence emission spectra of structure-sensitive dyes bound to amyloid plaques in tissue from sporadic or familial AD patients (48, 49).Solid-state NMR spectroscopy is a powerful method for investigating fibril polymorphism because even small, localized changes in molecular conformation or structural environment produce measurable changes in ¹³C and ¹⁵N NMR chemical shifts (i.e., in NMR frequencies of individual carbon and nitrogen sites). Full molecular structural models for amyloid fibrils can be developed from large sets of measurements on structurally homogeneous samples (21, 25, 26, 29, 38, 50). Alternatively, simple two-dimensional (2D) solid-state NMR spectra can serve as structural fingerprints, allowing assessments of polymorphism and comparisons between samples from different sources (22, 51).Solid-state NMR requires isotopic labeling and milligram-scale quantities of fibrils, ruling out direct measurements on amyloid fibrils extracted from brain tissue. However, Aβ fibril structures from autopsied brain tissue can be amplified and isotopically labeled by seeded fibril growth, in which fibril fragments (i.e., seeds) in a brain tissue extract are added to a solution of isotopically labeled peptide (21, 22, 52). Labeled “daughter” fibrils that grow from the seeds retain the molecular structures of the “parent” fibrils, as demonstrated for Aβ (19, 21, 24, 53) and other (54, 55) amyloid fibrils. Solid-state NMR measurements on the brain-seeded fibrils then provide information about molecular structures of fibrils that were present in the brain tissue at the time of autopsy. Using this approach, Lu et al. (21) developed a full molecular structure for Aβ40 fibrils derived from one AD patient with an atypical clinical history (patient 1), showed that Aβ40 fibrils from a second patient with a typical AD history (patient 2) were qualitatively different in structure, and showed that the predominant brain-derived Aβ40 polymorph was the same in multiple regions of the cerebral cortex from each patient. Subsequently, Qiang et al. (22) prepared isotopically labeled Aβ40 and Aβ42 fibrils from frontal, occipital, and parietal lobe tissue of 15 patients in three categories, namely typical long-duration Alzheimer''s disease (t-AD), the posterior cortical atrophy variant of Alzheimer''s disease (PCA-AD), and rapidly progressing Alzheimer''s disease (r-AD). Quantitative analyses of 2D solid-state NMR spectra led to the conclusions that Aβ40 fibrils derived from t-AD and PCA-AD tissue were indistinguishable, with both showing the same predominant polymorph; that Aβ40 fibrils derived from r-AD tissue were more structurally heterogeneous (i.e., more polymorphic); and that Aβ42 fibrils derived from all three categories were structurally heterogeneous, with at least two prevalent Aβ42 polymorphs (22).In this paper, we address the question of whether Aβ fibrils that develop in cortical tissue of nondemented elderly individuals with high amyloid loads are structurally distinguishable from fibrils that develop in cortical tissue of AD patients. As described below, quantitative analyses of 2D solid-state NMR spectra of brain-seeded samples indicate statistically significant differences for both Aβ40 and Aβ42 fibrils. Differences in the 2D spectra are subtle, however, indicating that nondemented individuals and AD patients do not develop entirely different Aβ fibril structures. Instead, data and analyses described below suggest overlapping distributions of fibril polymorphs, with different relative populations on average.     

Sequential in vivo labeling of insulin secretory granule pools in INS-SNAP transgenic pigs

β cells produce, store, and secrete insulin upon elevated blood glucose levels. Insulin secretion is a highly regulated process. The probability for insulin secretory granules to undergo fusion with the plasma membrane or being degraded is correlated with their age. However, the molecular features and stimuli connected to this behavior have not yet been fully understood. Furthermore, our understanding of β cell function is mostly derived from studies of ex vivo isolated islets in rodent models. To overcome this translational gap and study insulin secretory granule turnover in vivo, we have generated a transgenic pig model with the SNAP-tag fused to insulin. We demonstrate the correct targeting and processing of the tagged insulin and normal glycemic control of the pig model. Furthermore, we show specific single- and dual-color granular labeling of in vivo–labeled pig pancreas. This model may provide unprecedented insights into the in vivo insulin secretory granule behavior in an animal close to humans.

Dysfunction of pancreatic islet β cells is a key contributor to type 2 diabetes mellitus (T2DM) (1, 2), starting in the early onset of the disease (3). Each β cell contains several thousand insulin secretory granules (SGs) (4, 5). However, only a small percentage of insulin SGs undergo exocytosis upon glucose stimulation (6). Insulin is secreted in two phases: a rapid first and a sustained second phase (7–9). On the level of insulin SGs, our understanding of insulin secretion has been shaped by two basic concepts: 1) the recruitment of SG pools defined by their spatial confinement in the cell and 2) the higher probability of young SGs for exocytosis. In model 1), the so-called, readily releasable pool consists of SGs that are already docked with the plasma membrane and are released immediately upon glucose stimulation, thereby creating the first rapid phase of insulin secretion (10). The second prolonged phase is then caused by the recruitment of SGs from the reserve pool, which is located deeper inside the β cell (6). The detailed properties of SGs of the different pools have been debated and refined recently (11). Additionally, data obtained by radio-labeling experiments suggest that young insulin SGs are preferentially secreted (12, 13). A method that allows for the visualization of age-defined pools of the desired protein is to fuse it with the SNAP-tag, a 20-kDa protein tag that reacts covalently in a bioorthogonal manner with fluorescent benzylguanine (BG)-fused substrates in living cells and organisms (14). By using a pulse–chase-labeling approach to track SGs containing an insulin-SNAP chimera, we could confirm the preferential exocytosis of young SGs and also show the preferential intracellular degradation of old SGs (15–17). Furthermore, we found that in insulinoma INS-1 cells a pool of young SGs travels fast on microtubules, while this property is lost for old SGs (18). Young insulin SGs additionally have a more acidic luminal pH compared to old ones (19). Addressing the heterogeneity of insulin SGs and their differential reaction to stimuli and pharmaceutical intervention poses possibilities for the treatment of T2DM. Genetically modified mouse models have been the method of choice to investigate intracellular signaling, as well as metabolism, in diabetes research. Recently, transgenic pigs have been made available that allow for conducting β cell research in a context even closer to humans (20).Here, we describe the generation and characterization of a transgenic pig model with the SNAP-tag fused to insulin, called the Study OF Insulin granule Aging (SOFIA) pig. We demonstrate the correct targeting and processing of insulin-SNAP to insulin SGs. Finally, we show successful in vivo labeling with one and two SNAP-substrates staining pancreatic islets and distinct insulin SG pools. In summary, our pig model is a valuable system enabling the imaging-based investigation of insulin SG turnover in a large living mammal.     

Synaptotagmin-1 is a bidirectional Ca2+ sensor for neuronal endocytosis

Exocytosis and endocytosis are tightly coupled. In addition to initiating exocytosis, Ca²⁺ plays critical roles in exocytosis–endocytosis coupling in neurons and nonneuronal cells. Both positive and negative roles of Ca²⁺ in endocytosis have been reported; however, Ca²⁺ inhibition in endocytosis remains debatable with unknown mechanisms. Here, we show that synaptotagmin-1 (Syt1), the primary Ca²⁺ sensor initiating exocytosis, plays bidirectional and opposite roles in exocytosis–endocytosis coupling by promoting slow, small-sized clathrin-mediated endocytosis but inhibiting fast, large-sized bulk endocytosis. Ca²⁺-binding ability is required for Syt1 to regulate both types of endocytic pathways, the disruption of which leads to inefficient vesicle recycling under mild stimulation and excessive membrane retrieval following intense stimulation. Ca²⁺-dependent membrane tubulation may explain the opposite endocytic roles of Syt1 and provides a general membrane-remodeling working model for endocytosis determination. Thus, Syt1 is a primary bidirectional Ca²⁺ sensor facilitating clathrin-mediated endocytosis but clamping bulk endocytosis, probably by manipulating membrane curvature to ensure both efficient and precise coupling of endocytosis to exocytosis.

