Chromosome segregation during female meiosis in C. elegans: A tale of pushing and pulling

The role of the kinetochore during meiotic chromosome segregation in C. elegans oocytes has been a matter of controversy. Danlasky et al. (2020. J. Cell. Biol. https://doi.org/10.1083/jcb.202005179) show that kinetochore proteins KNL-1 and KNL-3 are required for early stages of anaphase during female meiosis, suggesting a new kinetochore-based model of chromosome segregation.

Meiosis consists of two consecutive chromosome segregation events preceded by a single round of DNA replication. Homologous chromosomes are separated in meiosis I, which is followed by sister chromatid separation in meiosis II to produce haploid gametes. Both of these stages require chromosomes/chromatids to align during metaphase before separating to opposite poles during anaphase. During mitosis, microtubules emanating from centrosomes at opposite poles of the cell bind chromosomes through a multiprotein complex called the kinetochore, allowing chromosomes to be pulled apart (1, 2). This segregation event takes place in two stages: anaphase A, where chromosomes are pulled toward spindle poles due to microtubule depolymerization, and anaphase B, where spindle poles themselves move farther apart, taking the attached chromosomes with them (3, 4). In many organisms, including mammals, oocytes lack centrosomes, and it has been of great interest to clarify the mechanisms used to ensure chromosomes are properly segregated during female meiosis (5, 6). Caenorhabditis elegans has served as a model for studying both mitosis and meiosis, but the mechanisms operating during female meiosis have been a matter of debate and controversy.In 2010, Dumont et al. showed that the kinetochore is required for chromosome alignment and congression during metaphase (7). However, they suggested that chromosome segregation was the result of microtubule polymerization between the segregating chromosomes (Fig. 1), resulting in a pushing force exerted onto chromosomes toward the spindle poles in a largely kinetochore-independent manner (7). This mechanism was also supported by the finding that CLIP-associated protein (CLASP)–dependent microtubule polymerization between the segregating chromosomes is essential for chromosome separation (8). An alternative model suggested that chromosomes are transported through microtubule-free channels toward the spindle poles by the action of dynein (9). Later evidence put in doubt a role for dynein and favored a model in which chromosomes initially separate when the spindle shortens and the poles overlap with chromosomes in an anaphase A–like mechanism. This is then followed by separation of chromosome-bound poles by outward microtubule sliding in an anaphase B–like fashion (10). However, because microtubules emanating from the spindle poles are not required to separate the homologous chromosomes but microtubules between the separating chromosomes are (8), this model is unlikely, at least as an explanation for mid-/late-anaphase movement. Furthermore, although lateral microtubule interactions with chromosomes predominate during metaphase of C. elegans oocyte meiosis, cryo-electron tomography data described end-on attachments between the separating chromosomes as anaphase progresses (11). This led to the suggestion that lateral microtubule interactions with chromosomes are responsible for the initial separation, but microtubule polymerization between the separating chromosomes is required for the later stages of segregation (11). The mechanisms involved in this initial separation have remained obscure. In this issue, Danlasky et al. show that the kinetochore is in fact required for the initial stages of chromosome segregation during female meiosis—an important step forward in our understanding of the mechanisms governing acentrosomal chromosome segregation (12).Open in a separate windowFigure 1.Some of the key findings in Danlasky et al. Kinetochore proteins surround the outer surface of the chromosomes, resulting in a characteristic cup shape. As anaphase progresses, chromosomes come into close contact to the spindle poles (anaphase A). Chromosome stretching is provided by KNL-1, MIS-12 (KNL-3), and NDC-80 (KMN)–dependent forces. Once the spindle starts elongating (anaphase B), central spindle microtubules provide the pushing forces for chromosome segregation. At this stage, kinetochore proteins also occupy the inward face of separating chromosomes. Upon KMN network depletion, bivalents flatten, and chromosome congression and alignment are defective. Anaphase A chromosome movement is almost absent, which leads to error-prone anaphase B.By simultaneously depleting kinetochore proteins KNL-1 and KNL-3 in C. elegans, Danlasky et al. observed the meiotic chromosome congression and alignment defects described in previous studies (7). However, this double-depletion phenotype displayed three key characteristics that suggested a role for kinetochores in chromosome segregation, which are discussed below.The kinetochore is required for bivalent stretching. It was previously shown that the bivalent chromosomes stretch before the initiation of segregation (10). Danlasky et. al found that this stretching of the chromosomes did not occur when KNL-1,3 were depleted, indicating that the kinetochore is required for this process (Fig. 1). Together with the observation that kinetochore proteins appear to extend toward the spindle poles, this finding suggested that pulling forces resulting from the interaction between the kinetochore and spindle microtubules are occurring during metaphase/preanaphase (Fig. 1).The kinetochore is required for anaphase A. In C. elegans female meiosis, anaphase A occurs when homologous chromosomes begin to separate during spindle shortening, and anaphase B when the chromosomes separate alongside the spindle poles (10). Danlasky et al. observed that KNL-1,3 depletion drastically reduced the velocity of anaphase A, as chromosomes only separated when spindle poles began to move apart. This indicated that pulling forces caused by the interaction between the kinetochore and spindle microtubules are also important for the initial separation of homologous chromosomes in anaphase A.The kinetochore is required for proper separation of homologous chromosomes. In KNL-1,3 depletion strains, 60% of bivalents failed to separate before segregation began, resulting in intact bivalents being pulled to the same spindle pole (Fig. 1). This failure of homologous chromosomes to separate was not thought to be a result of KNL-1,3 depletion interfering with the cleavage of cohesin that holds the two homologous chromosomes together because (a) separase and AIR-2^AuroraB, both of which are required for cohesin cleavage, localized normally during metaphase and anaphase, and (b) bivalents separated by metaphase II. This leaves the possibility open that the failure of bivalents to separate was due to the disrupted pulling forces thought to be important in bivalent stretching and anaphase A.Altogether, these data strongly indicate that the kinetochore is required not only for chromosome congression and alignment but also for the early stages of homologue separation. Anaphase B occurred successfully in the absence of KNL-1,3 but was more error prone, likely as a result of the earlier congression and anaphase A defects. While it is clear that chromosome masses do segregate in the absence of the kinetochore, this segregation is highly erroneous as a result of defects during the earlier stages of segregation in anaphase A (Fig. 1).The findings of Danlasky et al. raise testable hypotheses that could significantly enhance our understanding of acentrosomal chromosome segregation. Further investigation of the proposed pulling forces required during metaphase and early anaphase will be of great interest. Additionally, a more detailed analysis of the dynamic localization of separase and Securin, as well as assessing successful cohesin cleavage when KNL-1,3 are depleted, would back up the assertion that the failure of homologous chromosomes to separate was not due to the kinetochore impacting cohesin cleavage. It has previously been shown that the CLASP orthologue CLS-2 in C. elegans localizes to the kinetochore surrounding the bivalent chromosomes during metaphase before relocalizing to the central spindle during anaphase (7, 8, 13). It will be interesting to examine whether this key microtubule-stabilizing protein contributes to anaphase A pulling forces alongside its essential role in microtubule polymerization between chromosomes in anaphase B (8).While the regulation of proper chromosome segregation during acentrosomal meiosis in C. elegans is not yet fully understood, Danlasky et al.’s results represent a significant step forward in this endeavor by showing that the kinetochore is in fact required for the early stages of chromosome segregation.     

