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Spines are tiny nanoscale protrusions from dendrites of neurons. In the cortex and hippocampus, most of the excitatory postsynaptic sites reside in spines. The bulbous spine head is connected to the dendritic shaft by a thin membranous neck. Because the neck is narrow, spine heads are thought to function as biochemically independent signaling compartments. Thus, dynamic changes in the composition, distribution, mobility, conformations, and signaling properties of molecules contained within spines can account for much of the molecular basis of postsynaptic function and regulation. A major factor in controlling these changes is the diffusional properties of proteins within this small compartment. Advances in measurement techniques using fluorescence microscopy now make it possible to measure molecular diffusion within single dendritic spines directly. Here, we review the regulatory mechanisms of diffusion in spines by local intra-spine architecture and discuss their implications for neuronal signaling and synaptic plasticity.IntroductionNeurons communicate with each other through synapses that organize to create functional circuits. Most excitatory synapses in the central nervous system are formed on dendritic spines, tiny protrusions that extend from dendrites (Bourne and Harris, 2008; Fig. 1 a). The spine typically has a head of 200–1,000-nm diameter, which is connected to the dendritic shaft via a neck of 100–200-nm width (Arellano et al., 2007; Fig. 1 b). The head contains postsynaptic density (PSD) proteins, the actin cytoskeleton, membrane structures, and organelles (Sheng and Hoogenraad, 2007; Fig. 1 c). The molecular composition of spine heads is different from that of the shaft. Because of its characteristic morphology, spines are thought to function as biochemically independent compartments by limiting molecular movement between the spine head and the rest of the dendrite (Adrian et al., 2014; Tønnesen and Nägerl, 2016). Clarifying this regulation is key to understanding how this unitary site of synaptic transmission is controlled. This is particularly crucial to our understanding about how changes in the postsynaptic site lead to synaptic plasticity.Open in a separate windowFigure 1.The shape and internal architecture of dendritic spines. (a) A super-resolution SIM image of a hippocampal neuron dendrite expressing GFP. (b) A surface image of a spine (arrow in a) reconstructed from a SIM image. (c) Schematic representation of a spine containing the PSD, actin cytoskeleton, recycling endosome, and SER.The control of spine architecture is critical at excitatory synapses in the brain (Alvarez and Sabatini, 2007; Forrest et al., 2018). Excitatory synapses exhibit synaptic plasticity, which changes the strength of synaptic transmission through mechanisms at both pre- and postsynaptic sides (Citri and Malenka, 2008). This process is generally thought to be a basis for changes in neural circuits controlled by experiences—i.e., learning and memory (Humeau and Choquet, 2019; Magee and Grienberger, 2020). Here, the size and shape of spines are strongly correlated with the strength of synaptic transmission (Kasai et al., 2010). Spine volume is proportional to PSD area (Harris and Stevens, 1989) and the number of α-amino-3-hydroxyl-5-methyl-4-isoxazole-propionate–type glutamate receptors (AMPARs; Nusser et al., 1998; Matsuzaki et al., 2001). Recently, combinational analysis of electrophysiology and correlative light and EM (CLEM) revealed the linear relationship between PSD area and synaptic strength (Holler et al., 2021). Also, longer spine necks attenuate somatic potentials to a greater degree (Araya et al., 2006). Thus, structural and functional plasticity of spines is tightly regulated. Specifically, when synaptic transmission is strengthened (e.g., long-term potentiation [LTP]), spines grow (Matsuzaki et al., 2004). In turn, when synaptic transmission weakens (e.g., long-term depression), spines shrink (Zhou et al., 2004; Oh et al., 2013).While many molecules involved in the plasticity of spine synapses have been identified (Sala and Segal, 2014), their mechanisms and regulations can only be discovered by monitoring the regulated changes in the composition and signaling properties of these factors within the confined space of the spine’s cytoplasm. To this end, the development of local photolysis of caged-glutamate played an important role (Matsuzaki et al., 2001). This method made it possible to induce structural plasticity locally at a single spine. Spine enlargement is induced by uncaging of caged-glutamate in the absence of Mg2+ or with postsynaptic depolarization in the presence of Mg2+ to activate N-methyl-D-aspartate–type glutamate receptors (NMDARs; Matsuzaki et al., 2004). Conversely, spine shrinkage is induced by low-frequency uncaging of caged-glutamate in the absence of Mg2+ or with postsynaptic depolarization (Oh et al., 2013). Shrinkage can also be induced by glutamate uncaging temporally coupled with back propagation action potential and uncaging of caged–γ-aminobutyric acid (GABA; Hayama et al., 2013). Glutamate uncaging–induced structural plasticity has also been seen to occur in vivo (Noguchi et al., 2019).These methods have vastly improved our understanding of the molecular mechanisms of structural plasticity (Nishiyama and Yasuda, 2015). In particular, stimulus-dependent increases in spine size (structural LTP [sLTP]), which are thought to be associated with functional LTP, have been studied extensively as a model of LTP (Nakahata and Yasuda, 2018; Fig. 2). Strong synaptic input causes an influx of Ca2+ through NMDARs that activates Ca2+/calmodulin-dependent protein kinase II (CaMKII) and the downstream signaling cascades. This modification of signaling cascades can affect cytoskeletal organization and membrane trafficking, which are responsible for two subsequent cellular events. First, spine morphology is modulated through cytoskeletal changes (Borovac et al., 2018). Second, synaptic transmission is enhanced by increased AMPAR insertion into the plasma membrane and movement to the PSD (Huganir and Nicoll, 2013). However, the molecular mechanisms that link these two phenomena are not fully understood (Herring and Nicoll, 2016). As sLTP progresses, the molecular composition within spines changes (Bosch et al., 2014; Meyer et al., 2014). Specifically, immediately after sLTP induction, actin-related molecules such as cofilin and actin-related protein 2/3 (Arp2/3) complex accumulate within a stimulated spine. On the other hand, scaffold proteins such as PSD-95 slowly accumulate over tens of minutes. SynGAP, which is localized at the PSD through interaction with PSD-95, escapes from spines immediately after stimulation and contributes to the expression of sLTP (Araki et al., 2015). These changes in molecular compositions immediately after stimulation may be due not only to molecule-specific binding but also to the physical regulation of diffusion (Obashi et al., 2019).Open in a separate windowFigure 2.Molecular motion important in sLTP. Strong synaptic input causes an influx of Ca2+ through NMDARs that activates CaMKII and the downstream signaling cascades. This modification of signaling cascades can affect cytoskeletal organization and membrane trafficking, which regulate spine morphology. Spine morphology affects the molecular exchange between the spine head and the dendritic shaft and lateral diffusion of membrane proteins including AMPARs. Regulation of molecular movements through the spine neck affects the molecular composition within spines. This change affects signal propagation into nearby spines. For example, cofilin and Arp2/3 complex accumulate within spines. SynGAP and activated RhoA escape from spines. Reorganization of the actin cytoskeleton affects movement of large molecules and the formation of a large signaling complex containing CaMKII and Tiam1. Also, the structure of the PSD affects membrane protein diffusion and alters the synaptic trafficking of AMPARs.In addition to molecular localization, fluorescence lifetime imaging of FRET-based biosensors has made it possible to measure spatiotemporal changes in the activity of signaling molecules involved in sLTP (Yasuda, 2012). These studies have demonstrated a critical relationship between the time that a molecule spends within a spine and the rate of signal inactivation. This relationship determines whether an activated signaling molecule is confined within a single spine or escapes from the spine and interacts with effectors present in the adjacent dendritic shaft or nearby spines (Yasuda, 2017). The signal propagation into nearby spines is most likely related to heterosynaptic plasticity, where activated synapses influence neighbor synapses within the same dendritic segments (Oh et al., 2015; Colgan et al., 2018; Chater and Goda, 2021). Thus, diffusion is a central feature of the regulation of spine structural plasticity. However, because the size of spines is small relative to the spatial resolution of diffraction-limited fluorescence microscopy and measuring methods are limited, elucidation of the mechanism regulating diffusion within spines has been challenging.To address this gap in understanding, researchers advanced fluorescence microscopy techniques, which enabled us to measure changes in the nanoscale localization, diffusion, and signal activities inside spines. These studies allow us to directly understand how spine structures physically limit molecular diffusion and reveal fundamental mechanisms that control the localization and biochemical signal transduction pathways in neurons. Here, we summarize recent findings that have revealed physical barriers within spines using super-resolution microscopy (Sigal et al., 2018; Table 1) and molecular dynamics measurements (Fig. 3 and Table 2), and we discuss how these barriers serve as a fundamental feature controlling neuronal signaling and synaptic plasticity.Table 1.List of super-resolution microscopy techniques
Open in a separate windowCLSM, confocal laser scanning microscopy; SMLM, single-molecule localization microscopy; STORM, stochastic optical reconstruction microscopy; TPLSM, two-photon laser scanning microscopy.Open in a separate windowFigure 3.Imaging techniques to measure diffusion inside dendritic spines. (a) FRAP. Fluorescence intensity change is measured after photobleaching fluorescent molecules in a spine head. Fluorescence recovery rate is mostly determined by the exchange rate between spine and dendrite. (b) FCS. The fluctuation of fluorescence intensity from the detection volume fixed inside a spine head (blue region in left panel) is recorded as a function of time (center panel). Since the fluorescence intensity fluctuates as the molecules enter and leave the fixed detection volume, the characteristics of intensity fluctuation essentially contain information about local diffusion speed. To estimate the diffusion coefficient, the autocorrelation function of fluorescence intensity fluctuation is calculated (right panel). (c) SPT. In SPT, molecular trajectory is directly measured with video microscopy. To analyze the speed and pattern of molecular motion, mean squared displacement (MSD) is calculated. For diffusion without barrier, MSD increases linearly against time. On the other hand, for diffusion within the compartment, MSD converges to a certain value, which corresponds to compartment size. (d) Comparison of three measurement techniques.Table 2.List of fluorescence molecular dynamic measurement techniques
Open in a separate windowSpine structures and diffusionThe complex physical structures of spines can impact the diffusion of molecules inside the spine cytoplasm and between spines and their parental dendritic shafts (Fig. 1 c and Fig. 2). For example, consider the diffusional translocation of molecules between the PSD and dendritic shaft. For cytoplasmic proteins, because a spine is connected to the dendritic shaft through a narrow neck, proteins must pass through the neck by diffusion or slow active transport. A spine neck functions as a diffusion barrier because of its narrow width (Svoboda et al., 1996). Molecular complexes with actin filaments and related proteins, such as synaptopodin and ankyrin-G, that maintain this characteristic neck morphology may also affect diffusion. Furthermore, the cytoplasm within spines is likely to be not homogeneous but organized with multiple nanoscale domains with different biophysical properties (Frost et al., 2010a; MacGillavry and Hoogenraad, 2015). Thus, these locally dense cytoskeletal and membranous structures can limit the molecular path of diffusion within a spine by specific binding interactions or nonspecific local steric effects. These factors will change the residence time of proteins within spines.Besides cytosolic proteins, membrane protein diffusion can be regulated by structures on and near the plasma membrane. For example, the cortical cytoskeleton affects the movement of membrane proteins (Kusumi et al., 2012). Furthermore, specialized membrane domains with a high density of membrane-associated structures, such as synaptic contact sites, accumulate many relatively immobile molecules and limit membrane protein diffusion (Trimble and Grinstein, 2015). Lastly, spines are not simply spherical. Boundaries between the spine shaft and neck—and also the spine head and neck—can contain high curvatures. Also, large spine heads contain a concave surface (Kashiwagi et al., 2019). Thus, local concavities, undulations, and convexities may affect the possible path a molecule can take (Simon et al., 2014; Klaus et al., 2016). From all these factors, the shape and internal architecture of spines can have strong effects on diffusion for both cytosolic and membranous proteins.Influences of spine morphology on diffusional coupling between spines and dendritesAlthough the cytoplasm of spines is directly connected to the cytoplasm of dendritic shafts, a narrow neck is thought to limit diffusion of both cytosolic and membrane molecules between two compartments (Holcman and Schuss, 2011; Kusters et al., 2013; Ramirez et al., 2015). FRAP is a method that can be used to measure the diffusional speeds from an exchange rate between nonbleached and bleached molecules after bleaching fluorescent molecules in a small region (Lippincott-Schwartz et al., 2018; Fig. 3, a and d). Local photoactivation or photoconversion and subsequent measurements of fluorescence intensity is another technique comparable to FRAP (Bancaud et al., 2010). When fluorescence bleaching is performed in a spine head, the speed of fluorescence recovery mostly reflects the rate of molecular exchange between the head and the connected dendrite. Because the diffusion of small molecules in a head is faster than the rate of molecular exchange between spines and dendrites, the fast component of intra-spine diffusion is more difficult to detect in FRAP recovery curves (Svoboda et al., 1996). FRAP or photoactivation experiments of cytoplasmic and membrane-anchored fluorescent proteins showed that diffusional coupling between spines and dendrites varies between spines (Bloodgood and Sabatini, 2005; Ashby et al., 2006). Since the shape of spines is diverse, it has been proposed that this diversity underlies variability in spine–dendrite coupling. However, because the details of spine morphology cannot be analyzed with the spatial resolution of diffraction-limited fluorescence microscopy, a relationship between the shape of spines and diffusional coupling had not been directly demonstrated.Recently, however, super-resolution microscopy has made it possible to analyze spine shape in living neurons with a spatial resolution of ∼50 nm (Nägerl et al., 2008). Influences of spine morphology on diffusional coupling were verified experimentally for the first time by directly comparing the morphological features of spines and diffusional coupling. This comparison was achieved by stimulated emission depletion (STED) microscopy of spines combined with FRAP of YFP or Alexa dyes applied to the same spine (Takasaki and Sabatini, 2014; Tønnesen et al., 2014). These direct comparisons indicated that the diversity in diffusional couplings could be explained solely by the diversity of spine shapes for more than half of the measured spines. In other words, it was shown that for many spines, the exchange rate (τ) of small molecules within spines could be explained by a single-compartment model (Svoboda et al., 1996) described by the shape of the spine:where V is the head volume, L is the length of the neck, A is the cross-sectional area of the neck, and D is the diffusion coefficient of molecules. Also, sLTP induction made a spine neck thicker and shorter (Tønnesen et al., 2014). This change in the spine neck complements the decrease in the coupling rate associated with the increase in the spine head volume. This coordinated morphological change appears to maintain molecular concentration in a spine.Besides the work focusing on cytoplasmic proteins, the influence of spine shape on the diffusional coupling of membrane molecules has also been investigated (Adrian et al., 2017). The spine–dendrite diffusional coupling was tested by photoactivated localization microscopy (PALM) and photoconversion experiments using membrane-anchored mEos3.2 as a probe. This study showed that even if spines have the same surface area and neck width, the diffusional coupling varies between different spine shapes. Therefore, a model spine was created based on the experimentally measured spine shape parameters, and a simulation was conducted on the model spine and compared with the experiment. As a result, although experimental results tended to provide slower diffusion kinetics than simulation values, experiments showed a good correlation with simulations based on the spine shape parameters alone.Experiments have confirmed that spine morphology is a major factor determining the diffusional coupling for both cytoplasmic and membrane-bound molecules in