The tubulin code: Molecular components,readout mechanisms,and functions

Microtubules are cytoskeletal filaments that are dynamically assembled from α/β-tubulin heterodimers. The primary sequence and structure of the tubulin proteins and, consequently, the properties and architecture of microtubules are highly conserved in eukaryotes. Despite this conservation, tubulin is subject to heterogeneity that is generated in two ways: by the expression of different tubulin isotypes and by posttranslational modifications (PTMs). Identifying the mechanisms that generate and control tubulin heterogeneity and how this heterogeneity affects microtubule function are long-standing goals in the field. Recent work on tubulin PTMs has shed light on how these modifications could contribute to a “tubulin code” that coordinates the complex functions of microtubules in cells.

Introduction

Microtubules are key elements of the eukaryotic cytoskeleton that dynamically assemble from heterodimers of α- and β-tubulin. The structure of microtubules, as well as the protein sequences of α- and β-tubulin, is highly conserved in evolution, and consequently, microtubules look alike in almost all species. Despite the high level of conservation, microtubules adapt to a large variety of cellular functions. This adaptation can be mediated by a large panel of microtubule-associated proteins (MAPs), including molecular motors, as well as by mechanisms that directly modify the microtubules, thus either changing their biophysical properties or attracting subsets of MAPs that convey specific functions to the modified microtubules. Two different mechanism can generate microtubule diversity: the expression of different α- and β-tubulin genes, referred to as tubulin isotypes, and the generation of posttranslational modifications (PTMs) on α- and β-tubulin (Figs. 1 and ​and2).2). Although known for several decades, deciphering how tubulin heterogeneity controls microtubule functions is still largely unchartered. This review summarizes the current advances in the field and discusses new concepts arising.Open in a separate windowFigure 1.Tubulin heterogeneity generated by PTMs. (A) Schematic representation of the distribution of different PTMs of tubulin on the α/β-tubulin dimer with respect to their position in the microtubule lattice. Acetylation (Ac), phosphorylation (P), and polyamination (Am) are found within the tubulin bodies that assemble into the microtubule lattice, whereas polyglutamylation, polyglycylation, detyrosination, and C-terminal deglutamylation take place within the C-terminal tubulin tails that project away from the lattice surface. The tubulin dimer represents TubA1A and TubB2B (Fig. 2), and modification sites for polyglutamylation and polyglycylation have been randomly chosen. (B) Chemical structure of the branched peptide formed by polyglutamylation and polyglycylation, using the γ-carboxyl groups of the modified glutamate residues as acceptor sites for the isopeptide bonds. Note that in the case of polyglutamylation, the elongation of the side chains generates classical peptide bonds (Redeker et al., 1991).Open in a separate windowFigure 2.Heterogeneity of C-terminal tails of tubulin isotypes and their PTMs. The amino acid sequences of all tubulin genes found in the human genome are indicated, starting at the last amino acid of the folded tubulin bodies. Amino acids are represented in single-letter codes and color coded according to their biochemical properties. Known sites for polyglutamylation are indicated (Eddé et al., 1990; Alexander et al., 1991; Rüdiger et al., 1992). Potential modification sites (all glutamate residues) are indicated. Known C-terminal truncation reactions of α/β-tubulin (tub) are indicated. The C-terminal tails of the yeast Saccharomyces cerevisiae are shown to illustrate the phylogenetic diversity of these domains.

Tubulin isotypes

The cloning of the first tubulin genes in the late 1970’s (Cleveland et al., 1978) revealed the existence of multiple genes coding for α- or β-tubulin (Ludueña and Banerjee, 2008) that generate subtle differences in their amino acid sequences, particularly in the C-terminal tails (Fig. 2). It was assumed that tubulin isotypes, as they were named, assemble into discrete microtubule species that carry out unique functions. This conclusion was reinforced by the observation that some isotypes are specifically expressed in specialized cells and tissues and that isotype expression changes during development (Lewis et al., 1985; Denoulet et al., 1986). These high expectations were mitigated by a subsequent study showing that all tubulin isotypes freely copolymerize into heterogeneous microtubules (Lewis et al., 1987). To date, only highly specialized microtubules, such as ciliary axonemes (Renthal et al., 1993; Raff et al., 2008), neuronal microtubules (Denoulet et al., 1986; Joshi and Cleveland, 1989), and microtubules of the marginal band of platelets (Wang et al., 1986; Schwer et al., 2001) are known to depend on some specific (β) tubulin isotypes, whereas the function of most other microtubules appears to be independent of their isotype composition.More recently, a large number of mutations in single tubulin isotypes have been linked to deleterious neurodevelopmental disorders (Keays et al., 2007; Fallet-Bianco et al., 2008; Tischfield et al., 2010; Cederquist et al., 2012; Niwa et al., 2013). Mutations of a single tubulin isotype could lead to an imbalance in the levels of tubulins as a result of a lack of incorporation of mutant isoforms into the microtubule lattice or to incorporation that perturbs the architecture or dynamics of the microtubules. The analysis of tubulin disease mutations is starting to reveal how subtle alterations of the microtubule cytoskeleton can lead to functional aberrations in cells and organisms and might provide novel insights into the roles of tubulin isotypes that have so far been considered redundant.

Tubulin PTMs

Tubulin is subject to a large range of PTMs (Fig. 1), from well-known ones, such as acetylation or phosphorylation, to others that have so far mostly been found on tubulin. Detyrosination/tyrosination, polyglutamylation, and polyglycylation, for instance, might have evolved to specifically regulate tubulin and microtubule functions, in particular in cilia and flagella, as their evolution is closely linked to these organelles. The strong link between those modifications and tubulin evolution has led to the perception that they are tubulin PTMs; however, apart from detyrosination/tyrosination, most of them have other substrates (Regnard et al., 2000; Xie et al., 2007; van Dijk et al., 2008; Rogowski et al., 2009).

Tubulin acetylation.

Tubulin acetylation was discovered on lysine 40 (K40; Fig. 1 A) of flagellar α-tubulin in Chlamydomonas reinhardtii (L’Hernault and Rosenbaum, 1985) and is generally enriched on stable microtubules in cells. Considering that K40 acetylation per se has no effect on the ultrastructure of microtubules (Howes et al., 2014), it is rather unlikely that it directly stabilizes microtubules. As a result of its localization at the inner face of microtubules (Soppina et al., 2012), K40 acetylation might rather affect the binding of microtubule inner proteins, a poorly characterized family of proteins (Nicastro et al., 2011; Linck et al., 2014). Functional experiments in cells have further suggested that K40 acetylation regulates intracellular transport by regulating the traffic of kinesin motors (Reed et al., 2006; Dompierre et al., 2007). These observations could so far not be confirmed by biophysical measurements in vitro (Walter et al., 2012; Kaul et al., 2014), suggesting that in cells, K40 acetylation might affect intracellular traffic by indirect mechanisms.Enzymes involved in K40 acetylation are HDAC6 (histone deacetylase family member 6; Hubbert et al., 2002) and Sirt2 (sirtuin type 2; North et al., 2003). Initial functional studies used overexpression, depletion, or chemical inhibition of these enzymes. These studies should be discussed with care, as both HDAC6 and Sirt2 deacetylate other substrates and have deacetylase-independent functions and chemical inhibition of HDAC6 is not entirely selective for this enzyme (Valenzuela-Fernández et al., 2008). In contrast, acetyl transferase α-Tat1 (or Mec-17; Akella et al., 2010; Shida et al., 2010) specifically acetylates α-tubulin K40 (Fig. 3), thus providing a more specific tool to investigate the functions of K40 acetylation. Knockout mice of α-Tat1 are completely void of K40-acetylated tubulin; however, they show only slight phenotypic aberrations, for instance, in their sperm flagellum (Kalebic et al., 2013). A more detailed analysis of α-Tat1 knockout mice demonstrated that absence of K40 acetylation leads to reduced contact inhibition in proliferating cells (Aguilar et al., 2014). In migrating cells, α-Tat1 is targeted to microtubules at the leading edge by clathrin-coated pits, resulting in locally restricted acetylation of those microtubules (Montagnac et al., 2013). A recent structural study of α-Tat1 demonstrated that the low catalytic rate of this enzyme, together with its localization inside the microtubules, caused acetylation to accumulate selectively in stable, long-lived microtubules (Szyk et al., 2014), thus explaining the link between this PTM and stable microtubules in cells. However, the direct cellular function of K40 acetylation on microtubules is still unclear.Open in a separate windowFigure 3.Enzymes involved in PTM of tubulin. Schematic representation of known enzymes (mammalian enzymes are shown) involved in the generation and removal of PTMs shown in Fig. 1. Note that some enzymes still remain unknown, and some modifications are irreversible. (*CCP5 preferentially removes branching points [Rogowski et al., 2010]; however, the enzyme can also hydrolyze linear glutamate chains [Berezniuk et al., 2013]).Recent discoveries have brought up the possibility that tubulin could be subject to multiple acetylation events. A whole-acetylome study identified >10 novel sites on α- and β-tubulin (Choudhary et al., 2009); however, none of these sites have been confirmed. Another acetylation event has been described at lysine 252 (K252) of β-tubulin. This modification is catalyzed by the acetyltransferase San (Fig. 3) and might regulate the assembly efficiency of microtubules as a result of its localization at the polymerization interface (Chu et al., 2011).

Tubulin detyrosination.

Most α-tubulin genes in different species encode a C-terminal tyrosine residue (Fig. 2; Valenzuela et al., 1981). This tyrosine can be enzymatically removed (Hallak et al., 1977) and religated (Fig. 3; Arce et al., 1975). Mapping of tyrosinated and detyrosinated microtubules in cells using specific antibodies (Gundersen et al., 1984; Geuens et al., 1986; Cambray-Deakin and Burgoyne, 1987a) revealed that subsets of interphase and mitotic spindle microtubules are detyrosinated (Gundersen and Bulinski, 1986). As detyrosination was mostly found on stable and long-lived microtubules, especially in neurons (Cambray-Deakin and Burgoyne, 1987b; Robson and Burgoyne, 1989; Brown et al., 1993), it was assumed that this modification promotes microtubule stability (Gundersen et al., 1987; Sherwin et al., 1987). Although a direct stabilization of the microtubule lattice was considered unlikely (Khawaja et al., 1988), it was found more recently that detyrosination protects cellular microtubules from the depolymerizing activity of kinesin-13–type motor proteins, such as KIF2 or MCAK, thus increasing their longevity (Peris et al., 2009; Sirajuddin et al., 2014).Besides kinesin-13 motors, plus end–tracking proteins with cytoskeleton-associated protein glycine-rich (CAP-Gly) domains, such as CLIP170 or p150/glued, specifically interact with tyrosinated microtubules (Peris et al., 2006; Bieling et al., 2008) via this domain (Honnappa et al., 2006). In contrast, kinesin-1 moves preferentially on detyrosinated microtubules tracks in cells (Liao and Gundersen, 1998; Kreitzer et al., 1999; Konishi and Setou, 2009). The effect of detyrosination on kinesin-1 motor behavior was recently measured in vitro, and a small but significant increase in the landing rate and processivity of the motor has been found (Kaul et al., 2014). Such subtle changes in the motor behavior could, in conjunction with other factors, such as regulatory MAPs associated with cargo transport complexes (Barlan et al., 2013), lead to a preferential use of detyrosinated microtubules by kinesin-1 in cells.Despite the early biochemical characterization of a detyrosinating activity, the carboxypeptidase catalyzing detyrosination of α-tubulin has yet to be identified (Hallak et al., 1977; Argaraña et al., 1978, 1980). In contrast, the reverse enzyme, tubulin tyrosine ligase (TTL; Fig. 3; Raybin and Flavin, 1975; Deanin and Gordon, 1976; Argaraña et al., 1980), has been purified (Schröder et al., 1985) and cloned (Ersfeld et al., 1993). TTL modifies nonpolymerized tubulin dimers exclusively. This selectivity is determined by the binding interface between the TTL and tubulin dimers (Szyk et al., 2011, 2013; Prota et al., 2013). In contrast, the so far unidentified detyrosinase acts preferentially on polymerized microtubules (Kumar and Flavin, 1981; Arce and Barra, 1983), thus modifying a select population of microtubules within cells (Gundersen et al., 1987).In most organisms, only one unique gene for TTL exists. Consequently, TTL knockout mice show a huge accumulation of detyrosinated and particularly Δ2-tubulin (see next section). TTL knockout mice die before birth (Erck et al., 2005) with major developmental defects in the nervous system that might be related to aberrant neuronal differentiation (Marcos et al., 2009). TTL is strictly tubulin specific (Prota et al., 2013), indicating that all observed defects in TTL knockout mice are directly related to the deregulation of the microtubule cytoskeleton.

Δ2-tubulin and further C-terminal modification.

A biochemical study of brain tubulin revealed that ∼35% of α-tubulin cannot be retyrosinated (Paturle et al., 1989) because of the lack of the penultimate C-terminal glutamate residue of the primary protein sequence (Fig. 2; Paturle-Lafanechère et al., 1991). This so-called Δ2-tubulin (for two C-terminal amino acids missing) cannot undergo retyrosination as a result of structural constraints within TTL (Prota et al., 2013) and thus is considered an irreversible PTM.Δ2-tubulin accumulates in long-lived microtubules of differentiated neurons, axonemes of cilia and flagella, and also in cellular microtubules that have been artificially stabilized, for instance, with taxol (Paturle-Lafanechère et al., 1994). The generation of Δ2-tubulin requires previous detyrosination of α-tubulin; thus, the levels of this PTM are indirectly regulated by the detyrosination/retyrosination cycle. This mechanistic link is particularly apparent in the TTL knockout mice, which show massive accumulation of Δ2-tubulin in all tested tissues (Erck et al., 2005). Loss of TTL and the subsequent increase of Δ2-tubulin levels were also linked to tumor growth and might contribute to the aggressiveness of the tumors by an as-yet-unknown mechanism (Lafanechère et al., 1998; Mialhe et al., 2001). To date, no specific biochemical role of Δ2-tubulin has been determined; thus, one possibility is that the modification simply locks tubulin in the detyrosinated state.The enzymes responsible for Δ2-tubulin generation are members of a family of cytosolic carboxypeptidases (CCPs; Fig. 3; Kalinina et al., 2007; Rodriguez de la Vega et al., 2007), and most of them also remove polyglutamylation from tubulin (see next section; Rogowski et al., 2010). These enzymes are also able to generate Δ3-tubulin (Fig. 1 A; Berezniuk et al., 2012), indicating that further degradation of the tubulin C-terminal tails are possible; however, the functional significance of this event is unknown.

Polyglutamylation.

Polyglutamylation is a PTM that occurs when secondary glutamate side chains are formed on γ-carboxyl groups of glutamate residues in a protein (Fig. 1, A and B). The modification was first discovered on α- and β-tubulin from the brain (Eddé et al., 1990; Alexander et al., 1991; Rüdiger et al., 1992; Mary et al., 1994) as well as on axonemal tubulin from different species (Mary et al., 1996, 1997); however, it is not restricted to tubulin (Regnard et al., 2000; van Dijk et al., 2008). Using a glutamylation-specific antibody, GT335 (Wolff et al., 1992), it was observed that tubulin glutamylation increases during neuronal differentiation (Audebert et al., 1993, 1994) and that axonemes of cilia and flagella (Fouquet et al., 1994), as well as centrioles of mammalian centrosomes (Bobinnec et al., 1998), are extensively glutamylated.Enzymes catalyzing polyglutamylation belong to the TTL-like (TTLL) family (Regnard et al., 2003; Janke et al., 2005). In mammals, nine glutamylases exist, each of them showing intrinsic preferences for modifying either α- or β-tubulin as well as for initiating or elongating glutamate chains (Fig. 3; van Dijk et al., 2007). Two of the six well-characterized TTLL glutamylases also modify nontubulin substrates (van Dijk et al., 2008).Knockout or depletion of glutamylating enzymes in different model organisms revealed an evolutionarily conserved role of glutamylation in cilia and flagella. In motile cilia, glutamylation regulates beating behavior (Janke et al., 2005; Pathak et al., 2007; Ikegami et al., 2010) via the regulation of flagellar dynein motors (Kubo et al., 2010; Suryavanshi et al., 2010). Despite the expression of multiple glutamylases in ciliated cells and tissues, depletion or knockout of single enzymes often lead to ciliary defects, particularly in motile cilia (Ikegami et al., 2010; Vogel et al., 2010; Bosch Grau et al., 2013; Lee et al., 2013), suggesting essential and nonredundant regulatory functions of these enzymes in cilia.Despite the enrichment of polyglutamylation in neuronal microtubules (Audebert et al., 1993, 1994), knockout of TTLL1, the major polyglutamylase in brain (Janke et al., 2005), did not show obvious neuronal defects in mice (Ikegami et al., 2010; Vogel et al., 2010). This suggests a tolerance of neuronal microtubules to variations in polyglutamylation.Deglutamylases, the enzymes that reverse polyglutamylation, were identified within a novel family of CCPs (Kimura et al., 2010; Rogowski et al., 2010). So far, three out of six mammalian CCPs have been shown to cleave C-terminal glutamate residues, thus catalyzing both the reversal of polyglutamylation and the removal of gene-encoded glutamates from the C termini of proteins (Fig. 3). The hydrolysis of gene-encoded glutamate residues is not restricted to tubulin, in which it generates Δ2- and Δ3-tubulin, but has also been reported for other proteins such as myosin light chain kinase (Rusconi et al., 1997; Rogowski et al., 2010). One enzyme of the CCP family, CCP5, preferentially removes branching points generated by glutamylation, thus allowing the complete reversal of the polyglutamylation modification (Kimura et al., 2010; Rogowski et al., 2010). However, CCP5 can also hydrolyze C-terminal glutamate residues from linear peptide chains similar to other members of the CCP family (Berezniuk et al., 2013).CCP1 is mutated in a well-established mouse model for neurodegeneration, the pcd (Purkinje cell degeneration) mouse (Mullen et al., 1976; Greer and Shepherd, 1982; Fernandez-Gonzalez et al., 2002). The absence of a key deglutamylase leads to strong hyperglutamylation in brain regions that undergo degeneration, such as the cerebellum and the olfactory bulb (Rogowski et al., 2010). When glutamylation levels were rebalanced by depletion or knockout of the major brain polyglutamylase TTLL1 (Rogowski et al., 2010; Berezniuk et al., 2012), Purkinje cells survived. Although the molecular mechanisms of hyperglutamylation-induced degeneration remain to be elucidated, perturbation of neuronal transport, as well as changes in the dynamics and stability of microtubules, is expected to be induced by hyperglutamylation. Increased polyglutamylation levels have been shown to affect kinesin-1–mediated transport in cultured neurons (Maas et al., 2009), and the turnover of microtubules can also be regulated by polyglutamylation via the activation of microtubule-severing enzymes such as spastin (Lacroix et al., 2010).Subtle differences in polyglutamylation can be seen on diverse microtubules in different cell types. The functions of these modifications remain to be studied; however, its wide distribution strengthens the idea that it could be involved in fine-tuning a range of microtubule functions.