Straightening up is required to nucleate new microtubules

In this issue, Ayukawa, Iwata, Imai, and colleagues (2021. J. Cell Biol. https://doi.org/10.1083/jcb.202007033) use rapid temporal and high-spatial-resolution electron microscopy imaging to examine the earliest stages of new microtubule nucleation. They discover that straightening of curved tubulin oligomers increases the efficiency of microtubule nucleation.

Microtubules are long cytoskeletal filaments that provide structure to cells, allow for transport within the cells, and participate in cell division by acting together with molecular motors to build a mitotic spindle. While microtubules act as the stiff “bones” of the cell, they also have a unique and important ability to rapidly restructure their length and organization in response to cellular cues. Therefore, large arrays of microtubules, such as in a mitotic spindle, can rapidly depolymerize and disappear as needed. However, in order to rebuild these microtubule networks, the nucleation of new microtubules is required. This nucleation of new microtubules, whether from existing templates such as centrosomes or spindle poles, or via the de novo organization of the tubulin subunits that make up microtubules, remains a poorly understood process.In this issue, Ayukawa, Iwata, Imai, and colleagues used rapid temporal and high-spatial-resolution imaging to study the earliest stages of microtubule nucleation (1). First, the authors purified αβ-tubulin heterodimers with a Y222F mutation in the β-tubulin subunit. This Y222F β-tubulin mutation increased the rate of microtubule assembly and, importantly, greatly accelerated the nucleation rate of new microtubules. Thus, a comparison between wild-type and mutant tubulin allowed the authors to dissect differences in the early nucleation process that could explain the increased nucleation rate for the mutant tubulin. Importantly, a “rapid flush method” was used to capture high-resolution transmission electron microscopy images of tubulin subunits very early in the nucleation process. The rapid flush method revealed “oligomers” of tubulin subunits: chains of tubulin subunits linked together along their long axis.The authors reasoned that these oligomers are likely on-pathway intermediates that are crucial for new microtubule nucleation. Examination of the oligomers revealed differences in curvature between wild-type and mutant tubulin: the mutant tubulin that nucleated more readily had oligomers that were straighter and less curled than the wild-type tubulin. Further, a comparison of GTP tubulin and GDP tubulin early nucleation intermediates revealed that, for both wild-type and mutant tubulin, the GDP tubulin oligomers, which did not nucleate efficiently, were more curved than the GTP tubulin oligomers that nucleated more efficiently (Fig. 1 A, left, red versus blue). Importantly, the fraction of nearly straight oligomers in each sample directly corresponded to the respective nucleation rate for each tubulin type. Thus, the degree of curvature of the oligomers predicted the overall nucleation rate, such that an increase in the fraction of straight oligomers was directly correlated to an increase in the microtubule nucleation rate (Fig. 1 A).Open in a separate windowFigure 1.Long, straight oligomers promote microtubule nucleation. (A) The nucleation of new microtubules is limited by the availability of critical-length, straight GTP tubulin (blue) oligomers. GDP tubulin oligomers (red) are more curved, with reduced nucleation efficiency. (B) Straight GTP tubulin protofilaments (blue) that are attached to templates (gray) could also be required to facilitate the nucleation of new microtubules from templates such as in centrosomes. Curved GDP tubulin protofilaments (red) attached to templates (gray) would not efficiently facilitate nucleation of new microtubules.The authors then examined the role of oligomer length in the nucleation rate, to determine whether there was a critical minimum oligomer length that could predict efficient microtubule nucleation. They fit their bulk microtubule growth curves (turbidity) to a standard nucleation-and-growth model (2) to estimate the minimum size of the oligomers that were likely to grow into microtubules, i.e., the critical length. For wild-type tubulin, ∼4 tubulin dimers were required in order for oligomers to grow into microtubules. However, this critical length was common, and indeed prevalent, within the early nucleation mixtures. Therefore, it seems likely that oligomer curvature, rather than length, is the limiting factor that slows the microtubule nucleation rate (Fig. 1 A).Why would oligomer curvature limit microtubule nucleation rate? While oligomers of sufficient length were readily observed in early nucleation mixtures, lateral association of new oligomers with existing oligomers was rare. Further, when multiple oligomers had indeed associated laterally to form a new, multi-protofilament assembly, the length of the longest strand greatly exceeded the maximum size of the single-stranded oligomers. Thus, it is likely that straight oligomers facilitate the lateral association of new tubulin subunits or oligomers along their length, stabilizing the nascent microtubules and allowing for their stable growth as a multistranded filament.While this work sheds light on the nucleation of new microtubules using purified tubulin, the described results provide interesting insights into potential nucleation mechanisms inside of cells. In cells, microtubules predominately grow from templates, such as centrosomes, and various microtubule-associated proteins may also influence the nucleation process. A recent study of microtubule nucleation from templates found that there was a significant time lag between the arrival of new tubulin subunits to a template and the growth of a new microtubule from the template (3). That study concluded that GTP hydrolysis inhibits microtubule nucleation by destabilizing the nascent microtubule. These results are consistent with the conclusion from Ayukawa, Iwata, Imai, and colleagues that GDP tubulin oligomers are curved and therefore unable to efficiently make lateral associations with other oligomers to stabilize the new microtubule and allow for growth (Fig. 1 B; 1). However, recent electron microscopy studies have also revealed that the GTP tubulin–containing ends of growing microtubules show extended protofilaments with a gentle curvature (4, 5, 6, 7, 8). Thus, one additional barrier to the nucleation of new microtubules on templates may be the straightening of GTP tubulin oligomers. Here, the straightening of template-attached, gently curved GTP tubulin oligomers would then allow for lateral binding of new tubulin dimers and oligomers (Fig. 1 B). The transient straightening of gently curved GTP tubulin oligomers could potentially be accomplished via thermal forces and the resulting protofilament curvature fluctuations. However, this straightening process could also be facilitated by microtubule-associated proteins that promote nucleation, such as TPX2 and XMAP215 (9, 10). In support of this idea, Ayukawa, Iwata, Imai, and colleagues found that the distribution of oligomer curvatures appeared similar to what has been reported for the protofilaments at the growing ends of microtubules (5).In light of these results, interesting future work could explore whether microtubule-associated proteins act to regulate the curvature of the oligomers involved in microtubule nucleation. While many aspects of the mechanisms of microtubule nucleation remain unknown, this new work sheds light on the very earliest stages of microtubule nucleation, which may have wide-ranging implications in future studies of microtubule nucleation and growth in cells.     