dendrites. However, for some spines, the simulated and experimental results diverge. One possibility is that the effects of local intra-spine architectures on molecular diffusion vary for each spine. Another possibility is that there was insufficient spatial resolution for reconstructing the spine morphology. Although the above studies used rotationally symmetric shapes as model spines, actual spines are not rotationally symmetrical structures and generally have a more complicated morphology and surface features (Nägerl et al., 2008; Berning et al., 2012; Kashiwagi et al., 2019; Zaccard et al., 2020; Fig. 1, a and b). Thus, it is possible that estimations of spine shape were insufficient or that the fine structure of spines affects diffusion. In this regard, developing an analysis method for spine morphology from both the experimental and computational sides is key (Okabe, 2020a; Tamada et al., 2020). Recently, Kashiwagi et al. (2019) developed a 3-D structured illumination microscopy (SIM)–based nanoscale analysis of spine morphology. Direct comparison of SIM images and serial-section EM images revealed that the basic morphological features were highly correlated among these images. This indicates the high precision of SIM-based nanoscale spine analysis. To analyze spines computationally, SIM images were converted into a computational geometry, and morphological features were calculated. Then, these features were analyzed by principal component analysis. By mapping the temporal changes of spine morphology obtained by live-cell SIM imaging in the dimension-reduced feature space, the authors revealed that the spine population can be categorized based on different simplified morphological dynamics.Also, expansion microscopy (ExM) is another new and important imaging technique for spine structural analysis (Wassie et al., 2019). However, it can only be applied to fixed samples. Since ExM samples are transparent, 3-D super-resolution imaging is available for thick samples with large volumes (Gao et al., 2019). With recent developments in sample preparation technology, ExM has the potential to investigate spine morphology and localization of multiple biomolecules and organelles within a single sample (Chozinski et al., 2016; Tillberg et al., 2016; Karagiannis et al., 2019 Preprint; Sun et al., 2021). Minimizing the distortion of isotropy during expansion will be important for nanoscale morphological analysis. In the future, combining dynamic fluorescence measurements and structural measurements gained from EM (CLEM) will be a powerful approach to evaluate the effects of spine ultrastructure on molecular diffusion in greater nanoscale detail (Maco et al., 2013; Taraska, 2015; Luckner et al., 2018).Along with biochemical compartmentalization, dendritic spines have been proposed to be important for electrical compartmentalization (Yuste, 2013; Araya, 2014; Tønnesen and Nägerl, 2016). Spine morphology, particularly spine neck morphology, is thought to be critical for this effect (Cartailler et al., 2018). Several studies have sought to measure neck resistance based on morphological analysis using EM (Harris and Stevens, 1989; Tamada et al., 2020), super-resolution microscopy (Tønnesen et al., 2014), FRAP of small molecules (Svoboda et al., 1996; Tønnesen et al., 2014), glutamate uncaging (Araya et al., 2006; Takasaki and Sabatini, 2014), calcium imaging (Grunditz et al., 2008; Harnett et al., 2012), voltage imaging (Popovic et al., 2015; Acker et al., 2016; Kwon et al., 2017), and intracellular recordings directly from spine heads (Jayant et al., 2017). However, results were not completely consistent, and the degree of electrical compartmentalization is still unclear. Thus, the relationship between spine morphology and electrical signaling of the synapse is still an open question. Likewise, how morphological changes in the neck induced by LTP affect dendritic computation will be an important area of future study (Araya et al., 2014; Tazerart et al., 2020).Actin cytoskeletonThe cytoskeleton in spines is primarily composed of actin (Hotulainen and Hoogenraad, 2010; Okabe, 2020b). Actin is present in high densities in both the head and neck regions (Korobova and Svitkina, 2010). Actin polymers are essential in controlling the localization of PSD molecules and in changing and maintaining spine morphology (Frost et al., 2010a; Bertling and Hotulainen, 2017). In addition to these functions, dense actin polymers in spines may regulate synaptic functions by controlling diffusion because the intracellular cytoskeleton and membrane structures influence diffusion (Novak et al., 2009). If this regulation occurs in spines, variations in the distribution of intra-spine structures can be a factor in the large deviations between the measured values of diffusional coupling and the value predicted from models. A ratio of the spine FRAP recovery time of Alexa dyes to that of YFP was comparable to that of hydrodynamic radii (Tønnesen et al., 2014). This suggests that the suppression of diffusion by actin polymers is weak for molecules with the size of GFP. However, suppressive effects on molecular diffusion by the cytoskeleton, such as actin polymers, is dependent on the size of molecules (Baum et al., 2014; Katrukha et al., 2017). Thus, diffusion of larger molecules may be influenced to a greater degree by actin polymers.Because the shape of spines affects the recovery time of FRAP measurement, it is difficult to investigate the effects of intra-spine structure on molecular diffusion using FRAP alone. Therefore, there is a need for a method capable of measuring diffusion directly in confined spaces. Lu et al. (2014) measured the motion of mEOS2-fused CaMKIIα in spines by single-particle tracking (SPT)–PALM. SPT can directly evaluate diffusional speed in spines because it analyzes the molecular movement trajectory of single molecules (Fig. 3, c and d). The SPT measurement showed that CaMKIIα exhibited at least three different diffusion modes within spines: (1) a free diffusion component, (2) a component bound to immobile molecules, and (3) a component moving at an intermediate velocity. Depolymerization of actin polymers by latrunculin A reduced the proportion of molecules with intermediate velocities in spines while concomitantly increasing the free diffusion component. Also, diffusional speeds of CaMKIIα were slower and the ratio of the intermediate component was larger in spines than in dendrites. Because the transition between free and bound states would occur rarely during the measurement period due to the slow unbinding rate of CaMKII from actin polymers, transient binding alone does not explain the mechanism for the intermediate velocity. Although the details are unclear, CaMKII motion is restricted by actin polymers through a mechanism distinct from direct binding, including a molecular sieve effect or transient binding to actin-associated molecules.Obashi et al. (2019) used fluorescence correlation spectroscopy (FCS) and raster image correlation spectroscopy (RICS) to measure the diffusion of biologically inert probes within spines. FCS is a method for estimating diffusion speed from the time taken for fluorescent molecules to pass through the detection volume excited by a high numerical aperture objective and is capable of measuring fast diffusion within a small cellular compartment (Elson, 2011; Fig. 3, b and d). RICS is another method for estimating diffusion speed from the spatial similarity of fluorescence intensity in a scanned image (Digman and Gratton, 2011). Since FCS and RICS are affected by the small size of spines due to the boundary effect, it is not possible to measure the diffusion coefficient accurately (Jiang et al., 2020). Still, by averaging, values proportional to the actual values can be obtained. Diffusion of GFP and GFP tandem pentamer (GFP5) were compared, and only diffusion of GFP5 within spines was enhanced by depolymerizing actin with latrunculin A treatment. Molecular dynamics simulation confirmed that the diffusion of molecules over the size of GFP5 was suppressed by actin polymers with a density (380 µM) estimated from the values in the literature and experiments.Together, these experiments support the idea that a meshwork of dense actin polymers in spines acts as a physical barrier to the diffusion of larger (>100 kD) molecules (Fig. 4 a). Photoactivation experiments of intra-spine photoactivatable GFP (PA-GFP)–actin showed that there are at least three groups of actin polymers with different reorganization rates (Honkura et al., 2008). In addition, experiments with SPT-PALM of PA-GFP–actin showed that a rate of actin filament polymerization increased near PSDs (Frost et al., 2010b). PALM analysis also revealed that actin-related molecules within spines are arranged in a manner specific for each molecule (Chazeau et al., 2014). These results suggest that the diffusional control by actin polymers in spines may differ between each subcompartment.Open in a separate windowFigure 4.Diffusion within network of actin polymers and PSD. (a) Comparison of the size of actin polymer network and diffusion molecules. Average distance between actin polymers is estimated for actin polymers with 380 µM (Obashi et al., 2019). GFP is represented as a diameter of 3 nm and CaMKII is represented as a diameter of 20 nm (Myers et al., 2017). (b) A schematic model of AMPAR diffusion within a crowded PSD. Different localization patterns of molecules cause different diffusion patterns. Such mechanisms will occur within the PSD. Density and size of molecules are based on the literature (Okabe, 2007; Li et al., 2016).It was also shown that reorganization of the actin cytoskeleton immediately after sLTP induction (Chazeau and Giannone, 2016; Mikhaylova et al., 2018) enhanced the diffusion of larger molecules within the spine head (Obashi et al., 2019). Further, FRAP experiments showed that diffusional coupling and synaptic translocation of large synaptic molecules, such as CaMKII and T cell lymphoma invasion and metastasis-inducing protein 1 (Tiam1), were facilitated at the initial phase of sLTP. Thus, the reorganization of actin polymers regulates molecular translocation between dendrites and the PSD in coordination with morphological changes of the spine neck (Tønnesen et al., 2014). The enhancement of molecular diffusion by actin may also be related to the formation of a large signaling complex containing CaMKII and Tiam1 and may be an important physical mechanism responsible for the initiation of sLTP (Saneyoshi et al., 2019).Membranous organellesAlong with the cytoskeleton, dendrites also contain many membranous organelles and compartments, and some are present in spines (Bourne and Harris, 2008). Smooth ER (SER) and recycling endosomes are present in <50% of spines. Spine apparatus, which is composed of stacked SER, is present in 10–20% of spines. Localizations of these organelles change after LTP induction, and spines containing SER are larger than those without SER (Chirillo et al., 2019; Kulik et al., 2019; Perez-Alvarez et al., 2020). While mitochondria are abundant in dendritic shafts but rarely present in spines (Wu et al., 2017), synaptic activation relocates mitochondria into spines (Li et al., 2004). Therefore, variations in the diffusional coupling between spines and dendrites could be due to the heterogeneous localization of these organelles (Cugno et al., 2019). Holbro et al. (2009) compared the diffusional coupling of ER-containing spines and ER-free spines by using an ER-targeted GFP probe. FRAP recovery times of RFP were not different among ER-containing and ER-free spines, indicating that the ER does not block cytoplasmic diffusion between spines and dendritic shafts. Understanding both spine morphology and the volume of ER within spines in the future will clarify the effects of excluded volume by the ER and other organelles in more detail.Structures around spine necksMolecules present in spine necks may physically control diffusion by forming a complex higher-order structure. Platinum replica EM showed the presence of Arp2/3 complex within necks and a longitudinal network of branched and linear actin filaments (Korobova and Svitkina, 2010). SPT-PALM of PA-GFP–actin also showed that actin polymers in necks are dynamically reorganized and that they are arranged in many orientations (Frost et al., 2010b). Thus, actin polymers in spine necks may affect molecular diffusion. Synaptopodin, for example, is an actin-binding protein located predominantly in spine necks. It is colocalized with the spine apparatus (Vlachos, 2012). Wang et al. (2016) used SPT to measure metabotropic glutamate receptor 5 (mGluR5) diffusion around necks. They compared the diffusion of mGluR5 around the necks of spines containing (or not containing) synaptopodin. The diffusion of mGluR5 decreases around spine necks near synaptopodin clusters. Further, latrunculin A treatment specifically enhanced the diffusion around spine necks near synaptopodin clusters. These results suggest that synaptopodin regulates the actin polymer network around spine necks. This actin complex can act as a diffusion barrier for membrane proteins.Another protein that has been implicated in diffusional control of membrane proteins is ankyrin-G. Ankyrin-G forms nanodomain at perisynaptic membranes and in spine necks (Smith et al., 2014). AMPARs accumulated in spines with ankyrin-G clusters and showed slower spine–dendrite coupling. Ankyrin-G is the major cytoskeletal scaffold of the axon initial segment (AIS; Leterrier, 2018). Ankyrin-G and actin scaffolds densely accumulate at the AIS and inhibit diffusion of the membrane and cytoplasmic molecules (Winckler et al., 1999; Nakada et al., 2003; Song et al., 2009). Also, super-resolution microscopy recently revealed the presence of membrane-associated periodic skeleton composed of actin rings, spectrin, and accompanying proteins in the axon including the AIS (Xu et al., 2013; Zhong et al., 2014). At the AIS, the actin rings and associated structures act as a diffusion barrier to membrane proteins (Albrecht et al., 2016). Adding to the axon, membrane-associated periodic skeleton was also observed in the dendrites and spine necks (Bär et al., 2016; Sidenstein et al., 2016). Therefore, it is interesting to postulate that a molecular complex with actin filaments similar to the AIS is also present in spine necks and could regulate molecular diffusion in this small compartment.Another cytoskeletal component, septin 7, localizes to the base of spines and acts as a diffusion barrier for membrane-bound molecules (Ewers et al., 2014). Recently, actin patches were found at the base of spines and were shown to be remodeled by synaptic activity. These structures modulate microtubule entry into spines and the transport of lysosomes (Schätzle et al., 2018; van Bommel et al., 2019). It is interesting to ask whether actin patches at spine bases affect molecular diffusion. There are still many unknown features at the spine neck, and how these structures limit the diffusion of cytoplasmic and membrane molecules to control neuronal functions remains unclarified.Molecular crowding in the PSDThe PSD is a membrane-associated structure containing densely packed postsynaptic molecules (Sheng and Hoogenraad, 2007). It was originally identified as an electron-dense structure in EM (Okabe, 2007). The number and location of receptors and adhesion molecules in PSDs are directly related to synaptic function (Chen et al., 2018). SPT studies indicate that AMPARs diffuse laterally into and out of PSDs and regulate synaptic function by controlling the number and location of AMPARs (Choquet and Hosy, 2020). Because there are many scaffold proteins in PSDs, membrane proteins including AMPARs accumulate in PSDs due to intermolecular binding. Furthermore, because the molecular density in PSDs is high, the accumulation of membrane proteins may be regulated by the suppression of mobility within the PSD and molecular exchange at the boundary of PSDs (Gerrow and Triller, 2010; Kokolaki et al., 2020).To check this possibility, Li et al. (2016) combined FRAP, SPT, and Monte Carlo simulation to investigate the effect of molecular crowding of PSDs on the lateral diffusion of membrane molecules. When the intracellular domain size of membrane proteins was large, diffusion within the PSD and the exchange rate between the inside and outside of the PSD decreased. Super-resolution microscopy showed that the distribution of PSD-95, a major scaffolding protein of the PSD, within PSDs is not uniform (Fukata et al., 2013; MacGillavry et al., 2013; Nair et al., 2013; Broadhead et al., 2016; Gwosch et al., 2020). Interestingly, the simulation showed that the residence time of membrane proteins within PSDs was longer in the condition of experimentally measured PSD-95 distribution, while the residence time decreased with a random distribution of PSD-95 (Li et al., 2016).Recently, the shape of PSDs inside spines induced by sLTP was analyzed by CLEM (Sun et al., 2019 Preprint). It was shown that rearrangements of PSD shape occurred immediately after induction of sLTP (<3 min), and the PSD took more complex morphology. This increased structural complexity persisted in the late phase (120 min). PSD size and the accumulation of PSD-95 increased slowly over several tens of minutes after sLTP induction (Meyer et al., 2014), whereas synaptic transmission efficiency increased immediately (Matsuzaki et al., 2004). This difference in time may be explained by a mechanism in which the acute ultrastructural changes of the PSD without net growth of the molecular assembly alter the mobility of AMPARs by changing the distribution of a physical barrier, leading to alternations in the number and localization of AMPARs (Fig. 4 b). In future studies, it will be necessary to clarify how coordination between intermolecular binding and physical diffusion barriers in PSDs supports both acute accumulation of AMPARs and their subsequent stabilization in stimulated spines. Further, 3-D SIM imaging