Polyglycylation.

Tubulin polyglycylation or glycylation, like polyglutamylation, generates side chains of glycine residues within the C-terminal tails of α- and β-tubulin (Fig. 1, A and B). The modification sites of glycylation are considered to be principally the same as for glutamylation, and indeed, both PTMs have been shown to be interdependent in cells (Rogowski et al., 2009; Wloga et al., 2009). Initially discovered on Paramecium tetraurelia tubulin (Redeker et al., 1994), glycylation has been extensively studied using two antibodies, TAP952 and AXO49 (Bressac et al., 1995; Levilliers et al., 1995; Bré et al., 1996). In contrast to polyglutamylation, glycylation is restricted to cilia and flagella in most organisms analyzed so far.Glycylating enzymes are also members of the TTLL family, and homologues of these enzymes have so far been found in all organisms with proven glycylation of ciliary axonemes (Rogowski et al., 2009; Wloga et al., 2009). In mammals, initiating (TTLL3 and TTLL8) and elongating (TTLL10) glycylases work together to generate polyglycylation (Fig. 3). In contrast, the two TTLL3 orthologues from Drosophila melanogaster can both initiate and elongate glycine side chains (Rogowski et al., 2009).In mice, motile ependymal cilia in brain ventricles acquire monoglycylation upon maturation, whereas polyglycylation is observed only after several weeks (Bosch Grau et al., 2013). Sperm flagella, in contrast, acquire long glycine chains much faster, suggesting that the extent of polyglycylation could correlate with the length of the axonemes (Rogowski et al., 2009). Depletion of glycylases in mice (ependymal cilia; Bosch Grau et al., 2013), zebrafish (Wloga et al., 2009; Pathak et al., 2011), Tetrahymena thermophila (Wloga et al., 2009), and D. melanogaster (Rogowski et al., 2009) consistently led to ciliary disassembly or severe ciliary defects. How glycylation regulates microtubule functions remains unknown; however, the observation that glycylation-depleted axonemes disassemble after initial assembly (Rogowski et al., 2009; Bosch Grau et al., 2013) suggests a role of this PTM in stabilizing axonemal microtubules. Strikingly, human TTLL10 is enzymatically inactive; thus, humans have lost the ability to elongate glycine side chains (Rogowski et al., 2009). This suggests that the elongation of the glycine side chains is not an essential aspect of the function of this otherwise critical tubulin PTM.

Other tubulin PTMs.

Several other PTMs have been found on tubulin. Early studies identified tubulin phosphorylation (Eipper, 1974; Gard and Kirschner, 1985; Díaz-Nido et al., 1990); however, no specific functions were found. The perhaps best-studied phosphorylation event on tubulin takes place at serine S172 of β-tubulin (Fig. 1 A), is catalyzed by the Cdk1 (Fig. 3), and might regulate microtubule dynamics during cell division (Fourest-Lieuvin et al., 2006; Caudron et al., 2010). Tubulin can be also modified by the spleen tyrosine kinase Syk (Fig. 3; Peters et al., 1996), which might play a role in immune cells (Faruki et al., 2000; Sulimenko et al., 2006) and cell division (Zyss et al., 2005; Sulimenko et al., 2006).Polyamination has recently been discovered on brain tubulin (Song et al., 2013), after having been overlooked for many years as a result of the low solubility of polyaminated tubulin. Among several glutamine residues of α- and β-tubulin that can be polyaminated, Q15 of β-tubulin is considered the primary modification site (Fig. 1 A). Polyamination is catalyzed by transglutaminases (Fig. 3), which modify free tubulin as well as microtubules in an irreversible manner, and most likely contribute to the stabilization of microtubules (Song et al., 2013).Tubulin was also reported to be palmitoylated (Caron, 1997; Ozols and Caron, 1997; Caron et al., 2001), ubiquitinated (Ren et al., 2003; Huang et al., 2009; Xu et al., 2010), glycosylated (Walgren et al., 2003; Ji et al., 2011), arginylated (Wong et al., 2007), methylated (Xiao et al., 2010), and sumoylated (Rosas-Acosta et al., 2005). These PTMs have mostly been reported without follow-up studies, and some of them are only found in specific cell types or organisms and/or under specific metabolic conditions. Further studies will be necessary to gain insights into their potential roles for the regulation of the microtubule cytoskeleton.

Current advances and future perspectives

The molecular heterogeneity of microtubules, generated by the expression of different tubulin isotypes and by the PTM of tubulin has fascinated the scientific community for ∼40 years. Although many important advances have been made in the past decade, the dissection of the molecular mechanisms and a comprehensive understanding of the biological functions of tubulin isotypes and PTMs will be a challenging field of research in the near future.

Direct measurements of the impact of tubulin heterogeneity.

The most direct and reliable type of experiments to determine the impact of tubulin heterogeneity on microtubule behavior are in vitro measurements with purified proteins. However, most biophysical work on microtubules has been performed with tubulin purified from bovine, ovine, or porcine brains, which can be obtained in large quantities and with a high degree of purity and activity (Vallee, 1986; Castoldi and Popov, 2003). Brain tubulin is a mixture of different tubulin isotypes and is heavily posttranslationally modified and thus inept for investigating the functions of tubulin heterogeneity (Denoulet et al., 1986; Cambray-Deakin and Burgoyne, 1987b; Paturle et al., 1989; Eddé et al., 1990). Thus, pure, recombinant tubulin will be essential to dissect the roles of different tubulin isoforms and PTMs.Attempts to produce recombinant, functional α- and β-tubulin in bacteria have failed so far (Yaffe et al., 1988), most likely because of the absence of the extensive tubulin-specific folding machinery (Yaffe et al., 1992; Gao et al., 1993; Tian et al., 1996; Vainberg et al., 1998) in prokaryotes. An alternative source of tubulin with less isotype heterogeneity and with almost no PTMs is endogenous tubulin from cell lines such as HeLa, which in the past has been purified using a range of biochemical procedures (Bulinski and Borisy, 1979; Weatherbee et al., 1980; Farrell, 1982; Newton et al., 2002; Fourest-Lieuvin, 2006). Such tubulin can be further modified with tubulin-modifying enzymes, such as polyglutamylases, either by expressing those enzymes in the cells before tubulin purification (Lacroix and Janke, 2011) or in vitro with purified enzymes (Vemu et al., 2014). Despite some technical limitations of these methods, HeLa tubulin modified in cells has been successfully used in an in vitro study on the role of polyglutamylation in microtubule severing (Lacroix et al., 2010).Naturally occurring variants of tubulin isotypes and PTMs can be purified from different organisms, organs, or cell types, but obviously, only some combinations of tubulin isotypes and PTMs can be obtained by this approach. The recent development of an affinity purification method using the microtubule-binding TOG (tumor overexpressed gene) domain of yeast Stu2p has brought a new twist to this approach, as it allows purifying small amounts of tubulin from any cell type or tissue (Widlund et al., 2012).The absence of tubulin heterogeneity in yeast has made budding and fission yeast potential expression systems for recombinant, PTM-free tubulin (Katsuki et al., 2009; Drummond et al., 2011; Johnson et al., 2011). However, the expression of mammalian tubulin in this system has remained impossible. This problem was then partially circumvented by expressing tubulin chimeras that consist of a yeast tubulin body fused to mammalian C-terminal tubulin tails, thus mimicking different tubulin isotypes (Sirajuddin et al., 2014). Moreover, detyrosination can be generated by deleting the key C-terminal residue from endogenous or chimeric α-tubulin (Badin-Larçon et al., 2004), and polyglutamylation is generated by chemically coupling glutamate side chains to specifically engineered tubulin chimeras (Sirajuddin et al., 2014). These approaches allowed the first direct measurements of the impact of tubulin isotypes and PTMs on the behavior of molecular motors in vitro (Sirajuddin et al., 2014) and the analysis of the effects of tubulin heterogeneity on microtubule behavior and interactions inside the yeast cell (Badin-Larçon et al., 2004; Aiken et al., 2014).Currently, the most promising development has been the successful purification of fully functional recombinant tubulin from the baculovirus expression system (Minoura et al., 2013). Using this system, defined α/β-tubulin dimers can be obtained using two different epitope tags on α- and β-tubulin, respectively. Although these epitope tags are essential for separating recombinant from the endogenous tubulin, they could also affect tubulin assembly or microtubule–MAP interactions. Thus, future developments should focus on eliminating these tags.Current efforts have brought the possibility of producing recombinant tubulin into reach. Further improvement and standardization of these methods will certainly provide a breakthrough in understanding the mechanisms by which tubulin heterogeneity contributes to microtubule functions.

Complexity of tubulin—understanding the regulatory principles.

The diversity of tubulin genes (isotypes) and the complexity of tubulin PTMs have led to the proposal of the term “tubulin code” (Verhey and Gaertig, 2007; Wehenkel and Janke, 2014), in analogy to the previously coined histone code (Jenuwein and Allis, 2001). Tubulin molecules consist of a highly structured and thus evolutionarily conserved tubulin body and the unstructured and less conserved C-terminal tails (Nogales et al., 1998). As PTMs and sequence variations within the tubulin body are expected to affect the conserved tubulin fold and therefore the properties of the microtubule lattice, they are not likely to be involved in generating the tubulin code. In contrast, modulations of the C-terminal tails could encode signals on the microtubule surface without perturbing basic microtubule functions and properties (Figs. 1 A and ​and4).4). Indeed, the highest degree of gene-encoded diversity (Fig. 2) and the highest density and complexity of PTMs (Fig. 1) are found within these tail domains.Open in a separate windowFigure 4.Molecular components of the tubulin code. Schematic representation of potential coding elements that could generate specific signals for the tubulin code. (A) The length of the C-terminal tails of different tubulin isotypes differ significantly (Fig. 2) and could have an impact on the interactions between microtubules and MAPs. (B) Tubulin C-terminal tails are rich in charged amino acid residues. The distribution of these residues and local densities of charges could influence the electrostatic interactions with the tails and the readers. (C) Although each glutamate residue within the C-terminal tails could be considered a potential modification site, only some sites have been found highly occupied in tubulin purifications from native sources. This indicates selectivity of the modification reactions, which can participate in the generation of specific modification patterns (see D). Modification sites might be distinguished by their neighboring amino acid residues, which could create specific modification epitopes. (D) As a result of the large number of modification sites and the variability of side chains, a large variety of modification patterns could be generated within a single C-terminal tail of tubulin. (E) Modification patterns as shown in D can be distinct between α- and β-tubulin. These modification patterns could be differentially distributed at the surface of the microtubule lattice, thus generating a higher-order patterning. Tub, tubulin. For color coding, see Fig. 2.Considering the number of tubulin isotypes plus all potential combinations of PTMs (e.g., each glutamate residue within the C-terminal tubulin tail could be modified by either polyglutamylation or polyglycylation, each of them generating side chains of different lengths; Fig. 4), the number of distinct signals generated by the potential tubulin code would be huge. However, as many of these potential signals represent chemical structures that are similar and might not be reliably distinguished by readout mechanisms, it is possible that the tubulin code generates probabilistic signals. In this scenario, biochemically similar modifications would have similar functional readouts, and marginal differences between those signals would only bias biological processes but not determine them. This stands in contrast to the concept of the histone code, in which precise patterns of different PTMs on the histone proteins encode distinct biological signals.The concept of probabilistic signaling is already inscribed in the machinery that generates the tubulin code. Polyglutamylases and polyglycylases from the TTLL family have preferential activities for either α- or β-tubulin and for generating different lengths of the branched glutamate or glycine chains. Although under conditions of low enzyme concentrations, as found in most cells and tissues, the enzymes seem to selectively generate their preferential type of PTM, higher enzyme concentrations induce a more promiscuous behavior, leading, for instance, to a loss of selectivity for α- or β-tubulin (van Dijk et al., 2007). Similarly, the modifying enzymes might prefer certain modification sites within the C-terminal tails of tubulin but might be equally able to modify other sites, which could be locally regulated in cells. For example, β-tubulin isotypes isolated from mammalian brain were initially found to be glutamylated on single residues (Alexander et al., 1991; Rüdiger et al., 1992), which in the light of the comparably low sensitivity of mass spectrometry at the time might rather indicate a preferential than a unique modification of these sites. Nevertheless, the neuron-specific polyglutamylase for β-tubulin TTLL7 (Ikegami et al., 2006) can incorporate glutamate onto many more modification sites of β-tubulin in vitro (Mukai et al., 2009), which clearly indicates that not all of the possible modification events take place under physiological conditions.Several examples supporting a probabilistic signaling mode of the tubulin code are found in the recent literature. In T. thermophila, a ciliate without tubulin isotype diversity (Gaertig et al., 1993) but with a huge repertoire of tubulin PTMs and tubulin-modifying enzymes (Janke et al., 2005), tubulin can be easily mutagenized to experimentally eliminate sites for PTMs. Mutagenesis of the most commonly occupied glutamylation/glycylation sites within the β-tubulin tails did not generate a clear decrease of glycylation levels nor did it cause obvious phenotypic alterations. This indicates that the modifying enzymes can deviate toward alternative modification sites and that similar PTMs on different sites can compensate the functions of the mutated site. However, when all of the key modification sites were mutated, glycylation became prominently decreased, which led to severe phenotypes, including lethality (Xia et al., 2000). Most strikingly, these phenotypes could be recovered by replacing the C-terminal tail of α-tubulin with the nonmutated β-tubulin tail. This α–β-tubulin chimera became overglycylated and functionally compensated for the absence of modification sites on β-tubulin. The conclusion of this study is that PTM- and isotype-generated signals can fulfill a biological function within a certain range of tolerance.But how efficient is such compensation? The answer can be found in a variety of already described deletion mutants for tubulin-modifying enzymes in different model organisms. Most single-gene knockouts for TTLL genes (glutamylases or glycylases) did not result in prominent phenotypic alterations in mice, even for enzymes that are ubiquitously expressed. Only some highly specialized microtubule structures show functional aberrations upon the deletion of a single enzyme. These “tips of the iceberg” are usually the motile cilia and sperm flagella, which carry very high levels of polyglutamylation and polyglycylation (Bré et al., 1996; Kann et al., 1998; Rogowski et al., 2009). It thus appears that some microtubules are essentially dependent on the generation of specific PTM patterns, whereas others can tolerate changes and appear to function normally. How “normal” these functions are remains to be investigated in future studies. It is possible that defects are subtle and thus overlooked but could become functionally important under specific conditions.A tubulin code also requires readout mechanisms. The most likely “readers” of the tubulin code are MAPs and molecular motors. Considering the probabilistic signaling hypothesis, the expected effects of the signals would be in most cases rather gradual changes, for instance, to fine-tune molecular motor traffic and/or to bias motors toward defined microtubule tracks but not to obliterate motor activity or MAP binding to microtubules. An in vitro study using recombinant tubulin chimeras purified from yeast confirmed this notion (Sirajuddin et al., 2014). By analyzing which elements of the tubulin code can regulate the velocity and processivity of the molecular motors kinesin and dynein, these researchers found that the C-terminal tails of α- and β-tubulin differentially influence the kinetic parameters of the tested motors; however, the modulation was rather modest. One of their striking observations was that a single lysine residue, present in the C-terminal tails of two β-tubulin isotypes (Figs. 2 and ​and4),4), significantly affected motor traffic and that this effect can be counterbalanced by polyglutamylation. These observations are the first in vitro evidence for the interdependence of different elements of the tubulin code and provide another indication for its probabilistic mode of signaling.

Future directions.

One of the greatest technological challenges to understanding the function of the tubulin code is to detect and interpret subtle and complex regulatory events generated by this code. It will thus be instrumental to further develop tools to better distinguish graded changes in PTM levels on microtubules in cells and tissues (Magiera and Janke, 2013) and to reliably measure subtle modulations of microtubule behavior in reconstituted systems.The current advances in the field and especially the availability of whole-organism models, as well as first insights into the pathological role of tubulin mutations (Tischfield et al., 2011), are about to transform our way of thinking about the regulation of microtubule cytoskeleton. Tubulin heterogeneity generates complex probabilistic signals that cannot be clearly attributed to single biological functions in most cases and that are not essential for most cellular processes. Nevertheless, it has been conserved throughout evolution of eukaryotes and can hardly be dismissed as not important. To understand the functional implications of these processes, we might be forced to reconsider how we define biologically important events and how we measure events that might encode probabilistic signals. The answers to these questions could provide novel insights into how complex systems, such as cells and organisms, are sustained throughout difficult and challenging life cycles, resist to environmental stress and diseases, and have the flexibility needed to succeed in evolution.     

