A Two-Hybrid-Receptor Assay Demonstrates Heteromer Formation as Switch-On for Plant Immune Receptors

Receptor kinases sense extracellular signals and trigger intracellular signaling and physiological responses. However, how does signal binding to the extracellular domain activate the cytoplasmic kinase domain? Activation of the plant immunoreceptor Flagellin sensing2 (FLS2) by its bacterial ligand flagellin or the peptide-epitope flg22 coincides with rapid complex formation with a second receptor kinase termed brassinosteroid receptor1 associated kinase1 (BAK1). Here, we show that the receptor pair of FLS2 and BAK1 is also functional when the roles of the complex partners are reversed by swapping their cytosolic domains. This reciprocal constellation prevents interference by redundant partners that can partially substitute for BAK1 and demonstrates that formation of the heteromeric complex is the molecular switch for transmembrane signaling. A similar approach with swaps between the Elongation factor-Tu receptor and BAK1 also resulted in a functional receptor/coreceptor pair, suggesting that a “two-hybrid-receptor assay” is of more general use for studying heteromeric receptor complexes.Cell surface receptors are chemical sensors, often with an exquisite specificity and sensitivity, which detect extracellular signals and initiate corresponding intracellular response programs. Many of these receptors are transmembrane proteins with an extracellular ligand-binding domain and an intracellular protein kinase domain. Higher plants, such as Arabidopsis (Arabidopsis thaliana), have several hundred genes encoding receptor like kinases (Shiu and Bleecker, 2001; Shiu and Li, 2004). How are these receptor like kinases activated by their ligands, and how do they initiate a subsequent intracellular signaling cascade? In our work, we used the leucine-rich repeat receptor kinase (LRR-RK) Flagellin sensing2 (FLS2), which specifically detects bacterial flagellin or its peptide epitope flg22 at subnanomolar concentrations (Gomez-Gomez and Boller, 2000; Gómez-Gómez et al., 2001; Chinchilla et al., 2006). FLS2 undergoes heteromeric complex formation with brassinosteroid receptor1 associated kinase1 (BAK1) within seconds after application of the flagellin-derived peptide ligand flg22 (Chinchilla et al., 2007; Heese et al., 2007; Schulze et al., 2010). Thus, BAK1 might act as a coreceptor of FLS2. However, as previously observed (Chinchilla et al., 2007; Roux et al., 2011), FLS2 is still functional in the absence of BAK1, although with a reduced efficiency. This raises the question whether the ligand-induced heteromeric complex has merely an enhancing effect or whether association with BAK1 or a functional substitute acts as the essential switch-on for transmembrane signaling of FLS2. BAK1 is one of the five members that form the somatic embryogenesis receptor kinase (SERK) family (Albrecht et al., 2008), and other members of this family might partially substitute for BAK1 (Roux et al., 2011). However, a rigorous genetic approach to delineate the role of these potential substitutes is not feasible because triple mutants (serk1 serk3 serk4) and quadruple mutants (serk1 serk2 serk3 serk4) exhibit severe general phenotypes of dwarfing or even lethality at the early embryo stage (He et al., 2007; Gou et al., 2012) that might be due to the important role of SERKs in plant developmental processes (Li et al., 2002; Nam and Li, 2002).To address the role of the heteromeric complex with BAK1 in the absence of other interfering SERKs, we took a two-hybrid-receptor approach based on the premise that the apoplastic and cytoplasmic domains of FLS2 and BAK1 function in a modular manner. A heteromeric complex might thus also form and function when the roles of FLS2 and BAK1 are reversed by reciprocal swapping of their cytoplasmic protein kinase domains (Fig. 1A, schematic view).Open in a separate windowFigure 1.Heteromeric complex formation of FLS with BAK1 switches on flagellin-dependent transmembrane signaling. A, Model for the flg22-dependent heteromeric receptor complex. Schematic representation of FLS2 (blue), BAK1 (red), and the chimeric receptor constructs Ftm-B and Btm-F. Ftm-B comprises the extracellular and the transmembrane domains of FLS2 (blue) and the cytoplasmic domain of BAK1 (red). The receptor chimera Btm-F represents the reciprocal construct with the cytoplasmic part of BAK1 replaced by that of FLS2. The cytoplasmic domain of FLS2 was C-terminally tagged with a GFP. B and C, Functional comparison of native FLS2 and BAK1 (B) with the two hybrid receptors Ftm-B and Btm-F (C). The experiments show luciferase activity in Arabidopsis fls2 bak1-4 double mutant protoplasts cotransformed with pFRK1::Luciferase as a reporter and the receptor constructs indicated. At 0 h (dashed line) protoplasts were treated with 100 nm of flg22 (black diamonds) or 100 nm of the inactive analog flg22^Atum (white circles). Light emission of the protoplasts was measured with a luminometer. Values represent averages and sds of three replicates. Data shown are representative for at least three independent repetitions of the experiments with all constructs. D and E, Dose-response relationship for flg22-dependent induction of pFRK1::Luciferase in protoplasts expressing the receptor constructs indicated. Values represent increase in luciferase activity after 5 h of treatment as percentage of the increase observed with FLS2 plus BAK1 treated with saturating doses of greater than or equal to 10 nm flg22. Comparison of (half-maximal stimulation values in D and E shows that cells coexpressing Ftm-B plus Btm-F responded at least as sensitive to flg22 as cells coexpressing FLS2 plus BAK1. A combination of Ftm-B with the kinase dead version Btm-F_KD was not functional, even when treated with 1,000 nm flg22. LU, Light units.     

Ethylene Signaling Regulates Accumulation of the FLS2 Receptor and Is Required for the Oxidative Burst Contributing to Plant Immunity

Reactive oxygen species (ROS) are potent signal molecules rapidly generated in response to stress. Detection of pathogen-associated molecular patterns induces a transient apoplastic ROS through the function of the NADPH respiratory burst oxidase homologs D (RbohD). However, little is known about the regulation of pathogen-associated molecular pattern-elicited ROS or its role in plant immunity. We investigated ROS production triggered by bacterial flagellin (flg22) in Arabidopsis (Arabidopsis thaliana). The oxidative burst was diminished in ethylene-insensitive mutants. Flagellin Sensitive2 (FLS2) accumulation was reduced in etr1 and ein2, indicating a requirement of ethylene signaling for FLS2 expression. Multiplication of virulent bacteria was enhanced in Arabidopsis lines displaying altered ROS production at early but not late stages of infection, suggesting an impairment of preinvasive immunity. Stomatal closure, a mechanism used to reduce bacterial entry into plant tissues, was abolished in etr1, ein2, and rbohD mutants. These results point to the importance of flg22-triggered ROS at an early stage of the plant immune response.A rapid and transient increase in reactive oxygen species (ROS), termed an “oxidative burst,” is often associated with responses to abiotic and biotic stresses and could trigger changes in stomatal aperture or programmed cell death in defense against pathogens (Kwak et al., 2003; Torres and Dangl, 2005). ROS production can occur extracellularly through activities of plasma membrane-resident NADPH oxidases (Kangasjärvi et al., 2005; Torres and Dangl, 2005). In plants, Rboh proteins, which are homologs of mammalian NADPH oxidase 2, were shown to be the predominant mediators of apoplastic ROS production (Torres et al., 1998; Galletti et al., 2008). Respiratory burst oxidase homologs D and F (RbohD and RbohF) were identified by mutation to be the responsible oxidases in Arabidopsis (Arabidopsis thaliana) defense responses (Torres et al., 2002). While most ROS generated in response to avirulent Pseudomonas syringae bacteria and Hyaloperonospora oomycete pathogens depend on RbohD function, the induced cell death response by these pathogens appears to be mostly regulated by RbohF. Cell death provoked upon infection with the necrotizing fungus Alternaria, however, is under the control of RbohD (Pogány et al., 2009). The contribution of NADPH oxidases to plant immunity was also described in barley (Hordeum vulgare) and tobacco (Nicotiana benthamiana), where resistance to powdery mildew fungi and the oomycete Phytophthora infestans, respectively, was dependent on Rboh functions (Yoshioka et al., 2003; Trujillo et al., 2006).An early layer of active plant defense is mediated by pattern recognition receptors, which sense microbes according to conserved constituents, so-called pathogen-associated molecular patterns (PAMPs). These initiate a plethora of defense responses referred to as PAMP-triggered immunity (Boller and Felix, 2009). The Arabidopsis receptor kinase Flagellin Sensitive2 (FLS2) recognizes and physically interacts with flg22, the elicitor-active epitope of bacterial flagellin (Felix et al., 1999; Gomez-Gomez and Boller, 2000; Chinchilla et al., 2006). FLS2 is plasma membrane localized and expressed throughout the plant (Robatzek et al., 2006). FLS2 requires the receptor kinase BRI1-Associated Kinase1 (BAK1), which forms a heteromeric complex upon flg22 binding (Chinchilla et al., 2007). Subsequently, a rapid and transient flg22-stimulated oxidative burst occurs that is dependent on RbohD (Zhang et al., 2007). In addition, flg22 triggers early responses, such as ethylene biosynthesis, activation of mitogen-activated protein (MAP) kinase cascades, and changes in gene expression (Felix et al., 1999; Asai et al., 2002; Zipfel et al., 2004). Late flg22 responses include the accumulation of salicylic acid (SA), callose deposition, and an arrest of seedling growth (Gomez-Gomez et al., 1999; Mischina and Zeier, 2007). This collectively contributes to plant immunity (Zipfel et al., 2004; Melotto et al., 2006).Little is known about the regulatory components of FLS2-activated early flg22 responses and their relevance in plant resistance to pathogens. Here, we investigated flg22-triggered ROS production in Arabidopsis seedlings and have identified ethylene signaling as a critical component of the oxidative burst in response to flg22, partly through promoting the accumulation of FLS2. We further provide evidence that the flg22-triggered oxidative burst is required for resistance to bacterial infection at the point of pathogen entry through stomata.     

Concerted actions of DnaA complexes with DNA-unwinding sequences within and flanking replication origin oriC promote DnaB helicase loading

Unwinding of the replication origin and loading of DNA helicases underlie the initiation of chromosomal replication. In Escherichia coli, the minimal origin oriC contains a duplex unwinding element (DUE) region and three (Left, Middle, and Right) regions that bind the initiator protein DnaA. The Left/Right regions bear a set of DnaA-binding sequences, constituting the Left/Right-DnaA subcomplexes, while the Middle region has a single DnaA-binding site, which stimulates formation of the Left/Right-DnaA subcomplexes. In addition, a DUE-flanking AT-cluster element (TATTAAAAAGAA) is located just outside of the minimal oriC region. The Left-DnaA subcomplex promotes unwinding of the flanking DUE exposing TT[A/G]T(T) sequences that then bind to the Left-DnaA subcomplex, stabilizing the unwound state required for DnaB helicase loading. However, the role of the Right-DnaA subcomplex is largely unclear. Here, we show that DUE unwinding by both the Left/Right-DnaA subcomplexes, but not the Left-DnaA subcomplex only, was stimulated by a DUE-terminal subregion flanking the AT-cluster. Consistently, we found the Right-DnaA subcomplex–bound single-stranded DUE and AT-cluster regions. In addition, the Left/Right-DnaA subcomplexes bound DnaB helicase independently. For only the Left-DnaA subcomplex, we show the AT-cluster was crucial for DnaB loading. The role of unwound DNA binding of the Right-DnaA subcomplex was further supported by in vivo data. Taken together, we propose a model in which the Right-DnaA subcomplex dynamically interacts with the unwound DUE, assisting in DUE unwinding and efficient loading of DnaB helicases, while in the absence of the Right-DnaA subcomplex, the AT-cluster assists in those processes, supporting robustness of replication initiation.