Endocytosis and subsequent vesicle recycling are spatiotemporally coupled to exocytosis, which is critical for neurons and endocrinal cells to maintain the integrity of plasma membrane architecture, intracellular homeostasis, and sustained neurotransmission (1–3). In addition to triggering vesicular exocytosis, neural activity/Ca²⁺ also play an executive role in the coupling of endocytosis to exocytosis (1, 2, 4–6). Following a pioneering study 40 y ago (7), extensive studies have been conducted and showed that Ca²⁺ triggers and facilitates vesicle endocytosis in neurons and nonneuronal secretory cells (1, 8–11). Accumulating evidence also shows that intracellular Ca²⁺ may inhibit endocytosis (12–15), which has been challenged greatly due to the apparently lower occurrences in few preparations and the missing underlining mechanisms, making the endocytic role of Ca²⁺ a four-decades–long dispute (1, 2, 4, 6).Machineries and regulators involved in exocytosis–endocytosis coupling have been extensively studied for over 30 y. The soluble N-ethylmaleimide–sensitive factor attachment protein receptors (SNAREs) and synaptophysin play critical dual roles in exocytosis and endocytosis during neurotransmission (2, 3, 16, 17). Calmodulin and synaptotagmin-1 (Syt1) are currently known primary Ca²⁺ sensors facilitating endocytosis (1, 9, 16, 18, 19). Ca²⁺/calmodulin activate calcineurin, which dephosphorylates endocytic proteins (e.g., dynamin, synaptojanin, and amphiphysin) to facilitate clathrin-mediated endocytosis (CME) and clathrin-independent fast endocytosis (1, 2). Syt1 is a dual Ca²⁺ sensor for both exocytosis and endocytosis (5, 16, 18–20). It promotes CME through binding with the endocytic adaptors adaptor protein-2 (AP-2) and stonin-2 (21–24). In contrast to the well-established Ca²⁺ sensors that promote endocytosis, the mechanism of Ca²⁺-dependent inhibition in endocytosis remains unknown.CME is the classical but slow endocytosis pathway for vesicle retrieval under resting conditions or in response to mild stimulation, while the accumulated Ca²⁺ also triggers calmodulin/calcineurin-dependent bulk endocytosis, which takes up a large area of plasma membrane to fulfill the urgent requirement for high-speed vesicle exocytosis (1–3). They cooperate with kiss-and-run and ultrafast endocytosis to ensure both sufficient and precise membrane retrieval following exocytosis (3, 25–27). These endocytic pathways are all initiated from membrane invagination and are critically controlled by neural activity. However, how the switch between different endocytic modes is precisely determined remains largely unknown.Here, by combining electrophysiological recordings, confocal live imaging, superresolution stimulated emission depletion (STED) imaging, in vitro liposome manipulation, and electron microscope imaging of individual endocytic vesicles, we define Syt1 as a primary and bidirectional Ca²⁺ sensor for endocytosis, which promotes CME but inhibits bulk endocytosis, probably by mediating membrane remodeling. The balance between the facilitatory and inhibitory effects of Syt1 on endocytosis offers a fine-tuning mechanism to ensure both efficient and precise coupling of endocytosis to exocytosis. By including a non-Ca²⁺–binding Syt as the constitutive brake, this work also explains the four-decades–long puzzle about the positive and negative Ca²⁺ effects on endocytosis.     

Synaptotagmin-7 phosphorylation mediates GLP-1–dependent potentiation of insulin secretion from β-cells

Glucose stimulates insulin secretion from β-cells by increasing intracellular Ca²⁺. Ca²⁺ then binds to synaptotagmin-7 as a major Ca²⁺ sensor for exocytosis, triggering secretory granule fusion and insulin secretion. In type-2 diabetes, insulin secretion is impaired; this impairment is ameliorated by glucagon-like peptide-1 (GLP-1) or by GLP-1 receptor agonists, which improve glucose homeostasis. However, the mechanism by which GLP-1 receptor agonists boost insulin secretion remains unclear. Here, we report that GLP-1 stimulates protein kinase A (PKA)-dependent phosphorylation of synaptotagmin-7 at serine-103, which enhances glucose- and Ca²⁺-stimulated insulin secretion and accounts for the improvement of glucose homeostasis by GLP-1. A phospho-mimetic synaptotagmin-7 mutant enhances Ca²⁺-triggered exocytosis, whereas a phospho-inactive synaptotagmin-7 mutant disrupts GLP-1 potentiation of insulin secretion. Our findings thus suggest that synaptotagmin-7 is directly activated by GLP-1 signaling and may serve as a drug target for boosting insulin secretion. Moreover, our data reveal, to our knowledge, the first physiological modulation of Ca²⁺-triggered exocytosis by direct phosphorylation of a synaptotagmin.Glucose-stimulated insulin secretion (GSIS) from pancreatic β-cells follows a biphasic time course consisting of an initial, transient first phase lasting 5–10 min followed by a slowly developing, sustained second phase (1). Type 2 diabetes (T2D) is associated with partial or complete loss of the first insulin secretion phase and a reduction in the second insulin secretion phase (2, 3). Incretins, especially GLP-1, boost GSIS in T2D patients, thereby improving glucose homeostasis (4). GLP-1 exerts its action by activating GLP-1R, a G-protein–coupled receptor expressed on the surface of β-cells, which leads to an increase of adenylate cyclase activity and production of cAMP. Elevated cAMP levels in β-cells enhance GSIS through PKA-dependent and -independent (mediated by Epac2) mechanisms (5, 6). Mouse models with constitutively increased PKA activity have established PKA’s predominant role in the GLP-1–induced potentiation of β-cell GSIS (7, 8), but the downstream effectors remain unidentified.Insulin is secreted in response to glucose by regulated exocytosis of insulin-containing secretory granules. Electrical activity leads to opening of plasmalemmal voltage-gated Ca²⁺ channels (VGCCs) on the β-cell plasma membrane; the resulting increase in [Ca²⁺]_i then triggers Ca²⁺-dependent exocytosis (9). Insulin granule exocytosis is mediated by a multiprotein complex composed of soluble SNAP-receptor (SNARE) proteins (SNAP-25, Syntaxin, and synaptobrevin-2) and Sec1/Munc18-like (SM) proteins (Munc18-1) by a process that shares similarities with synaptic vesicle exocytosis in neurons (10). To date, numerous SNARE isoforms have been implicated in GSIS (11, 12), including Syntaxin-1, Syntaxin-4, SNAP-25 or SNAP-23, and synaptobrevin-2/3 (or VAMP2/3), whereas VAMP8, a nonessential SNARE for GSIS, may be involved in the regulation of GLP-1 potentiation of insulin secretion (13).In addition to SNARE and SM proteins, a Ca²⁺ sensor is required to initiate membrane fusion during exocytosis. Synaptotagmins, expressed mainly in neurons and endocrine cells, share a similar domain structure: a short N-terminal domain, followed by a transmembrane domain, a linker region with variable length, and two tandem Ca²⁺-binding C₂ domains (C₂A and C₂B) at the C terminus (14, 15). Some synaptotagmins bind to phospholipids in a Ca²⁺-dependent manner and have been identified as major Ca²⁺ sensors for regulated exocytosis (14, 16). Synaptotagmin-1, -2, -7, and -9 function as Ca²⁺ sensors for neurotransmitter release, whereas synaptotagmin-1, -7 (Syt7), and -10 regulate hormone secretion and neuropeptide release (9, 17, 18). Specifically, Syt7 regulates insulin granule exocytosis in insulin-secreting cell lines (19, 20). Syt7 is highly expressed in human pancreatic β-cells, and Syt7 KO mice exhibit reduced insulin secretion and consequently impaired glucose tolerance following glucose stimulation (21–23). Collectively, these studies demonstrate that Syt7 is a major Ca²⁺ sensor mediating GSIS in β-cells.Given that GLP-1 potentiates insulin secretion in a glucose-dependent manner, it is highly likely that its insulinotropic action is exerted distally to the initiation of electrical activity, possibly at the level of Ca²⁺ sensing and membrane fusion. Here we report that Syt7 is a stoichiometric substrate for PKA and functions as a downstream target of PKA activated by GLP-1 signaling. Compared with wild-type mice, Syt7 KO mice showed reduced insulin secretion ex vivo and in vivo in response to treatment with the GLP-1 analog exendin-4 in a manner that depended on Syt7 phosphorylation at serine-103. Our data not only provide a mechanism by which GLP-1 stimulates insulin secretion, but also report the physiological regulation of a synaptotagmin by phosphorylation.     