Identification of a Novel Abscisic Acid-Regulated Farnesol Dehydrogenase from Arabidopsis

In Arabidopsis (Arabidopsis thaliana), farnesylcysteine is oxidized to farnesal and cysteine by a membrane-associated thioether oxidase called farnesylcysteine lyase. Farnesol and farnesyl phosphate kinases have also been reported in plant membranes. Together, these observations suggest the existence of enzymes that catalyze the interconversion of farnesal and farnesol. In this report, Arabidopsis membranes are shown to possess farnesol dehydrogenase activity. In addition, a gene on chromosome 4 of the Arabidopsis genome (At4g33360), called FLDH, is shown to encode an NAD⁺-dependent dehydrogenase that oxidizes farnesol more efficiently than other prenyl alcohol substrates. FLDH expression is repressed by abscisic acid (ABA) but is increased in mutants with T-DNA insertions in the FLDH 5′ flanking region. These T-DNA insertion mutants, called fldh-1 and fldh-2, are associated with an ABA-insensitive phenotype, suggesting that FLDH is a negative regulator of ABA signaling.Isoprenylated proteins are modified at the C terminus via cysteinyl thioether linkage to either a 15-carbon farnesyl or a 20-carbon geranylgeranyl group (Clarke, 1992; Zhang and Casey, 1996; Rodríguez-Concepción et al., 1999; Crowell, 2000; Crowell and Huizinga, 2009). These modifications mediate protein-membrane and protein-protein interactions and are necessary for the proper localization and function of hundreds of proteins in eukaryotic cells. In Arabidopsis (Arabidopsis thaliana), the PLURIPETALA (PLP; At3g59380) and ENHANCED RESPONSE TO ABA1 (At5g40280) genes encode the α- and β-subunits of protein farnesyltransferase (PFT), respectively (Cutler et al., 1996; Pei et al., 1998; Running et al., 2004). These subunits form a heterodimeric zinc metalloenzyme that catalyzes the efficient transfer of a farnesyl group from farnesyl diphosphate to protein substrates with a C-terminal CaaX motif, where “C” is Cys, “a” is an aliphatic amino acid, and “X” is usually Met, Gln, Cys, Ala, or Ser (Fig. 1). The PLP and GERANYLGERANYL-TRANSFERASE BETA (At2g39550) genes encode the α- and β-subunits of protein geranylgeranyltransferase type 1 (PGGT1), respectively (Running et al., 2004; Johnson et al., 2005). These subunits form a distinct heterodimeric zinc metalloenzyme that catalyzes the efficient transfer of a geranylgeranyl group from geranylgeranyl diphosphate to protein substrates with a C-terminal CaaL motif, where “C” is Cys, “a” is an aliphatic amino acid, and “L” is Leu. A third protein prenyltransferase, called protein geranylgeranyltransferase type II or RAB geranylgeranyltransferase, catalyzes the dual geranylgeranylation of RAB proteins with a C-terminal XCCXX, XXCXC, XXCCX, XXXCC, XCXXX, or CCXXX motif, where “C” is Cys and “X” is any amino acid. However, RAB proteins must be associated with the RAB ESCORT PROTEIN to be substrates of RAB geranylgeranyltransferase. Plant protein prenylation has received considerable attention in recent years because of the meristem defects of Arabidopsis PFT mutants and the abscisic acid (ABA) hypersensitivity of Arabidopsis PFT and PGGT1 mutants (Cutler et al., 1996; Pei et al., 1998; Running et al., 1998, 2004; Johnson et al., 2005).Open in a separate windowFigure 1.Proposed metabolism of farnesal and farnesol as it relates to protein prenylation. The portion of the cycle shown in red is the subject of this article.Proteins that are prenylated by either PFT or PGGT1 undergo further processing in the endoplasmic reticulum (Crowell, 2000; Crowell and Huizinga, 2009). First, the aaX portion of the CaaX motif is removed by proteolysis (Fig. 1). This reaction is catalyzed by one of two CaaX endoproteases, which are encoded by the AtSTE24 (At4g01320) and AtFACE-2 (At2g36305) genes (Bracha et al., 2002; Cadiñanos et al., 2003). Second, the prenylated Cys residue at the new C terminus is methylated by one of two isoprenylcysteine methyltransferases (Fig. 1), which are encoded by the AtSTE14A (At5g23320) and AtSTE14B (ICMT; At5g08335) genes (Crowell et al., 1998; Crowell and Kennedy, 2001; Narasimha Chary et al., 2002; Bracha-Drori et al., 2008). A specific isoprenylcysteine methylesterase encoded by the Arabidopsis ICME (At5g15860) gene has also been described, demonstrating the reversibility of isoprenylcysteine methylation (Deem et al., 2006; Huizinga et al., 2008).Like all proteins, prenylated proteins have a finite half-life. However, unlike other proteins, prenylated proteins release farnesylcysteine (FC) or geranylgeranylcysteine (GGC) upon degradation. Mammals possess a prenylcysteine lyase enzyme that catalyzes the oxidative cleavage of FC and GGC (Zhang et al., 1997; Tschantz et al., 1999; Tschantz et al., 2001; Beigneux et al., 2002; Digits et al., 2002). This FAD-dependent thioether oxidase consumes molecular oxygen and generates hydrogen peroxide, Cys, and a prenyl aldehyde product (i.e. farnesal or geranylgeranial). In Arabidopsis, a similar lyase exists. However, the Arabidopsis enzyme, which is encoded by the FCLY (At5g63910) gene, is specific for FC (Fig. 1; Crowell et al., 2007; Huizinga et al., 2010). GGC is metabolized by a different mechanism.Plant membranes have been shown to contain farnesol kinase, geranylgeraniol kinase, farnesyl phosphate kinase, and geranylgeranyl phosphate kinase activities (Fig. 1; Thai et al., 1999). These membrane-associated kinases differ with respect to nucleotide specificity, suggesting that they are distinct enzymes (i.e. farnesol kinase and geranylgeraniol kinase can use CTP, UTP, or GTP as a phosphoryl donor, whereas farnesyl phosphate kinase and geranylgeranyl phosphate kinase exhibit specificity for CTP as a phosphoryl donor). However, it remains unclear if farnesol kinase is distinct from geranylgeraniol kinase or if farnesyl phosphate kinase is distinct from geranylgeranyl phosphate kinase. Nonetheless, it is clear that these kinases convert farnesol and geranylgeraniol to their monophosphate and diphosphate forms for use in isoprenoid biosynthesis, including sterol biosynthesis and protein prenylation.Because plants have the metabolic capability to generate farnesal from FC and farnesyl diphosphate from farnesol, we considered the possibility that plant membranes also contain an oxidoreductase capable of catalyzing the reduction of farnesal to farnesol and/or the oxidation of farnesol to farnesal (Fig. 1; Thai et al., 1999; Crowell et al., 2007). To date, the only reports of such an oxidoreductase are from the corpora allata glands of insects, where it participates in juvenile hormone synthesis, and black rot fungus-infected sweet potato (Ipomoea batatas; Baker et al., 1983; Inoue et al., 1984; Sperry and Sen, 2001; Mayoral et al., 2009). Insect farnesol dehydrogenase is an NADP⁺-dependent oxidoreductase that is encoded by a subfamily of short-chain dehydrogenase/reductase (SDR) genes (Mayoral et al., 2009). Farnesol dehydrogenase from sweet potato is a 90-kD, NADP⁺-dependent homodimer with broad specificity for prenyl alcohol substrates and is induced by wounding and fungus infection of potato roots (Inoue et al., 1984).Here, we extended previous work in which [1-³H]FC was shown to be oxidized to [1-³H]farnesal, and [1-³H]farnesal reduced to [1-³H]farnesol, in the presence of Arabidopsis membranes (Crowell et al., 2007). The reduction of [1-³H]farnesal to [1-³H]farnesol was abolished by pretreatment of Arabidopsis membranes with NADase, suggesting that sufficient NAD(P)H is present in Arabidopsis membranes to support the enzymatic reduction of farnesal to farnesol. In this report, we demonstrate the presence of farnesol dehydrogenase activity in Arabidopsis membranes using [1-³H]farnesol as a substrate. Moreover, we identify a gene on chromosome 4 of the Arabidopsis genome (At4g33360), called FLDH, that encodes an NAD⁺-dependent dehydrogenase with partial specificity for farnesol as a substrate. FLDH expression is repressed by exogenous ABA, and fldh mutants exhibit altered ABA signaling. Taken together, these observations suggest that ABA regulates farnesol metabolism in Arabidopsis, which in turn regulates ABA signaling.     

Live to cheat another day: bacterial dormancy facilitates the social exploitation of β-lactamases

The breakdown of antibiotics by β-lactamases may be cooperative, since resistant cells can detoxify their environment and facilitate the growth of susceptible neighbours. However, previous studies of this phenomenon have used artificial bacterial vectors or engineered bacteria to increase the secretion of β-lactamases from cells. Here, we investigated whether a broad-spectrum β-lactamase gene carried by a naturally occurring plasmid (pCT) is cooperative under a range of conditions. In ordinary batch culture on solid media, there was little or no evidence that resistant bacteria could protect susceptible cells from ampicillin, although resistant colonies could locally detoxify this growth medium. However, when susceptible cells were inoculated at high densities, late-appearing phenotypically susceptible bacteria grew in the vicinity of resistant colonies. We infer that persisters, cells that have survived antibiotics by undergoing a period of dormancy, founded these satellite colonies. The number of persister colonies was positively correlated with the density of resistant colonies and increased as antibiotic concentrations decreased. We argue that detoxification can be cooperative under a limited range of conditions: if the toxins are bacteriostatic rather than bacteridical; or if susceptible cells invade communities after resistant bacteria; or if dormancy allows susceptible cells to avoid bactericides. Resistance and tolerance were previously thought to be independent solutions for surviving antibiotics. Here, we show that these are interacting strategies: the presence of bacteria adopting one solution can have substantial effects on the fitness of their neighbours.A cooperative trait is a behaviour by one individual that can benefit another (West et al., 2006). In bacteria, cooperative traits often come in the form of ‘public goods'' released into the environment and available to all. Many virulence factors, including siderophore production, Cry proteins and quorum-regulated traits are cooperative (West and Buckling, 2003; Diggle et al., 2007; Sandoz et al., 2007; Raymond et al., 2012; Zhou et al., 2014). Antibiotic resistance conferred by the enzymatic breakdown of drugs is potentially a cooperative trait as it can detoxify the environment for all cells, and the production of β-lactamases, which cleave and deactivate penicillins, is often cited as a social trait in bacteria (West et al., 2006; Diggle et al., 2007; Brown et al., 2009). Clinical studies have suggested that protective clearance is mediated by the release of β-lactamase enzymes into the environment by producing cells (Brook, 2004), and packaging of β-lactamases into extracellular vesicles has been demonstrated in Pseudomonas aeruginosa (Ciofu et al., 2000). Secretion of enzymes could increase the area of antibiotic clearance, benefiting all cells in a local population.The phenomenon of protective clearance of antibiotics by resistant cells is commonly seen by microbiologists in the presence of ‘satellite'' colonies on transformation plates (Figure 1a). These non-resistant colonies are able to grow on ampicillin plates where resistant colonies are already established. A plausible hypothesis that explains this phenomenon is that resistant transformants clear the antibiotic from their immediate vicinity, creating an antibiotic-free space where susceptible ‘satellite'' colonies can then grow. Previous studies have demonstrated the survival of antibiotic-sensitive Escherichia coli and Salmonella sp. in the presence of resistant strains at high concentrations of antibiotic in vitro (Dugatkin et al., 2005; Clark et al., 2009; Perlin et al., 2009). Cross-species protection of susceptible bacteria by β-lactamase producers has also been seen in vivo (Tacking, 1954; Hackman and Wilkins, 1975; Brook et al., 1983), suggesting that the benefits of β-lactamase enzymes may spread to entire communities.Open in a separate windowFigure 1(a) Satellite colonies around a successful pUC19 transformant on ampicillin agar. Transformation of E. coli DH10B with pUC19 confers resistance to ampicillin and also restores the lac operon, resulting in a blue colony on agar containing X-Gal and IPTG. The successful transformants appeared after ~16 h, white susceptible colonies appeared after >24 h incubation and can be seen growing around the resistant transformants. (b) Antibiotic clearance bioassay. Susceptible colonies grow on ampicillin plates in the presence of resistant colonies (bottom row), but not alone (top row). Resistant colonies were incubated for 48 h on 100 μg ml⁻¹ ampicillin plates before the addition of susceptible colonies. Susceptible strains can grow in the region around the resistant colony, but do not grow outside the central zone or in the absence of a resistant colony (top row). These assays were repeated seven times.Although social evolution theory has done much to alter and improve modern microbial ecology, there is some justification for being cautious about claiming whether specific traits are cooperative or not. A recent controversy has highlighted two important points when studying cooperation: first, we need to demonstrate that behaviours have real fitness benefits for populations, and second that it is desirable to study social evolution with realistic models (Zhang, 2004; Ghoul et al., 2014), something we have endeavoured to do in previous studies (Raymond et al., 2012; Zhou et al., 2014). For example, demonstrating that a microbial product is secreted is not sufficient evidence for cooperation; spatial structure, for example, can prevent secreted products from being publicly available (Raymond and Bonsall, 2013; Zhou et al., 2014), or metabolic products may not be beneficial in all contexts (Zhang and Rainey, 2013; Ghoul et al., 2014). Conversely, secretion may not be necessary for cooperation in the case of detoxification. Active removal of toxins and the ensuing diffusion gradients or lowered concentration of toxin may be all that is required to protect susceptible bacteria (Lenski and Hattingh, 1986).There are some grounds for being sceptical about any claim that β-lactamases are generally cooperative. Previous experiments have used model bacteria with altered sites of expression and potentially increased secretion (Dugatkin et al., 2005; Perlin et al., 2009) and these data might not reflect social interactions in more natural conditions. It is important therefore to consider whether antibiotic resistance genes in the more realistic context of naturally-occurring plasmids can lead to cooperative detoxification of antibiotics. An additional consideration is whether the antibiotic in question is bacteriostatic or bactericidal. When β-lactams are bacteriostatic, that is, if they suspend growth but do not rapidly kill bacteria, then the potential for social interactions may be increased as susceptible bacteria may survive until detoxification by neighbours can reduce concentrations of antibiotics to below inhibitory doses. However, β-lactam antibiotics can be bactericidal, that is, rapidly lethal to bacteria, under a range of conditions (Rolinson et al., 1977; Cozens et al., 1986). Any bactericidal activity is expected to substantially restrict the conditions for coexistence and social exploitation to a narrow range that may depend on initial dosage, as well as the frequency of resistant and susceptible bacteria (Lenski and Hattingh, 1986; Levin, 1988).However, bacteria do have mechanisms that enable them to escape or tolerate the effects of bactericidal antibiotics, one being a ‘persister'' state in which dormant cells can survive exposure to antibiotics (Lewis, 2010). Persister cells were identified early in the clinical life of penicillin (Bigger, 1944), but a recent resurgence in interest has been fuelled by a wider appreciation of their clinical importance, especially in the light of the current antibiotic resistance crisis (Lewis, 2007). Persister cells are natural variants present at low frequency in the bacterial population (Lewis, 2010). The phenotypic switch between persistence and active growth appears to occur at random, although it is affected by growth phase (Balaban, 2004). The presence of persisters in biofilms is thought to contribute to increased antimicrobial tolerance and the maintenance of chronic infections (Brooun et al., 2000; Lewis, 2001; Harrison et al., 2005). Since satellite colonies on ampicillin plates are characterized by a delayed growth pattern, appearing after 24–72 h of cultures, and emerge as rare individuals from a high density of bacteria, they have many of the characteristics of cells that have passed through a persister state. We therefore hypothesised that phenotypically susceptible bacteria might only rarely be able to benefit from detoxification by others while persisters may be more likely to exploit the β-lactamases of their neighbours.The primary aim of this work was to examine cooperative β-lactam resistance using a naturally occurring resistance plasmid, pCT, (Cottell et al., 2011) and to investigate both the environmental and demographic conditions under which cooperative resistance occurs. In line with social evolution theory, we expected that ‘cheating'' or exploitation of detoxification by susceptible cells should increase with the density and frequency of resistant bacteria (Ross-Gillespie et al., 2007, 2009; Raymond et al., 2012). To this end, competition experiments were conducted between the pCT-carrying strain and an otherwise isogenic plasmid-free E. coli under a variety of conditions. The results of these experiments showed little or no cooperative resistance, in other words phenotypically susceptible bacteria did not tend to have an increased ability to survive antibiotics in the presence of resistant cells. We then tested whether dormancy has a role in the ability of E. coli to exploit the β-lactamase activity of neighbouring cells.     

A Real-Time Fluorogenic Assay for the Visualization of Glycoside Hydrolase Activity in Planta

There currently exists a diverse array of molecular probes for the in situ localization of polysaccharides, nucleic acids, and proteins in plant cells, including reporter enzyme strategies (e.g. protein-glucuronidase fusions). In contrast, however, there is a paucity of methods for the direct analysis of endogenous glycoside hydrolases and transglycosidases responsible for cell wall remodeling. To exemplify the potential of fluorogenic resorufin glycosides to address this issue, a resorufin β-glycoside of a xylogluco-oligosaccharide (XXXG-β-Res) was synthesized as a specific substrate for in planta analysis of XEH activity. The resorufin aglycone is particularly distinguished for high sensitivity in muro assays due to a low pK_a (5.8) and large extinction coefficient (ε 62,000 m⁻¹cm⁻¹), long-wavelength fluorescence (excitation 571 nm/emission 585 nm), and high quantum yield (0.74) of the corresponding anion. In vitro analyses demonstrated that XXXG-β-Res is hydrolyzed by the archetypal plant XEH, nasturtium (Tropaeolum majus) NXG1, with classical Michaelis-Menten substrate saturation kinetics and a linear dependence on both enzyme concentration and incubation time. Further, XEH activity could be visualized in real time by observing the localized increase in fluorescence in germinating nasturtium seeds and Arabidopsis (Arabidopsis thaliana) inflorescent stems by confocal microscopy. Importantly, this new in situ XEH assay provides an essential complement to the in situ xyloglucan endotransglycosylase assay, thus allowing delineation of the disparate activities encoded by xyloglucan endotransglycosylase/hydrolase genes directly in plant tissues. The observation that XXXG-β-Res is also hydrolyzed by diverse microbial XEHs indicates that this substrate, and resorufin glycosides in general, may find broad applicability for the analysis of wall restructuring by polysaccharide hydrolases during morphogenesis and plant-microbe interactions.The development and application of molecular probes for the localization of biomolecules in planta continues to have a profound impact on the field of plant physiology. A number of elegant techniques have been devised for the detection of nucleic acids, polypeptides, and polysaccharides in situ, including DNA/RNA hybridization (Jin and Lloyd, 1997), reporter protein fusions (Jefferson et al., 1987; Ehrhardt, 2003; Chapman et al., 2005; Stewart, 2005; Berg and Beachy, 2008; Nelson et al., 2008), immunohistochemical methods (Walker et al., 2001; Chapman et al., 2005), applications of natural carbohydrate-binding proteins (Knox, 2008), and direct spectroscopy (Vicente et al., 2007). While there now exists a considerable toolbox to identify the location to which biomolecules are directed in the cell, elucidation of specific biochemical function at the site of localization often remains challenging.Presently, there is a growing interest in the roles of glycoside hydrolases (GHs) and transglycosylases in plant cell wall biogenesis, remodeling, and degradation (Minic and Jouanin, 2006; Vicente et al., 2007; Gilbert et al., 2008; Lopez-Casado et al., 2008). A technical limitation of many studies, however, is that enzyme activities can only be measured for crude whole-tissue extracts, or purified or recombinant enzymes, and thus cannot be directly correlated with the high-resolution in situ localization of other biomacromolecules. As such, the in situ analysis of GH activities responsible for the degradation of plant cell wall polysaccharides has received comparatively little attention, primarily due to a paucity of convenient assay methods (Vicente et al., 2007). Some notable exceptions include the use of commercially available X (5-bromo-4-chloro-3-indolyl glycoside) substrates for the detection of exoglycosidase activity (Monroe et al., 1999; Chantarangsee et al., 2007; Macquet et al., 2007; Wen et al., 2008). Likewise, transglycosylase activity has been visualized in higher plant and yeast cell walls using sulforhodamine-oligosaccharide acceptor substrates (Vissenberg et al., 2000; Bourquin et al., 2002; Nishikubo et al., 2007; Cabib et al., 2008). Both types are examples of end point, or stopped, assays, in which precipitated indigoid dyes or incorporated fluorescent oligosaccharide conjugate, respectively, are observed after a terminal incubation time.In this study, we have developed the use of resorufin glycosides as substrates for the real-time, continuous observation of GH activity in situ (Fig. 1). Enzymatic hydrolysis of such substrates releases the resorufin aglycone, which is distinguished by a low pK_a value (5.8) and a large extinction coefficient (ε 62,000 m⁻¹cm⁻¹), long-wavelength fluorescence (excitation/emission maxima, 571 nm/585 nm), and high quantum yield (0.74) of the resorufinyl anion (Bueno et al., 2002). The pK_a value and spectral properties make resorufin glycosides particularly suitable for high sensitivity in muro enzyme activity assays due to significant ionization of resorufin at typical apoplastic pH values (Felle, 2005). To highlight the potential of this class of substrates in cell wall morphological studies, we have chemically synthesized a xylogluco-oligosaccharide (XGO) resorufin β-glycoside (XXXG-β-Res; Fig. 1 [1]; XGO nomenclature according to Fry et al., 1993) and demonstrated its use for the real-time imaging of xyloglucan endohydrolase (XEH) activity in plant tissues from nasturtium (Tropaeolum majus) and Arabidopsis (Arabidopsis thaliana) by confocal fluorescence microscopy.Open in a separate windowFigure 1.Use of resorufin glycosides as fluorogenic substrates for glycosidases. R = saccharide or hydrogen; [1] and [2], substrates for determination of (xylo)glucanase activity. Oligosaccharide nomenclature is according to Fry et al. (1993). [See online article for color version of this figure.]     