The initiation of bacterial DNA replication requires local duplex unwinding of the chromosomal replication origin oriC, which is regulated by highly ordered initiation complexes. In Escherichia coli, the initiation complex contains oriC, the ATP-bound form of the DnaA initiator protein (ATP–DnaA), and the DNA-bending protein IHF (Fig. 1, A and B), which promotes local unwinding of oriC (1, 2, 3, 4). Upon this oriC unwinding, two hexamers of DnaB helicases are bidirectionally loaded onto the resultant single-stranded (ss) region with the help of the DnaC helicase loader (Fig. 1B), leading to bidirectional chromosomal replication (5, 6, 7, 8). However, the fundamental mechanism underlying oriC-dependent bidirectional DnaB loading remains elusive.Open in a separate windowFigure 1Schematic structures of oriC, DnaA, and the initiation complexes. A, the overall structure of oriC. The minimal oriC region and the AT-cluster region are indicated. The sequence of the AT-cluster−DUE (duplex-unwinding element) region is also shown below. The DUE region (DUE; pale orange bars) contains three 13-mer repeats: L-DUE, M-DUE, and R-DUE. DnaA-binding motifs in M/R-DUE, TT(A/G)T(T), are indicated by red characters. The AT-cluster region (AT cluster; brown bars) is flanked by DUE outside of the minimal oriC. The DnaA-oligomerization region (DOR) consists of three subregions called Left-, Middle-, and Right-DOR. B, model for replication initiation. DnaA is shown as light brown (for domain I–III) and darkbrown (for domain IV) polygons (right panel). ATP–DnaA forms head-to-tail oligomers on the Left- and Right-DORs (left panel). The Middle-DOR (R2 box)-bound DnaA interacts with DnaA bound to the Left/Right-DORs using domain I, but not domain III, stimulating DnaA assembly. IHF, shown as purple hexagons, bends DNA >160° and supports DUE unwinding by the DnaA complexes. M/R-DUE regions are efficiently unwound. Unwound DUE is recruited to the Left-DnaA subcomplex and mainly binds to R1/R5M-bound DnaA molecules. The sites of ssDUE-binding B/H-motifs V211 and R245 of R1/R5M-bound DnaA molecules are indicated (pink). Two DnaB homohexamer helicases (light green) are recruited and loaded onto the ssDUE regions with the help of the DnaC helicase loader (cyan). ss, single stranded.The minimal oriC region consists of the duplex unwinding element (DUE) and the DnaA oligomerization region (DOR), which contains specific arrays of 9-mer DnaA-binding sites (DnaA boxes) with the consensus sequence TTA[T/A]NCACA (Fig. 1A) (3, 4). The DUE underlies the local unwinding and contains 13-mer AT-rich sequence repeats named L-, M-, and R-DUE (9). The M/R-DUE region includes TT[A/G]T(A) sequences with specific affinity for DnaA (10). In addition, a DUE-flanking AT-cluster (TATTAAAAAGAA) region resides just outside of the minimal oriC (Fig. 1A) (11). The DOR is divided into three subregions, the Left-, Middle-, and Right-DORs, where DnaA forms structurally distinct subcomplexes (Fig. 1A) (8, 12, 13, 14, 15, 16, 17). The Left-DOR contains high-affinity DnaA box R1, low-affinity boxes R5M, τ1−2, and I1-2, and an IHF-binding region (17, 18, 19, 20). The τ1 and IHF-binding regions partly overlap (17).In the presence of IHF, ATP–DnaA molecules cooperatively bind to R1, R5M, τ2, and I1-2 boxes in the Left-DOR, generating the Left-DnaA subcomplex (Fig. 1B) (8, 17). Along with IHF causing sharp DNA bending, the Left-DnaA subcomplex plays a leading role in DUE unwinding and subsequent DnaB loading. The Middle-DOR contains moderate-affinity DnaA box R2. Binding of DnaA to this box stimulates DnaA assembly in the Left- and Right-DORs using interaction by DnaA N-terminal domain (Fig. 1B; also see below) (8, 12, 14, 16, 21). The Right-DOR contains five boxes (C3-R4 boxes) and cooperative binding of ATP–DnaA molecules to these generates the Right-DnaA subcomplex (Fig. 1B) (12, 18). This subcomplex is not essential for DUE unwinding and plays a supportive role in DnaB loading (8, 15, 17). The Left-DnaA subcomplex interacts with DnaB helicase, and the Right-DnaA subcomplex has been suggested to play a similar role (Fig. 1B) (8, 13, 16).In the presence of ATP–DnaA, M- and R-DUE adjacent to the Left-DOR are predominant sites for in vitro DUE unwinding: unwinding of L-DUE is less efficient than unwinding of the other two (Fig. 1B) (9, 22, 23). Deletion of L-DUE or the whole DUE inhibits replication of oriC in vitro moderately or completely, respectively (23). A chromosomal oriC Δ(AT-cluster−L-DUE) mutant with an intact DOR, as well as deletion of Right-DOR, exhibits limited inhibition of replication initiation, whereas the synthetic mutant combining the two deletions exhibits severe inhibition of cell growth (24). These studies suggest that AT-cluster−L-DUE regions stimulate replication initiation in a manner concerted with Right-DOR, although the underlying mechanisms remain elusive.DnaA consists of four functional domains (Fig. 1B) (4, 25). Domain I supports weak domain I–domain I interaction and serves as a hub for interaction with various proteins such as DnaB helicase and DiaA, which stimulates ATP–DnaA assembly at oriC (26, 27, 28, 29, 30). Two or three domain I molecules of the oriC–DnaA subcomplex bind a single DnaB hexamer, forming a stable higher-order complex (7). Domain II is a flexible linker (28, 31). Domain III contains AAA+ (ATPase associated with various cellular activities) motifs essential for ATP/ADP binding, ATP hydrolysis, and DnaA–DnaA interactions in addition to specific sites for ssDUE binding and a second, weak interaction with DnaB helicase (1, 4, 8, 10, 19, 25, 32, 33, 34, 35). Domain IV bears a helix-turn-helix motif with specific affinity for the DnaA box (36).As in typical AAA+ proteins, a head-to-tail interaction underlies formation of ATP–DnaA pentamers on the DOR, where the AAA+ arginine-finger motif Arg285 recognizes ATP bound to the adjacent DnaA protomer, promoting cooperative ATP–DnaA binding (Fig. 1B) (19, 32). DnaA ssDUE-binding H/B-motifs (Val211 and Arg245) in domain III sustain stable unwinding by directly binding to the T-rich (upper) strand sequences TT[A/G]T(A) within the unwound M/R-DUE (Fig. 1B) (8, 10). Val211 residue is included in the initiator-specific motif of the AAA+ protein family (10). For DUE unwinding, ssDUE is recruited to the Left-DnaA subcomplex via DNA bending by IHF and directly interacts with H/B-motifs of DnaA assembled on Left-DOR, resulting in stable DUE unwinding competent for DnaB helicase loading; in particular, DnaA protomers bound to R1 and R5M boxes play a crucial role in the interaction with M/R-ssDUE (Fig. 1B) (8, 10, 17). Collectively, these mechanisms are termed ssDUE recruitment (4, 17, 37).Two DnaB helicases are thought to be loaded onto the upper and lower strands of the region including the AT-cluster and DUE, with the aid of interactions with DnaC and DnaA (Fig. 1B) (25, 38, 39). DnaC binding modulates the closed ring structure of DnaB hexamer into an open spiral form for entry of ssDNA (40, 41, 42, 43). Upon ssDUE loading of DnaB, DnaC is released from DnaB in a manner stimulated by interactions with ssDNA and DnaG primase (44, 45). Also, the Left- and Right-DnaA subcomplexes, which are oriented opposite to each other, could regulate bidirectional loading of DnaB helicases onto the ssDUE (Fig. 1B) (7, 8, 35). Similarly, recent works suggest that the origin complex structure is bidirectionally organized in both archaea and eukaryotes (1, 46). In Saccharomyces cerevisiae, two origin recognition complexes containing AAA+ proteins bind to the replication origin region in opposite orientations; this, in turn, results in efficient loading of two replicative helicases, leading to head-to-head interactions in vitro (46). Consistent with this, origin recognition complex dimerization occurs in the origin region during the late M-G1 phase (47). The fundamental mechanism of bidirectional origin complexes might be widely conserved among species.In this study, we analyzed various mutants of oriC and DnaA in reconstituted systems to reveal the regulatory mechanisms underlying DUE unwinding and DnaB loading. The Right-DnaA subcomplex assisted in the unwinding of oriC, dependent upon an interaction with L-DUE, which is important for efficient loading of DnaB helicases. The AT-cluster region adjacent to the DUE promoted loading of DnaB helicase in the absence of the Right-DnaA subcomplex. Consistently, the ssDNA-binding activity of the Right-DnaA subcomplex sustained timely initiation of growing cells. These results indicate that DUE unwinding and efficient loading of DnaB helicases are sustained by concerted actions of the Left- and Right-DnaA subcomplexes. In addition, loading of DnaB helicases are sustained by multiple mechanisms that ensure robust replication initiation, although the complete mechanisms are required for precise timing of initiation during the cell cycle.     

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.     

Interplay between Velocity and Travel Distance of Kinesin-based Transport in the Presence of Tau