In vivo synaptic recovery following optogenetic hyperstimulation

Local recycling of synaptic vesicles (SVs) allows neurons to sustain transmitter release. Extreme activity (e.g., during seizure) may exhaust synaptic transmission and, in vitro, induces bulk endocytosis to recover SV membrane and proteins; how this occurs in animals is unknown. Following optogenetic hyperstimulation of Caenorhabditis elegans motoneurons, we analyzed synaptic recovery by time-resolved behavioral, electrophysiological, and ultrastructural assays. Recovery of docked SVs and of evoked-release amplitudes (indicating readily-releasable pool refilling) occurred within ∼8–20 s (τ = 9.2 s and τ = 11.9 s), whereas locomotion recovered only after ∼60 s (τ = 20 s). During ∼11-s stimulation, 50- to 200-nm noncoated vesicles (“100nm vesicles”) formed, which disappeared ∼8 s poststimulation, likely representing endocytic intermediates from which SVs may regenerate. In endophilin, synaptojanin, and dynamin mutants, affecting endocytosis and vesicle scission, resolving 100nm vesicles was delayed (>20 s). In dynamin mutants, 100nm vesicles were abundant and persistent, sometimes continuous with the plasma membrane; incomplete budding of smaller vesicles from 100nm vesicles further implicates dynamin in regenerating SVs from bulk-endocytosed vesicles. Synaptic recovery after exhaustive activity is slow, and different time scales of recovery at ultrastructural, physiological, and behavioral levels indicate multiple contributing processes. Similar processes may jointly account for slow recovery from acute seizures also in higher animals.Efficient chemical synaptic neurotransmission requires synaptic vesicle (SV) biogenesis, transmitter loading, membrane approximation and docking, priming, fusion, and release of transmitter (1–3). These processes are followed by retrieval of membrane and proteins from the plasma membrane (PM) via endocytosis (4, 5). Particularly, sustained SV release relies on a tight coupling of exocytosis and endocytosis (5–8). During high-frequency or long-term neuronal activity, SVs need to be efficiently recycled, because, otherwise, the readily releasable pool and the (mobilized) resting pool of SVs would be depleted and transmission would seize (9, 10). After fusion, SV membranes and proteins are recycled (11). Coupling SV exocytosis with local recycling largely eliminates the dependence of chemical transmission on somatic de novo SV synthesis and transport. Thus far, these processes have been studied in dissected preparations or cultured cells and tissues; how and at which time scales this occurs within a live, nondissected animal (e.g., during seizures) is currently unclear. For example, patients suffering from a seizure often remain unconscious for minutes to hours (12, 13). Although fatigue at different levels of circuits and brain systems is likely to contribute, also physiological changes in chemical synapses may play a role in this slow recovery.Depending on the SV fusion rate, endocytosis occurs via different pathways: (i) clathrin-mediated endocytosis (14), supposedly accounting for most recycled SVs; (ii) fast “kiss-and-run” recycling, where SVs do not fully fuse but open a transient pore for transmitter release (15, 16); and (iii) clathrin-independent bulk-phase endocytosis, going along with high neuronal activity (17–20). Following membrane invagination, or to close the fusion pore, the GTPase dynamin finalizes, or at least speeds up, the process of membrane severing (21–25). Before scission by dynamin, the phospholipid phosphatase synaptojanin, via the membrane-binding Bin–amphiphysin–Rvs (BAR) domain protein endophilin, binds to the phospholipid enriched PM at endocytic sites and modifies lipids to promote scission (6, 26–28). Synaptojanin is also required after scission, particularly to uncoat endocytosed vesicles: by dephosphorylating the lipid head groups, synaptojanin releases the interaction of clathrin adaptors with the endocytosed membranes (27, 29). By recruiting clathrin adaptors, synaptotagmin/SNT-1 is also involved in SV recycling (30, 31).Across systems, different modes of endocytosis appear to be in effect: for example, in dynamin knockout mice, spontaneous activity induced endocytosed synaptic membrane, which appeared trapped as invaginations, tubulated and capped by clathrin-coated pits (21). Upon excessive stimulation, in inhibitory neurons, bulk-endocytosed, endosome-like structures resulted, which were severed from the PM despite the absence of dynamin (25). In neuron terminals of Drosophila in which clathrin function was acutely inhibited, SV recycling was impaired, whereas bulk endocytosis was still observed (22), and inhibition of dynamin uncovered different modes of SV recycling (32). Furthermore, when dynamin dephosphorylation was stalled, clathrin-mediated SV endocytosis was functional, but bulk endocytosis was affected (23). Thus, during moderate activity, SV recycling may occur by clathrin-mediated endocytosis (CME), to allow SV proteins to be retrieved and release sites to be “cleared” from integral membrane proteins. Upon prolonged or high-frequency activation, bulk endocytosis may follow in a clathrin-independent and, depending on synapse type, dynamin-dependent or -independent fashion, whereas resolution of the invaginated membrane structures may again be clathrin-dependent. However, in retinal bipolar cells, clathrin was required for a slow (τ ≈ 10–20 s) but not a fast (τ ≈ 1–2 s) phase of endocytosis (33), the latter of which was shown to depend on endophilin (34). In Caenorhabditis elegans, clathrin inactivation, surprisingly, had no effect on normal chemical transmission, and yet SV size was altered (35). Thus, it is unclear whether CME is required for SV endocytosis in C. elegans or is needed at a later step (e.g., following a different endocytic pathway) to shape new SVs. Bulk endocytosis can rapidly remove membrane material from the PM after excessive fusion of many SVs in vitro and was observed in experimental paradigms involving long-term electrical or chemical stimulation and pharmacological treatment (36–42), and yet it has not been studied in an intact animal.How the diverse endocytic events differentially contribute to the dynamic refilling of different SV pools is only partially understood. Because virtually all processes at active zones (AZs) occur at scales below the diffraction limit of light microscopy, it is difficult to study their dynamic behavior during and after stimulation. Therefore, electron microscopy (EM) has been the method of choice to analyze SV pools and AZ morphology at high resolution. Although previous work could visualize triggered SV exocytosis and endocytosis, these dynamic processes are difficult to analyze at high temporal resolution using classical EM (36–39). The dependence on slow chemical-fixation techniques precluded the capture of precise time points during dynamic events and limited the preservation of synaptic structures. Both problems may be overcome using cryofixation by high-pressure freeze (HPF)-EM (43, 44). The requirement of endophilin, synaptojanin, clathrin, and other proteins for SV endocytosis has been studied to some extent in C. elegans, also by EM (27, 28, 35, 43, 45–47). However, this was not done in a temporally resolved fashion relative to a stimulus (i.e., only steady-state “snapshots” were analyzed), and also HPF-EM has not yet been used in this context. How C. elegans synapses regulate endocytosis during and following periods of extreme activity, possibly by different modes of endocytosis, and which proteins are required for this, is unknown.Using a combination of channelrhodopsin-2 (ChR2)-mediated photostimulation of neurons and electrophysiological analysis in dissected animals (48, 49), as well as photostimulation followed by HPF-EM in intact animals, we monitored dynamic processes at AZs in three dimensions at EM resolution, and in a time-dependent manner. We studied the kinetics of docked SV depletion and recovery, as well as the generation and decomposition of bulk-endocytosed vesicles, during and following prolonged photostimulation. Whereas behavioral recovery required 60 s, synapses became fully competent to release transmitter only after ∼20 s, in line with morphological recovery of most docked SVs, whereas spontaneous release occurred at normal rates right after the stimulus. In addition, we found formation and disintegration of large (50–200 nm) bulk-endocytosed vesicles, within 11 and 8 s, respectively, the disassembly of which was largely delayed in animals expressing mutant endophilin, synaptojanin, and dynamin proteins.     