The role of molecular diffusion within dendritic spines in synaptic function

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

Genetics of Sexual Development: An Evolutionary Playground for Fish

Teleost fishes are the most species-rich clade of vertebrates and feature an overwhelming diversity of sex-determining mechanisms, classically grouped into environmental and genetic systems. Here, we review the recent findings in the field of sex determination in fish. In the past few years, several new master regulators of sex determination and other factors involved in sexual development have been discovered in teleosts. These data point toward a greater genetic plasticity in generating the male and female sex than previously appreciated and implicate novel gene pathways in the initial regulation of the sexual fate. Overall, it seems that sex determination in fish does not resort to a single genetic cascade but is rather regulated along a continuum of environmental and heritable factors.IN contrast to mammals and birds, cold-blooded vertebrates, and among them teleost fishes in particular, show a variety of strategies for sexual reproduction (Figure 1), ranging from unisexuality (all-female species) to hermaphroditism (sequential, serial, and simultaneous, including outcrossing and selfing species) to gonochorism (two separate sexes at all life stages). The underlying phenotypes are regulated by a variety of sex determination (SD) mechanisms that have classically been divided into two main categories: genetic sex determination (GSD) and environmental sex determination (ESD) (Figure 2).Open in a separate windowFigure 1Reproductive strategies in fish. Fish can be grouped according to their reproductive strategy into unisexuals, hermaphrodites, and gonochorists. Further subdivisions of these three categories are shown with pictures of species exemplifying the strategies. Fish images: Amphiprion clarkii courtesy of Sara Mae Stieb; Hypoplectrus nigricans courtesy of Oscar Puebla; Scarus ferrugineus courtesy of Moritz Muschick; Astatotilapia burtoni courtesy of Anya Theis; Poecilia formosa and Kryptolebias marmoratus courtesy of Manfred Schartl; Trimma sp. courtesy of Rick Winterbottom [serial hermaphroditism has been described in several species of the genus Trimma (Kuwamura and Nakashima 1998; Sakurai et al. 2009; and references therein)].Open in a separate windowFigure 2Sex-determining mechanisms in fish. Sex-determining systems in fish have been broadly classified into environmental and genetic sex determination. For both classes, the currently described subsystems are shown.Environmental factors impacting sex determination in fish are water pH, oxygen concentration, growth rate, density, social state, and, most commonly, temperature (for a detailed review on ESD see, e.g., Baroiller et al. 2009b and Stelkens and Wedekind 2010). As indicated in Figure 2, GSD systems in fish compose a variety of different mechanisms and have been reviewed in detail elsewhere (e.g., Devlin and Nagahama 2002; Volff et al. 2007).The GSD systems that have received the most scientific attention so far are those involving sex chromosomes, which either may be distinguishable cytologically (heteromorphic) or appear identical (homomorphic). In both cases, one sex is heterogametic (possessing two different sex chromosomes and hence producing two types of gametes) and the other one homogametic (a genotype with two copies of the same sex chromosome, producing only one type of gamete). A male-heterogametic system is called an XX-XY system, and female-heterogametic systems are denoted as ZZ-ZW. Both types of heterogamety exist in teleosts and are even found side by side in closely related species [e.g., tilapias (Cnaani et al. 2008), ricefishes (Takehana et al. 2008), or sticklebacks (Ross et al. 2009)]; for more details on the phylogenetic distribution of GSD mechanisms in teleost fish, see Mank et al. (2006). Note that sex chromosomes in fish are mostly homomorphic and not differentiated (Ohno 1974), which is in contrast to the degenerated Y and W chromosomes in mammals (Graves 2006) and birds (Takagi and Sasaki 1974), respectively. This is one possible explanation for the viable combination of different sex chromosomal systems within a single species or population of fish (Parnell and Streelman 2013) and could be a mechanistic reason why sex chromosome turnovers occur easily and frequently in this group (Mank and Avise 2009). Additionally, fish can have more complex sex chromosomal systems involving more than one chromosome pair (see Figure 2). Even within a single fish species, more than two sex chromosomes may occur at the same time, or more than two types of sex chromosomes may co-exist in the same species (Schultheis et al. 2006; Cioffi et al. 2013), which can sometimes be due to chromosome fusions (Kitano and Peichel 2012).Detailed insights on the gene level for GSD/sex chromosomal systems are currently available for only a limited number of fish species, and all but one of these cases involve a rather simple genetic system with male heterogamety and one major sex determiner (see below). The only exception is the widely used model species zebrafish (Danio rerio), which has a polyfactorial SD system implicating four different chromosomes (chromosomes 3, 4, 5, and 16) (Bradley et al. 2011; Anderson et al. 2012) and also environmental cues (Shang et al. 2006).In this review, we focus on newly described genetic sex-determining systems and possible mechanisms allowing their emergence in fishes, which are the most successful group of vertebrates with ∼30,000 species.     

Synapses have autophagy under control

Regulation of autophagy in neurons remains unclear. In this issue, Kulkarni et al. (2021. J. Cell Biol. https://doi.org/10.1083/jcb.202002084) show with elegant live imaging that in dendrites, but not in axons, autophagosome motility and function is regulated by synaptic activity.