Although the disease-relevant microtubule-associated protein tau is known to severely inhibit kinesin-based transport in vitro, the potential mechanisms for reversing this detrimental effect to maintain healthy transport in cells remain unknown. Here we report the unambiguous upregulation of multiple-kinesin travel distance despite the presence of tau, via decreased single-kinesin velocity. Interestingly, the presence of tau also modestly reduced cargo velocity in multiple-kinesin transport, and our stochastic simulations indicate that the tau-mediated reduction in single-kinesin travel underlies this observation. Taken together, our observations highlight a nontrivial interplay between velocity and travel distance for kinesin transport, and suggest that single-kinesin velocity is a promising experimental handle for tuning the effect of tau on multiple-kinesin travel distance.Conventional kinesin is a major microtubule-based molecular motor that enables long-range transport in living cells. Although traditionally investigated in the context of single-motor experiments, two or more kinesin motors are often linked together to transport the same cargo in vivo (1–4). Understanding the control and regulation of the group function of multiple kinesins has important implications for reversing failure modes of transport in a variety of human diseases, particularly neurodegenerative diseases. Tau is a disease-relevant protein enriched in neurons (5,6). The decoration of microtubules with tau is known to strongly inhibit kinesin transport in vitro (7–9), but how kinesin-based transport is maintained in the presence of high levels of tau, particularly in healthy neurons, remains an important open question. To date, no mechanism has been directly demonstrated to reverse the inhibitory effect of tau on kinesin-based transport. Here we present a simple in vitro study that demonstrates the significant upregulation of multiple-kinesin travel distance with decreasing ATP concentration, despite the presence of tau.This investigation was motivated by our recent finding that single-kinesin velocity is a key controller for multiple-kinesin travel distance along bare microtubules (10). The active stepping of each kinesin motor is stimulated by ATP (11), and each kinesin motor remains strongly bound to the microtubule between successive steps (10,11). As demonstrated for bare microtubules (10), with decreasing ATP concentrations, each microtubule-bound kinesin experiences a decreased stepping rate per unit time and spends an increased fraction of time in the strongly bound state; additional unbound kinesins on the same cargo have more time to bind to the microtubule before cargo travel terminates. Thus, reductions in single-kinesin velocity increase the probability that at least one kinesin motor will remain bound to the microtubule per unit time, thereby increasing the travel distance of each cargo (10). Because this effect only pertains to the stepping rate of each individual kinesin and does not address the potential presence of roadblocks such as tau on the microtubules, we hypothesized in this study that single-kinesin velocity may be exploited to relieve the impact of tau on multiple-kinesin travel distance.We focused our in vitro investigation on human tau 23 (htau23, or 3RS tau), an isoform of tau that exhibits the strongest inhibitory effect on kinesin-based transport (7–9). Importantly, htau23 does not alter the stepping rate of individual kinesins (7,9), supporting our hypothesis and enabling us to decouple single-kinesin velocity from the potential effects of tau. We carried out multiple-kinesin motility experiments using polystyrene beads as in vitro cargos (8,10), ATP concentration as an in vitro handle to controllably tune single-kinesin velocity (10,11), and three input kinesin concentrations to test the generality of potential findings for multiple-kinesin transport. Combined with previous two-kinesin studies (10,12), our measurements of travel distance (Fig. 1 A) indicate that the lowest kinesin concentration employed (0.8 nM) corresponds to an average of ∼2–3 kinesins per cargo. Note that in the absence of tau, the observed decrease in bead velocity at the higher kinesin concentrations (Fig. 1 A) is consistent with a recent in vitro finding (13). At 1 mM ATP, htau23 reduced kinesin-based travel distance by a factor of two or more (Fig. 1, A and B). This observation is in good agreement with previous reports (7,8).Open in a separate windowFigure 1Distributions of multiple-kinesin travel distances measured at three experimental conditions, to verify the effect of tau (A and B) and to investigate the impact of single-kinesin velocity on the tau effect (B and C). Shaded bars at 8.7 μm indicate counts of travel exceeding the field of view. The mean travel distance (d; ± standard error of mean, SEM), sample size (n), and corresponding mean velocity (v; ± SEM) are indicated. MT denotes microtubule. Mean travel distance increased substantially at 20 μM ATP (C), despite the presence of htau23. This effect persisted across all three kinesin concentrations tested (left to right).Consistent with our hypothesis, reducing the available ATP concentration to 20 μM increased the multiple-kinesin travel distance by >1.4-fold for all three input kinesin concentrations (Fig. 1, B and C), despite the presence of htau23. The corresponding reduction in single-kinesin velocity with decreasing ATP concentration (10,11) is reflected in the ∼3.4-fold reduction in the measured bead velocities (Fig. 1, B and C). Therefore, the strong negative relationship between single-kinesin velocity and multiple-kinesin travel distance occurs not only for bare microtubules (10), but also for tau-decorated microtubules.What causes the observed increase in travel distance at the lower ATP concentration (Fig. 1, B and C)? In addition to the mechanism discussed above for the case of bare microtubules (10), an intriguing mechanism was suggested by recent studies of tau-microtubule interactions in which htau23 was observed to dynamically diffuse along microtubule lattices (14,15): reducing the stepping rate of a microtubule-bound kinesin may effectively increase the probability that a tau roadblock can diffuse away before the kinesin takes its next step.Perhaps surprisingly, although htau23 does not impact single-kinesin velocity (7,9), we observed a modest reduction in the average velocity of multiple-kinesin transport in experiments using tau-decorated microtubules (Fig. 1, A and B). This decreased velocity reflects a substantially larger variance in the instantaneous velocity for bead trajectories in the presence of htau23 (see Fig. S1 in the Supporting Material), as quantified by parsing each bead trajectory into a series of constant-velocity segments using a previously developed automatic software incorporating Bayesian statistics (16).To test the possibility that single-kinesin travel distance impacts multiple-kinesin velocity, we performed stochastic simulations (see the Supporting Material) that assumed N identical kinesin motors available for transport and included kinesin’s detachment kinetics (17). Previously, this model successfully captured multiple-dynein travel distances in vivo using single-dynein characteristics measured in vitro (18). In this study, we introduced one (and only one) free parameter to reflect the probability of each bound kinesin encountering tau at each step. When encountering tau, each kinesin has a 54% probability of detaching from the microtubule (interpolated from Fig. 2A of Dixit et al. (7)); the undetached kinesin is assumed to remain engaged in transport and completes its step along the microtubule despite the presence of tau.Remarkably, our simple simulation suggested that the tau-mediated reduction in single-kinesin travel is sufficient to reduce multiple-kinesin velocity (Fig. 2 A). The majority of the velocity decrease is predicted to occur at the transition from single-kinesin to two-kinesin transport (Fig. 2). Further decreases in cargo velocity with increasing motor number are predicted to be modest and largely independent of tau (Fig. 2 B). The results of our simulation remain qualitatively the same when evaluated at two bounds (40 and 65%) encompassing the interpolated 54% probability of kinesin detaching at tau (see Fig. S2).Open in a separate windowFigure 2Stochastic simulations predict a tau-dependent reduction in multiple-kinesin velocity, assuming that the only effect of tau protein is to prematurely detach kinesin from the microtubule (or, to reduce single-kinesin travel distance). (A) Average velocity of cargos carried by the indicated number of kinesins was evaluated at 1 mM ATP, and for four probabilities that a kinesin may encounter tau at each step. Mean velocity was evaluated using 600 simulated trajectories for all simulation conditions. Error bars indicate SEM. (B) Change in cargo velocity with each additional kinesin (ΔVel/kinesin) as a function of tau-encounter probability. These values were calculated from cargo velocities shown in panel A. Error bars indicate SEM.We note that our simple simulations do not consider the possibility that kinesin may pause in front of a tau roadblock, as previously reported in Dixit et al. (7). We omitted this consideration because the interaction strength between kinesin and the microtubule in such a paused state is unknown. In a multiple-motor geometry, could a paused kinesin be dragged along by the other motors bound to the same cargo? Could a tau roadblock be forcefully swept off the microtubule surface by the collective motion of the cargo-motor complex? Significant experimental innovations are necessary to specifically address these questions in future multiple-motor assays and to guide modeling efforts. Nonetheless, our simple simulation demonstrates that reducing single-kinesin travel distance is sufficient to decrease multiple-kinesin travel distance.Taken together, our observations highlight a nontrivial interplay between velocity and travel distance for kinesin-based transport in the presence of tau. We uncover a previously unexplored dual inhibition of tau on kinesin-transport: in addition to limiting cargo travel distance, the tau-mediated reduction in single-kinesin travel distance also leads to a modest reduction in multiple-kinesin velocity. We provide what we believe to be the first demonstration of the unambiguous upregulation of multiple-kinesin travel distance despite the presence of tau, via reducing single-kinesin velocity, suggesting a mechanism that could be harnessed for future therapeutic interventions in diseases that result from aberrant kinesin-based transport.     

Conformational States of Melittin at a Bilayer Interface

The distribution of peptide conformations in the membrane interface is central to partitioning energetics. Molecular-dynamics simulations enable characterization of in-membrane structural dynamics. Here, we describe melittin partitioning into dioleoylphosphatidylcholine lipids using CHARMM and OPLS force fields. Although the OPLS simulation failed to reproduce experimental results, the CHARMM simulation reported was consistent with experiments. The CHARMM simulation showed melittin to be represented by a narrow distribution of folding states in the membrane interface.Unstructured peptides fold into the membrane interface because partitioned hydrogen-bonded peptide bonds are energetically favorable compared to free peptide bonds (1–3). This folding process is central to the mechanisms of antimicrobial and cell-penetrating peptides, as well as to lipid interactions and stabilities of larger membrane proteins (4). The energetics of peptide partitioning into membrane interfaces can be described by a thermodynamic cycle (Fig. 1). State A is a theoretical state representing the fully unfolded peptide in water, B is the unfolded peptide in the membrane interface, C is the peptide in water, and D is the folded peptide in the membrane. The population of peptides in solution (State C) is best described as an ensemble of folded and unfolded conformations, whereas the population of peptides in State D generally is assumed to have a single, well-defined helicity, as shown in Fig. 1 A (5). Given that, in principle, folding in solution and in the membrane interface should follow the same basic rules, peptides in state D could reasonably be assumed to also be an ensemble. A fundamental question (5) is therefore whether peptides in state D can be correctly described as having a single helicity. Because differentiating an ensemble of conformations and a single conformation may be an impossible experimental task (5), molecular-dynamics (MD) simulations provide a unique high-resolution view of the phenomenon.Open in a separate windowFigure 1Thermodynamic cycles for peptide partitioning into a membrane interface. States A and B correspond to the fully unfolded peptide in solution and membrane interface, respectively. The folded peptide in solution is best described as an ensemble of unfolded and folded conformations (State C). State D is generally assumed to be one of peptides with a narrow range of conformations, but the state could actually be an ensemble of states as in the case of State C.Melittin is a 26-residue, amphipathic peptide that partitions strongly into membrane interfaces and therefore has become a model system for describing folding energetics (3,6–8). Here, we describe the structural dynamics of melittin in a dioleoylphosphatidylcholine (DOPC) bilayer by means of two extensive MD simulations using two different force fields.We extended a 12-ns equilibrated melittin-DOPC system (9) by 17 μs using the Anton specialized hardware (10) with the CHARMM22/36 protein/lipid force field and CMAP correction (11,12) (see Fig. S1 and Fig. S2 in the Supporting Material). To explore force-field effects, a similar system was simulated for 2 μs using the OPLS force field (13) (see Methods in the Supporting Material). In agreement with x-ray diffraction measurements on melittin in DOPC multilayers (14), melittin partitioned spontaneously into the lipid headgroups at a position below the phosphate groups at similar depth as glycerol/carbonyl groups (Fig. 2).Open in a separate windowFigure 2Melittin partitioned into the polar headgroup region of the lipid bilayer. (A) Snapshot of the simulation cell showing two melittin molecules (MLT1 and MLT2, in yellow) at the lipid-water interface. (B) Density cross-section of the simulation cell extracted from the 17-μs simulation. The peptides are typically located below the lipid phosphate (PO4) groups, in a similar depth as the glycerol/carbonyl (G/C) groups.To describe the secondary structure for each residue, we defined helicity by backbone dihedral angles (φ, ψ) within 30° from the ideal α-helical values (–57°, –47°). The per-residue helicity in the CHARMM simulation displays excellent agreement with amide exchange rates from NMR measurements that show a proline residue to separate two helical segments, which are unfolded below Ala⁵ and above Arg²² (15) (Fig. 3 A). In contrast, the OPLS simulation failed to reproduce the per-residue helicity except for a short central segment (see Fig. S3).Open in a separate windowFigure 3Helicity and conformational distribution of melittin as determined via MD simulation. (A) Helicity per residue for MLT1 and MLT2. (B) Corresponding evolution of the helicity. (C) Conformational distributions over the entire 17-μs simulation.Circular dichroism experiments typically report an average helicity of ∼70% for melittin at membrane interfaces (3,6,16,17), but other methods yield average helicities as high as 85% (15,18). Our CHARMM simulations are generally consistent with the experimental results, especially amide-exchange measurements (15); melittin helicity averaged to 78% for MLT1, whereas MLT2 transitioned from 75% to 89% helicity at t ≈ 8 μs, with an overall average helicity of 82% (Fig. 3 B). However, in the OPLS simulation, melittin steadily unfolds over the first 1.3 μs, after which the peptide remains only partly folded, with an average helicity of 33% (see Fig. S3). Similar force-field-related differences in peptide helicity were recently reported, albeit at shorter timescales (19). Although suitable NMR data are not presently available, we have computed NMR quadrupolar splittings for future reference (see Fig. S4).To answer the question asked in this article—whether the conformational space of folded melittin in the membrane interface can be described by a narrow distribution—the helicity distributions for the equilibrated trajectories are shown in Fig. 3 C. Whereas MLT1 in the CHARMM simulation produces a single, narrow distribution of the helicity, MLT2 has a bimodal distribution as a consequence of the folding event at t ≈ 8 μs (Fig. 3 C). We note that CHARMM force fields have a propensity for helix-formation and this transition might therefore be an artifact. We performed a cluster analysis to describe the structure of the peptide in the membrane interface. The four most populated conformations in the CHARMM simulation are shown in Fig. 4. The dominant conformation for both peptides was a helix kinked at G12 and unfolded at the last 5–6 residues of the C-terminus. The folding transition of MLT2 into a complete helix is visible by the 48% occupancy of a fully folded helix.Open in a separate windowFigure 4Conformational clusters of the two melittin peptides (MLT1 and MLT2) from the 17-μs CHARMM simulation in DOPC. Clustering is based on Cα-RMSD with a cutoff criterion of 2 Å.We conclude that the general assumption when calculating folding energetics holds: Folded melittin partitioned into membrane interfaces can be described by a narrow distribution of conformations. Furthermore, extended (several microsecond) simulations are needed to differentiate force-field effects. Although the CHARMM and OPLS simulations would seem to agree for the first few hundred nanoseconds, the structural conclusions differ drastically with longer trajectories, with CHARMM parameters being more consistent with experiments. However, as implied by the difference in substate distributions between MLT1 and MLT2, 17 μs might not be sufficient to observe the fully equilibrated partitioning process. The abrupt change in MLT2 might indicate that the helicity will increase to greater than experimentally observed in a sufficiently long simulation. On the other hand, it could be nothing more than a transient fluctuation. Increased sampling will provide further indicators of convergence of the helix partitioning process.     