Regulation of beta-amyloid production in neurons by astrocyte-derived cholesterol

Alzheimer’s disease (AD) is characterized by the presence of amyloid β (Aβ) plaques, tau tangles, inflammation, and loss of cognitive function. Genetic variation in a cholesterol transport protein, apolipoprotein E (apoE), is the most common genetic risk factor for sporadic AD. In vitro evidence suggests that apoE links to Aβ production through nanoscale lipid compartments (lipid clusters), but its regulation in vivo is unclear. Here, we use superresolution imaging in the mouse brain to show that apoE utilizes astrocyte-derived cholesterol to specifically traffic neuronal amyloid precursor protein (APP) in and out of lipid clusters, where it interacts with β- and γ-secretases to generate Aβ-peptide. We find that the targeted deletion of astrocyte cholesterol synthesis robustly reduces amyloid and tau burden in a mouse model of AD. Treatment with cholesterol-free apoE or knockdown of cholesterol synthesis in astrocytes decreases cholesterol levels in cultured neurons and causes APP to traffic out of lipid clusters, where it interacts with α-secretase and gives rise to soluble APP-α (sAPP-α), a neuronal protective product of APP. Changes in cellular cholesterol have no effect on α-, β-, and γ-secretase trafficking, suggesting that the ratio of Aβ to sAPP-α is regulated by the trafficking of the substrate, not the enzymes. We conclude that cholesterol is kept low in neurons, which inhibits Aβ accumulation and enables the astrocyte regulation of Aβ accumulation by cholesterol signaling.

Alzheimer’s disease (AD), the most prevalent neurodegenerative disorder, is characterized by the progressive loss of cognitive function and the accumulation of amyloid β (Aβ) peptide and phosphorylated tau (1). Amyloid plaques are composed of aggregates of Aβ peptide, a small hydrophobic protein excised from the transmembrane domain of amyloid precursor protein (APP) by proteases known as beta- (β-) and gamma- (γ-) secretases (SI Appendix, Fig. S1A). In high concentrations, Aβ peptide can aggregate to form Aβ plaques (2–4). The nonamyloidogenic pathway involves a third enzyme, alpha- (α-) secretase, which generates a soluble APP fragment (sAPP-α), helps set neuronal excitability in healthy individuals (5), and does not contribute to the generation of amyloid plaques. Therefore, by preventing Aβ production, α-secretase–mediated APP cleavage reduces plaque formation. Strikingly, both pathways are finely regulated by cholesterol (6) (SI Appendix, Fig. S1B).In cellular membranes, cholesterol regulates the formation of lipid clusters (also known as lipid rafts) and the affinity of proteins to lipid clusters (7), including β-secretase and γ-secretase (8–10). α-secretase does not reside in lipid clusters; rather, α-secretase is thought to reside in a region made up of disordered polyunsaturated lipids (11). The location of APP is less clear. In detergent-resistant membrane (DRM) studies, it primarily associates with lipid from the disordered region, although not exclusively (8, 10, 12–14). Endocytosis is thought to bring APP in proximity to β-secretase and γ-secretase, and this correlates with Aβ production. Cross-linking of APP with β-secretase on the plasma membrane also increases Aβ production, leading to a hypothesis that lipid clustering in the membrane contributes to APP processing (11, 14, 15) (SI Appendix, Fig. S1A). Testing this hypothesis in vivo has been hampered by the small size and transient nature of lipid clusters (often <100 nm), which is below the resolution of light microscopy.Superresolution imaging has emerged as a complimentary technique to DRMs, with the potential to interrogate cluster affinity more directly in a native cellular environment (16). We recently employed superresolution imaging to establish a membrane-mediated mechanism of general anesthesia (17). In that mechanism, cholesterol causes lipid clusters to sequester an enzyme away from its substrate. Removal of cholesterol then releases and activates the enzyme by giving it access to its substrate (SI Appendix, Fig. S1C) (7, 18). A similar mechanism has been proposed to regulate the exposure of APP to its cutting enzymes (11, 15, 19–21).Neurons are believed to be the major source of Aβ in normal and AD brains (22, 23). In the adult brain, the ability of neurons to produce cholesterol is impaired (24). Instead, astrocytes make cholesterol and transport it to neurons with apolipoprotein E (apoE) (25–27). Interestingly, apoE, specifically the e4 subtype (apoE4), is the strongest genetic risk factor associated with sporadic AD (28, 29). This led to the theory that astrocytes may be controlling Aβ accumulation through regulation of the lipid cluster function (11, 15, 19), but this has not yet been shown in the brain of an animal. Here, we show that astrocyte-derived cholesterol controls Aβ accumulation in vivo and links apoE, Aβ, and plaque formation to a single molecular pathway.     

Cross-talk between amyloidogenic proteins in type-2 diabetes and Parkinson’s disease

In type-2 diabetes (T2D) and Parkinson’s disease (PD), polypeptide assembly into amyloid fibers plays central roles: in PD, α-synuclein (aS) forms amyloids and in T2D, amylin [islet amyloid polypeptide (IAPP)] forms amyloids. Using a combination of biophysical methods in vitro we have investigated whether aS, IAPP, and unprocessed IAPP, pro-IAPP, polypeptides can cross-react. Whereas IAPP forms amyloids within minutes, aS takes many hours to assemble into amyloids and pro-IAPP aggregates even slower under the same conditions. We discovered that preformed amyloids of pro-IAPP inhibit, whereas IAPP amyloids promote, aS amyloid formation. Amyloids of aS promote pro-IAPP amyloid formation, whereas they inhibit IAPP amyloid formation. In contrast, mixing of IAPP and aS monomers results in coaggregation that is faster than either protein alone; moreover, pro-IAPP can incorporate aS monomers into its amyloid fibers. From this intricate network of cross-reactivity, it is clear that the presence of IAPP can accelerate aS amyloid formation. This observation may explain why T2D patients are susceptible to developing PD.Parkinson’s disease (PD) is the second most common neurological disorder and the most common movement disorder. It is characterized by widespread degeneration of subcortical structures of the brain, especially dopaminergic neurons in the substantia nigra. These changes are coupled with bradykinesia, rigidity, and tremor, resulting in difficulties in walking and abnormal gait in patients (1). The assembly process of the intrinsically unstructured 140-residue protein α-synuclein (aS) into amyloid fibers has been linked to the molecular basis of PD. aS is a major component of amyloid aggregates found in Lewy body inclusions, which are the pathological hallmark of PD, and duplications, triplications, and point mutations in the aS gene are related to familial PD cases (2, 3). The exact function of aS is unknown, but it is suggested to be involved in synaptic vesicle release and trafficking, regulation of enzymes and transporters, and control of the neuronal apoptotic response (4, 5). aS is present at presynaptic nerve terminals (6–8) and, intriguingly, also in many cells outside the brain (e.g., red blood cells and pancreatic β-cells). aS can assemble via oligomeric intermediates to amyloid fibrils under pathological conditions (9). Although soluble aS oligomers have been proposed to be toxic (10, 11), work with preformed aS fibrils has demonstrated that the amyloid fibrils themselves are toxic and can be transmitted from cell to cell and are also able to cross the blood–brain barrier (12–14).Type-2 diabetes (T2D) is another disease involving amyloid formation. Here, the primary pathological characteristic is islet amyloid of the hormone amylin, also known as islet amyloid polypeptide (IAPP), in pancreatic β-cells (15–18). The process of islet amyloid formation (19–21) leads to pancreatic β-cell dysfunction, cell death, and development of diabetes. IAPP (37 residues, natively unfolded) is cosecreted with insulin after enzymatic maturation of prohormones pro-IAPP (67 residues) and proinsulin in secretory granules. IAPP and insulin play roles in controlling gastric emptying, glucose homeostasis, and in the suppression of glucagon release. Although not understood on a mechanistic level, impairment of prohormone processing has been thought to play a role in initiation and progression of T2D (22, 23). Insulin and pro-IAPP (22, 24–26), but not proinsulin, can inhibit IAPP amyloid formation in vitro and in mice, suggesting that accumulation of unprocessed proinsulin may promote IAPP amyloid formation (22, 24). Insulin-degrading enzyme (IDE) is a conserved metallopeptidase that can degrade insulin and a variety of other small peptides including IAPP in the pancreas (27, 28). Genome-wide association studies have linked IDE to T2D (29, 30) and Ide mutant mice were found to have impaired glucose-stimulated insulin secretion as well as increased levels of IAPP, insulin, and, surprisingly, aS in pancreatic islets (31, 32). Here, aS may be associated with insulin biogenesis and exocytic release, as it was found to localize with insulin-secretory granules in pancreatic β-cells (33). We recently demonstrated in vitro that IDE readily inhibits aS amyloid formation via C-terminal binding and, in parallel, IDE activity toward insulin and other small substrates increases (34, 35).Together, the key role of aS in PD and the inverse correlation of impaired insulin secretion and increased aS levels in the pancreatic β-cells, imply that PD and T2D may be connected. In support, reports have suggested that patients with T2D are predisposed toward PD (36, 37). For Alzheimer’s disease (AD), a direct link with T2D was found (15, 38). Amyloid fiber seeds of the AD peptide, amyloid-β, were shown to efficiently accelerate amyloid formation of IAPP in vitro (39, 40) and IAPP was part of amyloid-β plaque found in mice brains (41). To address the unexplored question of cross-reactivity between the amyloidogenic peptides in PD and T2D, we here investigated cross-reactivity among aS, IAPP, and pro-IAPP using biophysical methods in vitro.     