Enzymatic Formation of β-Citraurin from β-Cryptoxanthin and Zeaxanthin by Carotenoid Cleavage Dioxygenase4 in the Flavedo of Citrus Fruit

In this study, the pathway of β-citraurin biosynthesis, carotenoid contents and the expression of genes related to carotenoid metabolism were investigated in two varieties of Satsuma mandarin (Citrus unshiu), Yamashitabeni-wase, which accumulates β-citraurin predominantly, and Miyagawa-wase, which does not accumulate β-citraurin. The results suggested that CitCCD4 (for Carotenoid Cleavage Dioxygenase4) was a key gene contributing to the biosynthesis of β-citraurin. In the flavedo of Yamashitabeni-wase, the expression of CitCCD4 increased rapidly from September, which was consistent with the accumulation of β-citraurin. In the flavedo of Miyagawa-wase, the expression of CitCCD4 remained at an extremely low level during the ripening process, which was consistent with the absence of β-citraurin. Functional analysis showed that the CitCCD4 enzyme exhibited substrate specificity. It cleaved β-cryptoxanthin and zeaxanthin at the 7,8 or 7′,8′ position. But other carotenoids tested in this study (lycopene, α-carotene, β-carotene, all-trans-violaxanthin, and 9-cis-violaxanthin) were not cleaved by the CitCCD4 enzyme. The cleavage of β-cryptoxanthin and zeaxanthin by CitCCD4 led to the formation of β-citraurin. Additionally, with ethylene and red light-emitting diode light treatments, the gene expression of CitCCD4 was up-regulated in the flavedo of Yamashitabeni-wase. These increases in the expression of CitCCD4 were consistent with the accumulation of β-citraurin in the two treatments. These results might provide new strategies to improve the carotenoid contents and compositions of citrus fruits.Carotenoids, a diverse group of pigments widely distributed in nature, fulfill a variety of important functions in plants and play a critical role in human nutrition and health (Schwartz et al., 1997; Cunningham and Gantt, 1998; Havaux, 1998; Krinsky et al., 2003; Ledford and Niyogi, 2005). The pathway of carotenoid biosynthesis has been well documented in various plant species, including Arabidopsis (Arabidopsis thaliana; Park et al., 2002), tomato (Lycopersicon esculentum; Isaacson et al., 2002), pepper (Capsicum annuum; Bouvier et al., 1998), citrus (Citrus spp.; Kato et al., 2004, 2006; Rodrigo et al., 2004; Rodrigo and Zacarías, 2007; Kato, 2012; Zhang et al., 2012a), and apricot (Prunus armenaica; Kita et al., 2007). Genes encoding the enzymes in the carotenoid biosynthetic pathway have been cloned, and their expression profiles have also been characterized (Fig. 1). As carotenoids contain a series of conjugated double bonds in the central chain, they can be oxidatively cleaved in a site-specific manner (Mein et al., 2011). The oxidative cleavage of carotenoids not only regulates their accumulation but also produces a range of apocarotenoids (Walter et al., 2010). In higher plants, many different apocarotenoids derive from the cleavage of carotenoids and have important metabolic functions, such as plant hormones, pigments, aroma and scent compounds, as well as signaling compounds (Fig. 1). A well-known example is abscisic acid, which is a C₁₅ compound derived from the cleavage of the 11,12 double bond of 9-cis-violaxanthin and 9′-cis-neoxanthin (Schwartz et al., 1997; Tan et al., 1997; Cutler and Krochko, 1999; Chernys and Zeevaart, 2000; Giuliano et al., 2003).Open in a separate windowFigure 1.Carotenoid and apocarotenoid metabolic pathway in plants. GGPP, Geranylgeranyl diphosphate. Enzymes, listed here from top to bottom, are named according to the designation of their genes: PSY, phytoene synthase; PDS, Phytoene desaturase; ZDS, ζ-carotene desaturase; ZISO, 15-cis-ζ-carotene isomerase; CRTISO, carotenoid isomerase; LCYb, lycopene β-cyclase; LCYe, lycopene ε-cyclase; HYe, ε-ring hydroxylase; HYb, β-ring hydroxylase; ZEP, zeaxanthin epoxidase; VDE, violaxanthin deepoxidase; NCED, 9-cis-epoxycarotenoid dioxygenase.Carotenoid cleavage dioxygenases (CCDs) are a group of enzymes that catalyze the oxidative cleavage of carotenoids (Ryle and Hausinger, 2002). CCDs are nonheme iron enzymes present in plants, bacteria, and animals. In plants, CCDs belong to an ancient and highly heterogenous family (CCD1, CCD4, CCD7, CCD8, and 9-cis-epoxycarotenoid dioxygenases [NCEDs]). The similarity among the different members is very low apart from four strictly conserved His residues and a few Glu residues (Kloer and Schulz, 2006; Walter et al., 2010). In Arabidopsis, the CCD family contains nine members (CCD1, NCED2, NCED3, CCD4, NCED5, NCED6, CCD7, CCD8, and NCED9), and orthologs in other plant species are typically named according to their homology with an Arabidopsis CCD (Huang et al., 2009). In our previous study, the functions of CitCCD1, CitNCED2, and CitNCED3 were investigated in citrus fruits (Kato et al., 2006). The recombinant CitCCD1 protein cleaved β-cryptoxanthin, zeaxanthin, and all-trans-violaxanthin at the 9,10 and 9′,10′ positions and 9-cis-violaxanthin at the 9′,10′ position. The recombinant CitNCED2 and CitNCED3 proteins cleaved 9-cis-violaxanthin at the 11,12 position to form xanthoxin, a precursor of abscisic acid (Kato et al., 2006). To date, information on the functions of other CCDs in citrus fruits remains limited, while the functions of CCD7 and CCD8, as well as NCED5, NCED6, and NCED9, in Arabidopsis have been characterized (Kloer and Schulz, 2006; Walter et al., 2010). In Arabidopsis, CCD7 cleaves all-trans-β-carotene at the 9′,10′ position to form all-trans-β-apo-10′-carotenal. All-trans-β-apo-10′-carotenal is further shortened by AtCCD8 at the 13,14 position to produce β-apo-13-carotenone (Alder et al., 2012). NCED5, NCED6, and NCED9 cleave 9-cis-violaxanthin at the 11,12 position to form xanthoxin (Tan et al., 2003). Compared with other CCDs, the function of CCD4 is poorly understood. In Chrysanthemum morifolium, CmCCD4a contributed to the white color formation by cleaving carotenoids into colorless compounds (Ohmiya et al., 2006). Recently, it has been reported that CsCCD4, CmCCD4a, and MdCCD4 could cleave β-carotene to yield β-ionone (Rubio et al., 2008; Huang et al., 2009).β-Citraurin, a C₃₀ apocarotenoid, is a color-imparting pigment responsible for the reddish color of citrus fruits (Farin et al., 1983). In 1936, it was first discovered in Sicilian oranges (Cual, 1965). In citrus fruits, the accumulation of β-citraurin is not a common event; it is only observed in the flavedos of some varieties during fruit ripening. The citrus varieties accumulating β-citraurin are considered more attractive because of their red-orange color (Ríos et al., 2010). Although more than 70 years have passed since β-citraurin was first identified, the pathway of its biosynthesis is still unknown. As its structure is similar to that of β-cryptoxanthin and zeaxanthin, β-citraurin was presumed to be a degradation product of β-cryptoxanthin or zeaxanthin (Oberholster et al., 2001; Rodrigo et al., 2004; Ríos et al., 2010; Fig. 1). To date, however, the specific cleavage reaction producing β-citraurin has not been elucidated. In this study, we found that the CitCCD4 gene was involved in the synthesis of β-citraurin, using two citrus varieties of Satsuma mandarin (Citrus unshiu), Yamashitabeni-wase, which accumulates β-citraurin predominantly, and Miyagawa-wase, which does not accumulate β-citraurin. To confirm the role of the CitCCD4 gene further, functional analyses of the CitCCD4 enzyme were performed in vivo and in vitro. Additionally, the regulation of β-citraurin content and CitCCD4 gene expression in response to ethylene and red light-emitting diode (LED) light treatments was also examined. This study, to our knowledge, is the first to investigate the biosynthesis of β-citraurin in citrus fruits. The results might provide new strategies to enhance the nutritional and commercial qualities of citrus fruits.     

Phenotypic Plasticity in Photosynthetic Temperature Acclimation among Crop Species with Different Cold Tolerances

While interspecific variation in the temperature response of photosynthesis is well documented, the underlying physiological mechanisms remain unknown. Moreover, mechanisms related to species-dependent differences in photosynthetic temperature acclimation are unclear. We compared photosynthetic temperature acclimation in 11 crop species differing in their cold tolerance, which were grown at 15°C or 30°C. Cold-tolerant species exhibited a large decrease in optimum temperature for the photosynthetic rate at 360 μL L⁻¹ CO₂ concentration [Opt (A₃₆₀)] when growth temperature decreased from 30°C to 15°C, whereas cold-sensitive species were less plastic in Opt (A₃₆₀). Analysis using the C₃ photosynthesis model shows that the limiting step of A₃₆₀ at the optimum temperature differed between cold-tolerant and cold-sensitive species; ribulose 1,5-bisphosphate carboxylation rate was limiting in cold-tolerant species, while ribulose 1,5-bisphosphate regeneration rate was limiting in cold-sensitive species. Alterations in parameters related to photosynthetic temperature acclimation, including the limiting step of A₃₆₀, leaf nitrogen, and Rubisco contents, were more plastic to growth temperature in cold-tolerant species than in cold-sensitive species. These plastic alterations contributed to the noted growth temperature-dependent changes in Opt (A₃₆₀) in cold-tolerant species. Consequently, cold-tolerant species were able to maintain high A₃₆₀ at 15°C or 30°C, whereas cold-sensitive species were not. We conclude that differences in the plasticity of photosynthetic parameters with respect to growth temperature were responsible for the noted interspecific differences in photosynthetic temperature acclimation between cold-tolerant and cold-sensitive species.The temperature dependence of leaf photosynthetic rate shows considerable variation between plant species and with growth temperature (Berry and Björkman, 1980; Cunningham and Read, 2002; Hikosaka et al., 2006). Plants native to low-temperature environments and those grown at low temperatures generally exhibit higher photosynthetic rates at low temperatures and lower optimum temperatures, compared with plants native to high-temperature environments and those grown at high temperatures (Mooney and Billings, 1961; Slatyer, 1977; Berry and Björkman, 1980; Sage, 2002; Salvucci and Crafts-Brandner, 2004b). For example, the optimum temperature for photosynthesis differs between temperate evergreen species and tropical evergreen species (Hill et al., 1988; Read, 1990; Cunningham and Read, 2002). Such differences have been observed even among ecotypes of the same species (Björkman et al., 1975; Pearcy, 1977; Slatyer, 1977).Temperature dependence of the photosynthetic rate has been analyzed using the biochemical model proposed by Farquhar et al. (1980). This model assumes that the photosynthetic rate (A) is limited by either ribulose 1,5-bisphosphate (RuBP) carboxylation (A_c) or RuBP regeneration (A_r). The optimum temperature for photosynthetic rate in C₃ plants is thus potentially determined by (1) the temperature dependence of A_c, (2) the temperature dependence of A_r, or (3) both, at the colimitation point of A_c and A_r (Fig. 1; Farquhar and von Caemmerer, 1982; Hikosaka et al., 2006).Open in a separate windowFigure 1.A scheme illustrating the shift in the optimum temperature for photosynthesis depending on growth temperature. Based on the C₃ photosynthesis model, the A₃₆₀ (white and black circles) is limited by A_c (solid line) or A_r (broken line). The optimum temperature for the photosynthetic rate is potentially determined by temperature dependence of A_c (A), temperature dependence of A_r (B), or the intersection of the temperature dependences of A_c and A_r (C). When the optimum temperature for the photosynthetic rate shifts to a higher temperature, there are also three possibilities determining the optimum temperature: temperature dependence of A_c (D), temperature dependence of A_r (E), or the intersection of the temperature dependences of A_c and A_r (F). Especially in the case that the optimum temperature is determined by the intersection of the temperature dependences of A_c and A_r, the optimum temperature can shift by changes in the balance between A_c and A_r even when the optimum temperatures for these two partial reactions do not change.In many cases, the photosynthetic rate around the optimum temperature is limited by A_c, and thus the temperature dependence of A_c determines the optimum temperature for the photosynthetic rate (Hikosaka et al., 1999, 2006; Yamori et al., 2005, 2006a, 2006b, 2008; Sage and Kubien, 2007; Sage et al., 2008). As the temperature increases above the optimum, A_c is decreased by increases in photorespiration (Berry and Björkman, 1980; Jordan and Ogren, 1984; von Caemmerer, 2000). Furthermore, it has been suggested that the heat-induced deactivation of Rubisco is involved in the decrease in A_c at high temperature (Law and Crafts-Brandner, 1999; Crafts-Brandner and Salvucci, 2000; Salvucci and Crafts-Brandner, 2004a; Yamori et al., 2006b). Numerous previous studies have shown changes in the temperature dependence of A_c with growth temperature (Hikosaka et al., 1999; Bunce, 2000; Yamori et al., 2005). Also, the temperature sensitivity of Rubisco deactivation may differ between plant species (Salvucci and Crafts-Brandner, 2004b) and with growth temperature (Yamori et al., 2006b), which may explain variation in the optimum temperature for photosynthesis (Fig. 1, A and D).A_r is more responsive to temperature than A_c and often limits photosynthesis at low temperatures (Hikosaka et al., 1999, 2006; Sage and Kubien, 2007; Sage et al., 2008). Recently, several researchers indicated that A_r limits the photosynthetic rate at high temperature (Schrader et al., 2004; Wise et al., 2004; Cen and Sage, 2005; Makino and Sage, 2007). They suggested that the deactivation of Rubisco at high temperatures is not the cause of decreased A_c but a result of limitation by A_r. However, it remains unclear whether limitation by A_r is involved in the variation in the optimum temperature for the photosynthetic rate (Fig. 1, B and E).A shift in the optimum temperature for photosynthesis can result from changes in the balance between A_r and A_c, even when the optimum temperatures for these two partial reactions do not change (Fig. 1, C and F; Farquhar and von Caemmerer, 1982). The balance between A_r and A_c has been shown to change depending on growth temperature (Hikosaka et al., 1999; Hikosaka, 2005; Onoda et al., 2005a; Yamori et al., 2005) and often brings about a shift in the colimitation temperature of A_r and A_c. Furthermore, recent studies have shown that plasticity in this balance differs among species or ecotypes (Onoda et al., 2005b; Atkin et al., 2006; Ishikawa et al., 2007). Plasticity in this balance could explain interspecific variation in the plasticity of photosynthetic temperature dependence (Farquhar and von Caemmerer, 1982; Hikosaka et al., 2006), although there has been no evidence in the previous studies that the optimum temperature for photosynthesis occurs at the colimitation point of A_r and A_c.Temperature tolerance differs between species and, with growth temperature, even within species from the same functional group (Long and Woodward, 1989). Bunce (2000) indicated that the temperature dependences of A_r and A_c to growth temperature were different between species from cool and warm climates and that the balance between A_r and A_c was independent of growth temperature for a given plant species. However, it was not clarified what limited the photosynthetic rate or what parameters were important in temperature acclimation of photosynthesis. Recently, we reported that the extent of temperature homeostasis of leaf respiration and photosynthesis, which is assessed as a ratio of rates measured at their respective growth temperatures, differed depending on the extent of the cold tolerance of the species (Yamori et al., 2009b). Therefore, comparisons of several species with different cold tolerances would provide a new insight into interspecific variation of photosynthetic temperature acclimation and their underlying mechanisms. In this study, we selected 11 herbaceous crop species that differ in their cold tolerance (Yamori et al., 2009b) and grew them at two contrasting temperatures, conducting gas-exchange analyses based on the C₃ photosynthesis model (Farquhar et al., 1980). Based on these results, we addressed the following key questions. (1) Does the plasticity in photosynthetic temperature acclimation differ between cold-sensitive and cold-tolerant species? (2) Does the limiting step of photosynthesis at several leaf temperatures differ between plant species and with growth temperature? (3) What determines the optimum temperature for the photosynthetic rate among A_c, A_r, and the intersection of the temperature dependences of A_c and A_r?     