Mitofusins: ubiquitylation promotes fusion

Mitochondrial genes including Mfn2 are at the center of many diseases, underscoring their potential as a therapeutical target. The Chen group now identified 15-oxospiramilactone as a chemical inhibitor of the mammalian deubiquitylase USP30, acting on Mfn1 and Mfn2.Mitofusins, Fzo1 in yeast and Mfn1 and Mfn2 in mammals, are ubiquitylated and this post-translational modification has both positive and negative consequences on mitochondrial fusion¹. The process of ubiquitylation requires enzymes belonging to three classes of proteins called E1, E2 and E3, which catalyze a cascade of successive steps leading to the covalent attachment of the modifier to its target protein². Deubiquitylating enzymes render this modification reversible, thus offering further possibilities for regulation². Ubiquitylation of mitofusins leads to their proteolyic breakdown, inhibiting fusion of mitochondria that consequently undergo fragmentation (Figure 1, left panel)^1,3. For example in response to mitochondrial depolarization or apoptotic stimuli, E3 ligases like Parkin and Huwe1 ubiquitylate and target Mfn1 and Mfn2 to the proteasome (Figure 1, left panel)^3,4. However, ubiquitylation of mitofusins is a dual process and a non-proteolytic role of mitofusin ubiquitylation that promotes mitochondrial fusion is now emerging¹. This opposing mechanism was first described in yeast, where the isopeptidases Ubp12 and Ubp2 that deubiquitylate Fzo1 have been identified⁵. Inhibition and activation of mitochondrial fusion by ubiquitylation enable different morphologies of mitochondria ranging from a multitude of small organelles to a hyperconnected network (Figure 1)⁵. In a recent paper published in Cell Research, Yue et al.⁶ reveal that a similar process is present in mammalian cells. The authors report that the isopeptidase USP30 acts on ubiquitylated forms of Mfn1 and Mfn2 that stimulate mitochondrial fusion (Figure 1, right panel). This discovery identifies for the first time in mammals a positive role of ubiquitylation in the regulation of Mfn1 and Mfn2 fusion activity⁶.Open in a separate windowFigure 1Dual roles of ubiquitylation and deubiquitylation of mitofusins Mfn1 and Mfn2, the key effectors for mitochondrial fusion, in regulating mitochondrial fusion. On one hand, ubiquitylation of Mfn1 and Mfn2 by E3 ligases like Parkin or Huwe1 targets their proteasomal degradation and inhibits mitochondrial fusion, which results in mitochondrial fragmentation due to unopposed fission events. On the other hand, ubiquitylation of Mfn1 and Mfn2 by an unknown E3 ligase enhances their activity and promotes mitochondrial fusion. This positive regulation is counteracted by the deubiquitylase USP30, targeted by the small molecule inhibitor 15-oxospiramilactone.Moreover, Yue et al.⁶ identified the first small molecule inhibitor of mitochondrial fusion, 15-oxospiramilactone, which targets USP30 in both human and mouse cell lines. 15-oxospiramilactone is a semi-synthetic diterpene alkaloid of 330 Da that can be chemically synthetized through an oxidation reaction from spiramines extracted from the roots of a Chinese herbal medicine Spiraea japonica (Rosaceae). Inhibition of USP30 increased ubiquitylation of Mfn1 and Mfn2 and led to an elongation of the mitochondrial network (Figure 1, right panel)^6,7. USP30 is a cysteine ubiquitin isopeptidase N-terminally anchored to the outer membrane of mitochondria, which was previously shown to regulate mitochondrial morphology dependent on Mfn1 and Mfn2⁷. USP30 knockdown leads to mitochondrial elongation, a phenotype rescued by ectopic expression of wild-type USP30, while the catalytically inactive mutant C77S USP30 failed to revert⁷. Yue et al.⁶ show that 15-oxospiramilactone directly interacts with USP30, which also depends on its catalytically active cysteine, and inhibits the DUB activity of USP30 on tetraubiquitin chains. Moreover, they demonstrate that inhibition of USP30 and subsequent mitochondrial elongation are due to stimulated mitochondrial fusion activity, apparently with no influence on mitochondrial fission⁶. Concomitantly, cells showed increased ubiquitylation of Mfn1 and Mfn2 without significant changes in protein turnover of these two proteins⁶. Therefore, in analogy to findings in yeast, ubiquitylation of Mfn1 and Mfn2 can either signal them to activate mitochondrial fusion or in contrast promote their proteasomal degradation, resulting in mitochondrial fission (Figure 1).Importantly, 15-oxospiramilactone reverts the mitochondrial fragmentation phenotype of single Mfn-knockout (Mfn1^−/− or Mfn2^−/−) cells, suggesting that mitochondrial fusion depends on the ubiquitylation of both mitofusin proteins⁶. In yeast, the importance of ubiquitylation was proven by directly attaching a deubiquitylase to Fzo1, which resulted in a non-ubiquitylated and non-functional Fzo1 protein⁵. In addition, the identification and the subsequent mutagenesis study of the ubiquitylation sites in Fzo1 confirmed an interplay between ubiquitylation and oligomerization in mitochondrial fusion in S. cerevisiae⁵. Impairing the yeast E3 ligase SCF^Mdm30 inhibited mitochondrial fusion and, conversely, ablation of UBP12 led to more fusion events^5,8. Given this new identification of USP30 as the functional orthologue of the yeast Ubp12, future studies will certainly aim at the identification of the E3 ligase counterpart of SCF^Mdm30 and ubiquitylation sites in Mfn1 and Mfn2. In addition to USP30 inhibition, other conditions leading to mitochondrial hyperfusion have been previously observed, such as mild stress conditions that increase reactive oxygen species (ROS)⁹. Importantly, oxidative stress and mitochondrial fusion are directly linked as ROS induces disulphide switching of Mfn2 to oligomeric forms that promote mitochondrial fusion⁹. It would be interesting to investigate whether 15-oxospiramilactone also affects the generation of disulphide-mediated mitofusin oligomers, thus activating mitochondrial fusion.Mutations in Mfn2 are causative for the Charcot-Marie-Tooth type 2A neuropathy, an autosomal dominant disorder of the peripheral nervous system that mainly affects axons and lower extremities¹. Deficiencies in Parkin and Mfn2 ubiquitylation were also linked to Parkinson''s disease³. In addition to neuropathies, Mfn2 is associated to other diseases like cardiomyophathies and diabetes¹. Yue et al.⁶ found that 15-oxospiramilactone reverted phenotypes arising from the lack of Mfn1 or Mfn2. It restored the normal distribution of mtDNA, allowed recovery of the ΔΨm and increased the ATP levels and OXPHOS capacity of the rebuilt mitochondrial network. Therefore, this study potentiates 15-oxospiramilactone for therapeutical benefit. The anti-cancer properties of 15-oxospiramilactone, also named S3 or NC043, have been previously reported^10,11. It inhibits Wnt/β-catenin signaling and colon cancer cell tumorigenesis in a xenograft model¹⁰. Moreover, 15-oxospiramilactone increases Bim expression and apoptosis to inhibit tumor growth from Bax^−/−/Bak^−/− cells implanted in mice¹¹. However, Yue et al.⁶ show that the effect of 15-oxospiramilactone in mitochondrial fusion is independent of apoptosis and suggest that the difference is due to drug concentration. Indeed, previous anti-cancer studies used 15-oxospiramilactone at a concentration range of 3.75-15 μM^10,11, whereas 2 μM suffice to inhibit USP30⁶. Further studies are needed to address the clinical relevance of 15-oxospiramilactone and USP30 in Mfn2-associated diseases.     

Mitochondrial Protein Synthesis,Import, and Assembly

The mitochondrion is arguably the most complex organelle in the budding yeast cell cytoplasm. It is essential for viability as well as respiratory growth. Its innermost aqueous compartment, the matrix, is bounded by the highly structured inner membrane, which in turn is bounded by the intermembrane space and the outer membrane. Approximately 1000 proteins are present in these organelles, of which eight major constituents are coded and synthesized in the matrix. The import of mitochondrial proteins synthesized in the cytoplasm, and their direction to the correct soluble compartments, correct membranes, and correct membrane surfaces/topologies, involves multiple pathways and macromolecular machines. The targeting of some, but not all, cytoplasmically synthesized mitochondrial proteins begins with translation of messenger RNAs localized to the organelle. Most proteins then pass through the translocase of the outer membrane to the intermembrane space, where divergent pathways sort them to the outer membrane, inner membrane, and matrix or trap them in the intermembrane space. Roughly 25% of mitochondrial proteins participate in maintenance or expression of the organellar genome at the inner surface of the inner membrane, providing 7 membrane proteins whose synthesis nucleates the assembly of three respiratory complexes.TO think about how mitochondrial proteins are synthesized, imported, and assembled, it is useful to have a clear picture of the organellar structures that they, along with membrane lipids, compose and the functions that they carry out. As almost every schoolchild learns, mitochondria carry out oxidative phosphorylation, the controlled burning of nutrients coupled to ATP synthesis. Since Saccharomyces cerevisiae prefers to ferment sugars, respiration is a dispensable function and nonrespiring mutants are viable [although they cannot undergo meiosis (Jambhekar and Amon 2008)]. However, mitochondria themselves are not dispensable. A substantial fraction of intermediary metabolism occurs in mitochondria (Strathern et al. 1982), and at least one of these pathways, iron–sulfur cluster assembly, is essential for growth (Kispal et al. 2005). Thus, any mutation that prevents the biogenesis of mitochondria by, for example, preventing the import of protein constituents from the cytoplasm, is lethal (Baker and Schatz 1991).The mitochondria of S. cerevisiae are tubular structures at the cell cortex. While the number of distinct compartments can range from 1 to ∼50 depending upon conditions (Stevens 1981; Pon and Schatz 1991), continual fusion and fission events among them effectively form a single dynamic network (Nunnari et al. 1997). The outer membrane surrounds the tubules. The inner membrane has a boundary domain closely juxtaposed beneath the outer membrane and cristae domains that project internally from the boundary into the matrix (Figure 1A). The matrix is the aqueous compartment surrounded by the inner membrane. The aqueous intermembrane space lies between the membranes and is continuous with the space within cristae.Open in a separate windowFigure 1 Overview of mitochondrial structure in yeast. (A) Schematic of compartments comprising mitochondrial tubules. The outer membrane surrounds the organelle. The inner membrane surrounds the matrix and consists of two domains, the inner boundary membrane and the cristae membranes, which are joined at cristae junctions. The intermembrane space lies between the outer membrane and inner membrane. (B) Electron tomograph image of a highly contracted yeast mitochondrion observed en face (a) with the outer membrane (red) and (b) without the outer membrane. Reprinted by permission from John Wiley & Sons from Mannella et al. (2001).Inner membrane cristae are often depicted as baffles emanating from the boundary domain. However, electron tomography of mitochondria from several species, including yeast, shows that cristae actually emanate from the boundary membrane as narrow tubular structures at sites termed “crista junctions” and expand as they project into the matrix (Frey and Mannella 2000; Mannella et al. 2001) (Figure 1B). It seems clear that the boundary and cristae domains of the inner membrane have distinct compositions with respect to the respiratory complexes that are embedded preferentially in the cristae membrane domains, as well as other components (Vogel et al. 2006; Wurm and Jakobs 2006; Rabl et al. 2009; Suppanz et al. 2009; Zick et al. 2009; Davies et al. 2011).The outer and inner boundary membranes are connected at multiple contact sites, at least some of which are involved in protein translocation and may be transient (Pon and Schatz 1991). In addition, there appear to be firm contact sites, not directly involved with protein translocation, preferentially colocalized with crista junctions (Harner et al. 2011a).Overall, there appear to be ∼1000 distinct proteins in yeast mitochondria (Premsler et al. 2009). One series of proteomic studies on highly purified organelles identified 851 proteins thought to represent 85% of the total number of species (Sickmann et al. 2003; Reinders et al. 2006; Zahedi et al. 2006). Another study identified an additional 209 candidates (Prokisch et al. 2004). A computationally driven search for candidates involved in yeast mitochondrial function, coupled with experiments to assay respiratory function and maintenance of mitochondrial DNA (mtDNA), identified 109 novel candidates, although many of these may not be mitochondrial per se (Hess et al. 2009). Taking the boundary and cristae domains together, the inner membrane is the most protein-rich mitochondrial compartment, followed by the matrix (Daum et al. 1982).Only eight of the yeast mitochondrial proteins detected in proteomic studies are encoded by mtDNA and synthesized within the organelle. They are hydrophobic subunits of respiratory complexes III (bc₁ complex or ubiquinol-cytochrome c reductase), IV (cytochrome c oxidase), and V (ATP synthase), as well as a hydrophilic mitochondrial small subunit ribosomal protein. The remaining ∼99% of yeast mitochondrial proteins are encoded by nuclear genes, synthesized in cytoplasmic ribosomes, and imported into the organelle.An overview of known nuclearly encoded mitochondrial protein functions (Figure 2) reveals that ∼25% of them are involved directly in genome maintenance and expression of the eight major mitochondrial genes (Schmidt et al. 2010). The functions of ∼20% of the proteins are not known. Fifteen percent are involved in the well-known processes of energy metabolism. Protein translocation, folding, and turnover functions occupy ∼10% of mitochondrial proteins.Open in a separate windowFigure 2 Classification of identified mitochondrial proteins according to function. Reprinted by permission from Nature Publishing Group from Schmidt et al. (2010).The following discussion reviews our understanding of the biogenesis of mitochondria starting on the outside, the cytoplasm, and working inward through the mitochondrial compartments.     