α-Synuclein assembles into higher-order multimers upon membrane binding to promote SNARE complex formation

Physiologically, α-synuclein chaperones soluble NSF attachment protein receptor (SNARE) complex assembly and may also perform other functions; pathologically, in contrast, α-synuclein misfolds into neurotoxic aggregates that mediate neurodegeneration and propagate between neurons. In neurons, α-synuclein exists in an equilibrium between cytosolic and membrane-bound states. Cytosolic α-synuclein appears to be natively unfolded, whereas membrane-bound α-synuclein adopts an α-helical conformation. Although the majority of studies showed that cytosolic α-synuclein is monomeric, it is unknown whether membrane-bound α-synuclein is also monomeric, and whether chaperoning of SNARE complex assembly by α-synuclein involves its cytosolic or membrane-bound state. Here, we show using chemical cross-linking and fluorescence resonance energy transfer (FRET) that α-synuclein multimerizes into large homomeric complexes upon membrane binding. The FRET experiments indicated that the multimers of membrane-bound α-synuclein exhibit defined intermolecular contacts, suggesting an ordered array. Moreover, we demonstrate that α-synuclein promotes SNARE complex assembly at the presynaptic plasma membrane in its multimeric membrane-bound state, but not in its monomeric cytosolic state. Our data delineate a folding pathway for α-synuclein that ranges from a monomeric, natively unfolded form in cytosol to a physiologically functional, multimeric form upon membrane binding, and show that only the latter but not the former acts as a SNARE complex chaperone at the presynaptic terminal, and may protect against neurodegeneration.α-Synuclein is an abundant presynaptic protein that physiologically acts to promote soluble NSF attachment protein receptor (SNARE) complex assembly in vitro and in vivo (1–3). Point mutations in α-synuclein (A30P, E46K, H50Q, G51D, and A53T) as well as α-synuclein gene duplications and triplications produce early-onset Parkinson''s disease (PD) (4–10). Moreover, α-synuclein is a major component of intracellular protein aggregates called Lewy bodies, which are pathological hallmarks of neurodegenerative disorders such as PD, Lewy body dementia, and multiple system atrophy (11–14). Strikingly, neurotoxic α-synuclein aggregates propagate between neurons during neurodegeneration, suggesting that such α-synuclein aggregates are not only intrinsically neurotoxic but also nucleate additional fibrillization (15–18).α-Synuclein is highly concentrated in presynaptic terminals where α-synuclein exists in an equilibrium between a soluble and a membrane-bound state, and is associated with synaptic vesicles (19–22). The labile association of α-synuclein with membranes (23, 24) suggests that binding of α-synuclein to synaptic vesicles, and its dissociation from these vesicles, may regulate its physiological function. Membrane-bound α-synuclein assumes an α-helical conformation (25–32), whereas cytosolic α-synuclein is natively unfolded and monomeric (refs. 25, 26, 31, and 32; however, see refs. 33 and 34 and Discussion for a divergent view). Membrane binding by α-synuclein is likely physiologically important because in in vitro experiments, α-synuclein remodels membranes (35, 36), influences lipid packing (37, 38), and induces vesicle clustering (39). Moreover, membranes were found to be important for the neuropathological effects of α-synuclein (40–44).However, the relation of membrane binding to the in vivo function of α-synuclein remains unexplored, and it is unknown whether α-synuclein binds to membranes as a monomer or oligomer. Thus, in the present study we have investigated the nature of the membrane-bound state of α-synuclein and its relation to its physiological function in SNARE complex assembly. We found that soluble monomeric α-synuclein assembles into higher-order multimers upon membrane binding and that membrane binding of α-synuclein is required for its physiological activity in promoting SNARE complex assembly at the synapse.     

Rab3-interacting molecules 2α and 2β promote the abundance of voltage-gated CaV1.3 Ca2+ channels at hair cell active zones

Ca²⁺ influx triggers the fusion of synaptic vesicles at the presynaptic active zone (AZ). Here we demonstrate a role of Ras-related in brain 3 (Rab3)–interacting molecules 2α and β (RIM2α and RIM2β) in clustering voltage-gated Ca_V1.3 Ca²⁺ channels at the AZs of sensory inner hair cells (IHCs). We show that IHCs of hearing mice express mainly RIM2α, but also RIM2β and RIM3γ, which all localize to the AZs, as shown by immunofluorescence microscopy. Immunohistochemistry, patch-clamp, fluctuation analysis, and confocal Ca²⁺ imaging demonstrate that AZs of RIM2α-deficient IHCs cluster fewer synaptic Ca_V1.3 Ca²⁺ channels, resulting in reduced synaptic Ca²⁺ influx. Using superresolution microscopy, we found that Ca²⁺ channels remained clustered in stripes underneath anchored ribbons. Electron tomography of high-pressure frozen synapses revealed a reduced fraction of membrane-tethered vesicles, whereas the total number of membrane-proximal vesicles was unaltered. Membrane capacitance measurements revealed a reduction of exocytosis largely in proportion with the Ca²⁺ current, whereas the apparent Ca²⁺ dependence of exocytosis was unchanged. Hair cell-specific deletion of all RIM2 isoforms caused a stronger reduction of Ca²⁺ influx and exocytosis and significantly impaired the encoding of sound onset in the postsynaptic spiral ganglion neurons. Auditory brainstem responses indicated a mild hearing impairment on hair cell-specific deletion of all RIM2 isoforms or global inactivation of RIM2α. We conclude that RIM2α and RIM2β promote a large complement of synaptic Ca²⁺ channels at IHC AZs and are required for normal hearing.Tens of Ca_V1.3 Ca²⁺ channels are thought to cluster within the active zone (AZ) membrane underneath the presynaptic density of inner hair cells (IHCs) (1–4). They make up the key signaling element, coupling the sound-driven receptor potential to vesicular glutamate release (5–7). The mechanisms governing the number of Ca²⁺ channels at the AZ as well as their spatial organization relative to membrane-tethered vesicles are not well understood. Disrupting the presynaptic scaffold protein Bassoon diminishes the numbers of Ca²⁺ channels and membrane-tethered vesicles at the AZ (2, 8). However, the loss of Bassoon is accompanied by the loss of the entire synaptic ribbon, which makes it challenging to distinguish the direct effects of gene disruption from secondary effects (9).Among the constituents of the cytomatrix of the AZ, RIM1 and RIM2 proteins are prime candidates for the regulation of Ca²⁺ channel clustering and function (10, 11). The family of RIM proteins has seven identified members (RIM1α, RIM1β, RIM2α, RIM2β, RIM2γ, RIM3γ, and RIM4γ) encoded by four genes (RIM1–RIM4). All isoforms contain a C-terminal C₂ domain but differ in the presence of additional domains. RIM1 and RIM2 interact with Ca²⁺ channels, most other proteins of the cytomatrix of the AZ, and synaptic vesicle proteins. They interact directly with the auxiliary β (Ca_Vβ) subunits (12, 13) and pore-forming Ca_Vα subunits (14, 15). In addition, RIMs are indirectly linked to Ca²⁺ channels via RIM-binding protein (14, 16, 17). A regulation of biophysical channel properties has been demonstrated in heterologous expression systems for RIM1 (12) and RIM2 (13).A role of RIM1 and RIM2 in clustering Ca²⁺ channels at the AZ was demonstrated by analysis of RIM1/2-deficient presynaptic terminals of cultured hippocampal neurons (14), auditory neurons in slices (18), and Drosophila neuromuscular junction (19). Because α-RIMs also bind the vesicle-associated protein Ras-related in brain 3 (Rab3) via the N-terminal zinc finger domain (20), they are also good candidates for molecular coupling of Ca²⁺ channels and vesicles (18, 21, 22). Finally, a role of RIMs in priming of vesicles for fusion is the subject of intense research (18, 21–27). RIMs likely contribute to priming via disinhibiting Munc13 (26) and regulating vesicle tethering (27). Here, we studied the expression and function of RIM in IHCs. We combined molecular, morphologic, and physiologic approaches for the analysis of RIM2α knockout mice [RIM2α SKO (28); see Methods] and of hair cell-specific RIM1/2 knockout mice (RIM1/2 cDKO). We demonstrate that RIM2α and RIM2β are present at IHC AZs of hearing mice, positively regulate the number of synaptic Ca_V1.3 Ca²⁺ channels, and are required for normal hearing.     