Macroautophagy is a type of autophagy that refers to the capacity to form double membrane compartments called autophagosomes that engulf large protein aggregates and defective organelles. Autophagosomes fuse with lysosomes, forming degradative autolysosomes (1). Autophagosome formation depends on the conjugation of LC3-I (cytosolic) to phosphatidylethanolamine, generating LC3-II, which remains bound to autolysosomes (1). In neurons, inactivation of autophagy genes impacts neurodevelopment, axon growth and guidance, synapse formation and pruning, ultimately leading to neurodegeneration. Particularly, in motor neurons and cerebellum Purkinje cells, autophagy gene knockout leads to the accumulation of intracellular protein aggregates and degeneration, impacting movement coordination (1). Interestingly, stimulation of memory up-regulates autophagy, and while reducing autophagy reduces memory, activating it has the opposite effect on memory (2). What triggers macroautophagy in neurons remains unclear. In this issue, Kulkarni et al. test whether synaptic activity regulates autophagy and detail the impact of synaptic activity on autophagosome motility (3).Kulkarni et al. used multiple strategies to manipulate synaptic activity. They stimulated synaptic activity by depolarizing neurons with high potassium, treating them with a cocktail of antagonists of voltage-gated potassium channels and inhibitory gamma-aminobutyric A receptors, and using uncaging of the excitatory neurotransmitter glutamate. To inhibit synaptic activity, the researchers treated neurons with antagonists of excitatory α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid and N-methyl-D-aspartate receptors (4). To image autophagosomes and autolysosomes (here globally termed autophagic vacuoles [AVs]) in live neurons, the authors expressed LC3 tagged with fluorescent proteins. They elegantly imaged the same neuronal compartment before and after depolarization, or under basal, increased, or reduced synaptic activity, and used kymograph analysis (via Kymoanalyser; 5) to quantify the mean speeds of AVs in both dendrites and axons. An increase in intracellular calcium measured with a genetically encoded calcium sensor, GCAMP3, indicated synaptic activity. Kulkarni et al. observed that, in dendrites, AVs stop with synaptic activity and move with synaptic inhibition (Fig. 1). This AV movement change was swift and unaltered by co-culture with astrocytes, and reversible. One key finding is that this change in AV movement occurred in dendrites, but not in axons. Interestingly, AVs stopped at or near synapses, which were identified with PSD-95-GFP.Open in a separate windowFigure 1.In dendrites, AVs stop at synapses upon synaptic activity.The authors further characterized the AVs in terms of acidity (lysotracker labelling of acidic organelles) and of degradative capacity (DQ-BSA fluorescence accumulation upon lysosomal degradation). Lysotracker motility changed similarly with synaptic activity. Interestingly, the lysotracker density increased with synaptic stimulation. The higher number of acidic organelles (likely autolysosomes) indicated increased autophagy or acidification with synaptic activity, which could underlie increased degradative activity. Indeed, about half of the LC3-positive AVs were degradative in dendrites, while in axons there was virtually no degradative AV, supporting the requirement for transport to the soma for degradation of autophagic cargo (6). Finally, Kulkarni et al. show that degradative AVs increase with synaptic activity, correlating with the reduced motility of LC3-positive AVs.An intriguing observation is that the autophagic vacuoles identified by LC3-mCherry were virtually all positive for LAMP1, a marker of late endosomes and lysosomes, indicating that dendrites mainly contain autolysosomes and no or very few autophagosomes (LC3-positive and LAMP1-negative) and late endosomes/lysosomes (LC3-negative and LAMP1-positive). One is left wondering if it results from LC3 overexpression and overflooding to interconnected organelles. An alternative possibility is that LC3 may not always label autophagosomes, in which case complementary electron microscopy is necessary for confirmation. Where are dendritic autolysosomes formed? In axons, a fraction of the LC3 autophagic vacuoles was LAMP1 negative, and the formation of autophagosomes at axon terminals has been well documented (7). Thus, do autophagosomes form in axons, fuse with LAMP1-positive late endosomes/lysosomes, and only after are they transported to dendrites? Alternatively, autophagosomes may form in dendrites and fuse with late endosomes/lysosomes, preventing their detection unless fusion is inhibited (8).Another interesting observation concerns the similar change in the motility of early endosomes, identified by Rab5, an early endosome GTPase, with synaptic activity. Other organelles, post-ER vesicles (4), and proteasomes (9) similarly display a change in motility in dendrites upon synaptic activity. In contrast, mitochondria stop moving in axons with synaptic activity (10). The significance of this arrest of several dendritic organelles with synaptic activity is an attractive area for research.Neuronal autophagy dysfunction is implicated in many neurodegenerative diseases (1). At least early in the disease, increasing autophagy improves neuronal function and synapse activity (1). Genetic risk factors include lysosomal proteins, whose defective function leads to the accumulation of nondegraded autophagic vacuoles and recapitulate neurodegenerative phenotypes (11). Lysosomal dysfunction is a mechanism of cellular aging. Moreover, synapses become dysfunctional with aging and lost in neurodegenerative diseases (12). Based on this study, synapse dysfunction and thus reduced synaptic activity could increase AV motility and reduce acidification and the degradative capacity of autolysosomes. Similarly, neuronal overexcitability, as in early Alzheimer''s disease patients with seizures, could cause excessive AV motility and degradative activity.What is the mechanism that stops AV movement? Do early endosomes, secretory vesicles, or proteasomes change motility using similar mechanisms? For post-ER vesicles, the CAMKII dependent phosphorylation of the microtubule motor Kif17 was sufficient to arrest movement (4). Alternatively, could it be the actin cytoskeleton that forms patches in the dendritic shaft at the base of postsynaptic glutamatergic synapses to halt microtubule-dependent transport of organelles (13)? More work is needed to tackle these questions and define the cell biological mechanisms by which synaptic activity controls AV function and dynamics in different neuronal compartments. Understanding the mechanisms underlying the regulation of autophagy and autophagosome maturation and degradation provides an exciting opportunity for therapeutic development in neurodegenerative diseases.     

Silencing the spindle assembly checkpoint: Let’s play Polo!

Silencing of the spindle assembly checkpoint involves two protein phosphatases, PP1 and PP2A-B56, that are thought to extinguish checkpoint signaling through dephosphorylation of a checkpoint scaffold at kinetochores. In this issue, Cordeiro et al. (2020. J. Cell Biol. https://doi.org/10.1083/jcb.202002020) now show that a critical function of these phosphatases in checkpoint silencing is removal of Polo kinase at kinetochores, which would otherwise autonomously sustain the checkpoint.