revealed that the concave surface of the spine head, which interacts with presynaptic membranes, is enlarged and stabilized by sLTP induction (Kashiwagi et al., 2019). In the future, it will be interesting to determine the relationships between concave membrane surfaces, PSD morphologies, and the dynamics of receptors and adhesion molecules at single spines.In addition, although AMPAR has been thought to be present as a tetramer (Greger et al., 2007), recent observations of SPT have shown that the majority of diffusive AMPARs are monomers or dimers (Morise et al., 2019). Molecular diffusion in the monomer form increases an exchange rate between the inside and the outside of PSDs, making it possible to efficiently change the AMPAR composition within synapses. It remains to be seen whether other molecular complexes, such as NMDARs and cell adhesion molecules, also modulate their diffusion within the molecularly dense PSD by changing their oligomeric state.Conclusion and outlookHere, we have highlighted key recent findings on the relationship between molecular diffusion and physical barriers within spines. The regulation of molecular diffusion is important for sLTP expression (Fig. 2). Spine structural changes during sLTP will affect synaptic function in a coordinated manner (Fig. 5). For example, after sLTP induction, the actin network is reorganized and diffusion of large molecules is enhanced (Obashi et al., 2019). This facilitates the formation of large signaling complexes and the rearrangement of protein complexes within spines. At the same time, spine necks become wider and shorter, and spine heads enlarge (Tønnesen et al., 2014). Changes in actin and spine morphology enhance the molecular movement between the PSD and the shaft and are important for the relocation of proteins (Fig. 2). These structural changes occur in the early phase of sLTP. Thus, the cooperative regulation of diffusion might act as a precise temporal switch of sLTP induction. Also, this enhancement of molecular exchange affects the relocation of activated signaling molecules into the shaft or nearby synapses, which leads to heterosynaptic plasticity (Yasuda, 2017). Potentiation of synaptic transmission requires synaptic trafficking of AMPARs (Choquet and Hosy, 2020). Although both spine morphology (Adrian et al., 2017) and PSD structure (Li et al., 2016) affect membrane protein diffusion, how structural changes associated with sLTP induction affect diffusion will be clarified in the future. Furthermore, the effects of transient SER visits (Perez-Alvarez et al., 2020) and structural changes around spine necks are an important area for future work. Although the relationship between structure and diffusion in sLTP is critical, the difficulty of measurements with a small single spine has made a comprehensive view difficult to obtain. Thus, future work will be necessary to clarify how structural changes affect diffusion and how this physical change to dendritic spines cooperatively modulates synaptic functions.Open in a separate windowFigure 5.Changes in the shape and internal architecture of spines after induction of sLTP. At the initial phase of sLTP, a spine head expands. In addition, the spine neck becomes wider and shorter (Tønnesen et al., 2014), and a concave surface area of spine head is increased (Kashiwagi et al., 2019). The actin polymer network is reorganized (Obashi et al., 2019), and the SER visits within a spine transiently (Perez-Alvarez et al., 2020). Also, PSD shape becomes more complex (Sun et al., 2019 Preprint). These physical changes should occur in concert and will affect molecular composition and biochemical signaling through diffusional regulation. These physical changes will act as a precise temporal switch of sLTP induction.Although new imaging techniques have demonstrated the connection between diffusion and physical barriers, little is known about how changes in the movement of molecules alter synaptic functions (Reshetniak et al., 2020b). Because of the small volume of the spine, very small molecules with high diffusivity, such as Ca2+, are expected to spread rapidly (∼1 ms) by diffusion (Chen and Sabatini, 2012). For large molecules such as signaling complexes, it remains to be seen whether spatially uniform diffusion takes place or whether local heterogeneity in the spine cytoplasm results in a more complex pattern of diffusion. It is also necessary to clarify whether such changes affect local biochemical signaling events and molecular localizations. The number of molecules per spine could influence the magnitude of functional changes (Okabe, 2007; Ribrault et al., 2011). Furthermore, the changes in diffusion induced by alterations in spine structure will affect the stability of the structure. This will subsequently change the molecule’s diffusivity. Thus, it will be interesting to investigate whether this type of mutual relationship exists within spines.New imaging techniques will help to answer these questions. By applying fast 3-D SPT to intra-spine measurements, it will be possible to investigate the spatial heterogeneity of diffusion in single spines of living neurons in detail (Hou et al., 2020; Xiang et al., 2020). STED-FCS/fluorescence cross-correlation spectroscopy can also detect changes in intermolecular interactions (Lanzanò et al., 2017). In addition to the development of new measurement techniques, molecular dynamics simulations based on experimental data will become increasingly important in the future (Okabe, 2020a; Reshetniak et al., 2020a; Vasan et al., 2020). Spine morphology and intra-spine structures, which affect diffusion, are closely related. Thus, it is difficult to investigate the effect of one without changing the other experimentally. Molecular dynamics simulation is a useful tool to examine how molecular motion is adjusted by combining elements that are difficult to verify experimentally (Bell et al., 2019). Furthermore, the shape of spines and intra-spine components, such as the actin cytoskeleton, which are the structural basis of spines, differ from spine to spine. Here, a combination of quantitative measurements and simulations based on experimental data will help us to understand molecular events more quantitatively.Although we reviewed work using fluorescence microscopy, details of spine morphology and intra-spine structures have also been revealed by EM at the nanoscale (Bourne and Harris, 2012; Tao et al., 2018). However, it is difficult to observe specific molecular localizations with EM. On the other hand, super-resolution microscopy is suitable for obtaining a nanoscale picture of molecular positions within spines. Yet, it is still difficult to observe dense structures such as actin polymers (Kommaddi et al., 2018). Therefore, in the future, it will be essential to combine the advantages of each technique, observing internal structures at the nanoscale using EM and measuring molecular localization with super-resolution microscopy (CLEM; Taraska, 2019; Hoffman et al., 2020). Of course, dynamic intracellular structures such as lipid rafts and biomolecular condensates are also likely to affect molecular mobility (Sezgin et al., 2017; Chen et al., 2020). Thus, it will be key to overlay molecular mobilities from living cells over the static structural information of CLEM. We believe that combinations of multiple imaging modalities, along with modeling, will allow for a more in-depth understanding of synapses at the molecular level. These data will reveal how the elaborate architecture, density, and compartmentalization of subcellular components influence the highly tuned, dynamic, and changeable actions of synapses in the brain. 相似文献
Technique | Principle | Resolution | Comments | Applications in spines and synapses | |
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Lateral (XY) | Axial (Z) | ||||
CLSM | 250 nm | 500 nm | |||
TPLSM | Two-photon excitation | 350 nm | 700 nm | Deeper tissue penetration; adaptive optics further improve | Helmchen and Denk, 2005; Ji, 2017 |
STED | Stimulated emission (Vicidomini et al., 2018) | 20–70 nm | 500 nm | 3-D STED increases axial resolution; chronic in vivo imaging is possible | Nägerl et al., 2008; Berning et al., 2012; Pfeiffer et al., 2018 |
SIM | Moiré effect with structured illumination (Wu and Shroff, 2018) | 100 nm | 250 nm | No need for special fluorophores; limited resolution improvement | Kashiwagi et al., 2019; Li et al., 2020 |
SMLM (PALM, STORM) | Photoactivation, photoconversion (Baddeley and Bewersdorf, 2018) | 10–30 nm | 30–60 nm | High spatial resolution; temporal resolution is relatively worse | Dani et al., 2010; Tang et al., 2016 |
ExM | Physical expansion of sample (Wassie et al., 2019) | 4–20-fold improvement | 4–20-fold improvement | Capable of combining with other imaging techniques, only for fixed samples | Gao et al., 2019; Sarkar et al., 2020 Preprint |
Technique | Principle | Applications in spines and synapses |
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FRAP | Fluorescent molecules in a small region are photobleached, and subsequent movement of surrounding nonbleached fluorescent molecules into the photobleached area is monitored (Lippincott-Schwartz et al., 2018). | Svoboda et al., 1996; Bloodgood and Sabatini, 2005 |
FCS | Fluctuation of fluorescence intensity from the detection volume fixed at a specific position is recorded, and a temporal correlation is analyzed (Elson, 2011). | Chen et al., 2015; Obashi et al., 2019 |
RICS | Spatial correlation is analyzed from raster-scanned images (Digman and Gratton, 2011). | Obashi et al., 2019 |
SPT | The movement of a single particle is tracked using time-lapse imaging, and a trajectory is made and analyzed. To detect single particles, the density of fluorescence particles should be kept low (Shen et al., 2017). | Borgdorff and Choquet, 2002; Varela et al., 2016 |
SPT-PALM | Only a small number of photoactivatable fluorescent proteins in the field of view are activated and tracked until they are bleached (Manley et al., 2010). | Frost et al., 2010b; Nair et al., 2013 |
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Tomas E. van den Berg Rameez Arshad Wojciech J. Nawrocki Egbert J. Boekema Roman Kouil Roberta Croce 《Plant physiology》2020,184(4):2040
PSI is an essential component of the photosynthetic apparatus of oxygenic photosynthesis. While most of its subunits are conserved, recent data have shown that the arrangement of the light-harvesting complexes I (LHCIs) differs substantially in different organisms. Here we studied the PSI-LHCI supercomplex of Botryococccus braunii, a colonial green alga with potential for lipid and sugar production, using functional analysis and single-particle electron microscopy of the isolated PSI-LHCI supercomplexes complemented by time-resolved fluorescence spectroscopy in vivo. We established that the largest purified PSI-LHCI supercomplex contains 10 LHCIs (∼240 chlorophylls). However, electron microscopy showed heterogeneity in the particles and a total of 13 unique binding sites for the LHCIs around the PSI core. Time-resolved fluorescence spectroscopy indicated that the PSI antenna size in vivo is even larger than that of the purified complex. Based on the comparison of the known PSI structures, we propose that PSI in B. braunii can bind LHCIs at all known positions surrounding the core. This organization maximizes the antenna size while maintaining fast excitation energy transfer, and thus high trapping efficiency, within the complex.The multisubunit-pigment-protein complex PSI is an essential component of the electron transport chain in oxygenic photosynthetic organisms. It utilizes solar energy in the form of visible light to transfer electrons from plastocyanin to ferredoxin.PSI consists of a core complex composed of 12 to 14 proteins, which contains the reaction center (RC) and ∼100 chlorophylls (Chls), and a peripheral antenna system, which enlarges the absorption cross section of the core and differs in different organisms (Mazor et al., 2017; Iwai et al., 2018; Pi et al., 2018; Suga et al., 2019; for reviews, see Croce and van Amerongen, 2020; Suga and Shen, 2020). For the antenna system, cyanobacteria use water-soluble phycobilisomes; green algae, mosses, and plants use membrane-embedded light-harvesting complexes (LHCs); and red algae contain both phycobilisomes and LHCs (Busch and Hippler, 2011). In the core complex, PsaA and PsaB, the subunits that bind the RC Chls, are highly conserved, while the small subunits PsaK, PsaL, PsaM, PsaN, and PsaF have undergone substantial changes in their amino acid sequences during the evolution from cyanobacteria to vascular plants (Grotjohann and Fromme, 2013). The appearance of the core subunits PsaH and PsaG and the change of the PSI supramolecular organization from trimer/tetramer to monomer are associated with the evolution of LHCs in green algae and land plants (Busch and Hippler, 2011; Watanabe et al., 2014).A characteristic of the PSI complexes conserved through evolution is the presence of “red” forms, i.e. Chls that are lower in energy than the RC (Croce and van Amerongen, 2013). These forms extend the spectral range of PSI beyond that of PSII and contribute significantly to light harvesting in a dense canopy or algae mat, which is enriched in far-red light (Rivadossi et al., 1999). The red forms slow down the energy migration to the RC by introducing uphill transfer steps, but they have little effect on the PSI quantum efficiency, which remains ∼1 (Gobets et al., 2001; Jennings et al., 2003; Engelmann et al., 2006; Wientjes et al., 2011). In addition to their role in light-harvesting, the red forms were suggested to be important for photoprotection (Carbonera et al., 2005).Two types of LHCs can act as PSI antennae in green algae, mosses, and plants: (1) PSI-specific (e.g. LHCI; Croce et al., 2002; Mozzo et al., 2010), Lhcb9 in Physcomitrella patens (Iwai et al., 2018), and Tidi in Dunaliela salina (Varsano et al., 2006); and (2) promiscuous antennae (i.e. complexes that can serve both PSI and PSII; Kyle et al., 1983; Wientjes et al., 2013a; Drop et al., 2014; Pietrzykowska et al., 2014).PSI-specific antenna proteins vary in type and number between algae, mosses, and plants. For example, the genomes of several green algae contain a larger number of lhca genes than those of vascular plants (Neilson and Durnford, 2010). The PSI-LHCI complex of plants includes only four Lhcas (Lhca1–Lhc4), which are present in all conditions analyzed so far (Ballottari et al., 2007; Wientjes et al., 2009; Mazor et al., 2017), while in algae and mosses, 8 to 10 Lhcas bind to the PSI core (Drop et al., 2011; Iwai et al., 2018; Pinnola et al., 2018; Kubota-Kawai et al., 2019; Suga et al., 2019). Moreover, some PSI-specific antennae are either only expressed, or differently expressed, under certain environmental conditions (Moseley et al., 2002; Varsano et al., 2006; Swingley et al., 2010; Iwai and Yokono, 2017), contributing to the variability of the PSI antenna size in algae and mosses.The colonial green alga Botryococcus braunii (Trebouxiophyceae) is found worldwide throughout different climate zones and has been targeted for the production of hydrocarbons and sugars (Metzger and Largeau, 2005; Eroglu et al., 2011; Tasić et al., 2016). Here, we have purified and characterized PSI from an industrially relevant strain isolated from a mountain lake in Portugal (Gouveia et al., 2017). This B. braunii strain forms colonies, and since the light intensity inside the colony is low, it is expected that PSI in this strain has a large antenna size (van den Berg et al., 2019). We provide evidence that B. braunii PSI differs from that of closely related organisms through the particular organization of its antenna. The structural and functional characterization of B. braunii PSI highlights a large flexibility of PSI and its antennae throughout the green lineage. 相似文献
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Hai-Kuan Liu Yu-Jie Li Shu-Jie Wang Ting-Lu Yuan Wei-Jie Huang Xin Dong Jia-Qi Pei Dong Zhang Sheila McCormick Wei-Hua Tang 《Plant physiology》2020,184(4):1853