Herpesvirus-mediated stabilization of ICP0 expression neutralizes restriction by TRIM23

Herpes simplex virus (HSV) infection relies on immediate early proteins that initiate viral replication. Among them, ICP0 is known, for many years, to facilitate the onset of viral gene expression and reactivation from latency. However, how ICP0 itself is regulated remains elusive. Through genetic analyses, we identify that the viral γ₁34.5 protein, an HSV virulence factor, interacts with and prevents ICP0 from proteasomal degradation. Furthermore, we show that the host E3 ligase TRIM23, recently shown to restrict the replication of HSV-1 (and certain other viruses) by inducing autophagy, triggers the proteasomal degradation of ICP0 via K11- and K48-linked ubiquitination. Functional analyses reveal that the γ₁34.5 protein binds to and inactivates TRIM23 through blockade of K27-linked TRIM23 autoubiquitination. Deletion of γ₁34.5 or ICP0 in a recombinant HSV-1 impairs viral replication, whereas ablation of TRIM23 markedly rescues viral growth. Herein, we show that TRIM23, apart from its role in autophagy-mediated HSV-1 restriction, down-regulates ICP0, whereas viral γ₁34.5 functions to disable TRIM23. Together, these results demonstrate that posttranslational regulation of ICP0 by virus and host factors determines the outcome of HSV-1 infection.

Herpes simplex viruses (HSV) are human pathogens that switch between lytic and latent infections intermittently (1, 2). This is a lifelong source of infectious viruses (1, 2), in which immediate early proteins drive the onset of HSV replication. Among them, ICP0 enables viral gene expression or reactivation from latency (2–4), which involves chromatin remodeling of the HSV genome, resulting in de novo virus production. In this process, the accessory factor γ₁34.5 of HSV is thought to govern viral protein synthesis (5, 6). It has long been known that γ₁34.5 precludes translation arrest mediated by double-stranded RNA–dependent protein kinase PKR (7–9). The γ₁34.5 protein has also been shown to dampen intracellular nucleic acid sensing, inhibit autophagy, and facilitate virus nuclear egress (10–17). In experimental animal models, wild-type HSV, but not HSV that lacks the γ₁34.5 gene, replicates competently, penetrates from the peripheral tissues to the nervous system and reactivates from latency (18–23). Despite these observations, active HSV replication or reactivation from latency is not readily reconciled by the currently known functions of the γ₁34.5 protein (8–13, 16, 17).Several lines of work demonstrate that tripartite motif (TRIM) proteins regulate innate immune signaling and cell intrinsic resistance to virus infections (24, 25). These host factors typically work as E3 ubiquitin ligases that can synthesize degradative or nondegradative ubiquitination on viral or host proteins. A number of TRIM proteins, for example TRIM5α, TRIM19, TRIM21, TRIM22, and TRIM43, act at different steps of virus replication and subsequently inhibit viral production (26–32). Recent evidence indicates that TRIM23 limits the replication of certain RNA viruses and DNA viruses, including HSV-1 (33). In doing so, TRIM23 recruits TANK-binding kinase 1 (TBK1) to autophagosomes, thus promoting TBK1-mediated phosphorylation and activation of the autophagy receptor p62 and ultimately leading to autophagy. It is unknown whether TRIM23 plays an additional role(s) in HSV infection.Here, we report that ICP0 expression is regulated by the γ₁34.5 protein and TRIM23 during HSV-1 infection. We show that TRIM23 facilitates the proteasomal degradation of ICP0, whereas viral γ₁34.5 maintains steady-state ICP0 expression by preventing K27-linked TRIM23 autoubiquitination that is required for TRIM23 activation. The γ₁34.5 protein also interacts with and stabilizes ICP0, enabling productive infection. Furthermore, we provide evidence that TRIM23 binds to ICP0 and induces its K11-linked polyubiquitination, which triggers K48-linked polyubiquitin-dependent proteasomal degradation of ICP0. These insights establish a model of posttranslational networks in which virus- and host-mediated mechanisms regulate immediate early protein ICP0 stability and thereby lytic HSV replication.     

Cyclodextrins increase membrane tension and are universal activators of mechanosensitive channels

The bacterial mechanosensitive channel of small conductance (MscS) has been extensively studied to understand how mechanical forces are converted into the conformational changes that underlie mechanosensitive (MS) channel gating. We showed that lipid removal by β-cyclodextrin can mimic membrane tension. Here, we show that all cyclodextrins (CDs) can activate reconstituted Escherichia coli MscS, that MscS activation by CDs depends on CD-mediated lipid removal, and that the CD amount required to gate MscS scales with the channel’s sensitivity to membrane tension. Importantly, cholesterol-loaded CDs do not activate MscS. CD-mediated lipid removal ultimately causes MscS desensitization, which we show is affected by the lipid environment. While many MS channels respond to membrane forces, generalized by the “force-from-lipids” principle, their different molecular architectures suggest that they use unique ways to convert mechanical forces into conformational changes. To test whether CDs can also be used to activate other MS channels, we chose to investigate the mechanosensitive channel of large conductance (MscL) and demonstrate that CDs can also activate this structurally unrelated channel. Since CDs can open the least tension-sensitive MS channel, MscL, they should be able to open any MS channel that responds to membrane tension. Thus, CDs emerge as a universal tool for the structural and functional characterization of unrelated MS channels.