The main goal of mitosis is to accurately segregate chromosomes, such that each daughter cell inherits a full complement of genetic information. To accomplish this delicate task, once each chromosome is faithfully duplicated through DNA replication, its identical sister chromatids must attach to spindle microtubules coming from opposite spindle poles through a process known as chromosome biorientation. Kinetochores are proteinaceous assemblies that reside at the centromeric region of chromosomes and are key to this process by capturing spindle microtubules (1). Microtubule capture, however, is inherently error prone, and several cycles of attachment/detachment are often required before chromosomes achieve biorientation. Obviously, chromosome segregation without error correction would be highly detrimental, leading to unbalanced chromosome numbers, referred to as aneuploidies, which are hallmarks of cancer and genetic diseases. Luckily, eukaryotic cells not only possess an error-correction machinery deputed to rectify faulty attachments (2), but they also have a safeguard device, called the spindle assembly checkpoint (SAC), that temporarily halts cells in mitosis to provide them with the necessary time window to fix the errors. SAC signaling fires at unattached kinetochores, which are continuously generated during error correction, and is extinguished once all chromosomes are bioriented, thus resuming mitotic progression and chromosome segregation (3).Prevailing models posit that a key trigger of SAC signaling is the phosphorylation of the kinetochore scaffold KNL1 by the SAC kinase MPS1. This creates a phospho-docking site at the MELT repeats (amino acid consensus Met-Glu-Leu-Thr) of KNL1 that recruits the heterotetrameric SAC complex BUB1:BUB3:BUB3:BUBR1 (referred to as BUB complex; Fig. 1), which in turn attracts to the kinetochore other SAC proteins that collectively prevent mitotic progression (3).Open in a separate windowFigure 1.The interplay of SAC kinases and phosphatases at kinetochores. When SAC is activated at an unattached kinetochore (SAC on), MPS1 phosphorylates the kinetochore scaffold KNL1, thereby recruiting the BUB complex. Contextually, CDK1-dependent phosphorylation of BUB1 and BUBR1 generates phospho-docking sites for recruitment of the Polo kinase PLK1, which on one side sustains KNL1 phosphorylation and on the other stimulates BUBR1 binding to PP2A-B56. The latter, in turn, counteracts PLK1 local activity by dislodging PLK1 from the kinetochore. During SAC silencing, local activity of MPS1 is shut off. Additionally, the PP1 phosphatase binds to KNL1 and, together with PP2A-B56, further evicts PLK1 from the kinetochore, possibly through dephosphorylation of its phospho-docking sites in BUB1 and BUBR1. This leads to KNL1 dephosphorylation and displacement of the BUB complex, thus extinguishing SAC signaling (SAC off). Whether PP1 and PP2A-B56, as opposed to other phosphatases, contribute directly to KNL1 dephosphorylation remains an open question.The protein phosphatases PP1 and PP2A-B56 are recruited to kinetochores through binding to KNL1 and BUBR1, respectively, and are thought to silence the SAC through dephosphorylation of the MELT repeats of KNL1, thus antagonizing MPS1 activity (Fig. 1). Additional mechanisms, such as MPS1 inhibition and stripping of SAC components from kinetochores, have been proposed to contribute to obliterate SAC signaling upon chromosome biorientation (4).In this issue, compelling evidence from Cordeiro et al. challenges the current view by showing that rather than dephosphorylating KNL1, PP1 and PP2A-B56 actually silence the SAC by down-regulating the activity of Polo kinase (PLK1 in human cells) at kinetochores (5). Polo kinase and MPS1 share a common substrate preference and both can phosphorylate the MELT repeats of KNL1. Additionally, Polo cooperates with MPS1 in SAC signaling in various species, while in organisms where MPS1 is absent, like nematodes, Polo functionally replaces MPS1 (6).Consistent with previous results (7, 8), Cordeiro et al. show that when kinetochore phosphatases are dampened, PLK1 levels increase at kinetochores through an unknown mechanism, which might involve dephosphorylation of the phosphoepitopes in the Polo-binding motifs generated on the BUB complex by CDK1 (BUB1-pT609 and BUBR1-pT620; 9, 10, 11). This implies that when PP1 and PP2A-B56 are low at kinetochores, PLK1 can amplify SAC signaling through a positive feedback loop by boosting KNL1 phosphorylation independently of Mps1, thereby recruiting the BUB complex and, in turn, increasing amounts of PLK1 (Fig. 1). In agreement with this view, in a sensitized setup where kinetochore phosphatases are crippled along with MPS1, concomitant inhibition of PLK1 is sufficient to bring about KNL1 dephosphorylation and restore SAC signaling. These data led the authors to the provocative conclusion that the primary role of PP1 and PP2A-B56 in SAC silencing is to harness PLK1 activity. This new model is appealing not only because it highlights a novel function for PP1 and PP2A-B56 in SAC silencing, but also because it explains the modest effects that are commonly observed on SAC signaling upon PLK1 inhibition alone. Indeed, kinetochore phosphatases, and primarily PP2A-B56 (12), are already partially active in a SAC-induced mitotic arrest (e.g., upon microtubule depolymerization), as shown here by the increased KNL1 phosphorylation upon their inactivation.Interestingly, sequence alignment of BUB1 and BUBR1 homologues across the phylogenetic tree reveals that, in metazoans, putative Polo-binding motifs are usually located in the vicinity of hypothetical PP2A-B56–binding motifs, suggesting that they coevolved. The physical proximity of Polo-binding and PP2A-B56–binding motifs in BUB1 and BUBR1 could position the Polo-binding motifs in a favorable arrangement for their PP2A-B56–driven dephosphorylation and, as a consequence, PLK1 clearance from kinetochores (Fig. 1).The data by Cordeiro et al. represent a paradigm shift in our understanding of SAC silencing for two main reasons. First, consistent with published data (13, 14), PP1 and PP2A-B56 might be involved in this process primarily by inactivating upstream SAC kinases (MPS1 and PLK1), rather than dephosphorylating their substrates. Second, since PLK1 is partially displaced from kinetochores by the above phosphatases already during a SAC arrest, MPS1 inactivation might be the main trigger of SAC silencing. Several mechanisms have been proposed to attenuate MPS1 activity once the SAC is satisfied, such as MPS1 displacement from kinetochores (6) and dephosphorylation of MPS1 in its activation loop (13, 14).The new model raises a burning question: If PP1 and PP2A do not dephosphorylate KNL1 at MELT repeats, what does? Other phosphatases, whose identity remains elusive, could be involved in KNL1 dephosphorylation. Alternatively, phosphorylated KNL1 might be actively turned over at kinetochores. Nevertheless, at present, the involvement of PP1 and PP2A-B56 in KNL1 dephosphorylation cannot be ruled out, as complete inhibition of kinetochore phosphatases in the experimental setup used here is likely very challenging. Further investigations will be required to solve this central issue.Another important question that deserves further scrutiny is, how exactly do PP1 and PP2A-B56 inhibit PLK1 activity at kinetochores? Cordeiro et al. propose that they could dephosphorylate the Polo-binding motifs in BUB1/BUBR1. Alternatively, the close proximity of PP2A- and Polo-binding motifs in metazoan BUBR1 homologues could make the association of PLK1 and PP2A with BUBR1 mutually exclusive.Finally, and most importantly, what is the physiological meaning of the complex interplay between SAC kinases and phosphatases described here? A crucial function of PLK1 bound to the BUB complex in human cells is to stabilize kinetochore-microtubule attachments in prometaphase by recruiting PP2A-B56 through phosphorylation of the PP2A-B56-binding motif in BUBR1 (15). In turn, eviction of PLK1 from kinetochores by PP2A-B56 will have two major outputs: (i) maintain microtubule dynamics at bioriented chromosomes (8) and (ii) stimulate binding of PP1 to KNL1, which primes the system for SAC silencing (16). As soon as MPS1 levels drop at kinetochores and/or other phosphatases intervene to dephosphorylate KNL1, SAC signaling is finally extinguished. The development of fluorescence-based biosensors combined with mathematical modeling will certainly provide in the future further mechanistic insights into such intricate network.     