The rapid and responsive growth of a pollen tube requires delicate coordination of membrane receptor signaling, Rho-of-Plants (ROP) GTPase activity switching, and actin cytoskeleton assembly. The tomato (Solanum lycopersicum) kinase partner protein (KPP), is a ROP guanine nucleotide exchange factor (GEF) that activates ROP GTPases and interacts with the tomato pollen receptor kinases LePRK1 and LePRK2. It remains unclear how KPP relays signals from plasma membrane-localized LePRKs to ROP switches and other cellular machineries to modulate pollen tube growth. Here, we biochemically verified KPP’s activity on ROP4 and showed that KPP RNA interference transgenic pollen tubes grew slower while KPP-overexpressing pollen tubes grew faster, suggesting that KPP functions as a rheostat for speed control in LePRK2-mediated pollen tube growth. The N terminus of KPP is required for self-inhibition of its ROPGEF activity, and expression of truncated KPP lacking the N terminus caused pollen tube tip enlargement. The C-terminus of KPP is required for its interaction with LePRK1 and LePRK2, and the expression of a truncated KPP lacking the C-terminus triggered pollen tube bifurcation. Furthermore, coexpression assays showed that self-associated KPP recruited actin-nucleating Actin-Related Protein2/3 (ARP2/3) complexes to the tip membrane. Interfering with ARP2/3 activity reduced the pollen tube abnormalities caused by overexpressing KPP fragments. In conclusion, KPP plays a key role in pollen tube speed and shape control by recruiting the branched actin nucleator ARP2/3 complex and an actin bundler to the membrane-localized receptors LePRK1 and LePRK2.The delivery of nonmotile sperm to the embryo sac via a pollen tube is a key innovation that allowed flowering plants to carry out sexual reproduction without the need for water (Friedman, 1993; Lord and Russell, 2002). Both the speed and signal responsiveness of pollen tube growth are critical for successful fertilization (Johnson et al., 2019). The typical shape of a growing pollen tube cell protruding from a pollen grain is a cylinder with a dome-shaped tip (Geitmann, 2010). Maintaining such a typical tube shape during pollen tube growth is fundamental to support its ability for fast growth (Michard et al., 2017), and a plasticity range of tubular growth rates allows a pollen tube to optimize directional growth along its journey from the stigma to the ovule (Luo et al., 2017). The pollen tube cell extends mainly by tip growth, requiring huge amounts of secretion/exocytosis at the tip (McKenna et al., 2009; Grebnev et al., 2017). The newly secreted cell wall at the tip is mainly composed of esterified pectin, which is expandable, whereas cell wall remodeling at the lateral region (including pectin deesterification and callose deposition) limits expansion (Grebnev et al., 2017). The tip width of a growing pollen tube actually reflects the size of the secretion zone capped by an expandable membrane and cell wall, as a collective result of multiple pollen tube growth machineries (Luo et al., 2017).The tip-localized exocytosis of a growing pollen tube is supported by a spatiotemporal tightly controlled actin cytoskeleton network (Hepler, 2016). The actin cytoskeleton configuration in a pollen tube includes highly dynamic fine actin filaments in the apical and subapical regions and parallel longitudinal actin bundles in the shank region (Qu et al., 2017). Various actin-binding proteins, such as actin nucleation factors, actin-severing proteins, and actin-bundling factors, are responsible for organizing the dynamic actin cytoskeleton network (Ren and Xiang, 2007). For example, the actin-bundling proteins fimbrin and LIM (Lin-1, isl1, Mec3) domain-containing proteins function in shank-localized actin bundles in pollen tubes (Zhang et al., 2019). For another example, the actin nucleator formin (formin3 in Arabidopsis [Arabidopsis thaliana] and formin1 in lily [Lilium longiflorum]) functions in actin polymerization in the pollen tube tip (Li et al., 2017; Lan et al., 2018). The branched actin nucleator Actin-Related Protein2/3 (ARP2/3) complex is an evolutionarily conserved, seven-subunit complex consisting of the actin-related proteins ARP2 and ARP3 (Machesky et al., 1994). The ARP2/3 complex initiates the formation of branches on the side of preexisting actin filaments, locally creating a force-generating branched actin network that underlies cellular protrusion and movement (Blanchoin et al., 2000; Amann and Pollard, 2001; Molinie and Gautreau, 2018). The phenotypes of mutants in ARP2/3 in the moss Physcomitrella patens (Harries et al., 2005; Perroud and Quatrano, 2006), in Arabidopsis (Le et al., 2003; Li et al., 2003; Mathur et al., 2003; Brembu et al., 2004; Deeks et al., 2004), in maize (Zea mays; Frank and Smith, 2002), and in tomato (Solanum lycopersicum; Chang et al., 2019) demonstrated the broad importance of the ARP2/3 complex and its activation during cellular morphogenesis, including tip-growing cells. Perhaps surprisingly, in Arabidopsis, null ARP2/3 alleles are transmitted normally through pollen and there is no obvious root hair phenotype (Le et al., 2003; Djakovic et al., 2006).These cell growth machineries are tightly coordinated by multiple signaling pathways, including membrane-localized receptor kinases and Rho-of-Plants (ROP) GTPases (Li et al., 2018). The tomato pollen-specific and membrane-localized receptor kinases LePRK1 and LePRK2 mediate signaling during pollen tube growth (Muschietti et al., 1998). LePRK2 perceives several extracellular growth-stimulating factors, including a Cys-rich extracellular protein (Late-Anther-Specific52 [LAT52]), a Leu-rich repeat protein from pollen, and two pistil/stigma molecules, Style Interactor for LePRKs and Stigma-Specific Protein1 (Tang et al., 2002, 2004; Wengier et al., 2003, 2010), which increase the speed of pollen tube growth (Zhang et al., 2008b; Huang et al., 2014). LePRK2 antisense and RNA interference (RNAi) pollen tubes grow slower (Zhang et al., 2008b), consistent with a positive role for LePRK2 in regulating the speed of pollen tube growth. LePRK1 binds LePRK2 (Wengier et al., 2003), but LePRK1 plays a negative role in pollen tube growth by controlling a switch from a fast tubular mode to a slow blebbing mode (Gui et al., 2014). LePRK1 RNAi pollen tubes burst more often than wild-type pollen tubes, implicating a role for LePRK1 in maintaining plasma membrane integrity (Gui et al., 2014). An Arabidopsis paralog of these LePRKs, PRK6, also localized on the tip membrane, perceives Arabidopsis attraction cues from the female, AtLURE1s, to guide pollen tube growth (Takeuchi and Higashiyama, 2016; Zhang et al., 2017).Rho family small guanine nucleotide-binding proteins called ROPs or RACs, which can switch between a GDP-bound inactive form and a GTP-bound active form, are regulators of polar growth in pollen tubes (Cheung and Wu, 2008; Yang, 2008). In Arabidopsis, ROP1-dependent signaling controls tip growth. Active ROP1 defines a cap region in the apical plasma membrane as an exocytosis zone (Luo et al., 2017). Overexpression of ROP1 or of a constitutively active version resulted in pollen tube tip swelling (i.e. increased tip width) and slower growth (i.e. reduced tube length), while overexpressing a dominant negative version of ROP1 inhibited pollen tube growth (i.e. shorter but normal width tubes). The size of the pollen tube tip reflects the aggregate activity of membrane-associated ROP at the tip (McKenna et al., 2009; Luo et al., 2017). Tomato ROPs have been reported to be associated with the LePRK1-LePRK2 complex (Wengier et al., 2003) and therefore presumably play similar roles as the Arabidopsis homologs in pollen tube growth, yet their biological roles have not been directly investigated.Guanine nucleotide exchange factors (GEFs) activate ROPs by promoting the conversion of ROP/RAC GTPases from a GDP-bound inactive form to a GTP-bound active form. Plants possess a plant-specific ROPGEF family whose members contain a highly conserved GEF catalytic domain, the PRONE (plant-specific ROP nucleotide exchanger) domain (Berken et al., 2005; Gu et al., 2006). The intracellular portions of LePRK1 and LePRK2 interact with Kinase Partner Protein (KPP; Kaothien et al., 2005), whose Arabidopsis homologs were later shown to belong to the PRONE-type ROPGEF family (Berken et al., 2005; Gu et al., 2006). Pollen tubes overexpressing nearly full-length KPP (missing eight amino acids at the N terminus) developed swollen tips with abnormal cytoplasmic streaming and F-actin arrangements (Kaothien et al., 2005). An Arabidopsis homolog of receptor kinase, AtPRK2a (also named AtPRK2), interacts with AtROPGEF12 (Zhang and McCormick, 2007) and with AtROPGEF1 (Chang et al., 2013) to affect ROP activity. Based on the in vitro catalytic activity of full-length and truncated AtROPGEF1, an autoinhibition conferred by the C-terminal variable region was proposed (Gu et al., 2006). AtROPGEF12 was also shown to interact with the guidance receptor kinase PRK6 (Takeuchi and Higashiyama, 2016).Increased expression of full-length KPP increased the speed of pollen tube growth without significantly affecting pollen tube shape. We show biochemically that the PRONE domain of KPP does have ROPGEF activity on several class I ROPs, with highest activity on ROP4. The N-terminal domain of KPP inhibits its own GEF activity, while its C-terminal domain enhances its own GEF activity. The C-terminal domain of KPP is also required for its interactions with LePRK1, LePRK2, and an actin-bundling protein, Pollen-expressed LIM2a (PLIM2a), while the C-terminal domain alone is sufficient to bind LePRK1 but insufficient to bind LePRK2. Furthermore, self-associated KPP colocalized with the actin nucleation proteins ARP2/3 complex during pollen tube growth and enriched the membrane localization of ARP2/3 in the pollen tube. Interfering with ARP2/3 activation by coexpressing a dominant negative version of ARP2 reduced the speed of pollen tube growth and alleviated the defects caused by the overexpression of truncated KPP. CK-666, a specific small molecule inhibitor of ARP2/3 activation, canceled the promotive effect of full-length KPP on the speed of pollen tube growth. These results indicate that during pollen germination and tube growth, KPP not only links pollen receptor kinase and ROP signaling but also links the actin network to the pollen tube plasma membrane, thereby directly affecting the cellular morphology and efficiency of pollen tube growth. 相似文献
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Pannexin 1 (Panx1) plays a decisive role in multiple physiological and pathological settings, including oxygen delivery to tissues, mucociliary clearance in airways, sepsis, neuropathic pain, and epilepsy. It is widely accepted that Panx1 exerts its role in the context of purinergic signaling by providing a transmembrane pathway for ATP. However, under certain conditions, Panx1 can also act as a highly selective membrane channel for chloride ions without ATP permeability. A recent flurry of publications has provided structural information about the Panx1 channel. However, while these structures are consistent with a chloride selective channel, none show a conformation with strong support for the ATP release function of Panx1. In this Viewpoint, we critically assess the existing evidence for the function and structure of the Panx1 channel and conclude that the structure corresponding to the ATP permeation pathway is yet to be determined. We also list a set of additional topics needing attention and propose ways to attain the large-pore, ATP-permeable conformation of the Panx1 channel.IntroductionInitially, the pannexin field got off to an inauspicious start. Because the three proteins of the pannexin family (Panx1, Panx2, and Panx3) were discovered on the basis of their limited sequence homology to the invertebrate innexin gap junction proteins, it was assumed that they, too, would form gap junctions (Panchin et al., 2000; Bruzzone et al., 2003). However, notwithstanding some disputed science, it soon became clear that the gap junction function of pannexins is not realized (Dahl and Locovei, 2006; Huang et al., 2007; Sosinsky et al., 2011). Gap junction formation by pannexins is moreover prevented by glycosylation of the Panx1 protein (Boassa et al., 2007; Penuela et al., 2007; Boassa et al., 2008). It is conceivable, however, that the nonglycosylated form of Panx1 could allow the docking of oligomers in apposing membranes to each other. Indeed, such docking was observed in cryo-EM preparations of a mutant Panx1, N255A (Ruan et al., 2020). However, because glycosylation of the protein is required for membrane trafficking (Boassa et al., 2007; Penuela et al., 2007), it is unlikely that this process occurs in vivo. This notion is supported by the observation that a glycosylation mutant of Panx1 failed to lead to junctional conductance between paired oocytes, while expression of wtPanx1 followed by glycosidase F treatment before pairing of the cells resulted in gap junction formation, albeit at a much reduced rate as compared with the rate observed with connexins (Boassa et al., 2008). Furthermore, the gap junction function also is unlikely because of theoretical reasons and experimental findings, including expression in single cells, such as erythrocytes (Locovei et al., 2006a), and exclusive expression in the apical membrane of polarized epithelial cells (Ransford et al., 2009; Hanner et al., 2012; Shum et al., 2019). Of the three pannexins, only Panx1 has been studied in detail, and a clear channel function is well established for it. Therefore, this Viewpoint will exclusively deal with Panx1.It is generally assumed that an important function of Panx1 is to form ATP release channels. For example, as of this writing, a PubMed search with “pannexin” and “ATP release” as search terms yielded 287 publications. However, the biophysical evidence for this function is sparse, and the supporting evidence is largely correlative and/or based on pharmacological or genetic interference with ATP release. The ATP release function of Panx1 channels has been contested based on evidence that the channels are highly selective for Cl− and the lack of detectable ATP release under the experimental conditions, i.e., activation by voltage, used in these studies (Ma et al., 2012; Romanov et al., 2012).It was long held that the oligomeric state of the Panx1 channel is that of a hexamer (Penuela et al., 2013; Dahl, 2015; Chiu et al., 2018). Recently, in a <6-mo period, six research groups independently published similar cryo-EM structures of the Panx1 membrane channel (Deng et al., 2020; Jin et al., 2020; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020). All these papers challenge the view of the oligomeric state of the channel. Instead of the hexameric arrangement of identical Panx1 subunits, the channel appears to be formed by a homomeric heptamer.It is uncontested that the Panx1 channel can operate in a conformation in which the channel exhibits a unitary conductance of <100 pS. However, it has also been reported that, under certain experimental conditions, the channel can exhibit several subconductances with very rare sojourns to a maximal conductance of ∼500 pS (Dahl 2015).Activation of the Panx1 channel can be obtained by various stimuli, some physiological or pathological, others in the form of experimental tools. The majority of the physiological stimuli involve ligands, including ATP, glutamate, α adrenergic agonists, and bradykinin, binding to their cognitive receptors and leading to opening of Panx1 channels and “secondary” ATP release (Dahl 2015, 2018). A nonreversible activation of the channel is induced by cleavage with caspase 3 or 8 (Chekeni et al., 2010), leading to cell death. Experimental stimulation can be obtained in some cells but not in others by increasing the extracellular K+ concentration (Silverman et al., 2009; Wang and Dahl 2018; Chiu et al., 2018).Here, we present a point of view on these contested issues in the Panx1 field and attempt to reconcile apparently contradictory data or their interpretations. In addition, we ask to what extent the new structural data support the functional data.The structure of Panx1 channels as revealed by cryo-EMThe excitement for the long-awaited Panx1 structure was rewarded with not one but six publications, all appearing within the first half of 2020 and using cryo-EM to image the channel (Deng et al., 2020; Jin et al., 2020; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020). Five of the six Panx1 publications showed human Panx1 structures (Deng et al., 2020; Jin et al., 2020; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020), and two showed frog Panx1 structures (Deng et al., 2020; Michalski et al., 2020). To stabilize the frog Panx1 structure, thus improving the resolution for cryo-EM, Michalski et al. (2020) truncated the C terminus by 71 amino acids and removed 24 amino acids from the intracellular loop between transmembrane (TM) helices 2 and 3; this truncation was the version reported. The other publications reported full-length Panx1 structures, although one used a truncated Panx1 variant (Ruan et al., 2020) and another used a mutation (Jin et al., 2020) as their principal references. Because the C terminus–truncated structure and the (combined) WT structure had the highest overall quality of the several Panx1 structures presented by Ruan et al. (2020), it was used for de novo model building, as the reference structure for analyses, and for discussion. For the same reason, Jin et al. (2020) used the double mutation D376E/D379E, which eliminated the caspase-cleavage site, for their de novo model building. The overall architecture and dimensions of the human and frog Panx1 structures were similar, and the extracellular domains (ECDs) and transmembrane domains (TMDs) were nearly identical, except that the N-terminal helix (NTH) of the frog Panx1 is positioned on the intracellular side and not within the TMD as reported in human Panx1. These structures are becoming an important and critical tool to guide functional experiments and open a new venue to perform molecular dynamics simulations.The most novel finding shared by all the Panx1 structures is that instead of the hexamer hitherto believed to represent the native state of the Panx1 channel, the cryo-EM data show unequivocally a homo-heptamer arranged around a central symmetry axis that constitutes the principal permeation pathway. Panx1’s structural envelope adopts an inverted pail shape in the view parallel to the membrane. The heptameric channel is ∼110 Å long and ∼100 Å wide, with a flat ECD protruding ∼35 Å above the cell membrane and an intracellular domain (ICD) extending ∼35 Å into the cytoplasm with the TMD in between (Fig. 1). Both the N and C termini reside on the cytoplasmic side of the channel. Each TMD has four membrane-spanning helices per protomer. The TMD pore is predominantly lined by hydrophobic amino acids. The NTH is short and lines the TMD pore (human Panx1) with the NTH of one subunit interacting with the TMD of the adjacent subunit. Thus, it has been proposed that the NTH helps to maintain a rigid TMD pore (Ruan et al., 2020). Both the extracellular and intracellular pore entrances are lined with positively charged amino acids, making them favorable for negatively charged cargos such as Cl− and ATP to enter and leave the pore.Open in a separate windowFigure 1.Structural model of Panx1. (A) Cartoon representation of Panx1 WT (PDB accession no. 6wbk) stabilized with detergent. The approximate position of the membrane is indicated by the bars. One protomer of the heptameric assembly is indicated in orange. The lipids resolved in the structure are colored in magenta. It is notable that many of the resolved lipids are found at the interface between subunits. CTH, C-terminal helix; CLH, cytoplasmic linker helix. (B) Overlay of