Modeling Lignin Polymerization. I. Simulation Model of Dehydrogenation Polymers

Lignin is a heteropolymer that is thought to form in the cell wall by combinatorial radical coupling of monolignols. Here, we present a simulation model of in vitro lignin polymerization, based on the combinatorial coupling theory, which allows us to predict the reaction conditions controlling the primary structure of lignin polymers. Our model predicts two controlling factors for the β-O-4 content of syringyl-guaiacyl lignins: the supply rate of monolignols and the relative amount of supplied sinapyl alcohol monomers. We have analyzed the in silico degradability of the resulting lignin polymers by cutting the resulting lignin polymers at β-O-4 bonds. These are cleaved in analytical methods used to study lignin composition, namely thioacidolysis and derivatization followed by reductive cleavage, under pulping conditions, and in some lignocellulosic biomass pretreatments.Lignins are aromatic polymers that are predominantly present in secondarily thickened cell walls. These polymers make the cell wall rigid and impervious, allowing transport of water and nutrients through the vascular system and protecting plants against microbial invasion. Lignins are heterogeneous polymers derived from phenylpropanoid monomers, mainly the hydroxycinnamyl alcohols coniferyl alcohol (G-monomer) and sinapyl alcohol (S-monomer) and minor amounts of p-coumaryl alcohol (H-monomer). These monolignols differ in their degree of aromatic methoxylation (-OCH₃ group; Fig. 1). The resulting units in the lignin polymer are the guaiacyl (G), syringyl (S), and p-hydroxyphenyl (H) units. They are linked by a variety of chemical bonds (Fig. 2) that have different chemical properties (Boerjan et al., 2003; Ralph et al., 2004; Vanholme et al., 2008).Open in a separate windowFigure 1.Chemical structures of three monolignols. A, H-monomer (p-coumaryl alcohol). B, G-monomer (coniferyl alcohol). C, S-monomer (sinapyl alcohol). G- and S-monomers are considered in our simulations. The G-monomer is methoxylated (-OCH₃ group) on position 3, and the S-monomer is methoxylated on positions 3 and 5.Open in a separate windowFigure 2.Chemical structures resulting from the possible bonding between two monomers (A) or a monomer and the bindable end of an oligomer (B). X and Y in the monomers denote the absence (for a G-unit) or presence (for an S-unit) of a methoxyl group at position 5 (see Fig. 1). The red line indicates the bonds generated by couplings of the B position and B, 4, or 5 position.Lignification is the process by which monomers and/or oligomers are polymerized via radical coupling reactions and typically occurs after the polysaccharides have been laid down in the cell wall. Lignin composition varies among cell types and can even be different in individual cell wall layers (Ruel et al., 2009). Lignin composition is also influenced by environmental conditions; for example, lignin in compression wood is enriched in H-units (Timell, 1986). Hence, both developmental and environmental parameters influence the composition and thus the structure of the lignin polymer (Boerjan et al., 2003; Ralph et al., 2004).Lignin is one of the main negative factors in the conversion of lignocellulosic plant biomass into pulp and bioethanol (Lynd et al., 1991; Hill et al., 2006). In these processes, lignin needs to be degraded by chemical or mechanical processes that are expensive and often environmentally polluting. Hence, major research efforts are devoted toward understanding lignin biosynthesis and structure. It has already been shown that reducing lignin content and modifying its composition in transgenic plants can result in dramatic improvements in pulping efficiency (Pilate et al., 2002; Baucher et al., 2003; Huntley et al., 2003; Leplé et al., 2007) and in the conversion of biomass into bioethanol (Stewart et al., 2006; Chen and Dixon, 2007; Custers, 2009). These altered biomass properties are related to the alterations in lignin composition and structure in terms of the frequencies of the lignin units and the bond types connecting them and possibly also their interaction with hemicelluloses (Ralph et al., 2004; Ralph, 2006).To study the parameters that influence lignin structure, lignin polymerization has been mimicked in vitro by experiments with dehydrogenation polymers (DHPs; Terashima et al., 1995). Indeed, lignification can be mimicked by oxidizing monolignols using a peroxidase, such as horseradish peroxidase (HRP), and supplying its cofactor hydrogen peroxide, producing synthetic DHP lignins. Monolignol oxidation can also be achieved without enzymes (e.g. by using transition metal one-electron oxidants, such as copper acetate). Some of these biomimetic DHPs have been suggested to be better models for wood lignins than HRP-generated DHPs (Landucci, 2000).In DHP experiments, the monolignols are either added in bulk (Zulauf experiment) or dropwise (Zutropf experiment) to the reaction mixture, yielding lignin polymers with very different bond frequencies (Freudenberg, 1956). Zutropf experiments approach the in vivo formation of lignin, which depends on the slow introduction of monolignols into the wall matrix via diffusion to the site of incorporation (Hatfield and Vermerris, 2001). Because the exact reaction conditions are known, such in vitro experiments have provided insight into the lignification process in planta. In this way, numerous factors were shown to influence lignin structure, including the relative supply of the monolignols, the pH, the presence of polysaccharides, hydrogen peroxide concentrations, and cell wall matrix elements in general (Grabber et al., 2003; Vanholme et al., 2008).Computer simulations of lignin polymerization can help explain and predict lignin structure from low-level chemical kinetic factors, including subunit-coupling probabilities and monolignol synthesis rates. Such models are helpful in explaining the mechanism behind a range of controlling factors identified in the experimental work, including (1) the ratio of coniferyl versus sinapyl alcohol monolignols, (2) the monolignol supply rate, and (3) the abundance of alternative monomers present during lignin biosynthesis in mutants and transgenics. Thus, computer models will also help in suggesting new targets for controlled lignin biosynthesis.Here, we propose a simulation model of synthetic lignin polymerization that is based upon an emerging consensus from a variety of observations and derives from a series of previous models of lignin polymerization (Glasser and Glasser, 1974; Glasser et al., 1976; Jurasek, 1995; Roussel and Lim, 1995). Our model uses a symbolic grammar to describe a constructive dynamical system (Fontana, 1992) or a rule-based system (Feret et al., 2009) in which it is not necessary to define all possible products in advance. We assume that G- and S-monomers and newly formed oligomers couple in a well-mixed medium, depending on coupling rules and experimentally measured coupling probabilities. To develop the model, we have used information from DHP experiments rather than natural lignins, as they are formed in a well-mixed medium and their reaction conditions are well known (e.g. the influx rate of monomers). Using information from natural lignin would have further complicated our model, as the structures of natural lignin polymers are influenced by many factors, including the possible involvement of dirigent proteins (Davin and Lewis, 2005), steric hindrance by polysaccharides, spatiotemporal regulation, and modifications during isolation procedures (Boerjan et al., 2003; Ralph et al., 2004).Using our simulation models, we study how putative controlling factors of lignin primary structure, including the influx rate of monomers and the relative amount of S-monomers, affect in silico lignin synthesis, and we compare our predictions with in vitro experiments. To predict the degradability of lignins formed in our simulations, we apply an in silico thioacidolysis, which cleaves the polymers at their β-O-4 positions. This simulates the molecular action of two of the most used methods to analyze lignin composition, thioacidolysis (Lapierre, 1993; Baucher et al., 2003) and derivatization followed by reductive cleavage (Lu and Ralph, 1997). The G+S-monomer yield is often taken as a reflection of the fraction of units bound by β-O-4 bonds. Cleavage of β-O-4 bonds is also the most important reaction in kraft pulping of wood (Baucher et al., 2003). The model predicts from first principles (1) that DHP lignins formed under Zutropf conditions have a higher β-O-4 content than those formed under Zulauf conditions, (2) that DHP lignins formed with high S content have a higher β-O-4 content than those formed with high G content, and (3) that a higher β-O-4 content does not necessarily reduce the average length of lignin fragments generated during in silico thioacidolysis.     