Bacterial mechanosensitive (MS) channels have been extensively used as models of ion channel–mediated mechanotransduction (1, 2). They have continually provided novel insights into the biophysical principles that govern ion-channel mechanosensitivity (3–6). While the structurally unrelated MS channels MscL (mechanosensitive channel of large conductance) (4) and MscS (mechanosensitive channel of small conductance) (7) both respond to changes in membrane tension (8–10), at the molecular level, they seem to employ different strategies to convert membrane forces into the conformational changes that underlie channel gating.Escherichia coli MscS is the archetypal member of a large structurally diverse family of ion channels that are expressed in bacteria (11, 12), archaea (13), some fungi (14), plants (15, 16), and eukaryotic parasites (17). This channel gates as a result of membrane tension (10) in accordance with the “force-from-lipids” gating mechanism (6). In response to increases in membrane tension, MscS exhibits complex adaptive gating kinetics (18–20). These kinetic responses may represent two separable processes, adaptation and inactivation (21, 22). In particular, point mutations within transmembrane domain 3 can instigate phenotypes in which adaptation and inactivation are affected differently (18, 23, 24). These complex kinetics are important for the role of this channel as an osmotic safety valve (25). However, since it is currently unknown whether these electrophysiologically separable processes correlate to structurally distinct states, we will refer to them collectively as “desensitization.” In addition, while some data suggest that MscS desensitization is sensitive to the lipid environment (26), this notion still awaits definitive proof.MscL was the first MS channel to be cloned and functionally characterized in a lipid-only environment (8). Members of the MscL family, unlike those of the MscS family, are almost exclusively expressed in archaea and bacteria. After X-ray crystallography revealed the structure of MscL (27), subsequent studies implicated membrane thinning in response to membrane tension as a major driver of MscL gating (3, 28).To fully understand the structural basis of the gating transitions in MscL and MscS, one must first find a way to apply a gating stimulus to the channels in a lipidic environment that is compatible with structural studies. This is, of course, less challenging when considering ligand-gated channels (29–31), for which the stimulus is a defined molecule that can readily be applied to visualize the resulting changes in protein conformation. For MS channels, until recently, only spectroscopic approaches, such as electron paramagnetic resonance spectroscopy (32, 33) and Förster resonance energy transfer spectroscopy (34, 35), were available to provide structural insights into their gating in response to changes in forces in their lipid environment. Other approaches had been confined to the use of activators (36) or mutations (37, 38). We recently demonstrated that lipid removal by β-cyclodextrin can mimic membrane tension in membrane-scaffold protein-based lipid nanodiscs, providing novel insights into the structural rearrangements that underlie MscS channel gating in response to membrane tension (39). The idea was that, as long as the surface area would not change and the lipids would not be replaced, β-cyclodextrin–mediated lipid removal from a membrane would result in the remaining lipids having to cover a larger surface area. This increase in “area-per-lipid” would result in a corresponding increase in membrane tension (40, 41) that would be experienced by integral membrane proteins incorporated in that membrane.Cyclodextrins (CDs) are a family of cyclic glucose oligomers with a cone-like three-dimensional architecture characterized by a polar external surface and a hydrophobic cavity (42, 43). α-, β-, and γ-CD contain six to eight glucose units, respectively. As the number of units increases, so does the diameter of the hydrophobic cavity (5 to 8 Å) (43). These compounds are of broad utility, as the hydrophobic cavity can chelate a plethora of small lipophilic molecules (44, 45). CDs can also form complexes with fatty acids and phospholipids (46). CDs have thus been widely used to remove lipids from native cell membranes (47, 48) and from model membranes (49, 50). This removal of lipids has already been directly linked to increases in membrane tension even in intact cellular environments (51). CDs also exhibit differential lipid selectivity. For example, α-CD has the selectivity profile of phosphatidylserine > phosphatidylethanolamine >> phosphatidylcholine (52). The methylated version of β-CD (mβ-CD) shows selectivity toward cholesterol at low concentrations and has been widely used to selectively remove or add cholesterol to cell membranes (48, 53–55). In addition to the headgroup, CDs also preferentially chelate unsaturated lipids and those containing shorter acyl chains (56, 57).Here, we show that all members of the CD family (α, β, and γ) can activate E. coli MscS in liposomal membranes. Even the methylated version of β-CD, which is widely used for its cholesterol selectivity, can activate E. coli MscS. Congruent with lipid removal increasing tension, the CD amount required for the activation of an MS channel depends on its tension sensitivity. Importantly, as a control, cholesterol-loaded CDs do not activate MscS. Our studies also clearly establish that MscS desensitization is modified by the lipid environment. Moreover, we show that CD-mediated lipid removal causes a concentration- and time-dependent increase in the tension in excised membrane patches and that the resulting tension can become sufficiently high to activate the structurally unrelated MS channel MscL that gates at membrane tensions immediately below the lytic limit of membranes. Two-dimensional (2D) class averages of nanodisc-embedded MscL obtained by cryo–electron microscopy (cryo-EM) indicate that β-CD treatment results in membrane thinning and channel expansion. The fact that CD activates MscL, which opens immediately below the lytic tension of the membrane, suggests that all other MS channels (which are all more sensitive to membrane tension) should also open in response to CDs. These data suggest that CDs will be of broad utility for the structural and functional characterization of structurally diverse MS channels, including Piezo channels (58, 59), two-pore domain K⁺ channels (40, 60), and OSCA channels (61, 62), all of which are known to respond to membrane forces.     

Structural insights into the interaction of IL-33 with its receptors

Interleukin (IL)-33 is an important member of the IL-1 family that has pleiotropic activities in innate and adaptive immune responses in host defense and disease. It signals through its ligand-binding primary receptor ST2 and IL-1 receptor accessory protein (IL-1RAcP), both of which are members of the IL-1 receptor family. To clarify the interaction of IL-33 with its receptors, we determined the crystal structure of IL-33 in complex with the ectodomain of ST2 at a resolution of 3.27 Å. Coupled with structure-based mutagenesis and binding assay, the structural results define the molecular mechanism by which ST2 specifically recognizes IL-33. Structural comparison with other ligand–receptor complexes in the IL-1 family indicates that surface-charge complementarity is critical in determining ligand-binding specificity of IL-1 primary receptors. Combined crystallography and small-angle X-ray–scattering studies reveal that ST2 possesses hinge flexibility between the D3 domain and D1D2 module, whereas IL-1RAcP exhibits a rigid conformation in the unbound state in solution. The molecular flexibility of ST2 provides structural insights into domain-level conformational change of IL-1 primary receptors upon ligand binding, and the rigidity of IL-1RAcP explains its inability to bind ligands directly. The solution architecture of IL-33–ST2–IL-1RAcP complex from small-angle X-ray–scattering analysis resembles IL-1β–IL-1RII–IL-1RAcP and IL-1β–IL-1RI–IL-1RAcP crystal structures. The collective results confer IL-33 structure–function relationships, supporting and extending a general model for ligand–receptor assembly and activation in the IL-1 family.Interleukin (IL)-33 has important roles in initiating a type 2 immune response during infectious, inflammatory, and allergic diseases (1–5). It was initially identified as a nuclear factor in endothelial cells and named NF-HEV (nuclear factor from high endothelial venules) (6, 7). In 2005, it was rediscovered as a new member of the IL-1 family and an extracellular ligand for the orphan IL-1 receptor family member ST2 (8). As an extracellular cytokine, IL-33 is involved in the polarization of Th2 cells and activation of mast cells, basophils, eosinophils, and natural killer cells (1–3). Recent studies also discovered that the type 2 innate lymphoid cells (ILC2s) are major target cells of IL-33 (9, 10). ILC2s express a high level of ST2 and secrete large amounts of Th2 cytokines, most notably IL-5 and IL-13, when stimulated with IL-33 (11–13). Activation of ILC2s is essential in the initiation of the type 2 immune response against helminth infection and during allergic diseases such as asthma (9, 10).IL-33 does not have a signal peptide and is synthesized with an N-terminal propeptide upstream of the IL-1–like cytokine domain. It is preferentially and constitutively expressed in the nuclei of structural and lining cells, particularly in epithelial and endothelial cells (14, 15). Tissue damage caused by pathogen invasion or allergen exposure may lead to the release of IL-33 into extracellular environment from necrotic cells, which functions as an endogenous danger signal or alarmin (14, 16). Full-length human IL-33 consists of 270 residues and is biologically active (17, 18). It is also a substrate of serine proteases released by inflammatory cells recruited to the site of injury (18, 19). The proteases elastase, cathespin G, and proteinase 3 cleave full-length IL-33 to release N-terminal–truncated mature forms containing the IL-1–like cytokine domain: IL-33_95–270, l-33_99–270, and IL-33_109–270 (18). These mature IL-33 forms process a 10-fold greater potency to activate ST2 than full-length IL-33 (18). Caspase-1 was also suggested to cleave IL-33 to generate an active IL-33_112–270 that is the commercially available mature IL-33 form (8). However, it was later demonstrated that this cleavage site does not exist and cleavage by caspases at other sites actually inactivates IL-33 (17, 20, 21).The signaling of IL-33 depends on its binding to the primary receptor ST2 and subsequent recruitment of accessory receptor IL-1RAcP (8, 22, 23). The ligand-binding–induced receptor heterodimerization results in the juxtaposition of the intracellular toll/interleukin-1 receptor (TIR) domains of both receptors, which is necessary and sufficient to activate NF-κB and MAPK pathways in the target cells (24). Previously, we determined the complex structure of IL-1β with its decoy receptor IL-1RII and accessory receptor IL-1RAcP (25). Based on this structure and other previous studies, we proposed a general structural model for the assembly and activation of IL-1 family of cytokines with their receptors (25). In this model, ligand recognition relies on interaction of IL-1 cytokine with its primary receptor: IL-1α and IL-1β with IL-1RI; IL-33 with ST2; IL-18 with IL-18Rα; and IL-36α, IL-36β, and IL-36γ with IL-1Rrp2 (26–28). The binding forms a composite surface to recruit accessory receptor IL-1RAcP shared by IL-1α, IL-1β, IL-33, IL-36α, IL-36β, and IL-36γ and IL-18Rβ by IL-18 (26, 27). This general structural model is further supported by the subsequent structural determination of IL-1β with IL-1RI and IL-1RAcP (29). However, there are still many key missing parts in the general structural model of ligand–receptor interaction in the IL-1 family. For example, the structural basis for specific recognition of IL-33 by ST2 and IL-18 by IL-18Rα, and promiscuous recognition of IL-36α, IL-36β, and IL-36γ by IL-1Rrp2 remains elusive. The proposed general model also needs further confirmation from structural studies of other signaling complexes in the IL-1 family. To address these issues, we studied the interaction of IL-33 with its receptors by a combination of X-ray crystallography and small-angle X-ray–scattering (SAXS) methods.     