The dynamic instability of actin filament barbed ends

The turnover of actin filament networks in cells has long been considered to reflect the treadmilling behavior of pure actin filaments in vitro, where only the pointed ends depolymerize. Newly discovered molecular mechanisms challenge this notion, as they provide evidence of situations in which growing and depolymerizing barbed ends coexist.

IntroductionIn cells, actin assembles into filament networks with diverse architectures and lifetimes, playing key roles in functions such as endocytosis, cell motility, and cell division. These filament networks are maintained and renewed by actin turnover, which implies that assembly and disassembly must take place simultaneously and in a controlled manner within the networks. Each actin filament end has the ability to either grow or shrink, depending on the concentration of actin and regulatory proteins, but pure actin treadmills at steady state: ATP-actin is added at the barbed end at a rate matching the departure of ADP-actin from the pointed end, and ATP hydrolysis takes place within the filament. This hallmark feature of actin dynamics has been known for decades (Wegner, 1976) and has been generalized to the cell context, in which it is commonly assumed that actin polymerization takes place at the barbed end, while depolymerization takes place only at the pointed end (whether it be the ends of filaments within the network or the ends of fragments that have detached from it). This notion is reinforced by the fact that the cytoplasm contains high concentrations of monomeric actin (G-actin) in complex with profilin (Funk et al., 2019), which is unable to bind to pointed ends and should drive the elongation of all noncapped barbed ends.Recently, however, in vitro studies have identified two seemingly independent mechanisms in which, in the presence of profilin-actin, filament barbed ends alternate between phases of growth and depolymerization. This behavior, referred to as “dynamic instability,” is widely observed for microtubules but was unexpected for actin filaments. It suggests that cells could use barbed ends for both elongation and disassembly.Driving the depolymerization of barbed ends with cofilin side-decorationProteins of the actin depolymerizing factor (ADF)/cofilin family (henceforth cofilin) are composed of a single ADF-homology (ADF-H) domain and are mostly known for their actin filament–severing activity (De La Cruz, 2009). Cofilin binds cooperatively to the sides of actin filaments, forming clusters where the conformation of the filament is locally altered, leading to its severing at cofilin cluster boundaries. In addition, the barbed ends of cofilin-decorated filaments steadily depolymerize, despite the presence of G-actin and profilin-actin (Fig. 1 A) and even capping protein (CP) in solution (Wioland et al., 2017, 2019). This unexpected result likely originates from the conformational change of actin subunits at the barbed end, induced by cofilin side-binding. As a consequence, filaments exposed to G-actin (with or without profilin), CP, and cofilin alternate between phases of barbed-end elongation and barbed-end depolymerization. In these conditions, actin filament barbed ends thus exhibit a form of dynamic instability.Open in a separate windowFigure 1.Two mechanisms that give rise to barbed-end depolymerization in elongation-promoting conditions. (A) When a cofilin side-decorated region reaches the barbed end, adding a new actin or profilin-actin becomes very difficult, and the barbed end depolymerizes. Not represented: Capping by CP can lead to depolymerization, as it allows the cofilin cluster to reach the barbed end, which then has a much weaker affinity for CP and steadily depolymerizes. Also, severing events occur at cofilin cluster boundaries, creating new barbed ends, either bare or cofilin-decorated. (B) Twinfilin binds to the barbed end, preventing its elongation and causing its depolymerization. Whether twinfilin remains processively attached to the depolymerizing barbed end or departs with the actin subunits is still unknown. Twinfilin has no impact on the elongation of mDia1-bearing barbed ends.Driving the depolymerization of barbed ends with twinfilin end-targetingTwinfilin has two ADF-H domains, but unlike cofilin, it binds poorly to the sides of actin filaments. Rather, twinfilin appears to mainly sequester ADP-actin monomers and target the barbed end to modulate its elongation and capping. Recent in vitro studies have shown that the interaction of twinfilin with actin filament barbed ends could drive their depolymerization, even in the presence of G-actin and profilin-actin (Johnston et al., 2015; Hakala et al., 2021; Shekhar et al., 2021). Very interestingly, the processive barbed-end elongator formin mDia1 is able to protect barbed ends from twinfilin, allowing them to sustain elongation (Shekhar et al., 2021). This leads to a situation in which, as filaments are exposed to profilin-actin and twinfilin, mDia1-bearing barbed ends elongate while bare barbed ends depolymerize (Fig. 1 B). It is safe to assume that, if filaments were continuously exposed to this protein mix including formin in solution, they would alternate between phases of growth and shrinkage over time, as formins come on and fall off the barbed end. This mix of proteins would therefore constitute another situation causing actin filament dynamic instability.From actin treadmilling to dynamic instability, in cells?This newly identified versatile behavior of actin filaments is reminiscent of microtubules. While dynamic instability is the hallmark behavior of microtubules, they can also be made to treadmill steadily by adding 4 microtubule-associated proteins (Arpağ et al., 2020). In cells, both microtubule dynamic instability and treadmilling have been clearly observed (Wittmann et al., 2003). In contrast, the disassembly of single actin filaments, either embedded in a network or severed from it, has not yet been directly observed in cells. Despite insights from techniques such as single-molecule speckle microscopy, it is still unclear from which end actin filaments depolymerize, even in networks that appear to globally treadmill, such as the lamellipodium. Pointed end depolymerization alone cannot account for what is observed in cells (Miyoshi et al., 2006) and alternative mechanisms have been proposed, including brutal filament-to-monomer transitions occurring in bursts, driven by cofilin, coronin, and Aip1 (Brieher, 2013; Tang et al., 2020).In cells, the high amounts of available G-actin (tens of micromolars; Funk et al., 2019) should limit barbed-end depolymerization. Based on the reported on-rate for ATP–G-actin at the barbed ends of cofilin-decorated filaments (Wioland et al., 2017, 2019), we can estimate that these barbed ends, under such conditions, would depolymerize for tens of seconds before being “rescued,” which is enough to remove tens of subunits from each filament. In contrast, twinfilin concentrations similar to those of G-actin appear necessary to drive barbed-end depolymerization (Hakala et al., 2021; Shekhar et al., 2021). As proteomics studies in HeLa cells report that twinfilin is 50-fold less abundant than actin, this may be difficult to achieve in cells (Bekker-Jensen et al., 2017). However, future studies may uncover proteins, or posttranslational modifications of actin, that enhance the ability of twinfilin to drive barbed-end depolymerization in the presence of high concentrations of profilin-actin.Molecular insights and possible synergiesWhile cofilin and twinfilin both interact with actin via ADF-H domains, they appear to drive barbed-end depolymerization through different mechanisms: twinfilin by directly targeting the barbed end, and cofilin by decorating the filament sides, thereby changing the conformation of the filament and putting its barbed end in a depolymerization-prone state.The two mechanisms, nonetheless, share clear similarities. For instance, cofilin side-binding and twinfilin end-targeting both slow down ADP-actin barbed-end depolymerization, compared with bare ADP-actin filaments (Wioland et al., 2017; Hakala et al., 2021; Shekhar et al., 2021). Strikingly, a crystal structure of the actin/twinfilin/CP complex indicates that the actin conformational change induced by twinfilin binding at the barbed end is similar to that induced by cofilin decorating the sides (Mwangangi et al., 2021). It is thus possible that the dynamic instability of actin filament barbed ends reflects the same conformation changes, triggered either by cofilin side-decoration or twinfilin end-targeting.In addition to decorating the filament sides, cofilin targets ADP-actin barbed ends. Unlike twinfilin, the direct interaction of cofilin with the barbed end cannot cause its depolymerization in the presence of ATP-actin monomers. Indeed, cofilin end-targeting accelerates the depolymerization of ADP-actin barbed ends in the absence of G-actin, but cofilin does not appear to interact with growing ATP-actin barbed ends (Wioland et al., 2017). This is in stark contrast with twinfilin end-targeting, which slows down ADP-actin depolymerization and accelerates ADP–Pi-actin depolymerization (Shekhar et al., 2021). These different behaviors regarding the nucleotide state of actin are intriguing and should be investigated further.Cofilin thus needs to decorate the filament sides in order to have an impact on barbed-end dynamics in elongation-promoting conditions. However, it is unknown whether cofilin side-decoration extends all the way to the terminal subunits and occupies sites that twinfilin would target. Thus, it is unclear whether cofilin and twinfilin would compete or synergize to drive barbed-end depolymerization.Synergies with other proteins are also worth further investigation, CP being an interesting candidate. Cofilin side-decoration drastically decreases the barbed-end affinity for CP, and capped filaments are thereby an efficient intermediate to turn growing barbed ends into depolymerizing barbed ends (Wioland et al., 2017). Twinfilin interacts with CP and the barbed end to enhance uncapping (Hakala et al., 2021; Mwangangi et al., 2021). Since CP can bind mDia1-bearing barbed ends and displace mDia1 (Bombardier et al., 2015; Shekhar et al., 2015), perhaps CP can also contribute to turn growing, mDia1-bearing barbed ends into depolymerizing barbed ends, by removing mDia1 from barbed ends and subsequently getting displaced from the barbed end by twinfilin.Finally, it is worth noting that profilin, which does not contain an ADF-H domain, also interacts with the barbed face of G-actin and with the barbed end of the filament. When profilin is in sufficient excess, it is able to promote barbed-end depolymerization in the presence of ATP–G-actin (Pernier et al., 2016). Unlike twinfilin, its depolymerization-promoting activity is not prevented by formin mDia1, and it thus does not lead to dynamic instability (bare and mDia1-bearing barbed ends all either grow or depolymerize). The coexistence of growing, mDia1-bearing barbed ends and depolymerizing, twinfilin-targeted barbed ends (Fig. 1 B) was observed in the presence of profilin (Shekhar et al., 2021), but profilin actually may not be required. Future studies should determine the exact role of profilin in this mechanism.ConclusionThe extent to which barbed-end dynamic instability contributes to actin turnover in cells is not known, but possible molecular mechanisms have now been identified. They should change the way we envision actin network dynamics, as we must now consider the possibility that cells also exploit the barbed end for disassembly. More work is needed to further document these mechanisms, but the idea of a “generalized treadmilling” has now been contradicted at its source: in vitro experiments.     