3D-SIM Super-resolution of FtsZ and Its Membrane Tethers in Escherichia coli Cells

FtsZ, a bacterial homolog of eukaryotic tubulin, assembles into the Z ring required for cytokinesis. In Escherichia coli, FtsZ interacts directly with FtsA and ZipA, which tether the Z ring to the membrane. We used three-dimensional structured illumination microscopy to compare the localization patterns of FtsZ, FtsA, and ZipA at high resolution in Escherichia coli cells. We found that FtsZ localizes in patches within a ring structure, similar to the pattern observed in other species, and discovered that FtsA and ZipA mostly colocalize in similar patches. Finally, we observed similar punctate and short polymeric structures of FtsZ distributed throughout the cell after Z rings were disassembled, either as a consequence of normal cytokinesis or upon induction of an endogenous cell division inhibitor.The assembly of the bacterial tubulin FtsZ has been well studied in vitro, but the fine structure of the cytokinetic Z ring it forms in vivo is not well defined. Super-resolution microscopy methods including photoactivated localization microscopy (PALM) and three-dimensional-structured illumination microscopy (3D-SIM) have recently provided a more detailed view of Z-ring structures. Two-dimensional PALM showed that Z rings in Escherichia coli are likely composed of loosely-bundled dynamic protofilaments (1,2). Three-dimensional PALM studies of Caulobacter crescentus initially showed that Z rings were comprised of loosely bundled protofilaments forming a continuous but dynamic ring (1–3). However, a more recent high-throughput study showed that the Z rings of this bacterium are patchy or discontinuous (4), similar to Z rings of Bacillus subtilis and Staphylococcus aureus using 3D-SIM (5). Strauss et al. (5) also demonstrated that the patches in B. subtilis Z rings are highly dynamic.Assembly of the Z ring is modulated by several proteins that interact directly with FtsZ and enhance assembly or disassembly (6). For example, FtsA and ZipA promote ring assembly in E. coli by tethering it to the cytoplasmic membrane (7,8). SulA is an inhibitor of FtsZ assembly, induced only after DNA damage, which sequesters monomers of FtsZ to prevent its assembly into a Z ring (9). Our initial goals were to visualize Z rings in E. coli using 3D-SIM, and then examine whether any FtsZ polymeric structures remain after SulA induction. We also asked whether FtsA and ZipA localized in patchy patterns similar to those of FtsZ.We used a DeltaVision OMX V4 Blaze microscope (Applied Precision, GE Healthcare, Issaquah, WA) to view the high-resolution localization patterns of FtsZ in E. coli cells producing FtsZ-GFP (Fig. 1). Three-dimensional views were reconstructed using softWoRx software (Applied Precision). To rule out GFP artifacts, we also visualized native FtsZ from a wild-type strain (WM1074) by immunofluorescence (IF).Open in a separate windowFigure 1Localization of FtsZ in E. coli. (A) Cell with a Z ring labeled with FtsZ-GFP. (B) Rotated view of Z ring in panel A. (C) Cell with a Z ring labeled with DyLight 550 (Thermo Fisher Scientific, Waltham, MA). (D) Rotated view of Z ring in panel C. (B1 and D1) Three-dimensional surface intensity plots of Z rings in panels B and D, respectively. (E) A dividing cell producing FtsZ-GFP. The cell outline is shown in the schematic. (Asterisk) Focus of FtsZ localization; (open dashed ovals) filamentous structures of FtsZ. Three-dimensional surface intensity plots were created using the software ImageJ (19). Scale bars, 1 μm.Both FtsZ-GFP (Fig. 1, A, B, and B1) and IF staining for FtsZ (Fig. 1, C, D, and D1) consistently localized to patches around the ring circumference, similar to the B. subtilis and C. crescentus FtsZ patterns (4,5). Analysis of fluorescence intensities (see Fig. S1, A and B, in the Supporting Material) revealed that the majority of Z rings contain one or more gaps in which intensity decreases to background levels (82% for FtsZ-GFP and 69% for IF). Most rings had 3–5 areas of lower intensity, but only a small percentage of these areas had fluorescence below background intensity (34% for FtsZ-GFP and 21% for IF), indicating that the majority of areas with lower intensity contain at least some FtsZ.To elucidate how FtsZ transitions from a disassembled ring to a new ring, we imaged a few dividing daughter cells before they were able to form new Z rings (Fig. 1 E). Previous conventional microscopy had revealed dynamic FtsZ helical structures (10), but the resolution had been insufficient to see further details. Here, FtsZ visualized in dividing cells by 3D-SIM localized throughout as a mixture of patches and randomly-oriented short filaments (asterisk and dashed oval in Fig. 1, respectively). These structures may represent oligomeric precursors of Z ring assembly.To visualize FtsZ after Z-ring disassembly another way, we overproduced SulA, a protein that blocks FtsZ assembly. We examined E. coli cells producing FtsZ-GFP after induction of sulA expression from a pBAD33-sulA plasmid (pWM1736) with 0.2% arabinose. After 30 min of sulA induction, Z rings remained intact in most cells (Fig. 2 A and data not shown). The proportion of cellular FtsZ-GFP in the ring before and after induction of sulA was consistent with previous data (data not shown) (1,11).Open in a separate windowFigure 2Localization of FtsZ after overproduction of SulA. (A) Cell producing FtsZ-GFP after 0.2% arabinose induction of SulA for 30 min. (B) After 45 min. (B1) Magnified cell shown in panel B. (C) Cell producing native FtsZ labeled with AlexaFluor 488 (Life Technologies, Carlsbad, CA) 30 min after induction; (D) 45 min after induction. (D1) Magnified cell shown in panel D. Scale bars, 1 μm. (Asterisk) Focus of FtsZ localization; (open dashed ovals) filamentous structures of FtsZ.Notably, after 45 min of sulA induction, Z rings were gone (Fig. 2, B and B1), replaced by numerous patches and randomly-oriented short filaments (asterisk and dashed ovals in Fig. 2), similar to those observed in a dividing cell. FtsZ normally rapidly recycles from free monomers to ring-bound polymers (11), but a critical concentration of SulA reduces the pool of available FtsZ monomers, resulting in breakdown of the Z ring (9). The observed FtsZ-GFP patches and filaments are likely FtsZ polymers that disassemble before they can organize into a ring.We confirmed this result by overproducing SulA in wild-type cells and detecting FtsZ localization by IF (Fig. 2, C, D, and D1). The overall fluorescence patterns in cells producing FtsZ-GFP versus cells producing only native FtsZ were similar (Fig. 2, B1 and D1), although we observed fewer filaments with IF, perhaps because FtsZ-GFP confers slight resistance to SulA, or because the increased amount of FtsZ in FtsZ-GFP producing cells might titrate the SulA more effectively.Additionally, we wanted to observe the localization patterns of the membrane tethers FtsA and ZipA. Inasmuch as both proteins bind to the same C-terminal conserved tail of FtsZ (12–14), they would be expected to colocalize with the circumferential FtsZ patches in the Z ring. We visualized FtsA using protein fusions to mCherry and GFP (data not shown) as well as IF using a wild-type strain (WM1074) (Fig. 3 A). We found that the patchy ring pattern of FtsA localization was similar to the FtsZ pattern. ZipA also displayed a similar patchy localization in WM1074 by IF (Fig. 3 B).Open in a separate windowFigure 3Localization of FtsA (A) and ZipA (B) by IF using AlexaFluor 488. (C) FtsA-GFP ring. (D) Same cell shown in panel C with ZipA labeled with DyLight 550. (C1 and D1) Three-dimensional surface intensity plots of FtsA ring from panel C or ZipA ring from panel D, respectively. (E) Merged image of FtsA (green) and ZipA (red) from the ring shown in panels C and D. (F) Intensity plot of FtsA (green) and ZipA (red) of ring shown in panel E. The plot represents intensity across a line drawn counterclockwise from the top of the ring around the circumference, then into its lumen. Red/green intensity plot and three-dimensional surface intensity plots were created using the software ImageJ (19). Scale bar, 1 μm.To determine whether FtsA and ZipA colocalized to these patches, we used a strain producing FtsA-GFP (WM4679) for IF staining of ZipA using a red secondary antibody. FtsA-GFP (Fig. 3 C) and ZipA (Fig. 3 D) had similar patterns of fluorescence, although the three-dimensional intensity profiles (Fig. 3, C1 and D1) reveal slight differences in intensity that are also visible in a merged image (Fig. 3 E). Quantitation of fluorescence intensities around the circumference of the rings revealed that FtsA and ZipA colocalized almost completely in approximately half of the rings analyzed (Fig. 3 F, and see Fig. S2 A), whereas in the other rings there were significant differences in localization in one or more areas (see Fig. S2 B). FtsA and ZipA bind to the same C-terminal peptide of FtsZ and may compete for binding. Cooperative self-assembly of FtsA or ZipA might result in large-scale differential localization visible by 3D-SIM.In conclusion, our 3D-SIM analysis shows that the patchy localization of FtsZ is conserved in E. coli and suggests that it may be widespread among bacteria. After disassembly of the Z ring either in dividing cells or by excess levels of the cell division inhibitor SulA, FtsZ persisted as patches and short filamentous structures. This is consistent with a highly dynamic population of FtsZ monomers and oligomers outside the ring, originally observed as mobile helices in E. coli by conventional fluorescence microscopy (10) and by photoactivation single-molecule tracking (15). FtsA and ZipA, which bind to the same segment of FtsZ and tether it to the cytoplasmic membrane, usually display a similar localization pattern to FtsZ and each other, although in addition to the differences we detect by 3D-SIM, there are also likely differences that are beyond its ∼100-nm resolution limit in the X,Y plane.As proposed previously (16), gaps between FtsZ patches may be needed to accommodate a switch from a sparse Z ring to a more condensed ring, which would provide force to drive ring constriction (17). If this model is correct, the gaps should close upon ring constriction, although this may be beyond the resolution of 3D-SIM in constricted rings. Another role for patches could be to force molecular crowding of low-abundance septum synthesis proteins such as FtsI, which depend on FtsZ/FtsA/ZipA for their recruitment, into a few mobile supercomplexes.How are FtsZ polymers organized within the Z-ring patches? Recent polarized fluorescence data suggest that FtsZ polymers are oriented both axially and circumferentially within the Z ring in E. coli (18). The seemingly random orientation of the non-ring FtsZ polymeric structures we observe here supports the idea that there is no strong constraint requiring FtsZ oligomers to follow a circumferential path around the cell cylinder. The patches of FtsZ in the unperturbed E. coli Z ring likely represent randomly oriented clusters of FtsZ filaments that are associated with ZipA, FtsA, and essential septum synthesis proteins. New super-resolution microscopy methods should continue to shed light on the in vivo organization of these protein assemblies.     

Cell biology in development: Cell division and the maintenance of epithelial order

Epithelia are polarized layers of adherent cells that are the building blocks for organ and appendage structures throughout animals. To preserve tissue architecture and barrier function during both homeostasis and rapid growth, individual epithelial cells divide in a highly constrained manner. Building on decades of research focused on single cells, recent work is probing the mechanisms by which the dynamic process of mitosis is reconciled with the global maintenance of epithelial order during development. These studies reveal how symmetrically dividing cells both exploit and conform to tissue organization to orient their mitotic spindles during division and establish new adhesive junctions during cytokinesis.The association of large numbers of cells in tightly organized epithelial layers is a unique and defining feature of Metazoa. Although classical studies of development once labeled distinct embryonic regions as territories, fields, layers, placodes, and primordia, we now know many of these structures to be primarily constructed from epithelial sheets. Epithelial structure and function are critically dependent on cell polarization, which is coupled to the targeted assembly of adhesive junctions along the apicolateral membranes of adjacent cells (Tepass et al., 2001; Cavey and Lecuit, 2009). In brief, the plasma membrane of epithelial cells is polarized into apical and basolateral domains, each enriched with distinct lipid and protein components (Fig. 1; Rodriguez-Boulan et al., 2005; St Johnston and Ahringer, 2010). At the molecular level, E-cadherins are the major class of adhesion proteins that establish cell–cell connections through homophilic interaction across cell membranes (Takeichi, 1991, 2011; Halbleib and Nelson, 2006; Harris and Tepass, 2010). Whereas E-cadherin is apically enriched in invertebrate epithelia, it is localized along the lateral domain of vertebrate epithelial cells. In both cases, E-cadherin interacts with cytoplasmic actin filaments via the catenin class of adaptor proteins, thus coupling intercellular adhesive contacts to the cytoskeleton (Cavey and Lecuit, 2009; Harris and Tepass, 2010; Gomez et al., 2011). Within this framework, the maintenance of both polarity and cell–cell adhesion are essential for epithelial barrier function and tissue architecture during growth and morphogenesis (Papusheva and Heisenberg, 2010; Guillot and Lecuit, 2013b).Open in a separate windowFigure 1.Architectural implications of orthogonal and planar spindle orientations during epithelial cell division. (A) Programmed orthogonal orientation of the mitotic spindle can promote epithelial stratification, although the remodeling of adhesion and polarity complexes during this process remains an important area for further study. (B) Planar spindle orientation is coordinated with the overall cell polarity machinery and thus facilitates conservation of monolayer organization during rapid cell proliferation.During development, epithelia expand by the combined effects of cell growth (increase in cell size) and cell division (increase in cell numbers). Division events are typically oriented either parallel or orthogonal to the plane of the layer and less frequently at oblique angles (Gillies and Cabernard, 2011). When cells divide orthogonally (perpendicular to the plane of the epithelium), the two daughters will be at least initially nonequivalent with respect to position within the cell layer (Fig. 1 A). Under normal conditions, such programmed orthogonal divisions can be used to effect asymmetric segregation of cell fates or to establish distinct cell types, such as in the developing cortex (Fietz et al., 2010; Hansen et al., 2010) or during morphogenesis of stratified epithelia (Lechler and Fuchs, 2005; Williams et al., 2011). Conversely, when cells divide parallel to the plane of the epithelium (planar orientation; Fig. 1 B), both daughter cells are equivalent with respect to mother cell polarity and tightly integrated in the growing monolayer (Morin and Bellaïche, 2011).During planar division, epithelial cells typically round up, constrict in the middle to form the cytokinetic furrow, and divide symmetrically with respect to the apicobasal axis to produce two equal daughter cells. These daughters construct new cell–cell junctions at their nascent interface, thus integrating into the monolayer (Fig. 2, A–G). Although the intricate relationship between cell polarity and cell division has been explored for many years in the context of asymmetric cell division (Rhyu and Knoblich, 1995; Siller and Doe, 2009; Williams and Fuchs, 2013), recent studies have also begun to explore how epithelia maintain their morphology, integrity, and barrier function during continuous rounds of planar cell division and junction assembly. In this review, we highlight recent findings that provide new insights into the problem of symmetric planar cell division in diverse polarized epithelia, with a focus on two crucial mitotic events: (1) the orientation of cell division and (2) the formation of new cell junctions.Open in a separate windowFigure 2.Progression of planar cell division in an epithelial monolayer. Apical cross section (xy, top row) and longitudinal (xz, bottom row) view of a dividing cell (red). (A) At the level of apical junctions, cells are packed in a polygonal cell arrangement during interphase. (B) In prophase, the dividing nucleus begins to translocate apically as the cell rounds up and maintains a thin basal projection enriched with nonmuscle myosin II and actin (light blue). Notably, this type of nuclear migration is typically observed in pseudostratified columnar epithelia and does not occur in cuboidal and squamous epithelial tissues. (C) Localized molecular landmarks (apical complexes marked as gray bars on cell sides) direct orientation of the mitotic spindle to the plane of the epithelium (arrows). (D) Within the plane of the cell layer, the spindle can be further oriented (arrows) in response to molecular cues, global tissue tension, and local cell geometry. (E and F) After chromosome segregation during anaphase, the cell constricts in the middle and cleaves orthogonal to the plane of the monolayer. (G) After cytokinesis, daughter nuclei move basally and daughter cells form new junctions at their nascent interface (white) while elongating along the apicobasal axis.