different Panx1 cryo-EM–based structures from different groups (PDB accession no. 6lto = gold, WT hPanx1; PDB accession no. 6wbk = blue, hPanx1 ΔC terminus, ΔN terminus; PDB accession no. 6v6d = lilac, WT hPanx1). The structures diverge very little in the ECD and the membrane domain. However, the variation is larger in the ICD. The approximate positions of F54 and I58 are indicated. (C) Comparison of the extracellular pore. Key residues are annotated using their one-letter abbreviation. All structures are shown as backbone traces except W74, which is part of the constriction ring of the pore. The narrowest constriction is indicated by the arrow.A cap structure is formed on the extracellular side from seven ECDs, one from each subunit. The ECDs are organized into the pore’s most constricted site, thus defining the maximum size of permeable molecules and establishing the extracellular entrance to the TMD. Each ECD contains one helix cross-linked through a disulfide bond to a three-stranded antiparallel β-sheet. The N-terminal end of each subunit helix protrudes inward toward the central pore axis contributing to the constriction site. The key residues involved in the gating of Panx1 are part of the W74-R75-D81 inter-subunit triplet (Fig. 1) from the N-terminal ends of the ECD helices and are proposed to provide rigidity to the extracellular entrance containing positive charges (Deng et al., 2020). This ring lining the wall of the constriction defines a hydrated pore diameter of ∼9 Å (Fig. 1) and is the crux of the argument that the extracellular constriction site is the effective selectivity filter discriminating cargoes on the basis of their charge and size. By contrast, the narrowest point of the pore in the TMD has a hydrated diameter of ∼13 Å.Other inter-subunit interactions that stabilize the pore arrangement in the TMD are (1) the F67-Y79 aromatic–aromatic and Q266-T252 hydrogen bonding in the ECD, (2) the TM helix TM1-TM1 and TM2-TM4 interfaces between adjacent subunits, (3) the N-terminal loop from the neighboring subunit extending into the TMD pore lining, and (4) the resolvable residues of C-terminal helix 3 association with the cytoplasmic linker helices 1 and 2 from the adjacent subunit (Fig. 1; Deng et al., 2020; Michalski et al., 2020). In addition, membrane lipids occupy the crevice between TM3 and an adjacent TM4 at the edge of the channel. Lipid head groups associate with positively charged residues from the N-terminal end of TM3 (K214) and the C-terminal domain of TM4 (R302 and K303; Deng et al., 2020). It may be that the lipid environment influences the assembly and possibly the activation of the Panx1 channel.The pore substantially widens toward the intracellular side. The ICDs extend away from the pore axis, producing a voluminous intracellular vestibule. Two helices in the ICD connect to two TMD helices to form a compact assembly that provides rigidity to the vestibule. The vestibule is slightly larger in the frog Panx1 structure, but the pore architecture is essentially the same as in the human version. The C-terminal segment consisting of ∼70 amino acids, including the caspase cleavage site located immediately before the C terminus, was not visualized in any of the published cryo-EM structures, suggesting that it is intrinsically flexible. C-terminal cleavage removes its autoinhibitory effect without inducing an overall conformational change to the pore (Jin et al., 2020).A novel feature presented by Ruan et al. (2020) is seven narrow side tunnels in the upper ICD running perpendicular to and ending at the pore, like a T-intersection. This tunnel network was proposed to allow passage of small anions because each tunnel contains several positively charged and other polar residues along its length. The flexible NTH-TM1 linker would function as a gate. They hypothesized that these side tunnels facilitate Cl− flux when the pore is blocked by the C terminus residing in the ICD vestibule. Movement or cleavage of the C terminus then would induce the ATP-permeable large-pore conformation. Further structural work is needed to determine whether the C terminus actually resides in the ICD vestibule. One complication with this tunnel hypothesis is that each tunnel has two constriction sites formed by R29 and A33, with diameters of 5.8 and 4.0 Å, respectively, meaning that partial dehydration of a Cl− ion is likely required for it to pass through the tunnel. The possibility of anion tunnels in the Panx1 structure raises the question: could there be multiple open (active) conformations, and does “open” mean open to ATP and Cl− ion or Cl− ion only? Three of the six papers (Michalski et al., 2020; Qu et al., 2020) assume that the Panx1 structure is in the closed or inactive conformation, whereas the other three report an open or active conformation. These discrepancies should be resolved with further work. Interestingly, the overlay of three of these structures, one presumably open and two closed, shows nearly identical conformations of the extracellular constriction of the pore (Fig. 1 C).The structures of Panx1 truncation and mutant variants, as well as its docked inhibitor, carbenoxolone (CBX), illuminate the roles of distinct structural elements. For example, cryo-EM revealed that CBX sits atop the ECD and plugs the W74-R75-D81 ring (Jin et al., 2020; Ruan et al., 2020), supporting pore blocking as its mechanism of Panx1 inhibition. Second, removal of the NTH did not much change the shape of the TMD. This truncation still generated a CBX-sensitive, voltage-dependent current similar to the full-length Panx1 current (Ruan et al., 2020), suggesting that the NTH may play a role in the assembly of the heptamer from the Panx1 protomers rather than being an essential element in the pore conduction pathway. Third, the Panx1 structures obtained in the presence of Ca2+ and K+ were indistinguishable from the untreated Panx1 structure, suggesting that neither Ca2+ nor K+ is likely to directly activate the Panx1 channel (Ruan et al., 2020). Fourth, the glycosylation-deficient mutant (N255A) generated gap junction structures as well as hemichannels as determined by cryo-EM (Ruan et al., 2020). The N255A gap junction is formed by two hemichannels docked by the interaction of the extracellular linker 2 on apposing ECDs. Ruan et al. (2020) noted that this Panx1 gap junction likely is not a normal physiological conformation.Comparing the Panx1 structure with other ATP-permeant pore structures offers a blueprint for dissecting pore properties of Panx1. The oligomeric configuration of the ATP-permeant channels does not appear to correlate with pore diameter, in particular when comparing the narrowest extent of the pore (Table 1). Like Panx1, CALHM1 was originally thought to adopt a hexameric configuration (Siebert et al., 2013) until the cryo-EM structure revealed an octameric configuration (Syrjanen et al., 2020). It should be noted that not all connexin channels transport ATP. For example, Cx26, Cx30, Cx43, and Cx46 transport ATP, but Cx32 does not (Harris, 2001; Hansen et al., 2014). An interesting comparison can be made between Panx1 and human Cx31.3. Cx31.3 has a pore with a constriction site diameter of ∼8 Å, similar to Panx1’s constriction of ∼9 Å, and also selectively transports Cl− and ATP (Lee et al., 2020). One difference between the two structures is that the constriction is in the ICD’s NTH for Cx31.3 and in the ECD for Panx1. Lee et al. (2020) pointed out that because Cx31.3’s pore constriction at the cytoplasmic entrance is smaller than the effective hydrated diameter of ATP (∼12 Å; Sabirov and Okada, 2004), the reported conformation would not allow ATP to pass through the pore without substantial conformational changes of the NTH. The same argument could be applied to Panx1’s ECD.Table 1.Comparison of oligomeric configuration with pore constriction diameter
Open in a separate windowIn a comparison of structural motifs, Panx1’s protomer shares a motif of four TMD helices with connexins, innexins, CALHM1 and 2, and volume-regulated anion channel (VRAC) protomers. They also share ECD topological similarities, but differences are manifested as well. For example, Panx1 and innexin6 have their ECD2 nestled between the inner and outer lobes of ECD1. CALHM1 and 2 and VRAC do not, and the outer lobe of ECD1 is absent in Cx46 (Myers et al., 2018; Deng et al., 2020; Flores et al., 2020; Qu et al., 2020). In Panx1 and VRAC, residues in the ECD1 helix form the pore constriction site, whereas in Cx46 and innexin6, the NTH forms the constriction in the middle of the TMD. Also, Panx1, innexin6, and CALHM2 have two ECD disulfide bonds, whereas Cx46 has three. The NTH of Panx1 is unique among ATP-permeant channels because it resides in the TMD pore close to the ECD, whereas the NTH in connexins and innexins is in the TMD closer to the ICD (Maeda et al., 2009; Myers et al., 2018; Flores et al., 2020), and the NTH in VRAC is in the ICD near the membrane face (Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018). In Panx1, connexins, innexins, and VRAC, TM1–TM4 are arranged anticlockwise as viewed from intracellular side, with TM1 nestled between TM2 and TM4, but the CALHM2 arrangement is clockwise and TM1 is attached to TM3. Finally, even though the ICD helix 1 is long, it is not involved in inter-subunit interactions, whereas in CALHM2, the equivalent cytoplasmic helix 1 is nearly parallel to the membrane and forms extensive inter-subunit interactions (Choi et al., 2019; Syrjanen et al., 2020).Biophysical properties and function of Panx1 channelsWhile the Panx1 structures obtained in six different laboratories are remarkably congruent, including the structures from different species, this is not the case for several functional aspects. It is now generally accepted that, contrary to original beliefs, pannexins do not form gap junctions despite their classification as “gap junction proteins” (Sosinsky et al., 2011). Instead, Panx1 is well documented to form a plasma membrane channel, which allows the exchange of solutes between the cytoplasm and the extracellular space.The first demonstration of Panx1 channel function involved the activation of the channel by positive membrane potentials in excess of +20 mV (Bruzzone et al., 2003). Robust Panx1-mediated membrane currents induced by positive membrane voltage were subsequently reported by various laboratories (Pelegrin and Surprenant, 2006; Iglesias et al., 2008; Iglesias et al., 2009b; Qiu and Dahl, 2009; Bunse et al., 2009; Prochnow et al., 2009; Silverman et al., 2009; Bunse et al., 2011; Gründken et al., 2011; Ma et al., 2012; Romanov et al., 2012; Zhan et al., 2012; Nomura et al., 2017). In oocytes expressing rodent Panx1, the currents are in the microampere range, and in these as in mammalian cells, the currents exhibit strong outward rectification.Weak Panx1 channel activation by positive membrane voltage was described, which could be boosted substantially by the insertion of a glycine-serine motif immediately after the first methionine (Michalski et al., 2018). Human and mouse Panx1 behaved similarly in these experiments, while mammalian cell lines (HEK and CHO) yielded different current densities for both pannexins. Frog Panx1 was subsequently found to exhibit similar voltage-sensitive properties (Michalski et al., 2020). In contrast, complete absence of voltage activation has been reported for human and frog Panx1, while mouse Panx1 was found to be weakly voltage sensitive (Chekeni et al., 2010; Chiu et al., 2018; Narahari et al., 2021).The first physiological role ascribed to Panx1 was that of a permeation pathway for ATP, allowing the efflux of ATP to act as an external signal to cells via activation of purinergic receptors (Bao et al., 2004). Subsequently, hundreds of publications affirmed the ATP release functions of Panx1 in multiple cell types based mainly on pharmacological and/or genetic interference. However, the role of Panx1 as an ATP-release channel has been challenged by data obtained with biophysical measurements. It has been shown that the voltage-activated Panx1 channel was highly selective for chloride ions and did not exhibit detectable ATP release (Ma et al., 2012; Romanov et al., 2012; Wang et al., 2014). On the other hand, several groups reported ATP release and/or activation of Panx1 currents after exposure of cells to high extracellular potassium ion (K+) concentrations (Silverman et al., 2009; Santiago et al., 2011; Heinrich et al., 2012; Suadicani et al., 2012; Michalski and Kawate, 2016; Qu et al., 2020). However, a lack of K+-mediated activation of Panx1 has also been reported (Chiu et al., 2018; Nielsen et al., 2020).Neither high positive membrane potentials nor high extracellular K+ concentrations can be considered physiological stimuli. A third stimulus for Panx1 activity, cleavage of C-terminal amino acids by caspase, occurs exclusively in apoptotic cells (Chekeni et al., 2010). A correlation of caspase cleavage with ATP release has been established (Chekeni et al., 2010; Boyd-Tressler et al., 2014; Imamura et al., 2020). However, whether caspase cleavage of Panx1 per se causes ATP release remains to be determined. The similarity of the electrophysiological profiles of the voltage-activated and the caspase-cleaved Panx1 channels (Chiu et al., 2017; Wang and Dahl, 2018) raises the question whether caspase by itself is sufficient for initiating ATP release.However, if we accept that physiological ATP release mediated by Panx1 occurs in a reversible fashion, as proposed in >200 publications, there must be an as-yet-unidentified gating mechanisms, since the activation mechanisms presently documented are either irreversible or unphysiological. Erythrocytes, for example, release ATP reversibly in a low oxygen environment but also in response to shear stress (Locovei et al., 2006a; Sridharan et al., 2010; Forsyth et al., 2011; Cinar et al., 2015; Zhang et al., 2018), two unrelated stimuli converging on the same ATP release channel Panx1.While it appears that the Panx1 field is riddled with controversies, the following paragraphs show that this is not necessarily so. It can safely be assumed that most, if not all, published data are correct. It may take only a little tuning of the interpretations to get a more coherent picture of the Panx1 field.Evidence for ATP release mediated by Panx1 channelsBased on the biophysical properties of the channel formed by Panx1, its function as an ATP release channel was proposed soon after its discovery (Bao et al., 2004; Dahl and Locovei, 2006; Locovei et al., 2006a). Ever since, supporting evidence has been accumulated by a large number of researchers, as documented in several reviews (Dahl and Locovei, 2006; Scemes et al., 2007; MacVicar and Thompson, 2010; Sosinsky et al., 2011; Penuela et al., 2013; Dahl, 2015). Key evidence includes the localization of Panx1 expression matching the site of ATP release at the apical membrane of polarized epithelial cells such as in the airways or renal tubules (Ransford et al., 2009; Hanner et al., 2012). Interestingly, in polarized cells releasing ATP at the basolateral membrane, this function is exerted by another release channel, CALHM (Taruno et al., 2013; Kashio et al., 2019). Additional evidence for the involvement of Panx1 in ATP release is that knockout of Panx1 expression attenuates ATP release (Iglesias et al., 2009a; Qiu et al., 2011; Qu et al., 2011; Suadicani et al., 2012). Furthermore, mutations of Panx1 and chemical modification of the protein affect ATP release (Wang and Dahl, 2010; Qiu et al., 2012).Evidence that Panx1 channels can be Cl− selective and lack ATP permeabilityThe ATP release function has been challenged, however, also based surprisingly on sound biophysical evidence (Ma et al., 2012; Romanov et al., 2012; Wang et al., 2014; Wang and Dahl, 2018). No ATP release was observed in voltage-clamped, Panx1-expressing HEK cells over the voltage range of −50 to +80 mV, despite robust CBX-sensitive currents at positive potentials (Romanov et al., 2012). Similarly, oocytes expressing Panx1 did not release ATP when the membrane potential was clamped at −60, 0, or +40 mV. However, CBX-sensitive ATP release was observed at the same holding potentials in the presence of high extracellular K+ concentrations (Fig. 2 A; Wang et al., 2014). Instead, the exclusively voltage-activated Panx1 channel was found to be highly selective for Cl−. Substitution of extracellular Cl− by larger anions resulted in attenuation of the membrane currents and a large shift of the reversal potential from negative to positive membrane potentials in both HEK cells and Xenopus laevis oocytes (Fig. 3, A, B, and D; Ma et al., 2012; Romanov et al., 2012; Chiu et al., 2014; Nomura et al., 2017; Li et al., 2018; Wang and Dahl, 2018). Thus, the Panx1 channel is highly selective for Cl− and lacks ATP permeability under the experimental conditions used in the referenced studies. Furthermore, in measuring the permeability of the Panx1 channel, it was found that substitution of extracellular Na+ by larger cations changed neither the reversal potential nor the amplitude of the currents in response to voltage steps or voltage ramps, indicating lack of cation permeability (Ma et al., 2012; Wang et al., 2018; Michalski et al., 2020).Open in a separate windowFigure 2.ATP release by oocytes expressing Panx1. (A) Oocytes expressing WT Panx1. All colored bars represent data obtained under voltage-clamp conditions; white bars are data from unclamped cells. ATP in the medium was measured as luciferase luminescence. ATP release from Panx1-expressing oocytes induced by potassium gluconate (KGlu) without and with holding the membrane potential under voltage-clamp conditions at −60, 0, or +40 mV, was determined 20 min after initiating the stimulus. Voltage-clamp conditions are indicated with teal lines below the graph and teal bars in the graph. The presence of 150 mM KGlu (K+ label) is indicated, and the data are displayed as hatched bars. Data are shown as means ± SD; n = 5 for each measurement. Adapted