Mechanisms of nonvesicular lipid transport

We have long known that lipids traffic between cellular membranes via vesicles but have only recently appreciated the role of nonvesicular lipid transport. Nonvesicular transport can be high volume, supporting biogenesis of rapidly expanding membranes, or more targeted and precise, allowing cells to rapidly alter levels of specific lipids in membranes. Most such transport probably occurs at membrane contact sites, where organelles are closely apposed, and requires lipid transport proteins (LTPs), which solubilize lipids to shield them from the aqueous phase during their transport between membranes. Some LTPs are cup like and shuttle lipid monomers between membranes. Others form conduits allowing lipid flow between membranes. This review describes what we know about nonvesicular lipid transfer mechanisms while also identifying many remaining unknowns: How do LTPs facilitate lipid movement from and into membranes, do LTPs require accessory proteins for efficient transfer in vivo, and how is directionality of transport determined?

IntroductionEukaryotic cells contain diverse membranes, each with a characteristic and carefully regulated protein and lipid content. Most membrane proteins are first inserted into the ER and then traffic among cellular compartments in vesicles (Mellman and Warren, 2000). The same is true of lipids—most are synthesized in the ER and are then exchanged between organelles in vesicles (Vance, 2015). Lipids, however, are also moved between organelles by nonvesicular pathways. This type of transport has a number of functions. One is the bulk transfer of lipids sufficient to sustain organelle biogenesis. For example, mitochondria membrane biogenesis requires the import of most lipids (Acoba et al., 2020). Nonvesicular lipid transport also allows cells to change the lipid composition more rapidly and precisely than is possible by vesicular trafficking or during stress conditions when vesicular trafficking is compromised. It can be used to enrich or deplete membranes of particular lipids, either to modulate the lipid composition of a membrane or to regulate levels of signaling lipids.Nonvesicular lipid exchange within cells could, in theory, occur by two mechanisms. One is lipid diffusion between membranes following hemifusion, where the outer leaflets of two bilayers merge. While hemifusion is thought to occur transiently during membrane fusion, there is no evidence that this mechanism is used to move lipids between cellular compartments that do not fuse. The second type of mechanism is transport by proteins known as lipid transport proteins (LTPs). These proteins have the ability to move lipids between membranes via hydrophobic cavities that shield the lipids from the aqueous environment during transport. There are many families of LTPs, with most eukaryotic cells expressing multiple members of each family (Chiapparino et al., 2016; Wong et al., 2019). Although most of the known transporters function as shuttles that carry single lipid molecules between membranes, others serve as bridges that allow lipids to flow between membranes (Li et al., 2020). Mutations in some LTPs are known to cause diseases (Table 1).Table 1.Diseases caused by mutations in LTPs

A Novel Single-Domain Na+-Selective Voltage-Gated Channel in Photosynthetic Eukaryotes

The evolution of Na⁺-selective four-domain voltage-gated channels (4D-Na_vs) in animals allowed rapid Na⁺-dependent electrical excitability, and enabled the development of sophisticated systems for rapid and long-range signaling. While bacteria encode single-domain Na⁺-selective voltage-gated channels (BacNa_v), they typically exhibit much slower kinetics than 4D-Na_vs, and are not thought to have crossed the prokaryote–eukaryote boundary. As such, the capacity for rapid Na⁺-selective signaling is considered to be confined to certain animal taxa, and absent from photosynthetic eukaryotes. Certainly, in land plants, such as the Venus flytrap (Dionaea muscipula) where fast electrical excitability has been described, this is most likely based on fast anion channels. Here, we report a unique class of eukaryotic Na⁺-selective, single-domain channels (EukCatBs) that are present primarily in haptophyte algae, including the ecologically important calcifying coccolithophores, Emiliania huxleyi and Scyphosphaera apsteinii. The EukCatB channels exhibit very rapid voltage-dependent activation and inactivation kinetics, and isoform-specific sensitivity to the highly selective 4D-Na_v blocker tetrodotoxin. The results demonstrate that the capacity for rapid Na⁺-based signaling in eukaryotes is not restricted to animals or to the presence of 4D-Na_vs. The EukCatB channels therefore represent an independent evolution of fast Na⁺-based electrical signaling in eukaryotes that likely contribute to sophisticated cellular control mechanisms operating on very short time scales in unicellular algae.