Mitotic spindle position and orientation in epithelial cells

Planar orientation of epithelial cell division requires coordinated interaction between the cell polarization machinery and the mitotic spindle itself (Morin and Bellaïche, 2011). In animal cells, the spindle is organized by two symmetrically positioned poles or centrosomes, which nucleate three forms of microtubules (Tanaka, 2010): kinetochore microtubules that attach to the chromosomes, polar microtubules that overlap in an antiparallel fashion over the midplane, and astral microtubules that extend to the cell cortex, which is the actin-rich layer beneath the cell membrane (Lancaster and Baum, 2014). Work in Drosophila melanogaster and vertebrates reveals that at least three factors influence the orientation of this spindle machinery with respect to polarized epithelial architecture: cytoskeletal forces, localized cortical cues, and tissue tension.

Cytoskeletal forces position mitotic nuclei near the apical cell membrane.

In columnar and pseudostratified epithelia where cells elongate along their apicobasal axes, mitotic events are typically restricted to the apical domain of the epithelium (corresponding to the apical membrane of each cell; Fig. 2, C–F). How does the mitotic nucleus achieve the correct apical position? In Drosophila wing discs and zebrafish neuroepithelia, mitotic nuclei and the bulk of the cell cytoplasm are driven apically by actomyosin-dependent cortical contractility at prophase entry (Norden et al., 2009; Leung et al., 2011; Meyer et al., 2011). These events are fundamentally similar to mitotic cell rounding in tissue culture cells (Kunda and Baum, 2009; Lancaster et al., 2013; Lancaster and Baum, 2014). In many epithelia, as the cell rounds up and the nucleus translocates apically, a thin actin-rich projection maintains contact with the basal lamina (Fig. 2, B and C). It remains poorly understood how this structure behaves during cleavage and whether this basal process plays any role in the correct reintegration of the postmitotic daughter cells into the monolayer. Although actomyosin may be the primary driver of apical rounding in many cases, evidence also supports a role for microtubule-based mechanisms in the positioning of premitotic nuclei. In chicken neural tube and mouse cerebral cortex, nuclei migrate apically on microtubules before actomyosin-dependent rounding (Spear and Erickson, 2012a). Centrosomes provide directionality to the microtubules on which the nucleus migrates and organize the spindle once the mitotic chromatin reaches the apical domain (Peyre et al., 2011; Spear and Erickson, 2012a; Nakajima et al., 2013). Collectively, current evidence suggests that both actomyosin- and microtubule-dependent forces conspire to effect mitotic nuclear translocation in a highly context- and species-specific manner. One possibility is that the varying physical dimensions of epithelial cells require varying mechanisms for apical nuclear translocation. For example, highly elongated radial glial cells require active transport of the nucleus on microtubules before mitotic rounding, whereas cortical actomyosin contractility may be sufficient in less elongated cells (Spear and Erickson, 2012b). A major outstanding problem is how cortical contractility triggers cell rounding that is polarized along the apicobasal axis of the cell. Whereas centrosomes function as an apical landmark for nuclei moving on microtubules, it remains unclear what provides the directionality for the basal-to-apical actomyosin contraction. One hypothesis is that certain proteins can restrict the localization of nonmuscle myosin II at the basal domain of epithelial cells. The microcephaly protein Asp interacts with myosin II and regulates its polarized localization along the apicobasal axis in the fly optic lobe neuroepithelium. In asp mutant flies, myosin II is enriched apically instead of basally. Many dividing nuclei fail to reach the apical domain and are thus broadly distributed along the apicobasal axis of the epithelium, leading to a disorganized tissue (Rujano et al., 2013). Interestingly, Asp also interacts with microtubules, associates with spindle poles, and is essential for positioning the spindle in fly and vertebrate epithelia (Saunders et al., 1997; do Carmo Avides and Glover, 1999; Wakefield et al., 2001; Fish et al., 2006). Elucidating the function of proteins such as Asp at the interface of microtubules and actomyosin will be essential to our understanding of how the cytoskeleton drives apical mitotic rounding.

Localized molecular landmarks direct planar spindle orientation.

In most animal cells, the mitotic spindle is anchored to the cell cortex by astral microtubules (Fig. 2, C–E; Théry and Bornens, 2006). Translocation of the dynein–dynactin motor toward the astral microtubule minus ends provides a pulling force on centrosomes and is essential for spindle orientation and pole separation during cell division (Dujardin and Vallee, 2002; Kotak et al., 2012). Molecular cues embedded in the cortex can thus determine spindle orientation by anchoring the dynein–dynactin complex in restricted domains. In cultured MDCK and chick neuroepithelia cells, the Gαi–LGN–nuclear mitotic apparatus (NuMA) complex serves this function (Busson et al., 1998; Hao et al., 2010; Zheng et al., 2010; Peyre et al., 2011). Knockdown or mislocalization of these factors leads to spindle orientation defects that ultimately lead to removal of cell progenitors from the monolayer (Peyre et al., 2011). LGN (Pins in Drosophila) localizes to the lateral cell cortex by binding to the membrane-bound Gαi and enforces spindle orientation by recruiting NuMA (Mud in Drosophila), which binds directly to the dynein–dynactin motor. In certain epithelia, including MDCK cells and Drosophila wing discs, LGN is excluded from the apical domain by atypical PKC (aPKC) phosphorylation, thus restricting it at the lateral cell cortex (Konno et al., 2008; Hao et al., 2010; Zheng et al., 2010; Guilgur et al., 2012). In chick neuroepithelia, however, LGN is restricted at the lateral cortex independently of aPKC, suggesting that other cues control its localization (Morin et al., 2007; Peyre et al., 2011). In the mouse embryonic neocortex, the actin–membrane linkers ERM (ezrin/radixin/moesin) promote the association of LGN with NuMA (Machicoane et al., 2014), indicating that organized cortical actin is critical for correct LGN localization.Cell–cell junctions have been implicated in planar cell division in mammalian epithelia, suggesting a possible direct link between the polarity apparatus and the spindle machinery (Reinsch and Karsenti, 1994; den Elzen et al., 2009). Interfering with E-cadherin function or reducing E-cadherin levels abolishes junctional localization of APC (adenomatous polyposis coli), a microtubule-interacting protein that is required for planar spindle orientation and chromosome alignment (Green et al., 2005; den Elzen et al., 2009). However, spindle orientation may not directly depend on E-cadherin or adherens junctions (AJs) in all cases. In Drosophila follicle cells and imaginal discs as well as Xenopus laevis embryonic epithelia, mitotic spindles exhibit planar orientation but do not align with the AJs (Woolner and Papalopulu, 2012; Bergstralh et al., 2013; Nakajima et al., 2013). Moreover, disruption of AJs in Drosophila follicle cells does not affect spindle position (Bergstralh et al., 2013).In Drosophila wing discs, the spindle poles localize in close proximity to septate junctions, which are positioned immediately basal to AJs (Nakajima et al., 2013). Septate junctions are enriched with many proteins, including the neoplastic tumor suppressors SCRIB (Scribbled) and DLG1 (Discs large 1; Bilder and Perrimon, 2000; Bilder et al., 2000). In asymmetrically dividing cells, such as Drosophila sensory organ precursors and neuroblasts, DLG1 interacts with LGN at the cortex and is required for proper spindle orientation (Bellaïche et al., 2001; Siegrist and Doe, 2005; Johnston et al., 2009). Recent findings indicate that DLG1 is also essential for planar spindle orientation in the symmetric division of epithelial cells. In wing discs, knockdown of scrib or dlg1 leads to randomized spindle orientations. scrib knockdown wing discs exhibit diffuse DLG1 localization but no obvious apicobasal polarity defect, suggesting that epithelial disorganization could be a consequence of aberrant spindle orientation (Nakajima et al., 2013). However, it is not clear whether the septate junctions themselves are important. In Drosophila follicle epithelial cells where septate junctions do not form until relatively late in development (Oshima and Fehon, 2011), DLG1 is localized at the lateral cell cortex and is essential for planar spindle orientation (Bergstralh et al., 2013). Interestingly, dlg1 mutant follicle cells display misoriented divisions yet normal epithelial polarity and tissue organization. In this case, planar spindle orientation appears to be independent of junctions per se but still depends on a DLG1–LGN–NuMA complex, similar to asymmetrically dividing cells (Bergstralh et al., 2013).

Global stress and local cell geometry influence mitotic spindle orientation within the plane of the epithelium.