Chloroplast DNA in Mature and Senescing Leaves: A Reappraisal

The fate of plastid DNA (ptDNA) during leaf development has become a matter of contention. Reports on little change in ptDNA copy number per cell contrast with claims of complete or nearly complete DNA loss already in mature leaves. We employed high-resolution fluorescence microscopy, transmission electron microscopy, semithin sectioning of leaf tissue, and real-time quantitative PCR to study structural and quantitative aspects of ptDNA during leaf development in four higher plant species (Arabidopsis thaliana, sugar beet [Beta vulgaris], tobacco [Nicotiana tabacum], and maize [Zea mays]) for which controversial findings have been reported. Our data demonstrate the retention of substantial amounts of ptDNA in mesophyll cells until leaf necrosis. In ageing and senescent leaves of Arabidopsis, tobacco, and maize, ptDNA amounts remain largely unchanged and nucleoids visible, in spite of marked structural changes during chloroplast-to-gerontoplast transition. This excludes the possibility that ptDNA degradation triggers senescence. In senescent sugar beet leaves, reduction of ptDNA per cell to ∼30% was observed reflecting primarily a decrease in plastid number per cell rather than a decline in DNA per organelle, as reported previously. Our findings are at variance with reports claiming loss of ptDNA at or after leaf maturation.In vascular plants, copy numbers of plastid genomes (plastomes) frequently range from <100 per cell in meristematic cells to several thousand per cell in fully developed diploid leaf parenchyma cells. Microscopy studies have shown that the multicopy organelle genomes are usually condensed in more or less distinct DNA regions (nucleoids) within the organelle matrix or stroma.During development, the ratio of nuclear to organelle genomes appears to be relatively stringently regulated (Herrmann and Possingham, 1980; Rauwolf et al., 2010). Disregarding greatly varying absolute values (summarized in Rauwolf et al., 2010; Liere and Börner, 2013), there is little dispute that the number of plastid genomes and nucleoids per organelle and cell increase during early leaf development in higher plants (Kowallik and Herrmann, 1972; Selldén and Leech, 1981; Baumgartner et al., 1989; Fujie et al., 1994; Li et al., 2006; Rauwolf et al., 2010). This increase is usually accompanied by an increase in both size and number of plastids per cell (Butterfass, 1979). By contrast, data about plastid DNA (ptDNA) amounts in chloroplasts and cells of mature, ageing, and senescent tissue differ and are highly controversial. Basically two patterns have been described: the maintenance of more or less constant amounts of ptDNA per cell and/or organelle (Li et al., 2006; Zoschke et al., 2007; Rauwolf et al., 2010; Udy et al., 2012) or a significant decrease in copy number brought about by either continued organelle and cell division without ptDNA replication (Lamppa and Bendich, 1979; Scott and Possingham, 1980; Tymms et al., 1983) or by ptDNA degradation (Baumgartner et al., 1989; Sodmergen et al., 1991). In a series of communications, Bendich and coworkers recently reported that ptDNA levels decline drastically before leaf maturation in several plant species. In Arabidopsis thaliana and maize (Zea mays), ptDNA levels were reported to decrease early and precipitously as leaves mature. It was concluded that, in fully expanded leaves, most chloroplasts contain no or only insignificant amounts of DNA long before the onset of leaf senescence (Oldenburg and Bendich, 2004; Rowan et al., 2004; Oldenburg et al., 2006; Shaver et al., 2006; Rowan et al., 2009). Retention of ptDNA was proposed to be dispensable after the photosynthetic machinery was established in that the plastome-encoded photosynthesis genes were no longer needed in adult leaves. Degradation or even entire loss of ptDNA was considered as an event during plastid and leaf development, common to all plants (Rowan et al., 2009). ptDNA degradation was also suggested to act as a signal inducing senescence (Sodmergen et al., 1991).A priori, there is no reason why different ptDNA patterns should not occur, and there is indeed evidence that organelle DNA can behave differently in different materials, both quantitatively and structurally (e.g., Selldén and Leech, 1981; Baumgartner et al., 1989). However, since contradictory data were reported for the same species that were grown under comparable, if not identical, conditions (Rowan et al., 2004, 2009; Li et al., 2006; Oldenburg et al., 2006; Shaver et al., 2006; Zoschke et al., 2007; Evans et al., 2010; Udy et al., 2012), it is apparent that some of them must reflect methodological insufficiencies of the experimental approaches employed.From a physiological point of view, the existence of DNA-deficient plastids in photosynthetically competent tissue seems unlikely. For instance, due to its susceptibility to photooxidative damage, the D1 protein (PsbA), a plastome-encoded core subunit of photosystem II, must be replaced continuously by a complex repair system to maintain photosynthesis (Prasil et al., 1992). This replacement requires de novo synthesis of the short-lived D1. There are no data available supporting an extreme mRNA stability, protein stability, or for another compensating biochemistry, preserving organelle functions for weeks or even months. The maximum mRNA half-life reported for psbA is in the range of 40 h (Kim et al., 1993).Resolving this controversy is of considerable scientific interest, both from a theoretical and an applied perspective. We therefore analyzed the fate of ptDNA in mature, ageing, and senescent leaves of four commonly studied higher plant species (Arabidopsis, sugar beet [Beta vulgaris], tobacco [Nicotiana tabacum], and maize; Figure 1) for which conflicting data have been reported. Four complementary methods were used for assessing the presence of ptDNA as well as its quantitative and morphological changes during leaf development: an improved 4′,6-diamidino-2-phenylindole (DAPI)–based fluorescence microscopy approach including deconvolution of fluorescence images, electron microscopy, semithin sectioning across leaf laminas, and real-time quantitative PCR (see Methods).Open in a separate windowFigure 1.Developmental Leaf Series of Sugar Beet, Tobacco, and Arabidopsis.(A) Sugar beet leaves, developmental stages II to VI (left to right; see text). Inset: leaf stages y1 and y3. Arrows indicate necrotic areas. Bar = 5 cm.(B) Tobacco leaves, developmental stages II and IV to VI. Inset: leaf stages y1 and y2. Bar = 5 cm; bar in inset = 1 cm.(C) Arabidopsis plants (left) from which leaves of developmental stages I to VI were taken. Bar = 4 cm.Figure 2, Supplemental Methods, and Supplemental Data Sets 1 to 4 present representative micrographs of developmental series of DAPI-stained chloroplasts in leaf spongy parenchyma cells of late ontogenetic stages from sugar beet, Arabidopsis, tobacco, and maize displaying clearly discernible nucleoid patterns. Figures 1A to 1C document some of the leaves from which samples were taken. Mesophyll cells of juvenile leaves investigated in our previous work (Li et al., 2006; Zoschke et al., 2007; Rauwolf et al., 2010) were included for comparison (Supplemental Data Sets 1 to 4, panels 1 to 37, 84 to 94, 112 to 117, and 123 to 128). The staining specificity of the fluorochrome was confirmed enzymatically. Treatment with DNase, but not DNase-free RNase or Proteinase K, either before or after staining with the fluorochrome, abolished the fluorescence but did not significantly affect chloroplast structure (compare with Rauwolf et al., 2010; see Methods).Open in a separate windowFigure 2.DAPI-DNA Fluorescence of Mature, Senescent, and Prenecrotic Leaf Mesophyll Cells or Cell Segments.Representative DAPI-stained squashed mesophyll cells of sugar beet ([A] to [C]), Arabidopsis ([D] to [F]), tobacco ([G] and [H]), and maize ([I] and [J]) leaflets or leaves (cell detail in [C], [E], [F], and [H]) of the developmental stages III/IV (I), IV ([A] and [D]), V ([B], [E], and [G]), and VI ([C], [F], [H], and [J]). Note that (E) represents a cell fragment of Supplemental Data Set 2, panel 102. Bar = 5 μm in (A), also for (B) to (J).     

Viunalikeviruses are environmentally common agents of horizontal gene transfer in pathogens and biocontrol bacteria

Bacteriophages have been used as natural biocontrol and therapeutic agents, but also as biotechnological tools for bacterial engineering. We showed recently that the transducing bacteriophage ϕMAM1 is a ViI-like phage and a member of the new genus, ‘Viunalikevirus''. Here, we show that four additional ViI-like phages and three new environmentally isolated viunalikeviruses, all infecting plant and human pathogens, are very efficient generalised transducers capable of transducing chromosomal markers at frequencies of up to 10⁻⁴ transductants per plaque-forming unit. We also demonstrate the interstrain transduction of plasmids and chromosomal markers, including genes involved in anabolism, genes for virulence and genes encoding secondary metabolites involved in biocontrol. We propose that all viunalikeviruses are likely to perform efficient horizontal gene transfer. Viunalikeviruses therefore represent useful agents for functional genomics and bacterial engineering, and for chemical and synthetic biology studies, but could be viewed as inappropriate choices for phage therapy.Combined morphological, genomic and phylogenetic analyses have recently led to the proposed creation of a new phage genus, ‘Viunalikevirus'', within the Myoviridae family (Adriaenssens et al., 2012a). The first member of this proposed genus, Salmonella phage ViI, was isolated in the 1930s (Craigie and Yen, 1938) and multiple viunalikeviruses have been sequenced and characterised since 2010 (Pickard et al., 2010; Anany et al., 2011; Hooton et al., 2011; Kutter et al., 2011; Matilla and Salmond, 2012; Park et al., 2012; Adriaenssens et al., 2012a, 2012b; Hsu et al., 2013; Luna et al., 2013; Shahrbabak et al., 2013). Viunalikeviruses are characterised as virulent (lytic) phages showing similar genome size, extensive DNA homology, strong gene synteny and a complex adsorption apparatus, which uses tail spike proteins as host-recognition determinants (Adriaenssens et al., 2012a).We recently isolated the ViI-like phage, ϕMAM1, that infects several environmental and clinical isolates belonging to Serratia and Kluyvera genera (Matilla and Salmond, 2012). During the characterisation of ϕMAM1, we showed that it mediates highly efficient generalised transduction (Matilla and Salmond, submitted for publication). These observations were consistent with a previous report, that the Salmonella phage ViI was also capable of transduction (Cerquetti and Hooke, 1993) and we have confirmed that phage ViI can transduce chromosomal markers and plasmids at frequencies of up to 4.6 × 10⁻⁵ transductants per plaque-forming unit (p.f.u.; Figure 1a; Supplementary Table 1).Open in a separate windowFigure 1Transduction capabilities of viunalikeviruses. (a) Transduction frequencies of LIMEstone1, LIMEstone2, ViI and CBA120 phages. The graph also shows transduction efficiencies of LIMEstone phages within and between Dickeya solani strains. Transduction efficiency was defined as the number of transductants obtained per p.f.u. In all cases, error bars represent the standard deviations (n=3). (b) Skimmed milk agar plates showing protease production in the wild-type (wt) Dickeya solani strains MK10, MK16 and IPO 2222. LIMEstone1- (LS1) and LIMEstone2- (LS2) mediated transduction of the spp::Km marker from the protease negative mutant strain MK10P1 to the wild-type strains MK10, MK16 and IPO 2222 result in a protease-negative phenotype. (c–e) LIMEstone-mediated transduction of the oocN::Km marker from the oocydin A-negative mutant strain MK10oocN to the wild-type strains MK10 (c), MK16 (d) and IPO 2222 (e) results in an oocydin A-negative phenotype and, consequently, in the generation of strains defective in their antimicrobial activity against the plant pathogenic oomycete, Pythium ultimum. The anti-oomycete assays were performed as described previously (Matilla et al., 2012).Most generalised transducers utilise a headful packing strategy where phage terminases recognise specific sequences (pac sites) in the DNA and perform cycles of packing that result in mature phage particles (Fineran et al., 2009a). Indeed, phage terminases with reduced specificity for pac sequences may lead to the evolution of efficient transducing phages (Schmeiger, 1972). Based on the high similarity between the terminases of ϕMAM1, ViI and those of other previously sequenced viunalikeviruses, we hypothesised that all of these ViI-like phages should be capable of transduction in their respective bacterial hosts. To test this hypothesis, we investigated three additional viunalikeviruses, Escherichia coli phage CBA120 (Kutter et al., 2011), and Dickeya phages LIMEstone1 and LIMEstone2 (Adriaenssens et al., 2012b). All the bacteriophages, bacterial strains, plasmids and primers used in this study are listed in the Supplementary Tables 2 and 3. Experimental procedures are presented as Supplementary Material.The LIMEstone phages specifically infect some strains of the emerging plant pathogen, Dickeya solani (Adriaenssens et al., 2012b), and here we showed that they also infect the recently sequenced D. solani strains MK10, MK16 and IPO 2222. As predicted, we confirmed that the LIMEstone phages effected efficient transduction of various auxotrophic markers between Dickeya solani strains (Figure 1a; Supplementary Table 4). To our knowledge, only one Dickeya transducing phage, ϕEC2, has been isolated previously (Resibois et al., 1984). Additional mutant strains were constructed and the generalised nature of the transduction was confirmed by transfer of multiple chromosomal markers, including mutations in the gene cluster encoding biosynthesis of the anti-oomycete haterumalide, oocydin A (Matilla et al., 2012) and in the locus for synthesis and secretion of protease virulence factors. Transduction frequency was higher at an multiplicity of infection (m.o.i.) of 0.1 and 0.01 with efficiencies of up to 10⁻⁴ transductants per p.f.u. (Figure 1a; Supplementary Tables 4 and 5).We also demonstrated transduction of a kanamycin resistance-marked plasmid pECA1039-Km3 between strains MK10, MK16 and IPO 2222 at frequencies of up to 8.6 × 10⁻⁵ (Supplementary Table 4). Plasmid pECA1039 (originally isolated from the phytopathogen, Pectobacterium atrosepticum) encodes a bifunctional type III Toxin-Antitoxin (TA) system, ToxIN, with abortive infection capacity. Although ToxIN aborts infection of various enterobacteria by diverse phages (Fineran et al., 2009b) it did not protect against infection by the tested viunalikeviruses, ϕMAM1, ViI, CBA120, LIMEstone1 or LIMEstone2 (not shown). Furthermore, another type III TA system, TenpIN, from the insect pathogen, Photorhabdus luminescens (Blower et al., 2012), failed to protect against any of the five ViI-like phages (not shown).In addition, we also tested the transduction capacity of the E. coli phage, CBA120, and confirmed transduction of plasmid-borne antibiotic resistances at a frequency of up to 10⁻⁴ transductants per p.f.u. (Figure 1a; Supplementary Table 6).We decided to test our hypothesis that the viunalikeviruses may all be generalised transducers by first isolating new viunalikeviruses from the environment. From treated sewage effluent, we isolated three new bacteriophages infecting Dickeya solani, ϕXF1, ϕXF3 and ϕXF4, as defined initially by their very characteristic ViI-like morphology in electron microscopy (Figures 2a–c). As predicted, all of these new phages were able to transduce chromosomal markers and plasmids at frequencies of up to 3 × 10⁻⁶ transductants per p.f.u. (Figure 2e; Supplementary Table 7). Sequencing of structural and non-structural protein-encoding genes of ϕXF1, ϕXF3 and ϕXF4 showed high nucleotide homology (between 80% and 100%) with the corresponding orthologs in LIMEstone1 (Supplementary Figure 1), indicating that these virgin environmental isolates also clade within the Viunalikevirus genus.Open in a separate windowFigure 2Environmental isolation and characterisation of new viunalikeviruses with generalised transduction functionality. Transmission electron micrographs of phages ϕXF1 (a), ϕXF3 (b), ϕXF4 (c) and ϕXF28 (d) are shown. As an internal control, ϕXF28 was an example of a new lytic phage isolated from the same environment but showing no transduction capabilities. Bars, 50 nm. (e) Transduction frequencies of the new viunalikeviruses ϕXF1, ϕXF3 and ϕXF4. Transduction experiments were performed using 10⁹ cells with ϕXF1, ϕXF3, ϕXF4 at an m.o.i. of 0.01. Transduction efficiency was defined as the number of transductants obtained per p.f.u. Error bars represent the standard deviations (n=3).Although we did not have access to other ViI-like Salmonella phages SFP10 (Park et al., 2012), ϕSH19 (Hooton et al., 2011) and Marshall (Luna et al., 2013), Escherichia phage PhaxI (Shahrbabak et al., 2013), Shigella phage ϕSboM-AG3 (Anany et al., 2011) and Klebsiella phage 0507-KN2-1 (Hsu et al., 2013), our results allow us to predict that all of these phages will mediate generalised transduction. Importantly, these phages would be expected to contribute to the horizontal gene transfer of virulence factors and antimicrobial-resistance determinants in diverse environments.Viunalikeviruses do not seem to be limited to the enterobacteria as bacteriophages showing ViI-like morphology have been isolated in Acinetobacter (Ackermann et al., 1994), Bordetella (Adriaenssens et al., 2012b) and Sinorhizobium (Werquin et al., 1988). Furthermore, another ViI-like morphotype phage (ϕM12 of Sinorhizobium meliloti) has also been shown to be an efficient transducer (Finan et al., 1984). Taken together, these results suggest that, even in the absence of strongly predictive comparative genomic detail, a characteristically discrete ViI-like morphology in electron microscopy may be sufficient to identify new phages as strong candidates for possession of generalised transduction capacity.The emergence and dissemination of antibiotic-resistant pathogens coupled with low discovery rates for new antimicrobials, plus increasing legal constraints on the use of chemical pesticides, have (re)focussed attention on the potential use of bacteriophages for ‘natural biocontrol'' of human, animal and plant pathogens. Several viunalikeviruses have been proposed as candidate therapeutic agents for the control of bacterial infections (Anany et al., 2011; Hooton et al., 2011; Park et al., 2012; Hsu et al., 2013; Shahrbabak et al., 2013) and the LIMEstone phages have been used in successful field trials for biocontrol of D. solani infections (Adriaenssens et al., 2012b). However, their efficient transduction capacities could provide a route for dissemination of virulence factors, such as proteases (Marits et al., 1999). In fact, we have demonstrated the interstrain transduction of plasmids and oocydin A, auxotrophy and protease markers between three different D. solani strains, at high frequencies (Figures 1 and ​and2;2; Supplementary Tables 4 and 7). Also, the irregular distribution of the oocydin A gene cluster within the Dickeya genus and the fact that its genomic context varies between strains raises the possibility of phage-mediated horizontal gene transfer between bacterial strains. These results emphasize strongly that when considering the genomics of phages for ‘phage therapy'' the absence of genes readily defined as playing roles in lysogeny or bacterial virulence may be insufficient to inspire confidence that use of a particular therapeutic phage presents no risk–particularly among the high efficiency-transducing viunalikeviruses.     