Electrical signals trigger rapid physiological events that underpin an array of fundamental processes in eukaryotes, from contractile amoeboid locomotion (Bingley and Thompson, 1962), to the action potentials of mammalian nerve and muscle cells (Hodgkin and Huxley, 1952). These events are mediated by voltage-gated ion channels (Brunet and Arendt, 2015). In excitable animal cells, Ca²⁺- or Na⁺-selective members of the four-domain voltage-gated cation channel family (4D-Ca_v/Na_v) underpin well-characterized signaling processes (Catterall et al., 2017). The 4D-Ca_v/Na_v family is broadly distributed across eukaryotes, contributing to signaling processes associated with motility in some unicellular protist and microalgal species (Fujiu et al., 2009; Lodh et al., 2016), although these channels are absent from land plants (Edel et al., 2017). It is likely that the ancestral 4D-Ca_v/Na_v channel was Ca²⁺-permeable, with Na⁺-selective channels arising later within the animal lineage (Moran et al., 2015). This shift in ion selectivity represented an important innovation in animals, allowing rapid voltage-driven electrical excitability to be decoupled from intracellular Ca²⁺ signaling processes (Moran et al., 2015).Na⁺-selective voltage-gated channels have not been described in other eukaryotes, although a large family of Na⁺-selective channels (BacNa_v) is present in prokaryotes (Ren et al., 2001; Koishi et al., 2004). BacNa_v are single-domain channels that form homotetramers, resembling the four-domain architecture of 4D-Ca_v/Na_v. Studies of BacNa_v channels have provided considerable insight into the mechanisms of gating and selectivity in voltage-dependent ion channels (Payandeh et al., 2012; Zhang et al., 2012). The range of activation and inactivation kinetics of native BacNa_v are generally slower than observed for 4D-Na_v, suggesting that the concatenation and subsequent differentiation of individual pore-forming subunits may have enabled 4D-Na_v to develop specific properties such as fast inactivation, which is mediated by the conserved intracellular Ile–Phe–Met linker between domains III and IV (Fig. 1A; Irie et al., 2010; Catterall et al., 2017).Open in a separate windowFigure 1.EukCatBs represent a novel class of single-domain channels. A, Schematic diagram of a single-domain EukCatB channel. The voltage-sensing module (S1–S4, blue), including conserved positively charged (++) residues of segment (S4) that responds to changes in membrane potential, is shown. The pore module (S5–S6, red) is also indicated, including the SF motif (Ren et al., 2001). The structure of a 4D-Na_v (showing the SF of rat 4D-Na_v1.4 with canonical “DEKA” locus of Na⁺-selective 4D-Na_v1s) is also displayed (right). The Ile–Phe–Met motif of the fast inactivation gate is indicated (West et al., 1992) B, Maximum likelihood phylogenetic tree of single-domain, voltage-gated channels including BacNa_v and the three distinct classes of EukCat channels (EukCatA–C). Representatives of the specialized family of single-domain Ca²⁺ channels identified in mammalian sperm (CatSpers) are also included. SF for each sequence is shown (right). “Position 0” of the high-field–strength site that is known to be important in determining Na⁺ selectivity (Payandeh et al., 2011), is colored red. Channel sequences selected for functional characterization in this study are shown in bold. EukCatA sequences previously characterized (Helliwell et al., 2019) are also indicated, as is NaChBac channel from B. halodurans (Ren et al., 2001). Maximum likelihood bootstrap values (>70) and Bayesian posterior probabilities (>0.95) are above and below nodes, respectively. Scanning electron micrographs of coccolithophores E. huxleyi (scale bar = 2 μm) and S. apsteinii, (scale bar = 10 μm) are shown.We recently identified several classes of ion channel (EukCats) bearing similarity to BacNa_v in the genomes of eukaryotic phytoplankton. Characterization of EukCatAs found in marine diatoms demonstrated that these voltage-gated channels are nonselective (exhibiting permeability to both Na⁺ and Ca²⁺) and play a role in depolarization-activated Ca²⁺ signaling (Helliwell et al., 2019). Two other distinct classes of single-domain channels (EukCatBs and EukCatCs) were also identified that remain uncharacterized. These channels are present in haptophytes, pelagophytes, and cryptophytes (EukCatBs), as well as dinoflagellates (EukCatCs; Helliwell et al., 2019). Although there is a degree of sequence similarity between the distinct EukCat clades, the relationships between clades are not well resolved, and there is not clear support for a monophyletic origin of EukCats. The diverse classes of EukCats may therefore exhibit significant functional differences. Characterization of these different classes of eukaryote single-domain channels is thus vital to our understanding of eukaryote ion channel structure, function, and evolution, and to our gaining insight into eukaryote membrane physiology more broadly.Notably, EukCatB channels were found in ecologically important coccolithophores, a group of unicellular haptophyte algae that represent major primary producers in marine ecosystems. Coccolithophores are characterized by their ability to produce a cell covering of ornate calcium carbonate platelets (coccoliths; Fig. 1B; Taylor et al., 2017). The calcification process plays an important role in global carbon cycling, with the sinking of coccoliths representing a major flux of carbon to the deep ocean. Patch-clamp studies of coccolithophores indicate several unusual aspects of membrane physiology, such as an inwardly rectifying Cl⁻ conductance and a large outward H⁺ conductance at positive membrane potentials, which may relate to the increased requirement for pH homeostasis associated with intracellular calcification. Here we report that EukCatB channels from two coccolithophore species (Emiliania huxleyi and Scyphosphaera apsteinii) act as very fast Na⁺-selective voltage-gated channels that exhibit many similarities to the 4D-Na_vs, which underpin neuronal signaling in animals. Thus, our findings demonstrate that the capacity for rapid Na⁺-based signaling has evolved in certain photosynthetic eukaryotes, contrary to previous widely held thinking.