During planar divisions, the mitotic spindle aligns to the plane of the epithelium (xz; Fig. 2 C) and also within the plane of the cell layer (xy; Fig. 2 D). Studies in gastrulating zebrafish embryos revealed a role for the Wnt–Frizzled–planar cell polarity signaling pathway in orienting cell divisions (Concha and Adams, 1998; Gong et al., 2004). Similarly, the atypical cadherins Fat and Dachsous are involved in orienting cell divisions in the Drosophila wing and in developing mouse kidneys (Baena-López et al., 2005; Saburi et al., 2008). Although both of these pathways have been reviewed elsewhere (Morin and Bellaïche, 2011), recent studies also point to at least two other mechanisms that may independently influence spindle orientation within the plane of the monolayer: (1) global tissue stress and (2) local epithelial cell geometry.Epithelial cell shape and spindle orientation are modulated by global stress that accumulates during tissue growth. In Drosophila wing discs, cells in the center of the wing blade primordium proliferate at a faster rate than in the periphery. Consequently, cells in the periphery are mechanically stretched, and cells in the center are compressed. As a result of stretching, peripheral cells localize myosin II at their cortex and align their mitotic spindle with the stretch axis (LeGoff et al., 2013; Mao et al., 2013). Similarly, epithelial cells of the enveloping cell layer in gastrulating zebrafish embryos elongate and orient their spindle along the direction of tension generated by spreading during epiboly (Campinho et al., 2013). It is unclear whether myosin II directly conveys cell tension to the mitotic apparatus, and it will be necessary to dissect whether cell elongation alone or additional mechanosensing pathways signal cell tension to the mitotic spindle. Keratinocytes from the mammalian epidermis reorient their mitotic spindle in response to mechanical stretch in a NuMA-dependent manner. The mitotic spindle aligns with the cortical NuMA-localized crescent upon stretch and fails to orient when NuMA levels are reduced (Seldin et al., 2013). In summary, global tension generated by growth and cell spreading impact division orientation, suggesting that shape changes in proximity to dividing cells may also lead to a similar effect.Although variations certainly exist, the apical surfaces of proliferating epithelia tend to feature a consistent percentage of hexagonal, pentagonal, heptagonal, and octagonal cell shapes (Gibson et al., 2006; Farhadifar et al., 2007; Aegerter-Wilmsen et al., 2010). In Drosophila imaginal discs, these local patterns of cell packing may systematically influence spindle orientation, as mitotic cells are biased toward cleaving their common interfaces with subhexagonal neighbors (less than six sides) and avoid cleaving their interfaces with superhexagonal neighbors (more than six sides; Gibson et al., 2011). Although the mechanisms underlying the effect of local cell geometry remain elusive, cell packing influences mitotic cell shape and the distribution of adhesive cues, both of which could, in turn, bias spindle orientation. Indeed, dividing cells maintain contacts with their neighbors, which can influence the cell cortex and direct spindle orientation (Goldstein, 1995; Wang et al., 1997). The distribution of adhesions between epithelial cells may also alter the position or action of cortical force generators that interact with spindle microtubules in the mitotic cell. In support of this idea, when single cells are placed on micropatterned substrates, they orient their spindle relative to the geometry of their adhesion pattern and not their cell shape (Théry et al., 2005, 2007). Alternatively, neighbors of different polygonal shapes could stretch the mitotic cell, thus imposing a bias on its long axis. Indeed, sea urchin embryos orient their spindles to divide their longest axis (Hertwig, 1884) and can even sense complex cell geometries to orient their spindles accordingly (Minc et al., 2011). Still, precisely how the interphase morphology of epithelial cells might impinge on mitotic spindle orientation remains an open question.

Genesis of nascent junctions during epithelial cell division

After spindle orientation, the essential processes of cytokinesis and abscission are driven by the assembly and contraction of an actomyosin ring positioned in the cleavage plane (Fededa and Gerlich, 2012). In epithelia, ring contraction accompanied by membrane invagination ultimately gives rise to a new junctional interface between nascent daughter cells. Precisely how this new interface forms remains poorly understood. Recent studies in Drosophila epithelia reveal that, during cytokinesis, (a) E-cadherin levels are reduced at the interface between the cleavage furrow of dividing cells and their neighbors (Fig. 3), and (b) neighbor tension and midbody position guide establishment of new AJs in context with local epithelial geometry (Fig. 4).Open in a separate windowFigure 3.Cytokinetic membrane dynamics in epithelial cells. (A) Cytokinesis of a dividing epithelial cell (yellow) presents several unique structural considerations not addressed by the analysis of single cells. Recent studies (Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013, 2014) report a local reduction of E-cadherin levels in proximity to the contractile ring in the dividing cell and its neighbor (red). How cytokinesis is resolved from there may vary in a context-dependent manner. (B) In Drosophila embryos, ring contraction leads to E-cadherin disengagement, and a gap forms between the mitotic cell and its neighbor (Guillot and Lecuit, 2013a). (C) In the Drosophila pupal notum, the contractile ring pulls the neighbor cell plasma membrane into the cleavage furrow, perhaps enabled by uncoupling of the membrane and the cortex in the neighbor (Herszterg et al., 2013, 2014).Open in a separate windowFigure 4.New AJ formation in dividing epithelial cells. Apical cross section (xy, top row) and longitudinal (xz, bottom row) view of a dividing epithelial cell (red). (A) Opposing forces (black vertical arrows) develop between the contractile ring in the dividing cell and the two neighboring cells (orange) in proximity to the cleavage furrow. E-cadherin clusters are reduced at the furrow/neighbor interface. (B) Myosin II and tension build up in the neighboring cell, causing the nascent daughter cells to juxtapose their plasma membranes at the presumptive site of junction assembly (black horizontal arrows). (C) Arp2/3 and Rac1 drive actin polymerization at the daughter cell interface around the midbody, stabilizing the nascent junction as the neighboring cell membrane withdraws. (D) The new junction is complete and of suitable length in context with the local epithelial geometry.

Mitotic cells remodel their adhesion junctions during cytokinesis.

Two kinds of forces are at work during cytokinesis: an active force in the dividing cell caused by ring contraction and a reactive force in contacting neighbors caused by their resistance to pulling to maintain their shape (Fig. 3 and Fig. 4 A; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013). Recent results indicate that these opposing forces can lead to a transient and partial reduction of cell adhesion after mitotic exit. In Drosophila epithelia, E-cadherin levels are reduced at the interface between the cleavage furrow of the dividing cell and its neighbors (Fig. 3, B and C; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013; Morais-de-Sá and Sunkel, 2013). Specifically in embryonic epithelia, the local reduction of E-cadherin facilitates membrane separation, and a gap appears between the dividing cell and its neighbors (Fig. 3 B; Guillot and Lecuit, 2013a). In the dorsal thorax, in contrast, the neighbor cell plasma membrane detaches from the cortex and is drawn into the cleavage furrow (Fig. 3 C; Herszterg et al., 2013, 2014). What triggers E-cadherin modulation in cells after mitotic exit? The loss of overall cell polarity is one possible mechanism. During mitosis in Drosophila, follicular epithelial cells lose cortical enrichment of some apical polarity proteins (aPKC, Crumbs, and Bazooka/Par3; Bergstralh et al., 2013; Morais-de-Sá and Sunkel, 2013), and embryonic cells lose localization of lethal giant larvae, a basolateral cortical protein (Huang et al., 2009). Contrasting with these observations, however, MDCK cells and Drosophila embryonic and dorsal thorax epithelial cells appear to maintain apicobasal polarity as they divide (Reinsch and Karsenti, 1994; Founounou et al., 2013; Guillot and Lecuit, 2013a; Herszterg et al., 2013). Furthermore, E-cadherin reduction is limited to the furrow/neighbor interface and is not observed in other areas of cell contact. Therefore, an alternative mechanism that explains local E-cadherin modulation is mechanical tension that arises precisely at the area between the contractile ring and the neighboring cell membrane (Founounou et al., 2013; Guillot and Lecuit, 2013a).Does E-cadherin modulation serve a functional role in mitotic cells? In Drosophila embryonic and dorsal thorax epithelia, E-cadherin decrease leads to a local adhesion disengagement proposed to facilitate the formation of new AJs between daughter cells (Founounou et al., 2013; Guillot and Lecuit, 2013a). It has been previously reported that cells maintain their AJs throughout division. For example, intercellular junctions are maintained in dividing cells of human colonic mucosal crypt cells and basal keratinocytes (Baker and Garrod, 1993). Similarly, mitotic MDCK cultured cells maintain tight junctions apically and E-cadherin basolaterally (Reinsch and Karsenti, 1994). The E-cadherin loss in certain Drosophila epithelia may be either a tissue-specific phenomenon or a highly dynamic process only observable with the temporal resolution of live-cell imaging. Moreover, dividing cells in the Drosophila dorsal thorax show decreased levels of E-cadherin yet maintain their cohesiveness (Herszterg et al., 2013). Interestingly, E-cadherin is internalized in mitotic MDCK cells (Bauer et al., 1998). It will therefore be important to investigate whether loss of E-cadherin leads to adhesion disengagement in other epithelial tissues and whether tension alone or in combination with biochemical pathways is responsible for E-cadherin modulation.

Epithelial neighbors exert tension on daughter cell membranes to facilitate new AJ formation.

How new junctional contacts form during mitosis is a poorly understood problem at the heart of epithelial cell biology. In Drosophila, new membrane interfaces between nascent daughter cells initially show only a weak level of E-cadherin clusters (Guillot and Lecuit, 2013a; Herszterg et al., 2013). Subsequently, the daughter cells assemble their AJs de novo. How is the length of these new junctions determined with respect to cell geometry? Recent evidence indicates that AJ length is a function of local cell packing within the epithelium. In dividing cells of the Drosophila dorsal thorax, the contractile ring triggers tension and accumulation of myosin II in neighbors at the furrow/neighbor interface (Fig. 4 B; Founounou et al., 2013; Herszterg et al., 2013). Myosin II in the neighboring cells in turn contracts and creates tension at the furrowing membrane of the nascent daughter cells, keeping them tightly pressed against each other (Fig. 4, B and C). This local membrane juxtaposition facilitates AJ formation. To allow expansion of the daughter cell interface and maintain AJ length, branched actin polymerization via Rac1 and Arp2/3 is oriented to the midbody, which serves as a positional landmark for new AJs (Fig. 4 C; Herszterg et al., 2013). The midbody is a narrow intercellular bridge that remains after the contracted cytokinetic ring has driven membrane invagination, and it recruits the abscission factors that will eventually separate the daughter cells (Fededa and Gerlich, 2012). Interestingly, the midbody is positioned apically as a result of the presence of AJs. In Drosophila follicular epithelia, the midbody also provides cues for the formation of the apical daughter cell interface, suggesting that it plays a role in both AJ and epithelial cell polarity establishment and maintenance in dividing epithelial cells (Morais-de-Sá and Sunkel, 2013). Thus, examples from Drosophila epithelia show that cohesion between dividing cells and their neighbors together with the apically positioned midbody provides a spatial template and polarized positional cue for de novo AJ assembly (Herszterg et al., 2013; Morais-de-Sá and Sunkel, 2013). Further work on other epithelial tissues may provide alternative mechanisms of junction biogenesis.

Growth and order in the epithelium: Thinking outside the cell

During development, epithelial monolayers have the remarkable capacity to maintain specialized morphologies and barrier functions during rapid cell proliferation. Mitotic cells remain adherent to their neighbors throughout cell division. Cell cohesion enables local geometry and global tissue tension to instruct mitotic cells where to position their cleavage plane and how to assemble their junctions. However, local tension may also lead to a transient disengagement of dividing cells from their neighbors after mitotic exit. How is global and local tension conveyed to protein complexes in mitotic cells so that different outcomes take place? Moreover, it is unclear whether and how tissue tension instructs synchronously dividing epithelial cells how to divide and reestablish their junctions after division. Clearly, this is a fundamental problem for the maintenance of epithelial order and may be linked to the origin of epithelial cancers, in which cells undergo rapid proliferation but fail to remain integrated into the monolayer.The selected studies discussed here hint at the remarkable level of coordination that occurs during epithelial cell division, recasting mitosis as a truly multicellular process. Looking ahead, understanding the interface between cells, proteins, and mechanical forces that each operate on different scales will require creative multidisciplinary approaches in diverse organismal systems. Indeed, epithelial organization is widespread in nature and is encountered among even the most basal animals, including sponges and cnidarians as well as the fruiting body of the nonmetazoan social amoeba Dictyostelium discoideum (Wood, 1959; Ereskovsky et al., 2009; Houliston et al., 2010; Dickinson et al., 2011; Meyer et al., 2011). Combined, future interdisciplinary studies and a fresh look at diverse animal models should yield new insight into epithelial cell division for many years to come.