Cell biology in development: The cell biology of planar cell polarity

Planar cell polarity (PCP) refers to the coordinated alignment of cell polarity across the tissue plane. Key to the establishment of PCP is asymmetric partitioning of cortical PCP components and intercellular communication to coordinate polarity between neighboring cells. Recent progress has been made toward understanding how protein transport, endocytosis, and intercellular interactions contribute to asymmetric PCP protein localization. Additionally, the functions of gradients and mechanical forces as global cues that bias PCP orientation are beginning to be elucidated. Together, these findings are shedding light on how global cues integrate with local cell interactions to organize cellular polarity at the tissue level.The collective alignment of cell polarity across the tissue plane is a phenomenon known as planar cell polarity (PCP). Exemplified by the uniform orientation of bristles covering the insect epidermis or of the hairs covering the mammalian body surface (Fig. 1 A), PCP patterns can align over thousands, even billions of cells. This phenomenon is controlled by the so-called PCP pathway, which integrates global directional cues to produce locally polarized cell behaviors. There has been a recent surge in interest in PCP after discoveries that various processes such as vertebrate gastrulation, mammalian ear patterning and hearing, and neural tube closure all require a conserved set of PCP genes (Heisenberg et al., 2000; Tada and Smith, 2000; Wallingford et al., 2000; Kibar et al., 2001; Murdoch et al., 2001; Curtin et al., 2003; Montcouquiol et al., 2003; Copley et al., 2013). Since that time, the PCP pathway has been found to coordinate cell behaviors in numerous diverse settings including polarized ciliary beating in the trachea and brain ventricles (Tissir et al., 2010; Vladar et al., 2012), oriented cell divisions (Gong et al., 2004; Baena-López et al., 2005; Ségalen et al., 2010; Mao et al., 2011), lung branching (Yates et al., 2010), and hair follicle alignment (Guo et al., 2004; Devenport and Fuchs, 2008), to name a few (Fig. 1). Genetic disruptions to PCP cause severe developmental abnormalities in vertebrates, notably neural tube defects, left/right patterning defects, and ciliopathies, which highlights the essential requirement for PCP in development (Kibar et al., 2001; Murdoch et al., 2001; Curtin et al., 2003; Wang et al., 2006a,b; Kim et al., 2010; Song et al., 2010).Open in a separate windowFigure 1.Planar cell polarity and the core PCP components. (A and B) The Drosophila wing blade and mammalian epidermis illustrate the phenomenon of PCP. In both cases, hairs point in a single direction along the tissue axis, where they align locally with neighboring hairs and globally across the tissue. Whereas Drosophila wing hairs are produced by single cells, mammalian hairs emerge from multicellular hair follicles, which orient as a unit. A conserved PCP pathway controls the collective alignment of both types of structures. (C) Core PCP components localize to the plasma membrane and asymmetrically segregate along the epithelial plane as indicated.Like many types of cell polarity, the establishment of PCP involves (1) a global orienting cue, (2) asymmetric segregation of dedicated polarity proteins, and (3) translation of polarity information into polarized outputs. But unlike other types of cell polarity, the PCP mechanisms we currently understand involve coupling between adjacent cells, allowing for the alignment of polarity over many cell distances.First described in insects and then genetically dissected in Drosophila melanogaster, PCP was long confined to the realm of experimental embryology and genetics until the discovery that the protein products of several PCP genes were localized asymmetrically within the cell, thrusting PCP into the domain of cell biology (for review see Strutt and Strutt, 2009). The challenge to understanding PCP on a molecular level is that long-range PCP is, in essence, an in vivo phenomenon that is difficult to recapitulate in a tissue culture dish. However, recent advances in imaging technology combined with increasingly sophisticated genetic tools are helping us to decipher the in vivo cell biology of PCP. In this review, I highlight some of the recent advances made toward understanding the cell biology underlying the establishment of coordinated polarized cell behaviors. For clarity, I limit discussions of PCP phenomena that meet the definition of PCP proposed by Goodrich and Strutt (2011): namely, that “cell–cell communication causes two or more cells to adopt coordinated polarity” in a process that is mechanistically “dependent upon planar polarity proteins.” Other aligned cellular patterns or examples of noncanonical Wnt signaling, sometimes described as “Wnt/PCP” signaling, will not be discussed.

PCP components and molecular asymmetries

Two molecular systems control PCP behavior, the “core” and the “Fat–Dachsous (Ft–Ds)” PCP pathways. A key feature of both is the asymmetric distribution of their constituents (Fig. 2). The core PCP pathway is composed of the multipass transmembrane proteins Frizzled (Fz), Van Gogh (Vang; also known as Strabismus/Stbm), and Flamingo (Fmi; also known as Starry night/Stan), and the cytosolic components Dishevelled (Dsh), Prickle (Pk), and Diego (Dgo). On one edge of the cell reside Fz, Dsh, and Dgo, and on the opposite side lie Vang and Pk (Figs. 1 C and 2 B; Axelrod, 2001; Strutt, 2001; Feiguin et al., 2001; Tree et al., 2002; Bastock et al., 2003). The atypical cadherin, Fmi, resides on both sides, where it forms homodimers between neighboring cells (Usui et al., 1999; Shimada et al., 2001). These molecular asymmetries are observed in sensory hair cells of the vertebrate inner ear (Wang et al., 2005, 2006a,b; Montcouquiol et al., 2006; Deans et al., 2007; Song et al., 2010), the mammalian epidermis (Devenport and Fuchs, 2008; Devenport et al., 2011), brain ventricles (Tissir et al., 2010), and trachea (Vladar et al., 2012). Mutations in core PCP components lead to a loss or randomization of polarity and misalignment of cellular structures along the tissues axis.Open in a separate windowFigure 2.Asymmetric localization of PCP components. (A) PCP asymmetry develops progressively from an initially uniform distribution of core PCP proteins. Fz, Dsh, and Dgo (red) localize to the distal/posterior edge, whereas Vang and Pk (turquoise) localize to the proximal/anterior side. Fmi (dark blue) localizes to both sides, where it forms homodimers between neighboring cells. (B) Feedback interactions between core PCP components. A Fz–Fmi complex interacts preferentially with a Vang–Fmi complex between cells, whereas proximal and distal complexes antagonize one another within the cell. (C) Ds and Fj are expressed in opposing gradients in the Drosophila wing blade. Fj positively modulates Ft activity, leading to a gradient of Ft activity across the wing (not depicted). (D) Graded expression of Ds and Fj leads to asymmetric cellular localization of Ds and Ft, which form heterodimers between adjacent cells. Dachs, a downstream component of the Ft–Ds pathway, also localizes asymmetrically in association with Ds.The Ft–Ds pathway includes the large protocadherins Ft and Ds and the Golgi resident transmembrane kinase, Four-jointed (Fj; for review see Matis and Axelrod, 2013; Thomas and Strutt, 2012). Similar to the core system, Ft–Ds also displays molecular asymmetries in flies. Ds and its ligand Ft accumulate on opposite cell edges, where they form intercellular heterophilic interactions (Fig. 2 D; Matakatsu and Blair 2004; Ambegaonkar et al., 2012; Brittle et al., 2012). Unlike the core components, Ds and Fj are expressed in complementary gradients in the Drosophila eye and developing wing, which contribute to the cellular asymmetries of Ds and Ft (Fig. 2, C and D). Whether Ft–Ds–Fj gradients and asymmetries are conserved in vertebrate systems has yet to be determined.

Segregation of cortical polarity proteins: Shaking hands with the enemy

The asymmetric segregation of Fz–Dsh–Fmi and Vang–Pk–Fmi complexes to opposite sides of the cell relies on their mutual exclusion intracellularly and their preferential binding between neighboring cells (Fig. 2 B; for review see Strutt and Strutt, 2009). There is mutual interdependence among core PCP components for their asymmetric localization. Depletion of any one core PCP component results in a loss of asymmetry of all the others. In addition, PCP asymmetry develops progressively from initially uniform distributions (Fig. 2 A). Thus, PCP asymmetry can be thought of not as a simple hierarchy of interactions, but the result of feedback amplification of an initial directional bias.

Intercellular interactions.

PCP requires cell–cell communication, mediated by the transmembrane components of the core system, where it is thought that Fz–Fmi on one cell interacts with Vang–Fmi on its neighbor. These interactions are best understood in the Drosophila wing blade, where PCP controls the alignment of wing hairs along the proximal–distal axis (Figs. 1 A and 2, A and B). In the wing, Vang–Pk localize to the proximal face of each cell, whereas Fz–Dsh–Dgo localize distally adjacent to the wing hair (Figs. 1 C and 2, A and B; Axelrod, 2001; Strutt, 2001; Tree et al., 2002; Bastock et al., 2003; Das et al., 2004). By generating mutant clones and examining PCP localization at the clone border, the intercellular interactions between neighboring cells can be assessed in vivo. For example, when Fz is lacking within a clone, leaving only Vang–Fmi available at cell junctions, then Fz–Fmi in adjacent wild-type cells is recruited to the clone border (Chen et al., 2008). Vang mutant clones produce a similar effect, but in this case the excess Fz recruits Vang to clone borders (Bastock et al., 2003). What mediates these intercellular asymmetric interactions? One possibility is that Vang and Fz interact directly, and in vitro binding assays between the Fz extracellular domain and Vang suggest that this mechanism is possible (Wu and Mlodzik, 2008). However, mutants of Fz or Vang lacking their extracellular domains can still recruit one another between cells, which suggests that something else must bridge the two proteins (Chen et al., 2008). The seven-pass transmembrane cadherin, Fmi, likely performs this function. Fmi is essential for the junctional recruitment of Fz and Vang, and Fmi homodimers appear to be functionally asymmetric (Chen et al., 2008; Strutt and Strutt, 2008; Struhl et al., 2012). Clonal overexpression of Fmi preferentially recruits Fz to the clone border, even in the absence of Vang, which suggests that excess or unpaired Fmi is in a configuration that has higher affinity for Fmi–Fz than Fmi–Vang (Chen et al., 2008; Strutt and Strutt, 2008). Thus, Fmi may exist in two forms depending on whether it is paired with Fz or Vang, but the molecular basis for this difference is not known (Chen et al., 2008; Strutt and Strutt, 2008; Struhl et al., 2012).

Amplification of asymmetry.

Intercellular Fz–Fmi and Vang–Fmi complexes can form between cells in any orientation, so how do they resolve into discrete and opposed asymmetric domains? One way is through clustering of Fz–Fmi and Vang–Fmi complexes of the same orientation, and the cytoplasmic PCP components are particularly important for this function. As PCP complexes grow increasingly asymmetric, they cluster into discrete puncta that are stably associated with the plasma membrane and are resistant to endocytosis (Strutt et al., 2011). FRAP analysis of Fz-containing puncta demonstrated that they are highly stable compared with diffuse Fz-GFP, and have limited lateral mobility within the membrane. In the absence of Dsh, Pk, or Dgo, the size, intensity, and stability of Fz-containing puncta are diminished (Strutt et al., 2011), whereas overexpression causes Fz accumulation and coalescence into larger puncta (Feiguin et al., 2001; Tree et al., 2002; Bastock et al., 2003). Although the precise mechanisms driving PCP puncta formation are not known, the cytoplasmic components do not affect endocytosis, which suggests that they contribute to puncta formation by clustering intercellular complexes (Strutt et al., 2011). Pk can interact homophilically (Jenny et al., 2003; Ayukawa et al., 2014), which might promote clustering of proximal Vang–Pk–Fmi complexes. It will also be interesting to determine whether the cytoskeleton is directly connected to PCP complexes to minimize lateral mobility within the membrane.A second mechanism contributing to PCP asymmetry is directed transport. Live imaging of fluorescently tagged PCP proteins in pupal wings showed that Fz- and Dsh-containing particles travel across the cell in a proximal-to-distal direction (Shimada et al., 2006; Matis et al., 2014; Olofsson et al., 2014). These particles most likely represent endosomes undergoing transcytosis, as they arise from the proximal cortex and are labeled by the endocytic tracer FM4-64. This mechanism could serve to amplify asymmetry or even provide the initial polarity bias by removing proximal Fz–Dsh–Fmi complexes and transporting them to the distal side. Directed PCP transport is mediated by an array of subapical, noncentrosomal microtubules (MTs) that align along the proximal–distal axis, with the plus ends oriented with a slight distal bias (Hannus et al., 2002; Shimada et al., 2006; Harumoto et al., 2010; Matis et al., 2014; Olofsson et al., 2014). Ft and Ds are required for proximal–distal MT alignment (Harumoto et al., 2010), which suggests that the Ft-Ds system may feed into the core PCP system by orienting cytoskeletal architecture to deliver Fz–Dsh–Fmi complexes to the distal edge of the cell.Directed transport of Vang-containing endosomes has not been reported in flies, but selective trafficking could target Vang to specific membrane domains. In mammalian cells, exit of the Vang homologue Vangl2 from the trans-Golgi network (TGN) requires Arfrp1 (an Arf-like GTPase) and the clathrin adaptor complex AP-1, neither of which are required for the transport of a mammalian Fz homologue Fz6 or Fmi/Celsr1, which suggests that the differential sorting of PCP complexes to opposite sides of the cell could initiate at TGN export (Guo et al., 2013). Whether newly synthesized Vang and Fz proteins are transported to opposing cell surfaces from the TGN has not yet been explored.Microtubule orientation also correlates with PCP asymmetry in mouse trachea epithelial cells, where PCP coordinates the alignment of motile cilia (Vladar et al., 2012). MTs are planar polarized with their plus ends oriented toward the Fz–Dvl domain, and disruption of MTs with nocodazole impairs core PCP localization. Similarly, MTs are needed to establish Pk asymmetry in gastrulating zebrafish embryos (Sepich et al., 2011). However, in the skin epithelium, MTs align perpendicular to the axis of PCP asymmetry (unpublished data). Thus, directed transport along MTs may not be required in all tissue types for the establishment of PCP asymmetry.

Negative regulation.

Repulsive interactions between Vang- and Fz-containing complexes may also contribute to the amplification of asymmetry, and cytoplasmic proteins have been proposed to perform this function. Pk and Dgo both bind to Dsh in vitro, interacting with the same domain on Dsh in a mutually exclusive manner (Jenny et al., 2005). In addition, overexpression of Pk can prevent Dsh translocation to the membrane (Tree et al., 2002; Carreira-Barbosa et al., 2003), which suggests that Pk binding to Dsh could displace it from the proximal side of the cell. On the distal side, Dgo binding to Dsh would prevent association with Pk, thus enhancing Dsh distal localization. This increase in Dsh and Pk asymmetry would then positively feed back by clustering the transmembrane components into stable membrane domains.Modulation of PCP protein levels by ubiquitin-mediated degradation also leads to feedback by restricting the amount of one PCP protein to antagonize another. In flies, regulation of Dsh by a Cullin-3-BTB E3 ubiquitin ligase complex limits its levels at cell junctions (Strutt et al., 2013a). Reduction of Cullin-3 leads to an increase in overall core PCP protein levels, a reduction of asymmetry, and defects in wing hair polarity, which is consistent with Dsh overexpression phenotypes (Strutt et al., 2013a). SkpA, a subunit of the SCF E3 ligase, regulates Pk levels by promoting its degradation in a Vang-dependent manner (Strutt et al., 2013b). In mice, Smurf E3 ligases ubiquitinate Pk and promote its local degradation by binding to phosphorylated Dvl2 (a mammalian homologue of Dsh; Narimatsu et al., 2009). Smurfs are required for Pk localization in the inner ear and floor plate, and their removal leads to defects in convergent extension (CE) and stereocilia alignment (Narimatsu et al., 2009). Thus, targeting Pk for degradation either balances total Pk protein levels or targets a specific pool of Pk for ubiquitination and proteasome degradation.

Tissue-level polarity cues: This way or that?

What provides the tissue-level polarity cue that biases core PCP asymmetry in one direction over another? This is perhaps the most fundamental, yet poorly understood, element of PCP. Current models propose that an upstream, graded cue provides an initial bias in PCP asymmetry by regulating the levels, localization, or activity of one or more of the core proteins. Gradients are attractive candidates for providing global polarity cues, as they can act across many cells and define the tissue boundaries over which polarity must be oriented.

Ft–Ds–Fj.

Unlike the core proteins, Ds and Fj are nonuniformly expressed in the Drosophila eye, wing, and abdominal segments, and as such, the Ft–Ds module has been proposed to provide a global polarity cue (Fig. 2, C and D; for review see Ma et al., 2003; Yang et al., 2002; Thomas and Strutt, 2012; Matis and Axelrod, 2013). Ft and Ds are heterodimeric cadherins, regulated by the Golgi kinase Fj (Ishikawa et al., 2008; Brittle et al., 2010; Simon et al., 2010). The complementary expression patterns of Ds and Fj are thought to give rise to asymmetric Ft and Ds protein localization, with Ft and Ds localizing to opposite sides of each cell (Fig. 2 D; Ambegaonkar et al., 2012; Bosveld et al., 2012; Brittle et al., 2012). Because Fj positively regulates the activity of Ft, a gradient of Ft activity is expressed across the wing complementary to that of its ligand, Ds (Simon et al., 2010). Mutations in the Ft–Ds system give rise to swirling wing hair patterns, and disrupt the global alignment of core PCP proteins, but not their asymmetric distributions.An appealing model for symmetry breaking in the early Drosophila wing is that cellular asymmetries of Ft–Ds polarize MT organization and promote the distal transport of Fz–Dsh–Fmi vesicles (Shimada et al., 2001, Harumoto et al., 2010; Matis and Axelrod, 2013; Matis et al., 2014). This would produce an initial bias in Fz–Dsh localization, which would then be amplified by feedback interactions. However, several pieces of evidence have prevented the model from gaining universal acceptance. First, Ds and Fj gradients are oriented in opposite directions with respect to the core PCP proteins in the wing compared with the eye and abdomen. This discrepancy has been rectified with the finding by two independent groups that cells interpret Ft–Ds–Fj gradients differently depending on which of two Pk isoforms is expressed (Ayukawa et al., 2014; Olofsson et al., 2014). Second, the Ft–Ds system can orient PCP independently of the core pathway, and thus the two systems orient polarity in parallel, as opposed to in a single, common pathway (Casal et al., 2006). Third, Ft–Ds mutations affect core PCP orientation only regionally in the wing, which suggests that, if Ft–Ds provides a global bias, other, redundant cues must also exist (Matakatsu and Blair, 2006; Matis et. al., 2014). Finally, the direction of Ft–Ds and core PCP asymmetry diverges late in wing development, where the two systems become completely uncoupled. Intriguingly, the extent of coupling depends on which isoform of Pk is expressed (Merkel et al., 2014). Perhaps the simplest explanation for Ft–Ds function is that it can both transmit polarity information independent of the core system and organize the cytoskeleton to provide an initial bias of core PCP asymmetry, but which mechanism predominates depends on the tissue and developmental stage.

Wnts.