from Wang et al., 2014. (B) Oocytes expressing the truncation mutant Panx1Δ378. Oocytes were not voltage clamped (white bar) or were clamped at −60 or 0 mV (red bars). The cells were exposed to oocyte Ringer solution or to a solution containing 85 mM KGlu (K+ label, hatched bars) as indicated, for 10 min. An aliquot of the supernatant was analyzed for the presence of ATP with the luciferase/luciferin assay. Means ± SE; n as indicated above each bar. Adapted from Wang and Dahl, 2018. Because oocytes expressing Panx1Δ378 have a shortened life span (Jackson et al., 2014), measurements had to be taken in a short time window after injection of mRNA at 80× lower concentration than wtPanx1. Thus, ATP release data cannot be compared between wtPanx1- and Panx1Δ378-expressing cells.Open in a separate windowFigure 3.Current–voltage relations of mouse WT Panx1 and Panx1Δ378. (A) Replacing extracellular Cl− by gluconate− in a voltage step protocol applied to HEK cells expressing Panx1 exogenously attenuated the membrane currents and shifted the reversal potential to a positive potential. Adapted with permission from Journal of Cell Science (Romanov et al., 2012). (B) Similarly, in oocytes expressing Panx1 exogenously, replacement of Cl− by gluconate− (NaGluc) resulted in a shift of the reversal potential to positive values and an attenuation of the currents induced by a voltage ramp from −100 to +100 mV. (C) Voltage ramp–induced currents of wtPanx1-expressing oocytes in KCl solution (black trace) were substantially larger than the currents in uninjected control cells under identical conditions (gray trace). Replacing Cl− by gluconate− in the bath solution of the same oocyte resulted in attenuation of the currents and a shift of the reversal potential to more negative potential (green trace). (D) Quantitative analysis of reversal potentials after anion replacement shows a shift from positive to negative potentials with K+ as the extracellular cation. Means ± SE; n = 5 (wtPanx1). (E) Voltage ramp–induced membrane currents of WT Panx1 channels (green trace) and of channels formed by the truncation mutant Panx1Δ378, where C-terminal amino acids after aspartate 378 are deleted. Uninjected oocytes served as control (magenta trace). Because Panx1Δ378-expressing cells had a short life span, mRNA was injected 80× diluted as compared with WT, and measurements were performed in a 24-h window. Thus, the current amplitudes of WT Panx1 and Panx1Δ378 cannot be compared. In contrast to WT Panx1 channels, those formed by Panx1Δ378 were active over a wide voltage range, yet currents through both types of channels reversed at the same membrane potential. (F) Similar to WT Panx1, currents through Panx1Δ378 channels reversed at negative membrane potential with extracellular chloride (blue trace) and at positive potential with gluconate solution (red trace). (G) Similar to WT Panx1 channels, replacement of chloride by gluconate ions resulted in the currents reversing at negative potentials when Panx1Δ378-expressing cells were exposed to high extracellular K+. (H) Quantitative analysis of reversal potentials of membrane currents carried by Panx1Δ378 after anion replacement shows a shift from positive to negative potentials with K+ as the extracellular cation. Means ± SE; n = 4. (B–H) Data from Wang and Dahl (2018).Lack of cation permeability, however, appears to be restricted to the voltage-activated Panx1 channel. The only direct measure of ATP permeability presently available indicates that the ATP-permeable conformation of the Panx1 channel also is permeable to cations (Bao et al., 2004; Locovei et al., 2006a). In an isolated membrane patch subjected to a K+-ATP gradient, the channel currents reversed neither at the K+ nor the ATP2− reversal potential, but in between the two (closer to the K+ equilibrium potential). Furthermore, uptake of positively charged dyes is considered to be an acceptable surrogate for channel-mediated ATP release (Dahl, 2015; Johnson et al., 2016; Chiu et al., 2018). Thus, under conditions of ATP release, the channel does not discriminate on the basis of charge. However, since dye uptake can be mediated by various other mechanisms, including other membrane channels such as gasdermin D (de Vasconcelos et al., 2019) or CALHM (Siebert et al., 2013), dye uptake per se cannot be used as evidence for the involvement of Panx1.Can Panx1 be both a Cl−-selective and a nonselective channel?The Panx1 channel was activated by voltage when it was shown to have high selectivity for Cl− and to lack both cation and ATP permeability (Romanov et al., 2012; Ma et al., 2012; Wang et al., 2014; Wang and Dahl 2018). This activation was unphysiological, however, for the membrane potential had to be at high positive potentials for Panx1-mediated currents to be detectable. In contrast, all the evidence supporting the ATP release function involved alternative activation mechanisms. Several stimuli, including oxygen deprivation and mechanical stress, trigger channel-mediated ATP release through Panx1 channels (Locovei et al., 2006a; Thompson et al., 2006) This “primary” ATP release is complemented by a “secondary” ATP release (Dahl, 2018), in which a ligand binding to its cognizant receptor opens Panx1 and releases ATP. This process initially explained ATP-induced ATP release (Locovei et al., 2006b) and has now been demonstrated to apply to many other ligands and receptors, including NMDA, angiotensin II, and α adrenergic receptors (Thompson et al., 2008; Billaud et al., 2011; Murali et al., 2014). The function of secondary ATP release is to boost the cytoplasmic signal, which in all known cases is [Ca2+]i. This amplification is achieved by ATP binding to P2Y receptors, triggering further Ca2+ release from intracellular stores.We propose that the Panx1 channel has at least two distinct open conformations with different permeabilities. When activated by high positive voltage or when truncated at the caspase cleavage site, the channel is highly selective for Cl− and has no cation or ATP permeability. In contrast, it has been inferred that when opened in physiological settings by various stimuli, the channel is permeant to ATP and allows the TM flux of cationic and anionic dyes. The latter permeability properties can experimentally be induced by high extracellular K+ concentrations, as verified by changes in the reversal potential and high rate of ATP release. However, since activation by extreme voltage or by K+ is unphysiological, it seems prudent to reexamine with biophysical measurements the permeability properties of the Panx1 channel when activated physiologically, i.e., by ligand binding to receptors known to activate Panx1.Activation of Panx1 by extracellular K+ATP release in a variety of cell types can be stimulated by raising the extracellular K+ concentration; based on pharmacological interference, the release has been attributed to Panx1 channels (Bao et al., 2004; Qiu and Dahl, 2009; Silverman et al., 2009; Santiago et al., 2011; Heinrich et al., 2012; Suadicani et al., 2012; Wang et al., 2013). The ATP release shown in one of the published cryo-EM studies also used K+ to stimulate ATP release (Qu et al., 2020). Since an increase in extracellular K+ depolarizes the plasma membrane, it is logical to assume that depolarization is the trigger for the release (Heinrich et al., 2012). However, the shift in membrane potential induced by K+ should not be sufficient to bring the Panx1 channel into the voltage range where it is fully active. Indeed K+-induced Panx1-mediated currents were observed under voltage-clamp conditions over a wide voltage range (Silverman et al., 2009). The slow time course of the K+-induced Panx1 activation also speaks against depolarization as the critical step. Substitution of several amino acids in the extracellular loops of Panx1 by alanine modulated the response (Wang et al., 2018). Among the substitutions attenuating the K+ effect were amino acids W74 and R75, which are part of the external constriction observed in all published cryo-EM structures. Alanine substitution of amino acid D241 in the second extracellular loop resulted in amplification of the K+ response. Although the alanine substitution effects are suggestive for a direct action of K+ on Panx1 through binding and/or gating, the very slow activation of Panx1-mediated currents indicates that the mechanism responsible could be more complex. Additional evidence for K+-activation of Panx1 channels comes from the observation that currents carried by Panx1 were inhibited in a cysteine replacement mutant (Panx1T62C, C426S) by the thiol reagent 2-(trimethylammonium)ethyl methanethiosulfonate regardless of whether voltage or K+ served as stimulus (Wang et al., 2014).It has been shown that unpaired connexons also exhibit sensitivity to extracellular K+. For example, replacement of extracellular Na+ with K+ resulted in a reversible >10-fold potentiation of Cx50 hemichannel currents (Srinivas et al., 2006). As observed for Panx1 channels, the effect of K+ on Cx50 channels was observed under voltage-clamp conditions, excluding depolarization as a mechanism.It appears that the K+-induced activation of Panx1 may not work in all cell types. For example, Panx1 currents were induced in CHO cells by K+ (Michalski and Kawate, 2016), while HEK cells expressing Panx1 exogenously did not respond to increased extracellular K+ with an induction of CBX-sensitive currents (Chiu et al., 2018). The discrepancy in cell responses to K+ could be due to the involvement of additional factors, which could be facilitating in some cell types or inhibitory in others. The observation of a lack of K+ effect on the cryo-EM structure is consistent with such a scenario (Ruan et al., 2020). However, the solution of this problem could be much more trivial. In the experiments where K+ failed to activate Panx1 in cells, the extracellular solution contained millimole concentrations of Ca2+ (Chiu et al., 2018; Nielsen et al., 2020). Because divalent cations, including Ca2+, interfere with the K+ effect on Panx1 (Wang et al., 2018), a response would not be expected under these conditions.Whatever the detailed mechanism of the Panx1 channel activation by K+ may be, the Cl− selectivity of the channel is not evident in high extracellular K+ (Fig. 3, C and D). Replacement of extracellular Cl− by gluconate resulted in Panx1-mediated currents, which reversed at negative potentials instead of the reversal at positive potentials observed in the absence of the K+ stimulus. In high extracellular K+, the Panx1 channels were active over the whole voltage range from −100 to +100 mV, yielding membrane currents with amplitudes exceeding the endogenous oocyte currents by far (see also Fig. 3 C; Silverman et al., 2009; Wang et al., 2018).ATP release studies under voltage clamp conditions have shown that Panx1 channel activity induced by membrane depolarization does not result in ATP release (Romanov et al., 2012; Wang et al., 2014). However, ATP release was detected when the extracellular solution contained high K+ concentrations at negative, zero, or positive membrane potentials (Fig. 2 A).The extracellular [K+] required to activate Panx1 channels exceeds by far the physiological concentration of the ion. However, like the activation by voltage, the K+ stimulus is an experimentally convenient tool. More importantly, under pathological conditions, such as epilepsy, stroke, or central nervous system trauma, the extracellular [K+] can exceed 50 mM (Somjen, 1979; Gidö et al., 1997; Sick et al., 1998), thereby activating Panx1. Moreover, at these concentrations, K+ interferes with the negative feedback of ATP on Panx1 channel function (Qiu and Dahl, 2009; Qiu et al., 2012; Jackson et al., 2014). Alanine scanning mutagenesis suggests partially overlapping amino acids in the extracellular loops being involved in both activation of Panx1 channels by K+ and inhibition of the channel by ATP (Qiu et al., 2012; Wang et al., 2018).Activation of Panx1 by caspase cleavageA completely different activation for Panx1 channels has been discovered in apoptotic cells and involves the cleavage of 48 (mouse) or 47 (human) C-terminal amino acids by caspase (Chekeni et al., 2010). Certainly, this is an irreversible process associated with cell death. Nevertheless, this mechanism has found general acceptance as a physiological function of Panx1, despite the fact that in normal cells ATP release is reversible and consequently cannot involve cleavage of the C terminus. Because a deletion mutant mimicking the caspase cleaved Panx1 has no cytoplasmic plug of the sort observed in the WT Panx1 structure, this has been said to provide a structural basis for the cleaved Panx1’s ATP permeability (Ruan et al., 2020).With caspase cleavage, the channel becomes constitutively active, with a high open probability not only at positive potentials, but also at potentials as low as −100 mV (Chekeni et al., 2010; Jackson et al., 2014; Chiu et al., 2017; Chiu et al., 2018; Wang and Dahl, 2018). Fig. 3 E shows the current–voltage relationship of WT Panx1 channels and that of Panx1Δ378, a truncation mutant mimicking the caspase-cleaved Panx1 (Wang and Dahl, 2018). Panx1Δ378 was active over the whole voltage range of −100 to + 100 mV. Thus, caspase cleavage has a profound effect on channel gating by increasing the open probability at negative membrane potentials. In their original report, Chekeni et al. (2010) reported that apoptosis, ATP release, and caspase cleavage of Panx1 were correlated. However, it was not clear whether the anti-Fas stimulus for apoptosis itself caused the large pore conformation, before or while the channel was rendered constitutively active by the cleavage, which sealed the apoptotic fate of the cells.Subsequent data on the caspase-cleaved Panx1 channel published by the same group raised doubts about the ATP permeability of the truncated channel in the absence of another stimulus. They reported that the cleaved channel has the same selectivity for Cl− as the exclusively voltage-activated channel (Chiu et al., 2014). The voltage-activated channel, however, has been reported not to be ATP or cation permeable (Ma et al., 2012; Romanov et al., 2012; Wang et al., 2014). Because neither cleavage nor truncation at the caspase site affected the reversal potential (Chiu et al., 2017), it can be expected that the selectivity of the caspase-cleaved channel did not change either compared with the voltage-activated WT Panx1 channel.The lack of effect on the reversal potential is particularly puzzling in the experiments involving concatemers of the Panx1 protein. The concatemers were designed to allow sequential removal of C-terminal tails in a hexameric channel. As expected, this resulted in a “quantal” activation of the channel, with each cleavage of a protomer leading to an incremental increase in single-channel conductance (Chiu et al., 2017). The data also show a continuous increase in current density in whole-cell recordings as more caspase sites within the concatemer were used. Yet, the ATP release was discontinuous, requiring removal of more than two C termini. Thus, there was selective (Cl−?) current in the absence of ATP release. However, this change in selectivity was not reflected in the voltage ramps applied to the channels with variable numbers of cleaved C termini; all currents reversed at exactly the same potential. In these experiments, collecting periods of 4 to 8 h were used to determine ATP released into the supernatant, which are orders of magnitude longer than typically used in ATP release studies, attenuating the confidence level for interpretation of the measured values. During this time period, many events of cell division and apoptosis could have been the source of extracellular ATP. In contrast, ATP release by airway epithelial cells was detectable within seconds after a sudden osmotic stimulus, was reversible, and was attenuated by the Panx1 inhibitors CBX or probenecid (Ransford et al., 2009). Similarly, ATP release by Schwann cells induced by hypotonicity was detectable within 2 min after the stimulus, occurred in the absence of the cytoplasmic marker lactate dehydrogenase, and was blocked by the same Panx1 inhibitors (Wei et al., 2021). Furthermore, ATP release by oocytes expressing WT Panx1 or Panx1Δ378 was detectable in the unstirred supernatant as early as 5 min after a K+ stimulus (Wang and Dahl, 2018). Even in apoptotic cells, ATP release and that of metabolites was observed in a shorter time scale when a complex signaling chain was initiated by anti-Fas or UV treatment (Chekeni et al., 2010; Medina et al., 2020). Intriguingly, different sets of metabolites were released depending on the apoptosis stimulus. UV radiation and anti-Fas stimulation had only a small subset of metabolites, including ATP, in common, while the majority of metabolites released in response to the two stimuli did not overlap (Medina et al., 2020). This observation suggests that different signaling modalities modify the selectivity of the Panx1 channel or that other release mechanisms were involved concurrently.Both voltage- and caspase-activated channels have different unitary conductances at negative potentials (∼15 pS) as compared with positive potentials (∼90 pS; Ma et al., 2012; Romanov et al., 2012; Chiu et al., 2017). Since physiological ATP release occurs at the normal or slightly depolarized membrane potential, i.e., at negative potentials, the ∼15-pS conformation rather than the ∼90-pS channel would need to be responsible for ATP release. Permeability measurements with Panx1 truncated at the caspase cleavage site (Panx1Δ378) and involving extracellular ion replacement confirmed the Cl− selectivity and lack of cation permeability like the WT Panx1 channel (Fig. 3, F and H; Wang and Dahl, 2018). However, in the presence of extracellular K+, Cl− selectivity was attenuated (Fig. 3, G and H), while cation permeability and ATP permeability became apparent just as in WT Panx1 channels (Wang and Dahl, 2018).As observed for WT Panx1 