From the Cover: PNAS Plus: Nicastrin functions to sterically hinder γ-secretase–substrate interactions driven by substrate transmembrane domain

γ-Secretase is an intramembrane-cleaving protease that processes many type-I integral membrane proteins within the lipid bilayer, an event preceded by shedding of most of the substrate’s ectodomain by α- or β-secretases. The mechanism by which γ-secretase selectively recognizes and recruits ectodomain-shed substrates for catalysis remains unclear. In contrast to previous reports that substrate is actively recruited for catalysis when its remaining short ectodomain interacts with the nicastrin component of γ-secretase, we find that substrate ectodomain is entirely dispensable for cleavage. Instead, γ-secretase–substrate binding is driven by an apparent tight-binding interaction derived from substrate transmembrane domain, a mechanism in stark contrast to rhomboid—another family of intramembrane-cleaving proteases. Disruption of the nicastrin fold allows for more efficient cleavage of substrates retaining longer ectodomains, indicating that nicastrin actively excludes larger substrates through steric hindrance, thus serving as a molecular gatekeeper for substrate binding and catalysis.Regulated intramembrane proteolysis (RIP) involves the cleavage of a wide variety of integral membrane proteins within their transmembrane domains (TMDs) by a highly diverse family of intramembrane-cleaving proteases (I-CLiPs) (1). I-CLiPs are found in all forms of life and govern many important biological functions, including but not limited to organism development (2), lipid homeostasis (3), the unfolded protein response (4), and bacterial quorum sensing (5). As the name implies, RIP must be tightly regulated to ensure that the resultant signaling events occur only when prompted by the cell and to prevent cleavage of the many nonsubstrate “bystander” proteins present within cellular membranes. Despite this, very little is known about the molecular mechanisms by which I-CLiPs achieve their exquisite specificity. Although traditional soluble proteases maintain substrate specificity by recognizing distinct amino acid sequences flanking the scissile bond, substrates for intramembrane proteases have little to no sequence similarity.Recent work on rhomboid proteases has demonstrated that this family of I-CLiPs achieves substrate specificity via a mechanism that is dependent on the transmembrane dynamics of the substrate rather than its sequence of amino acids (6, 7). Here, rhomboid possesses a very weak binding affinity for substrate and, in a rate-driven reaction, only cleaves those substrates that have unstable TMD helices that have had time to unfold into the catalytic active site, where they are cleaved before they can dissociate from the enzyme–substrate complex. Although it may be tempting to speculate that this is a conserved mechanism for all I-CLiPs, rhomboid is the only family of I-CLiPs that does not require prior activation of substrate through an initial cleavage by another protease (8). Specifically, site-2 protease substrates must be first cleaved by site-1 protease (9), signal peptide peptidase substrates are first cleaved by signal peptidase (10), and ectodomain shedding by α- or β-secretase is required before γ-secretase cleavage of its substrates (11, 12). These facts suggest that the diverse families of I-CLiPs likely have evolved fundamentally different mechanisms by which they recognize and cleave their substrates.Presenilin/γ-secretase is the founding member of the aspartyl family of I-CLiPs. The importance of γ-secretase function in biology and medicine is highlighted by its cleavage of the notch family of receptors, which is required for cell fate determination in all metazoans (2, 13–16), and of the amyloid precursor protein (APP), which is centrally implicated in Alzheimer’s disease (AD) (14, 17). In addition to APP and notch, γ-secretase has over 90 other reported substrates, many of which are involved in important signaling events (12, 18). Despite this, little is known about the mechanism by which γ-secretase binds and cleaves its substrates. Currently, the only known prerequisite for a substrate to be bound and hydrolyzed by γ-secretase is that it be a type-I integral membrane protein that first has most of its ectodomain removed by a sheddase, either α- or β-secretases (11, 12, 19). How γ-secretase selectively recognizes ectodomain-shed substrates and recruits them for catalysis while at the same time preventing cleavage of nonsubstrates remains unsettled.γ-Secretase is a multimeric complex composed of four integral membrane proteins both necessary and sufficient for full activity: presenilin, nicastrin, Aph-1, and Pen-2 (20–24). Presenilin is the proteolytic component, housing catalytic aspartates on TMDs 6 and 7 of its nine TMDs (17, 25, 26). After initial complex formation, the mature proteolytically active complex is formed when presenilin undergoes auto-proteolysis, resulting in N- and C-terminal fragments (NTF and CTF, respectively) (17, 27, 28), a process thought to be stimulated by the three-TMD component Pen-2 (29). The seven-TMD protein Aph-1 is believed to play a scaffolding role in complex formation (30, 31). Nicastrin is a type-I integral membrane protein with a large, heavily glycosylated ectodomain (32–34) that contains multiple stabilizing disulfide bridges (24, 34).The ectodomain of nicastrin is structurally homologous to a bacterial amino peptidase (34). Although nicastrin lacks the specific amino acids required for peptidase activity, it has been proposed to bind the N terminus of ectodomain-shed substrate, thereby directing substrate TMD to the γ-secretase active site for cleavage (35, 36). This mechanism has been suggested to depend on a key binding interaction between the free amine at the N terminus of the shortened substrate ectodomain and E333 of the vestigial amino peptidase domain of nicastrin (35, 36). However, the importance of nicastrin in substrate recognition has been questioned (37, 38), and although an initial high-resolution structure of γ-secretase suggested a role for nicastrin in substrate recognition (24), the most recent structures of the γ-secretase complex and the nicastrin ectodomain reveal that E333 is actually buried within the interior of nicastrin and resides on the opposite side of the complex relative to the active site (39, 40). Although this makes it unlikely that nicastrin is involved in direct substrate binding barring a large, energy-intensive conformational change, the basic mechanism of substrate recognition by γ-secretase remains controversial and requires resolution.Here, we demonstrate that nicastrin functions to sterically exclude substrates based on ectodomain size rather than actively recruit them for catalysis. This blocking mechanism allows γ-secretase to distinguish substrate from nonsubstrate and explains why substrate ectodomain shedding by α- or β-secretases is a prerequisite for γ-secretase catalysis. In contrast to rhomboid, γ-secretase apparently binds substrate TMD tightly, making the nicastrin steric hindrance mechanism necessary to prevent cleavage of nonectodomain-shed substrates and nonsubstrates alike.