Wnt proteins have long been considered attractive candidates to provide tissue-level polarity cues because Fz and Dsh are primary components of the Wnt–β-catenin signaling pathway. Wnts are secreted glycoproteins that bind to Fz and other receptors, and often display graded expression. In vertebrates, Wnts are clearly important regulators of PCP, but whether they act instructively or permissively is unclear. In zebrafish, Wnt5a and Wnt11 are required for CE movements during gastrulation, but uniform expression of Wnt11 rescues the mutant phenotype, which suggests that it is permissive rather than instructive (Heisenberg et al., 2000; Kilian et al., 2003). Wnt5a is expressed in a gradient along the axis of polarity in the mouse inner ear, where it interacts genetically with Vangl2 in cochlear hair cell orientation (Qian et al., 2007). In the mouse limb, Wnt5a and its atypical receptor Ror2 are required for limb elongation and the asymmetric localization of Vangl2 at the proximal face of converging and extending chondrocytes (Gao et al., 2011). Wnt5a is expressed in a distal-to-proximal gradient, which induces a gradient of Vangl2 phosphorylation. The functional consequences of Vangl2 phosphorylation are unknown but Vangl2 cellular asymmetry appears to be strongest distally, where Wnt5a and Vangl2 phosphorylation levels are highest (Gao et al., 2011).While several studies had argued against the involvement of Wnt proteins in Drosophila PCP (Lawrence et al., 2002; Chen et al., 2008), it was recently discovered that Wingless (Wg) and Wnt4a act redundantly to orient PCP in the wing, particularly near the wing margin (Wu et al., 2013). Misexpression of Wg or Wnt4a reorients wing hair polarity in a pattern reminiscent of Fz loss of function, which suggests that Wnt gradients may orient polarity by antagonizing Fz. Consistently, the ability of Fz and Vang to recruit one another between adjacent cells in culture was inhibited by the addition of Wg or Wnt4a, which suggests that Wnts could provide a polarizing cue by diminishing Fz–Vang interactions at the margin of the wing, where Wnt expression is highest (Wu et al., 2013). However, Wnt4a overexpression also reorients MT alignment, suggesting that Wnts may act as polarity cues thorough an effect on the cytoskeleton (Matis et. al., 2014). Alternatively, Sagner et al. (2012) suggest that Wg orients core PCP indirectly through its effects on wing patterning and growth. Although the evidence for Wnt gradients as global PCP cues is accumulating, the mechanisms by which they regulate core protein levels or activity remain to be elucidated.

Mechanical forces.

Anisotropic mechanical forces that accompany growth and morphogenesis can also provide global polarizing cues. During wing development, PCP reorients in response to extensive morphogenetic changes that elongate the wing along the proximal–distal axis. In early pupal wings, PCP aligns toward the wing margin and then reorients during wing elongation and contraction of the wing hinge (Aigouy et al., 2010). These morphogenetic changes have broad effects on cell behavior, inducing cell elongation, oriented divisions, and cell rearrangements with a concomitant reorientation of PCP. Severing the wing pouch from the hinge blocks cell flows and PCP reorientation, which suggests that the anisotropic tension from hinge contraction drives tissue flow and the reorientation of polarity (Aigouy et al., 2010). Although this model doesn’t explain what initially biases PCP, it does demonstrate how the morphogenetic processes that shape tissues can completely remodel global PCP alignment. This is an attractive model to explain how PCP aligns over very large tissues, like the mammalian skin, where hairs consistently reorient along regions of extensive tissue elongation such as the face, limbs, and ears.

Downstream effectors of PCP: Steering the wheel

If PCP is the cell’s compass, it is also the steering wheel, directing downstream, polarized cell behaviors in response to global directional cues. PCP can polarize a wide range of cell behaviors, which suggests that it can intersect with numerous downstream effectors. We focus here on three examples where the molecular mechanisms linking core PCP to their polarized outputs have recently been elucidated.

Distal positioning of wing hairs.

Each cell of the Drosophila wing blade emits a single actin-rich protrusion from its distal edge. The placement of the wing hair strongly correlates with the position of Fz–Dsh–Fmi, which suggests that core proteins may localize cytoskeletal regulators to distinct positions within the cell (Strutt and Warrington, 2008). On the proximal side, Vang recruits a group of proteins that negatively regulate actin prehair formation: Inturned, Fuzzy, and Fritz (Adler et al., 2004; Strutt and Warrington, 2008). These three proteins regulate Multiple Wing Hairs, a GTPase-binding/formin-homology 3 (GBD/FH3) domain protein thought to repress actin polymerization (Strutt and Warrington, 2008; Yan et al., 2008). This restricts actin nucleation to distal positions within the cell, and in the absence of Multiple Wing Hairs, ectopic actin bundles form across the apical surface (Wong and Adler, 1993). On the distal side, casein kinase 1 γ CK1g/gilamesh is required to further refine prehair nucleation to a single site through a parallel mechanism involving Rab11-dependent vesicle traffic to the site of prehair formation (Gault et al., 2012). Rho and Rho kinase (Drok) have also been implicated in wing hair formation, but their roles are difficult to dissect due to the numerous functions of Rho in cell shape and cell division (Winter et al., 2001; Yan et al., 2009).

Actomyosin contraction and convergent extension (CE).

CE was the first vertebrate process to be linked molecularly to PCP (Wallingford et al., 2000). During CE, mesenchymal cells elongate, form mediolateral-directed protrusions, and intercalate mediolaterally, narrowing the mediolateral axis while simultaneously lengthening the anterior–posterior (A-P) axis (Fig. 3 A; Keller, 2002). Mediolateral polarization, elongation, and intercalation are lost when core PCP components are disrupted, leading to a failure in CE (Tada and Smith, 2000; Wallingford et al., 2000; Goto and Keller, 2002; Jessen et al., 2002). While several PCP-dependent mechanisms have been proposed to mediate CE movements, two recent studies provide direct mechanistic links between asymmetrically localized core PCP components and CE behaviors. In neuroepithelial cells, PCP specifies the localization of myosin to the A-P faces of intercalating cells. Fmi/Celsr1 and Dvl recruit the formin DAAM1 to the A-P junction, which in turn binds and activates PDZ-RhoGEF. This likely activates RhoA and myosin contractility specifically at A-P junctions, resulting in medial-directed cell intercalation and neural plate bending (Nishimura et al., 2012). A similar mechanism was found to drive CE movements of mesenchymal cells during Xenopus gastrulation. In this case, Fritz and Dsh help to localize septins to mediolateral vertices, where they spatially restrict cortical actomyosin contractility and junctional shrinking to A-P cell edges, thus driving cell intercalation (Fig. 3 A; Kim et al., 2010; Shindo and Wallingford, 2014). Together these studies show how asymmetric PCP localization produces collectively polarized cell behaviors through spatial modulation of the cytoskeleton.Open in a separate windowFigure 3.Polarized cell behaviors controlled by PCP. (A) PCP drives convergent extension (CE). CE in vertebrates is driven by mediolateral intercalation, which narrows the mediolateral axis while simultaneously lengthening the A-P axis. Mediolateral intercalation is accompanied by cell polarization and elongation and the formation of mediolateral protrusions, all of which require core PCP function. Pk localizes anteriorly (Ciruna et. al., 2006; Yin et al., 2008), whereas Dsh localizes posteriorly (Yin et. al., 2008). In addition, PCP proteins recruit myosin to A-P cell borders, leading to actomyosin contractility and junctional shrinking. (B) Asymmetric cell division. Drosophila sensory organ precursors (SOPs) divide asymmetrically along the epithelial plane, giving rise to distinct anterior and posterior daughters. Spindle alignment along the A-P axis is PCP dependent. Dsh interacts with Mud/NuMA and the dynein complex posteriorly while Vang links Pins/LGN-Mud/NuMA-dynein on the anterior. This links astral MTs to the A-P cortex, bringing the spindle into register with the A-P axis. (C) Positioning of the kinocilium in the inner ear. The placement of kinocilium in sensory hair cells of the inner ear determines the position of V-shaped stereocilia bundles. Gαi and mPins/LGN localize on the abneural side on the hair cell, where they are required for abneural positioning the MT-based kinocilium. The collective alignment of kinocilia and stereocilia bundles across the epithelium requires the core PCP component Vangl2. Vangl2 (light green) localizes to the abneural side of supporting cells. Whether Fz (dark blue) associates on the opposite face is not yet clear (Ezan et. al., 2013).

Positioning of centrosomes and cilia.

PCP regulates the positioning of MT-based structures including the mitotic spindle and cilia. In Drosophila sensory organ precursors and early zebrafish embryos, PCP controls mitotic spindle orientation along the epithelial plane by interacting with the highly conserved spindle orientation complex, which links astral MTs to the cell cortex through Mud/NuMA-mediated recruitment of the dynein complex (Ségalen et al., 2010). To orient the spindle, posteriorly localized Dsh binds to Mud/NuMA, which recruits the dynein complex and astral MTs to the posterior cortex. On the anterior side, Pins/LGN recruits Mud/NuMA, bringing the spindle into A-P alignment (Fig. 3 B). Similarly, PCP was recently shown to interact with the spindle orientation machinery to position the kinocilium in nondividing cells of the inner ear (Ezan et al., 2013; Tarchini et al., 2013). In vestibular hair cells, Gαi and mPins/LGN localize to the abneural cortex, opposite Vangl2, where they are required for kinocilium positioning and subsequent alignment of stereocilia bundles (Fig. 3 C; Ezan et al., 2013). MT plus ends and dynein also show an abneural bias suggesting that Gαi-mPins/LGN induces pulling on MTs by a similar mechanism that orients the centrosome during spindle orientation. Vangl2 is required for the alignment of Gαi-Pins/LGN crescents between cells, coordinating kinocilia positioning and stereocilia polarity across the tissue (Fig. 3 C; Ezan et al., 2013). Thus the PCP pathway co-opts the spindle orientation machinery to specify not only the division plane but also cilia position in nondividing cells. As PCP is required for asymmetric cilia positioning in a wide range of cell types, including the node (Antic et al., 2010; Borovina et. al., 2010; Song et al., 2010; Hashimoto et. al., 2010), it will be interesting to determine whether this mechanism is conserved.

Concluding remarks

PCP is a fundamental and highly conserved process coordinating a vast number of polarized cell behaviors. While the number of functions ascribed to PCP continues to grow, an understanding of the mechanisms establishing PCP is still far from complete. The development of cellular asymmetry from uniform distributions is not well understood, and will benefit from recent advances in high-resolution, time-lapse imaging with photoconvertible fluorescent tags. Other important issues to resolve include deciphering the structural domains and biochemical interactions mediating intercellular communication, identifying the global cues that orient PCP especially in vertebrates, and deciphering the mechanisms by which complex multicellular structures, like lung branches and hair follicles, are oriented by the PCP machinery.     

Beyond the cell: The cell biology of renal filtration

The function of the kidney, filtering blood and concentrating metabolic waste into urine, takes place in an intricate and functionally elegant structure called the renal glomerulus. Normal glomerular function retains circulating cells and valuable macromolecular components of plasma in blood, resulting in urine with just trace amounts of proteins. Endothelial cells of glomerular capillaries, the podocytes wrapped around them, and the fused extracellular matrix these cells form altogether comprise the glomerular filtration barrier, a dynamic and highly selective filter that sieves on the basis of molecular size and electrical charge. Current understanding of the structural organization and the cellular and molecular basis of renal filtration draws from studies of human glomerular diseases and animal models of glomerular dysfunction.The mammalian kidney orchestrates the excretion of metabolic wastes found in blood, a function intimately related to its essential roles in general fluid homeostasis and osmoregulation. It is also important in the control of blood pressure, synthesis of vitamin D, bone mineralization, and the promotion of erythrocyte development. Despite its modest size (each is approximately the size of a human fist), a mammalian kidney is highly vascularized. A pair of kidneys receives and filters a remarkable volume of blood, estimated to be the equivalent of roughly 20% of total cardiac output (Stein and Fadem, 1978; Munger et al., 2011). In humans, blood filtration by the kidneys generates on average 1 liter of urine per day. Urine is produced and concentrated along the length of nephrons, the basic unit of kidneys (Fig. 1 A). An adult human kidney is known to contain an average of 1 million and up to as many as 2.5 million nephrons (Puelles et al., 2011).Open in a separate windowFigure 1.Anatomical overview of renal filtration. (A) Diagrammatic representation of nephron distribution in the kidney. Glomeruli, the filtration compartments of nephrons, are found within the kidney cortex. (B) Segmental structure of nephrons. The vascularized glomerulus is found at the proximal end and is connected through a series of renal tubules where urinary filtrate composition is refined through resorption and secretion. (C) Cellular organization of the glomeruli. GEC, glomerular endothelial cell; AA, afferent arteriole; EA, efferent arteriole; Pod, podocyte; MC, mesangial cell; PEC, parietal epithelial cell; PT, proximal tubule; DT, distal tubule; LOH, loop of Henle; CD, collecting duct; BS, Bowman’s space.A nephron is functionally subdivided into a filtration unit called the renal corpuscle or glomerulus and a segmented tubular resorption compartment (Fig. 1 B). The glomerulus is an assembly of four different cells: the glomerular endothelial cells (GECs), podocytes, mesangial cells (MCs), and parietal epithelial cells (PECs; Figs. 1 C and ​and2).2). The word glomerulus is a reference to its intricately tortuous inner capillary tuft formed by GECs, after the Latin word glomus for a ball of yarn (Fig. 2 C). Podocytes are specialized perivascular cells aptly named for their elaborate projections, known as foot processes (FPs) or pedicels, that are intimately wrapped around the exterior of glomerular capillaries (Fig. 2, A and B). GECs and podocytes share a common ECM known as the glomerular basement membrane (GBM). The GECs, the podocytes, and the GBM in between constitute the three distinctive layers of the glomerular filtration barrier (GFB; Fig. 2 E), an elegant sieve that selectively filters blood components, generating a dilute primary urinary filtrate. The mesangium, a stalk-like aggregate of MCs and their ECM called the mesangial matrix, provides the structural reinforcement for the glomerular vasculature. The PECs forms a watertight cuplike enclosure called the Bowman’s capsule. Primary urinary filtrate collects within the Bowman’s capsule and empties through a connected series of epithelial tubules starting from the proximal tubules, the loop of Henle, the distal tubules, and a final collecting duct. The renal tubules of the nephrons and the collecting ducts express various ion and water channels, as well as transporters that help concentrate and adjust the composition of the urinary filtrate by resorption and secretion. This last step is vital for fluid conservation, maintenance of electrolyte balance, and resorption of glucose.Open in a separate windowFigure 2.An ultrastructural overview of podocytes and the glomerular endothelium. (A) Scanning electron micrograph of an exposed glomerulus. In this image, the Bowman’s capsule is broken, permitting a striking view of podocytes (Pod) completely wrapped around the glomerular capillaries. (B) Higher magnification of a podocyte within the glomerulus revealing the interdigitated FPs. (C) A resin cast of the glomerular capillary tuft with the cells corroded to reveal its highly convoluted shape. Image courtesy of F. Hossler (East Tennessee State University, Johnson City, TN). (D) Scanning electron micrograph of an exposed glomerular capillary and its numerous perforations (fenestrae). (E) Simplified diagram of the GFB. The GEC and its fenestrae are lined by a filamentous glycocalyx enriched in negatively charged proteoglycans. The glycocalyx and adsorbed plasma components form the thicker ESL. The GBM is a stratified ECM in between podocytes and GECs. Podocytes form the final layer of the GFB. The interdigitating FPs of podocytes are linked by porous SDs where primary urinary filtrate passes through. Bars: (A) 20 µm; (B and D) 1 µm; (C) 50 µm.

Selective permeability in renal filtration

Water and small solutes (e.g., urea, glucose, amino acids, mineral ions) in blood plasma freely traverse the GFB while circulating cells such as erythrocytes and high-molecular-weight plasma components such as albumin are selectively retained in blood. Intriguingly, the glomerular permeability of proteins, particularly negatively charged proteins such as albumin, is well exceeded by those of neutral dextrans of comparable or even larger sizes (Chang et al., 1975). Additionally, the GFB strongly restricts passage of anionic macromolecules (Thomson and Blantz, 2010). Size and charge selectivity thus makes the GFB a formidable barrier for the bulk of plasma proteins and results in a urinary product that is virtually protein-free. While the role of tubular reuptake of proteins leaked into urine is well recognized, recent intravital imaging studies with fluorescent albumin conjugates validate the predominant role of the GFB in ensuring minimal loss of albumin in urine (Peti-Peterdi and Sipos, 2010).The diagnostic hallmark of a compromised GFB is the incidence of protein in urine, a condition called proteinuria, or more specifically albuminuria if measured in terms of urinary albumin content. Proteinuria manifests in a host of ailments ranging from congenital nephropathy, hypertension, and diabetes to chronic kidney diseases. Over the last two decades, multidisciplinary studies combining genetics, cell biology, physiology, and signal transduction analysis including extensive studies on proteinuric disease models have provided us with valuable insights regarding the physiological importance of each of the three distinctive layers of the GFB and the intricacies of how they may function as an integral unit. In this review, we summarize our current understanding of the cell and molecular basis of renal filtration, highlighting the development, organization, and properties of each compartment of the GFB, and how they contribute to selective permeability.