channels, Panx1Δ378 channels did not release ATP under voltage-clamp conditions despite robust membrane currents (Wang and Dahl, 2018). Extracellular K+ not only boosted the membrane currents of Panx1Δ378 channels but also induced ATP release with the membrane potential clamped at −60 or 0 mV (Fig. 2 B; Wang and Dahl, 2018).Recently, data obtained with purified Panx1 protein in lipid bilayers and liposomes were interpreted in support of the view that caspase cleavage is sufficient to induce the ATP-permeable conformation of the Panx1 channel (Narahari et al., 2021). Indeed, the data show properties expected from Panx1 channels in the ATP-permeable conformation, including cation permeability and flux of ATP and of dyes with a preference of anionic over cationic dyes. However, the data also show that the caspase-activated Panx1 channel in lipid bilayers is distinct from the caspase-activated Panx1 channel in mammalian cells. In bilayers, three populations of channels were observed. One group of channels exhibited a single channel conductance of 100 pS, which is similar to the one observed in mammalian cells. The second group has been shown to dwell mainly in a 189-pS state. The third group did not require caspase cleavage and was observed only transiently after insertion of the protein into the lipid bilayer. The channels in this group exhibited variable conductance states with a maximal conductance of ∼400 pS.Another distinction of the Panx1 channel in lipid bilayers from the channel in mammalian cells is the lack of rectification. In mammalian cells, the caspase-cleaved Panx1 channel exhibits a 12-pS conductance at negative membrane potentials and a conductance of 96 pS at positive potentials (Chiu et al., 2017). In contrast, in lipid bilayers the voltage–current relationship was linear over a voltage range of −200 to +200 mV (Narahari et al., 2021). Thus, the disposition of the purified and caspase-cleaved Panx1 channel to allow the flux of ATP and other molecules is much more favorable in lipids than in mammalian cells. A channel with a conductance of ∼200 pS at the resting membrane potential or a depolarized potential consistent with physiological events is not the same as the 12-pS channel in mammalian cells. In the lipid bilayer, the channel is not only exposed to a different lipid environment, but it is also missing the protein environment prevailing in intact cells. It will be intriguing to see whether lipids or proteins prevent the transition from the 12-pS to the ∼200-pS channel conformation and how this is regulated in cells. Overall, the data obtained with purified Panx1 in an exclusive lipid environment support the hypothesis of more than one open conformation of the Panx1 channel.Thus, although caspase cleavage of Panx1 may not be sufficient for ATP release, this process does not interfere with other stimuli for the release. For example, in oocytes expressing Panx1 truncated at the caspase cleavage site, the channel was constitutively active, yielding membrane currents, which were boosted further by extracellular K+. Furthermore, ATP release via the truncated Panx1 was observed only with the additional K+ stimulus (Wang and Dahl, 2018). The ATP release observed in apoptotic/pyroptotic cells, therefore, may be the consequence of other stimuli. This release, however, plays a key role in the attraction of macrophages or microglia to injured or sick cells, a process working in vertebrate and invertebrate organisms (Chekeni et al., 2010; Samuels et al., 2010; Samuels et al., 2013). In the invertebrate nervous system, the ATP signal, rather than being a “find me” signal (Chekeni et al., 2010), appears to serve as a mobilization signal for microglial cells (Duan et al., 2009; Dahl and Muller, 2014).What is the oligomeric state of the Panx1 channel?Until recently, it was held that Panx1 oligomerizes to form a homomeric hexameric channel (Penuela et al., 2013; Chiu et al., 2018). This conclusion was based on cross-linking studies (Boassa et al., 2007; Chiu et al., 2017; Epp et al., 2019) and EM with negative staining of the isolated protein (Wang et al., 2014; Chiu et al., 2017). Acceptance of these findings was facilitated because a hexameric oligomeric state of Panx1 matched that of connexin “hemichannels” (the half of gap junction channels in one of the two opposing plasma membranes). Cross-linking studies in general suffer from both under- and overestimation of protein interactions. If the cross-linking agent is applied in very low concentrations and/or for an insufficiently long period, native interactions may not be captured fully, and using high concentrations for excessive periods of time can result in fortuitous linkages, resulting in overestimation of the oligomeric state. Interestingly, in one of the cross-linking studies on Panx1 (Epp et al., 2019), monomeric Panx1 predominated, followed by intermediate oligomers (dimers and trimers), while the apparent hexameric form was barely detectable. In this setting, heptamers, even if present, would have escaped detection because of an insufficient amount of protein in that oligomeric state. These observations, although consistent with incomplete cross-linking, also could indicate that the majority of Panx1 molecules are not assembled to a functional channel. Instead, the intermediate oligomers may only acutely form a functional channel upon a still-mysterious stimulus. Given the existence of dimers and trimers, the functional channel may be a hexamer (2 + 2 + 2 or 3 + 3), but also could be a heptamer (2 + 2 + 3) or higher oligomer.While the visualization of single-particle Panx1 channels by negative stain appears to support the hexameric state of the channel, the imposed sixfold symmetry used in both studies (Wang et al., 2014; Chiu et al., 2017) could have biased the outcome. However, the support for the hexameric arrangement of the Panx1 channel culminated in the use of concatemers of Panx1 forcing the formation of hexamers. Not only were these concatemers shown to form functional channels, but these channels exhibited biophysical properties essentially identical to those of WT Panx1 channels (Chiu et al., 2017). Thus, the channel formed by WT Panx1 ought to have the same oligomeric state as the one formed by the concatemer. It is conceivable that instead of forming the expected hexamer, one protomer of a concatemer would incorporate into the channel to form a heptamer with other concatemers. However, Chiu et al. (2017) emphasized the hexameric state with Western blots and the exact number of quantal steps predicted for a hexamer. However, like Panx1, CALHM1 was originally thought to adopt a hexameric configuration based on concatemers (Siebert et al., 2013) until the cryo-EM structure revealed an octameric configuration (Syrjanen et al., 2020).The oligomeric state of the Panx1 channel has been challenged in six publications, all appearing within the first half of 2020 and using cryo-EM to image the channel (Deng et al., 2020; Jin et al., 2020; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020). Instead of the hexamer hitherto believed to represent the native state of the Panx1 channel, the cryo-EM data show unequivocally a homo-heptamer forming the Panx1 channel. However, it would be desirable to have the heptameric arrangement be supported by independent methods. So far, the heptameric assembly supports the closed and the small, Cl−-selective conformations of the Panx1 channel, but not the large-pore conformation. It needs to be seen whether the ATP-permeable Panx1 channel has the same heptameric arrangement or whether higher-order oligomers need to form to render the channel ATP permeable.Do cryo-EM structures support an ATP release function of Panx1?In the Panx1 structure publications (Deng et al., 2020; Jin et al., 2020; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020), the reported narrowest width of the pore, or narrowest exclusion width of the closed conformation for the Panx1 structures, is remarkably uniform—8–9.4 Å. It is worth noting that the extracellular constriction has the same dimensions for all the reported structures, irrespective of whether the C or N terminus is cleaved off (Fig. 1 C shows three overlaid structures). For Panx1 to function as an ATP release channel, the pore must be at least the size of an ATP molecule with an effective hydrated diameter of ATP of ∼12 Å (Sabirov and Okada, 2004) and a hydrodynamic diameter of ATP of 15.4 Å (Rostovtseva and Bezrukov, 1998). Indeed, a surface model of ATP is a poor fit in the cryo-EM map (Fig. 4 A). As seen in a side view of the pore, the ATP molecule resides below the external constriction. The diameter of ATP (mol wt 507) exceeds that of the constriction, making a passage unlikely. As shown in Table 1, other ATP release channels exhibit a considerably wider pore. The situation for the tracer molecule Yo-Pro (mol wt 375) appears to be somewhat more favorable (Fig. 4 B). It is conceivable that, despite the tight fit, thermal fluctuations of the pore and the tracer molecule could enable the Yo-Pro molecule to squeeze through the pore at a low rate.Open in a separate windowFigure 4.Space-filling model of Panx1 without the C terminus. Related to PDB accession no. 6wbg. (A) Top: Top view of the extracellular pore. The space-filling model of ATP (PubChem accession no. 5957, mol wt 507.2) is shown in cyan. Bottom: Side view of the extracellular pore showing the W 74 ring above the ATP. (B) Top: The same model as in A, top view of the extracellular pore. In magenta is the space-filling model of Yo-Pro-1 (PubChem accession no. 6913121, mol wt 375.5). Bottom: Side views of the extracellular pore with a model for Yo-Pro (magenta) placed at the level of the external constriction.For reference, the minimal pore diameter of other ATP release channels is ≥14 Å (Table 1). The CALHM 1 channel, for example, is an octamer with a pore diameter of 19.5 Å at its narrowest part. Interestingly, CALHM 1 originally also was considered to be a hexamer (Ma et al., 2016), but cryo-EM data show that the channels formed by various CALHM isoforms are all higher-order oligomers, including octamers, decamers, and undecamers (Choi et al., 2019; Drożdżyk et al., 2020; Syrjanen et al., 2020). CALHM 4 even has been shown to exhibit two oligomeric states with different pore diameters of 20 and 30 Å (Drożdżyk et al., 2020).The heptameric Panx1 channel with an 8–9.4-Å constriction is a poor candidate to represent a channel conformation with high ATP permeability. It does, however, provide a sound structural basis for a Cl−-selective channel, as it is observed when the channel is activated by voltage or caspase cleavage (Ma et al., 2012; Romanov et al., 2012; Chiu et al., 2014; Wang and Dahl, 2018; Michalski et al., 2020). The external restriction has a width to accommodate Cl− to pass through the channel when it is unobstructed by other channel components. Furthermore, the presence of positively charged amino acids could be the basis for anion selectivity. Indeed, mutations of R75 support this notion. The anion selectivity was preserved if R75 was changed to K, while a change to A not only diminished the Cl− selectivity but also promoted cation permeability (Michalski et al., 2020). Because overlay of the structures of WT Panx1 and the truncated version mimicking the caspase-cleaved Panx1 did not reveal changes in the external constriction (Mou et al., 2020), it can be expected that both exhibit the same exclusion limits and Cl− selectivity (Fig. 1).Taken at face value, the presently available Panx1 channel structures do not support an ATP-release function of Panx1. Abundant functional data, on the other hand, are consistent with such a function. Therefore, one may consider that Panx1 exhibits an additional conformation not captured in the cryo-EM studies so far. How could a structure of the ATP-permeable Panx1 conformation be obtained? To boost the chance to observe the large-pore conformation of the Panx1 channel, it may be advantageous to activate Panx1 through ligand binding to G protein–coupled receptors, known to activate Panx1 in the process of secondary ATP release (Locovei et al., 2006b; Billaud et al., 2012; Murali et al., 2014; Dahl, 2018). For example, there is ample evidence that activation of Panx1 through P2X7 and NMDA receptors involves the action of the Src gene product on Panx1, likely by tyrosine phosphorylation of the Panx1 protein itself (Weilinger et al., 2012; DeLalio et al., 2019; Lohman et al., 2019).To test whether Panx1 phosphorylation per se is sufficient to drive the Panx1 channel to the open conformation, a Panx1 channel in an excised patch could be exposed to the active form of Src in the presence of ATP. If successful, the cryo-EM sample could be treated likewise to enhance the chance to capture the ATP-permeable conformation.Other soluble potential activators of Panx1 include K+ and Ca2+. Although micromolar concentrations of Ca2+ were observed to activate large channels in inside-out membrane patches excised from oocytes expressing Panx1 (Locovei et al., 2006b), activation of Panx1 by Ca2+ was not observed in HEK cells (Ma et al., 2009). In contrast, secondary ATP release in carotid body type II cells was found to be attenuated by the Ca2+ chelator BAPTA (1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid; Murali et al., 2014), indicating Ca2+ involvement in Panx1 activation. Furthermore, it has been proposed that the activation of Panx1 by the mechanosensitive Piezo channels involves cytoplasmic [Ca2+] (Cinar et al., 2015; Wang et al., 2016; Diem et al., 2020). Although K+ and Ca2+ failed to alter the cryo-EM structure (Ruan et al., 2020), it is not clear whether all possible iterations of the experimental approach have been exhausted.Alternatively, the Panx1-P2X7 complex is particularly interesting because these proteins interact physically and functionally (Pelegrin and Surprenant, 2006; Locovei et al., 2007; Iglesias et al., 2008; Silverman et al., 2009). Cryo-EM of the complex of Panx1 and P2X7 with and without ATP not only may allow the capture of the large pore conformation but may also yield clues about the gating mechanism. In both approaches, the ATP concentration has to be carefully chosen, since high concentrations of ATP were found to inhibit Panx1-mediated currents (Qiu and Dahl, 2009; Qiu et al., 2012).Contribution of the C terminus to the pore structureNone of the cryo-EM structures published so far has resolved the C terminus. Only one paper shows an amorphous mass at the cytoplasmic entry to the channel (Ruan et al., 2020). It is noteworthy that all cryo-EM structures presently available were obtained with C terminus–tagged Panx1. It is conceivable that the C terminus was misfolded because of the tags and consequently escaped resolution. Resolution of the C terminus is important, since functional data strongly indicate a decisive role of the C terminus in channel function, in terms of both gating and permeability (Chekeni et al., 2010; Chiu et al., 2014; Jackson et al., 2014). In the voltage-activated and Cl−-selective channel, the terminal cysteine moiety is reactive to externally applied thiol reagents, as indicated by attenuation of Panx1-mediated membrane currents (Wang and Dahl, 2010). Furthermore, an analysis of pore-lining moieties by the substituted cysteine accessibility method showed that cysteine replacements of several amino acids in the C terminus exhibited reactivity to thiol reagents (Wang and Dahl, 2010), indicating that this terminus protrudes from the cytoplasmic site far into the channel permeation pathway. Further support for this conclusion comes from the observation that the endogenous C-terminal cysteine C426 interacted with Panx1F54C via a disulfide bond (Sandilos et al., 2012). The F54C position was identified as a pore lining moiety together with other positions in the first TM segment and the first extracellular loop of Panx1 in the substituted cysteine accessibility method analysis (Wang and Dahl, 2010). F54 is located in the proximity of I58 (Fig. 1 B), which is part of a second constriction (13 Å in diameter) of the permeation pathway (Deng et al., 2020). This observation suggests that the WT Panx1 cysteine C426 would be located a few angstroms from the extracellular surface in the voltage-activated channel.Given that a large stretch of C-terminal amino acids fills the pore from the intracellular vestibule up to close to the extracellular restriction, one would expect that removal of the C terminus by caspase cleavage should affect the unitary conductance of the Panx1 channel to some extent. However, the WT Panx1 and truncated or caspase-cleaved Panx1 channels exhibit similar unitary conductances (Ma et al., 2012; Romanov et al., 2012; Chiu et al., 2014; Wang et al., 2014; Chiu et al., 2017)Removal of the C terminus by caspase cleavage in apoptotic cells or genetically by placing a stop codon at the cleavage site renders the channel constitutively active, with sojourns to open and closed states (Chekeni et al., 2010; Chiu et al., 2014; Chiu et al., 2017). However, since nonapoptotic ATP release is reversible in most cells, cleavage cannot be an obligate step to transform the Cl−-selective and cation-impermeant conformation to the large-pore conformation, which allows the flux of ATP and is also cation permeant. Thus, for nonapoptotic ATP release, instead of being cleaved, the C terminus has to undergo a major rearrangement to move out of the permeation pathway for adopting the ATP-permeable large-pore conformation. Indeed, in the large-pore conformation, the terminal cysteine was not further reactive to externally applied thiol reagents, while engineered cysteines at the external pore entry remained reactive (Wang et al., 2014). Energetically, the most favorable scenario would be a swinging out of the intact C terminus into a gap between TM helices created by a gating mechanism for the physiological, i.e., reversible activation of the large pore conformation.An overlay of truncated and WT Panx1 cryo-EM structures did not reveal differences in the extracellular pore structure (Fig. 1; Mou et al., 2020). Thus, cleavage of the C terminus of Panx1 may not be sufficient to alter the pore conformation, which is consistent with identical selectivities of channels formed by WT and truncated Panx1. Consequently, the structure of the truncated Panx1 channel cannot account for ATP permeability of the full-length channel.Ruan et al. (2020) suggested an intriguing hypothesis for the existence of different selectivities of the Panx1 channel. According to this hypothesis, the pathway for selective Cl− flux involves a side tunnel, while the main pore is blocked by the C terminus residing in the intracellular vestibule. Movement or cleavage of the C terminus then would provide the ATP-permeable large-pore conformation. However, the functional data listed above, including thiol reaction of the terminal cysteine (Wang and Dahl, 2010) and cross-linking of it with an engineered cysteine in a position close to the external constriction (Sandilos et al., 2012), indicate that the C terminus extends deep into the channel pore and thus are not consistent with a role of a side tunnel. Furthermore, the reported restriction at the extracellular end of the pore still would exclude ATP molecules.At present, the available information about the C terminus is restricted to functional data, since none of the published cryo-EM structures has this part of the protein resolved. The functional observations listed above suggest that the C terminus reaches from the intracellular site deep into the channel pore, probably occluding it in the closed state (Fig. 5 A). It can be hypothesized that at positive potential, the C terminus is rearranged (Fig. 5 B) so that the C-terminal cysteine moiety can react with extracellularly applied thiol reagents. The rearrangement of the C terminus allows mainly Cl− to pass through the external restriction depicted in the cryo-EM structures, which imposes Cl− selectivity. Alternatively, the C terminus is cleaved by caspase with the same consequences, plus the channel becoming constitutively active at negative and positive potentials (Fig. 5 C). Physiological stimuli, such as mechanical stress, directly or via Piezo (Cinar et al., 2015; Diem et al., 2020), initiate a gating mechanism, which widens the external constriction to accommodate the passage of ATP and render the channel nonselective for ions. In addition, the C terminus undergoes a conformational change, so that the terminal cysteine becomes inaccessible to thiol reagents (or a reaction does not affect membrane currents in this conformation). Extracellular K+ can mimic the effect of the physiological stimuli (Fig. 5 D). Cleavage of the C terminus then solidifies the apoptotic fate of the cell (Fig. 5 E). Alternatively, the caspase-cleaved Panx1 channel can be “hyperactivated” to render the Cl−-selective conformation to the nonselective large-pore conformation (Fig. 5, C–E).Open in a separate windowFigure 5.Activation mechanisms of Panx1 channels. (A) Based on the presently available structural and functional data, we hypothesize that the closed channel exhibits the external restriction of ∼9 Å and that the C termini reach deep into the pore, occluding it. (B) Depolarization dislocates the C termini, so that the channel pore diameter equals or exceeds that of the external restriction and exposes the terminal cysteine (yellow C) to thiol reagents. The yellow dot indicates the position of Panx1F54 which, when mutated to a cysteine, can be disulfide bonded with the terminal cysteine (Sandilos et al., 2012). In this configuration, the external restriction limits entry by size and selects for chloride (orange dot) over other ions. (C) Cleavage of Panx1 at position 378 removes the “pore gate,” rendering the channel constitutively active. However, as indicated by the published cryo-EM data, caspase cleavage does not affect the external constriction, leaving a chloride-selective channel. (D) Various physiological stimuli initiate a gating mechanism at the external restriction, widening it to accept ATP (green dot) and other molecules in this size range. (E) This mechanism also may move charged amino acids within the external constriction, with the consequence that charge selectivity is attenuated. In the case of K+ stimulation, the terminal cysteine is no further reactive to thiol reagents, while engineered cysteines at the external end of the pore still are. Whether this terminal cysteine concealment also applies to the physiological stimuli remains to be determined. Cleavage of the C terminus by caspase “super stimulates” the channel and irreversibly seals the apoptotic fate of the cell.In theory, an alternative mechanism to switch the Cl−-selective channel to an ATP-permeable conformation would be a change of the oligomeric state of the channel. Such a mechanism may operate in CALHM 4 channels, for which cryo-EM data suggest the coexistence of decameric and undecameric channels, which exhibit different pore diameters (Drożdżyk et al., 2020). Furthermore, other large-pore channels such as gasdermin appear to have variable oligomeric states (Mulvihill et al., 2018). No experimental evidence for such a mechanism operating for Panx1 channels is presently in existence, though.Problems to be solvedThe cryo-EM structures of the Panx1 channel published in rapid succession have raised a number of crucial questions. While they are consistent with the chloride-selective conformation of Panx1, they do not support an ATP-release function of this channel despite the abundance of evidence for such a role from functional data. The following points need to be addressed in future studies.Although not likely, it is possible that the heptameric organization of the Panx1 channel could be a consequence of the procedures involved in cryo-EM. It would be desirable to have this oligomeric state verified by independent methods.The published cryo-EM structures exhibit a restriction with a diameter of ∼9 Å, which would allow the passage of Cl− but would be too small for the permeation of ATP. Considering, that innexin channels (innexons) are octamers and CALHM channels can be octamers, decamers, or undecamers (Choi et al., 2019; Drożdżyk et al., 2020; Syrjanen et al., 2020), one should not a priori rule out that the large-pore conformation is a higher-order oligomer.For structural support of the ATP-release function of Panx1, a large-pore conformation is required with a pore dimension fitting the effective hydrated diameter of ATP of ∼12 Å (Sabirov and Okada, 2004) or a hydrodynamic diameter of ∼15.4 Å (Rostovtseva and Bezrukov, 1998). The structures published so far fall short of that.Further support for the ATP-release function of Panx1 could come from cryo-EM structures of Panx1 in the presence of ATP. ATP molecules located in a properly sized pore would represent the strongest evidence for the ATP-release function of this protein. Furthermore, extracellular ATP triggers a negative feedback mechanism inhibiting the channel (Qiu and Dahl, 2009; Qiu et al., 2012). This process involves the critical extracellular loop amino acids R75 and W74. Thus, ATP may be found docked to the pore entry.Because functional data strongly suggest that the C terminus of Panx1 can protrude deep into the channel pore, any functional conclusions from cryo-EM structures may be premature until the detailed C-terminal structure is resolved.Because several receptors associate (at least functionally) with Panx1 to induce secondary ATP release, a cryo-EM structure of receptor-Panx1 complexes may not only reveal the type of physical interaction but may also enhance the chance of capturing the large-pore conformation by including the respective receptor ligands.The puzzling observation that the forced hexameric channel has the same biophysical properties as the presumably heptameric WT channel needs a plausible explanation.The channels formed by WT Panx1 and by caspase-cleaved or truncated Panx1 have been reported to have the same unitary conductances. Why does removal of a “pore plug” (the C terminus) not affect unitary conductance of the channel?It is accepted that the voltage-activated Panx1 channel has a unitary conductance of 50–90 pS at positive potentials (Ma et al., 2012; Romanov et al., 2012; Wang et al., 2014; Chiu et al., 2018). However, what is the “unitary” conductance of the Panx1 channel in its large-pore conformation? Several laboratories have reported a ∼500-pS conductance for this channel conformation (Bao et al., 2004; Thompson et al., 2006; Kienitz et al., 2011; Orellana et al., 2011; Kurtenbach et al., 2013; Wang et al., 2014). Typically, in most recordings this large unitary conductance is a rare occurrence since the channel mainly dwells in one of the several subconductance states for extended periods of time. Thus, the term unitary can be misleading, and “maximal” conductance describes the situation better. However, a conductance of <100 pS at high positive membrane potentials and ∼15 pS at negative potentials has been proposed to be correlated with ATP release (Chiu et al., 2017). It is unclear, however, whether this lower conductance is actually a stable subconductance state or a record of the chloride-selective conformation. The latter is suggested by the similarity to the exclusively voltage-activated channel (Ma et al., 2012; Romanov et al., 2012; Wang et al., 2014). Reconstitution of the Panx1 protein into lipid bilayers yielded channel conductances of ∼1,000 pS (Mou et al., 2020). Whether this value is truly a single-channel conductance or whether the bilayer contained at least two active 500-pS channels remains to be determined.In summary, the structural models from cryo-EM are consistent with each other and with electrophysiological recordings that describe a Panx1 channel that is selectively permeable to Cl−. However, the models’ small predicted pore sizes are insufficient to explain the considerable body of data showing that Panx1 is permeable to ATP and cations both physiologically and in apoptotic cells. Our previous experiments have suggested that Panx1 may work in more than one conformation, and this Viewpoint proposes ways to find a structural evidence for this hypothesis. 相似文献
ATP-permeant channel | Oligomeric configuration | Pore constriction diameter (Å) | Reference |
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VDAC | Monomer (β barrel) | 25 | Colombini, 2012 |
Cx26 | Hexamer | 14 (open), 6 (closed) | Maeda et al., 2009; Nielsen et al., 2012 |
Cx31.3 | Hexamer | 8 (closed?) | Lee et al., 2020 |
Cx43 | Hexamer | 25 (open), 18 (closed) | Nielsen et al., 2012 |
Cx46 | Hexamer | 14 | Myers et al., 2018; Flores et al., 2020 |
VRAC | Hexamer | 12–14 (open), 2–5 (closed) | Deneka et al., 2018; Kasuya et al., 2018; Kefauver et al., 2018; Osei-Owusu et al., 2018 |
Panx1 | Heptamer | 9 | Deng et al., 2020; Jin et al., 2020; Michalski et al., 2020; Mou et al., 2020; Qu et al., 2020; Ruan et al., 2020 |
Innexin6 | Octamer | 18 | Oshima et al., 2016 |
CALHM1 | Octamer | 19.5 | Syrjanen et al., 2020 |
CALHM2 | Undecamer | 50 (open), 23 (inhibited) | Choi et al., 2019 |
CALHM4 | Decamer, undecamer | 20 (without current), 30 (without current) | Drożdżyk et al., 2020 |
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Yang Li Xuwei Duan Chuanbao Wu Jie Yu Chunsheng Liu Jing Wang Xiaoming Zhang Guohua Yan Feng Jiang Tianzhong Li Kaichun Zhang Wei Li 《Plant physiology》2020,184(4):1702
Recent studies have shown that loss of pollen-S function in S4′ pollen from sweet cherry (Prunus avium) is associated with a mutation in an S haplotype-specific F-box4 (SFB4) gene. However, how this mutation leads to self-compatibility is unclear. Here, we examined this mechanism by analyzing several self-compatible sweet cherry varieties. We determined that mutated SFB4 (SFB4ʹ) in S4′ pollen (pollen harboring the SFB4ʹ gene) is approximately 6 kD shorter than wild-type SFB4 due to a premature termination caused by a four-nucleotide deletion. SFB4′ did not interact with S-RNase. However, a protein in S4′ pollen ubiquitinated S-RNase, resulting in its degradation via the 26S proteasome pathway, indicating that factors in S4′ pollen other than SFB4 participate in S-RNase recognition and degradation. To identify these factors, we used S4-RNase as a bait to screen S4′ pollen proteins. Our screen identified the protein encoded by S4-SLFL2, a low-polymorphic gene that is closely linked to the S-locus. Further investigations indicate that SLFL2 ubiquitinates S-RNase, leading to its degradation. Subcellular localization analysis showed that SFB4 is primarily localized to the pollen tube tip, whereas SLFL2 is not. When S4-SLFL2 expression was suppressed by antisense oligonucleotide treatment in wild-type pollen tubes, pollen still had the capacity to ubiquitinate S-RNase; however, this ubiquitin-labeled S-RNase was not degraded via the 26S proteasome pathway, suggesting that SFB4 does not participate in the degradation of S-RNase. When SFB4 loses its function, S4-SLFL2 might mediate the ubiquitination and degradation of S-RNase, which is consistent with the self-compatibility of S4′ pollen.In sweet cherry (Prunus avium), self-incompatibility is mainly controlled by the S-locus, which is located at the end of chromosome 6 (Akagi et al., 2016; Shirasawa et al., 2017). Although the vast majority of sweet cherry varieties show self-incompatibility, some self-compatible varieties have been identified, most of which resulted from the use of x-ray mutagenesis and continuous cross-breeding (Ushijima et al., 2004; Sonneveld et al., 2005). At present, naturally occurring self-compatible varieties are rare (Marchese et al., 2007; Wünsch et al., 2010; Ono et al., 2018). X-ray-induced mutations that have given rise to self-compatibility include a 4-bp deletion (TTAT) in the gene encoding an SFB4′ (S-locus F-box 4′) protein, located in the S-locus and regarded as the dominant pollen factor in self-incompatibility. This mutation is present in the first identified self-compatible sweet cherry variety, ‘Stellar’, as well as in a series of its self-compatible descendants, including ‘Lapins’, ‘Yanyang’, and ‘Sweet heart’ (Lapins, 1971; Ushijima et al., 2004). Deletion of SFB3 and a large fragment insertion in SFB5 have also been identified in other self-compatible sweet cherry varieties (Sonneveld et al., 2005; Marchese et al., 2007). Additionally, a mutation not linked to the S-locus (linked instead to the M-locus) could also cause self-compatibility in sweet cherry and closely related species such as apricot (Prunus armeniaca; Wünsch et al., 2010; Zuriaga et al., 2013; Muñoz-Sanz et al., 2017; Ono et al., 2018). Much of the self-compatibility in Prunus species seems to be closely linked to mutation of SFB in the S-locus (Zhu et al., 2004; Muñoz-Espinoza et al., 2017); however, the mechanism of how this mutation of SFB causes self-compatibility is unknown.The gene composition of the S-locus in sweet cherry differs from that of other gametophytic self-incompatible species, such as apple (Malus domestica), pear (Pyrus spp.), and petunia (Petunia spp.). In sweet cherry, in addition to a single S-RNase gene, the S-locus contains one SFB gene, which has a high level of allelic polymorphism, and three SLFL (S-locus F-box-like) genes with low levels of, or no, allelic polymorphism (Ushijima et al., 2004; Matsumoto et al., 2008). By contrast, the apple, pear, and petunia S-locus usually contains one S-RNase and 16 to 20 F-box genes (Kakui et al., 2011; Okada et al., 2011, 2013; Minamikawa et al., 2014; Williams et al., 2014a; Yuan et al., 2014; Kubo et al., 2015; Pratas et al., 2018). The F-box gene, named SFBB (S-locus F-box brother) in apple and pear and SLF (S-locus F-box) in petunia, exhibits higher sequence similarity with SLFL than with SFB from sweet cherry (Matsumoto et al., 2008; Tao and Iezzoni, 2010). The protein encoded by SLF in the petunia S-locus is thought to be part of an SCF (Skp, Cullin, F-box)-containing complex that recognizes nonself S-RNase and degrades it through the ubiquitin pathway (Kubo et al., 2010; Zhao et al., 2010; Chen et al., 2012; Entani et al., 2014; Li et al., 2014, 2016, 2017; Sun et al., 2018). In sweet cherry, a number of reports have described the expression and protein interactions of SFB, SLFL, Skp1, and Cullin (Ushijima et al., 2004; Matsumoto et al., 2012); however, only a few reports have examined the relationship between SFB/SLFL and S-RNase (Matsumoto and Tao, 2016, 2019), and none has investigated whether the SFB/SLFL proteins participate in the ubiquitin labeling of S-RNase.Although the function of SFB4 and SLFL in self-compatibility is unknown, the observation that S4′ pollen tubes grow in sweet cherry pistils that harbor the same S alleles led us to speculate that S4′ pollen might inhibit the toxicity of self S-RNase. In petunia, the results of several studies have suggested that pollen tubes inhibit self S-RNase when an SLF gene from another S-locus haplotype is expressed (Sijacic et al., 2004; Kubo et al., 2010; Williams et al., 2014b; Sun et al., 2018). For example, when SLF2 from the S7 haplotype is heterologously expressed in pollen harboring the S9 or S11 haplotype, the S9 or S11 pollen acquire the capacity to inhibit self S-RNase and break down self-incompatibility (Kubo et al., 2010). The SLF2 protein in petunia has been proposed to ubiquitinate S9-RNase and S11-RNase and lead to its degradation through the 26S proteasome pathway (Entani et al., 2014). If SFB/SLFL in sweet cherry have a similar function, the S4′ pollen would not be expected to inhibit self S4-RNase, prompting the suggestion that the functions of SFB/SLFL in sweet cherry and SLF in petunia vary (Tao and Iezzoni, 2010; Matsumoto et al., 2012).In this study, we used sweet cherry to investigate how S4′ pollen inhibits S-RNase and causes self-compatibility, focusing on the question of whether the SFB/SLFL protein can ubiquitinate S-RNase, resulting in its degradation. 相似文献
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