Fenestrated capillaries as primary portals of renal filtration

The glomerular vasculature consists of afferent and efferent arterioles and the glomerular capillary tuft (Fig. 1 C). Blood enters and exits the glomerulus via the afferent and efferent arterioles, respectively. Inside the glomerulus, the afferent arteriole immediately branches into the elaborate glomerular capillary tuft, a specialized region where blood filters through. Unlike the afferent and efferent arterioles, the glomerular capillaries are heavily perforated with transcellular pores and are not surrounded by smooth muscles. These glomerular capillary pores, known as fenestrae (the plural of fenestra, which means window in Latin), are 60–100 nm wide and comprise ∼20% of the endothelial surface, making glomerular capillaries efficient portals for the rapid passage of high volumes of fluid characteristic of renal filtration (Fig. 2 D; Levick and Smaje, 1987).The idea that the glomerular capillary is a bona fide filtering compartment was previously contentious due to the size of its fenestrae, which are seemingly wide enough to accommodate albumin with a molecular dimension of 8 × 8 × 3 nm (Sugio et al., 1999). Nevertheless, biophysical studies demonstrate that fenestrated and nonfenestrated capillaries have comparable permeabilities to macromolecules (Sarin, 2010). Recent studies indicate that the glomerular endothelium plays an active role in renal filtration on the basis of its negatively charged surface. The lumen of the glomerular capillaries and the fenestral surfaces are lined with a fibrous lattice of negatively charged glycoproteins called the glycocalyx (Fig. 2 E; Rostgaard and Qvortrup, 2002; Curry and Adamson, 2012). Additionally, plasma components are adsorbed within the glycocalyx, forming a broader coat >200 nm thick called the endothelial surface layer (ESL; Hjalmarsson et al., 2004). The filamentous structure and strongly negative charge of the ESL thus effectively make fenestrae narrower and more restrictive. Enzymatic destruction of different ESL components resulted in elevated albumin excretion accompanied by diminished ESL depth and loss of anionic sites on the endothelial surface (Gelberg et al., 1996; Jeansson and Haraldsson, 2003; Jeansson and Haraldsson, 2006; Meuwese et al., 2010; Dane et al., 2013). Similar loss of ESL charge density and increased passage of albumin across the GFB was observed when adsorbed ESL components are eluted by salt perfusion (Fridén et al., 2011).The role of the ESL in glomerular filtration has also been examined in proteinuric disease models. Renal perfusion of adriamycin, a drug used to induce proteinuria in mice, disrupted synthesis of glomerular proteoglycans and dramatically shriveled the glomerular ESL, impairing the size selectivity and charge density of the GFB (Jeansson et al., 2009). In rats, ageing-related proteinuria correlated with the loss of glomerular ESL (Salmon et al., 2012). In both animal models and in human patients, diabetes-induced proteinuria has also been strongly correlated with damage to the ESL (Salmon and Satchell, 2012). It has been proposed that the ESL could serve as a mechanosensor of fluid flow, an argument consistent with the loss of vasodilation upon removal of the ESL (Curry and Adamson, 2012; Fu and Tarbell, 2013). Altogether these findings align with the notion that the ESL is an essential feature of the glomerular endothelium and a crucial determinant of glomerular permeability.The establishment of a functional GFB is contingent on the proper development of the glomerular endothelium. Nascent podocytes secrete VEGFA, a potent chemoattractant and trophic factor for migratory angioblasts that become that glomerular endothelium. VEGFA binds the receptors VEGFR1 (Flt1), VEGFR2 (Flk1/Kdr), and neuropilin-1, which are expressed by these angioblasts (Robert et al., 2000). Homozygous ablation of Vegfa from podocytes results in arrest of glomerular development and failure to form a GFB. Haploinsufficiency for Vegfa, however, causes a latent and progressive hypertrophy of GECs with a concomitant disappearance of fenestrae, a phenomenon called endotheliosis (Eremina et al., 2003, 2006). This breakdown of the glomerular endothelium is seen when Vegfa ablation is induced in adult mouse podocytes or when its receptor is absent in GECs, which indicates that VEGFA acts in a paracrine manner via VEGFR2 (Eremina et al., 2008; Sison et al., 2010). These corroborate earlier findings showing that inhibition of VEGFA function causes rapid onset of endotheliosis and proteinuria (Sugimoto et al., 2003). Additionally, compound loss of the phospholipid-binding ATPases EHD3 and EHD4, which are expressed exclusively by GECs, strikingly resembles Vegfa haploinsufficiency and VEGFR2 deficiency (George et al., 2011). Their importance in vesicle trafficking suggests that EHD3 and EHD4 likely regulate the recycling of VEGFR2 based on the altered cell surface distribution of VEGFR2 in their absence.Interestingly, podocyte-specific overexpression of VEGFA₁₆₄, the predominant VEGFA isoform in the kidney, causes global collapse of the glomerular tuft, rapid depletion of GECs, and massive proteinuria (Eremina et al., 2003). Inducible overexpression of moderate levels of VEGFA₁₆₄ in postnatal and adult podocytes, however, causes a reversible disruption of glomerular structure and function (Veron et al., 2010a,b). As diabetic patients are known to have elevated levels of circulating VEGFA, these overexpression studies suggest that excessive VEGFA signaling could contribute to the progression of diabetic nephropathy (Chiarelli et al., 2000; Hovind et al., 2000). These findings further indicate that a delicately balanced dosage of VEGFA is necessary to coordinate the development and maintenance of the glomerular vasculature and the GFB.Signaling via secreted glycoproteins called angiopoietins intersects with the VEGFA-dependent pathway to balance stabilization and remodeling of renal and systemic vasculature (Augustin et al., 2009). The angiopoietin Angpt1 is produced by podocytes and MCs, whereas its cognate receptor Tie2 is expressed by GECs (Kolatsi-Joannou et al., 2001; Satchell et al., 2002). Inducible knockout of Angpt1 at mid-gestation (from mouse embryonic day 10.5) results in simplified and enlarged glomerular capillary tufts, and the delamination of GECs (Jeansson et al., 2011). Late gestation (embryonic day 16.5) deletion of Angpt1 does not cause overt glomerular maldevelopment but increased susceptibility to diabetic nephropathy. One factor that could contribute to impairment of the GFB is the loss of glomerular endothelial glycocalyx, which is caused by diabetic nephropathy and likely exacerbated by Angpt1 deficiency. In systemic vasculature, Angpt1 promotes barrier property and reduced permeability to albumin by stimulating the synthesis of glycocalyx and thickening of the ESL (Satchell et al., 2004; Salmon et al., 2009). Altogether, these studies underscore the importance of GECs and the ESL in renal filtration.

The GBM: A highly ordered ECM and filtration bed

The GBM derives from the fusion of the respective basement membranes of both podocytes and GECs (Abrahamson, 2012; Miner, 2012). Ultrastructure imaging by electron microscopy reveals a fibrous and stratified lattice with heterogeneous pores. Proteomic analysis identified 144 distinct proteins in purified human glomerular ECM including the GBM (Byron et al., 2014; Lennon et al., 2014), with the most abundant being collagens (types I, IV, VI, and XVIIII subunits), laminins (α5, β2, and γ1), nidogen-1, heparan sulfate proteoglycans (HSPGs, agrin, and perlecan), and tubulointerstitial nephritis antigen-like (TINAGL1) protein. The GBM is an integral component of the GFB acting as an intermediary sieving matrix. The GBM may also function as a sink for pro-angiogenic ligands and secreted factors that mediate cellular communication between podocytes and GECs. Lastly, the GBM cements podocytes and GEC in place by cell–ECM adhesive interactions, thus effectively stabilizing the GFB. Among the abundant components of the GBM, type IV collagens and laminins are the most indispensable.Alport syndrome is a hereditary disorder that targets the GBM, causing mild proteinuria during adolescence and progressing to end-stage renal failure. This ailment is linked to mutations in the genes COL4A3, COL4A4, and COL4A5, which encode the type IV collagen subunits α3, α4, and α5, respectively. Maturation of the GBM involves the substitution of the α1α1α2 (IV) collagen with the α3α4α5 (IV) collagen trimers as the predominant collagen complex, a developmental change that has been inferred to strengthen the GBM (Miner and Sanes, 1994). Mutations in Alport syndrome disrupt the assembly of α3α4α5 (IV) collagen trimers, leading to the persistent prominence of α1α1α2 (IV) collagen complexes. As α3α4α5 (IV) collagen trimers represent half the total proteins of a mature GBM (Candiello et al., 2010), it comes as no surprise that Alport GBMs are grossly perturbed in composition and are morphologically distorted. The importance of the collagen IV complex in the GBM is further highlighted by Goodpasture’s disease, an autoimmune disorder whereby self-reactive antibodies target the α3 subunit of collagen IV, resulting in glomerulonephritis (Cui and Zhao, 2011).Pierson syndrome is an autosomal recessive disorder presenting with congenital proteinuria and neuromuscular maldevelopment. Mutations linked to Pierson syndrome map to the gene LAMB2, which encodes the laminin β2, impairing the assembly of the laminin complex LM-521 (a heterotrimer formed among laminin-α5, -β2, and -γ1 subunits; Zenker et al., 2004; Matejas et al., 2010). Mice lacking Lamb2 also show the abnormal renal and neuromuscular phenotype of Pierson syndrome, and reveal a distinctive splitting of the GBM (Noakes et al., 1995). In mice, loss of Lama5 in podocytes or the expression of a hypomorphic allele of Lama5 (causing attenuated expression of laminin-α5) results in progressive proteinuria and ultrastructural deformation of the GBM (Kikkawa and Miner, 2006; Shannon et al., 2006; Goldberg et al., 2010). Loss of LM-521 in Pierson syndrome causes other laminin complexes (LM-111, -211, -332, and -511) to become more prevalent, although this apparent compensation is insufficient to restore normal GBM structure and GFB function.Scanning electron microscopy reveals that the GBM is not amorphous, but is rather a highly organized labyrinth of interconnected polygonal fibrils of varying thickness ranging from 4 to 10 nm (Kubosawa and Kondo, 1985; Hironaka et al., 1993). The fibrils are most densely packed within the core and have heterogeneous pores averaging 10 nm in diameter. In a proteinuric nephritis disease model in rodents, it was observed that the GBM fibril network was more loosely packed and had enlarged pores as big as 40 nm (Hironaka et al., 1996). Imaging analysis combining stochastic optical reconstruction microscopy (STORM) and correlative electron microscopy has revealed nanometer-resolution details of the highly stratified organization of the GBM, delineating the location and orientation of epitopes of major GBM components relative to the adhesion receptor integrin-β1 expressed by GECs and podocytes (Fig. 3 A; Suleiman et al., 2013). Collagen IV and nidogen-1 both map within the central region of the GBM. In contrast, laminin-α5, agrin, and integrin-β1 bimodally align within two distinct layers. Interestingly, the α3α4α5 and α1α1α2 (IV) collagen networks are particularly concentrated at the core, closer to the endothelial side, a distribution that is unexpectedly too distant from the extracellular domains of integrin-β1 at the surface of podocytes. This indicates that the physiologically important ligands of podocyte integrin-β1 are the agrin and laminin complexes. Remarkably, this imaging analysis correlates well with established domain–domain interactions of these GBM components.Open in a separate windowFigure 3.Molecular organization of the GBM and the SD. (A) Highly stratified assembly of GBM components. Laminin LM-521 and agrin are bimodally distributed, whereas collagen IV complexes are concentrated at the core of the GBM. The minor α1α1α2 collagen (Col) is notably biased toward the glomerular endothelium. Both the predominant α3α4α5 and the less abundant α1α1α2 type IV collagens are normally too distant from β1–integrin receptor (IR) complexes on the podocyte side. This suggests that LM-521 and agrin but not type IV collagens are the normal physiological ligands of IR complexes expressed by podocytes. (B) Simplified representation of major adhesion receptors (nephrin, Neph1, and Fat1) found in the SD. Lipid-raft localization of the SD is dependent on the cholesterol-binding podocin. The SD is coupled to both F-actin regulatory (Nck–N-WASP–Arp2/3 and CD2AP–Arp2/3) and cell polarity (Par6–aPKCλ/ι–Cdc42) complexes.Application of correlative STORM imaging to kidneys of Alport mice (Col4a3 mutant mice) demonstrates the dramatic redistribution of agrin and α1α1α2 (IV) collagen into a diffuse pattern throughout the width of the GBM. A likely implication of this is that podocytes might be inappropriately exposed to type IV collagens, thereby inducing a pathological transformation as observed in Alport disease. These imaging analyses bolster the argument that the ultrafine pore structure of the GBM is key to normal filtration, and that proteinuria results from perturbing the molecular and structural organization of the GBM.The abundance of HSPGs such as agrin, perlecan, and collagen XVIII confers a net negative charge to the GBM, which prompted a long-held assumption that the GBM is a critical determinant of the charge selectivity of the GFB (Rennke et al., 1975; Rennke and Venkatachalam, 1977; Harvey et al., 2007; van den Hoven et al., 2008; Goldberg et al., 2009). Genetic studies in mice aimed at minimizing the net negative charge of the GBM have disputed this argument and failed to result in overt proteinuria (Rossi et al., 2003; Harvey et al., 2007; Chen et al., 2008; Goldberg et al., 2009; Hamano et al., 2010). Similarly, treatment of the GBM with heparanase in order to strip glycosaminoglycan-associated anionic charges did not cause overt changes in glomerular morphology or induce proteinuria (van den Hoven et al., 2008). In light of these findings it is tempting to speculate that charge repulsion of circulating macromolecules in the GFB is primarily established within the glomerular compartment instead of the GBM.

The final gatekeepers: Renal podocytes and their slit diaphragms (SDs)

The defining feature of normal fully differentiated podocytes is their elaborate cytoarchitecture, which resembles the stellate body shape of an octopus, characterized by an arborized cell body with multiple projections subdivided into larger major processes and finer pedicels or FPs (Fig. 2, A and B). Major processes are reinforced by microtubules and intermediate filaments while FPs are actin-rich projections anchored to the GBM via focal adhesions (Ichimura et al., 2003). The podocyte cell bodies and their FPs wrap around the glomerular capillaries in a strikingly elaborate interdigitating pattern. Neighboring podocytes are physically adjoined through their FPs via unique intercellular junctions called the SD (Fig. 3 B). Unlike tight junctions, the SD lacks E-cadherin and is structurally porous (Tassin et al., 1994). The SD thereby serves as the exit port for primary urinary filtrate and is now well recognized as essential in the selective retention of high-molecular-weight plasma components. The seminal discoveries of the proteins nephrin and podocin as integral components of the SD are instrumental in proving that podocytes and their structural integrity are of key importance in the establishment and maintenance of the GFB (Kestilä et al., 1998; Holzman et al., 1999; Ruotsalainen et al., 1999; Boute et al., 2000; Schwarz et al., 2001; Huber et al., 2003). Inactivating mutations of NPHS1 and NPHS2, the respective genes encoding for nephrin and podocin, lead to congenital nephropathy characterized by the collapse of FPs and the absence of SDs (Kestilä et al., 1998; Boute et al., 2000). This stereotypical pathological transformation of podocytes called effacement is a distinctive hallmark of podocyte injury and is strongly correlated with the onset of proteinuria.Several genes, apart from NPHS1 and NPHS2, that encode for podocyte-specific proteins are strongly associated with the onset of proteinuric diseases including CD2AP (Shih et al., 1999; Kim et al., 2003), Kirrel/Neph1 (Donoviel et al., 2001), Fat1 (Ciani et al., 2003), TRPC6 (Reiser et al., 2005; Winn et al., 2005), ACTN4 (Kaplan et al., 2000), MYO1E (Krendel et al., 2009; Mele et al., 2011), ARHGAP24 (Akilesh et al., 2011), ARHGDIA (Togawa et al., 1999; Gee et al., 2013; Gupta et al., 2013), INF2 (Brown et al., 2010), COQ2 (Diomedi-Camassei et al., 2007), COQ6 (Heeringa et al., 2011), PLCE1 (Sadl et al., 2002; Hinkes et al., 2006), ANLN (Gbadegesin et al., 2014), PTPRO (Wharram et al., 2000; Ozaltin et al., 2011), and ADCK4 (Ashraf et al., 2013). Most of these genes encode for intrinsic SD components or their respective interacting partners while the rest encode for proteins needed for the survival, differentiation, cytoskeletal dynamics, and unique morphology of podocytes. The consequences of mutations of these genes highlight the important relationship between podocyte dysfunction and the disruption of the GFB.While many of the molecular constituents of the SDs have been identified, their topological assembly into a functional complex is poorly understood. The ectodomains of several adhesion receptors in the SD likely organize the bridge linking juxtaposed FPs via a combination of homophilic and heterophilic receptor–receptor interactions. By virtue of their large ectodomains, nephrin and Fat1 are excellent candidates to associate in trans to connect opposing FPs (Inoue et al., 2001; Ciani et al., 2003; Khoshnoodi et al., 2003; Wartiovaara et al., 2004). The smaller adhesion receptors such as Neph1 and Neph3, however, may interact in cis with nephrin and Fat1 (Gerke et al., 2003; Heikkilä et al., 2011). Consistent with its anatomical appearance as a junction between differentiated podocytes, other components of the SD are key molecules associated with adherens and tight junctions including ZO-1, CASK, spectrins, MAGI-2, JAMA-A, occludin, cingulin, and IQGAP1 (Lehtonen et al., 2005; Fukasawa et al., 2009). The huge scaffold protein ZO-1 is essential for the normal interdigitation of FPs and the formation of the SD (Itoh et al., 2014). Lack of ZO-1 triggers early onset proteinuria with podocyte effacement and the progressive scarring of the glomerulus (glomerulosclerosis). ZO-1 appears to be required to maintain the expression and correct spatial distribution of nephrin and podocin. Consistently, ZO-1 expression is significantly diminished in models of diabetic nephropathy.The distinctive morphology of podocytes underscores the importance of cell polarity signaling in podocyte biology. The SD marks the boundary between the apical and basolateral membrane domains of podocytes. Nephrin and Neph1 are known to interact with polarity proteins such as Par3, Par6, and aPKCλ/ι (Hartleben et al., 2008). The deletion of aPKCλ/ι and the small GTPase Cdc42, which regulates the activation of the Par3-Par6-aPKCλ/ι polarity complex, causes proteinuria and the formation of aberrant junctions between effaced FPs (Hirose et al., 2009; Huber et al., 2009; Scott et al., 2012; Blattner et al., 2013). Loss of aPKCλ/ι has been shown to interfere with cell surface localization of nephrin, podocin, and Neph1 (Satoh et al., 2014). Furthermore, studies in zebrafish demonstrate that the Crumbs (Crb) protein family member Crb2b is required for the differentiation of pronephric podocytes, whereas mutations in the human orthologue CRB2 have been linked to proteinuric disease (Ebarasi et al., 2009, 2015; Slavotinek et al., 2015). Crb proteins are part of the Crb–Patj–Pals1 polarity complex, which works alongside the Par3–Par6–aPKCλ/ι complex in directing the apical localization of particular membrane-bound proteins. Crb2b suppression in zebrafish leads to loss of polarized distribution of nephrin and the disruption of SD assembly (Ebarasi et al., 2009). In contrast, podocyte-specific ablation of Scribble, a determinant of basolateral trafficking, did not inhibit proper SD formation (Hartleben et al., 2012). These studies suggest that apical sorting predominates over basolateral sorting mechanisms to specify the polarized and dynamic assembly of the SD complex.Parallel bundles of actin filaments and a network of cortical actin form the backbone of terminal FPs, and the perturbation of this cytoskeletal assembly is thought to underlie FP effacement and the dismantling of the SD (Ichimura et al., 2003). Compound ablation of the adaptor molecules Nck1 and Nck2 in podocytes causes proteinuria, which demonstrates that the SD is intimately and dynamically coupled to the actin cytoskeleton (Jones et al., 2006). In vitro, oligomerized nephrin interacts with Nck adaptors, leading to the recruitment of N-WASP and Arp2/3 complex that mediates localized polymerization of actin filaments (Jones et al., 2006; Verma et al., 2006). Nck adaptors are not only required during podocyte maturation but are also needed to maintain preformed FPs (Jones et al., 2009). Since Nck proteins also interact with the PINCH–ILK–integrin complex, these adaptors could also help anchor podocytes to the GBM via actin-linked focal contacts (Tu et al., 1999; Dai et al., 2006; El-Aouni et al., 2006). Similarly, the Rho GTPase Cdc42 is required to mediate the linkage between the actin cytoskeleton and nephrin complexes (Scott et al., 2012). The cytoskeletal tethering of the SD is also dependent on the scaffold protein CD2AP, which promotes the stability of actin microfilament network of podocytes (Shih et al., 1999; Kim et al., 2003; Yaddanapudi et al., 2011; Tang and Brieher, 2013).Equally important to the assembly of the podocyte actin cytoskeleton are opposing events counteracting actin filament polymerization. Slit1-Robo2 signaling antagonizes nephrin-dependent actin polymerization yet is required to establish a normal pattern of FP interdigitation (Fan et al., 2012). Additionally, depletion of the actin-severing factor cofilin-1 has been shown to cause late-onset proteinuria and ultrastructural defects in podocytes in a manner akin to specific loss of Robo2 in podocytes (Ashworth et al., 2010; Garg et al., 2010). Mutations in ACTN4 (Kaplan et al., 2000), ARHGDIA (Togawa et al., 1999; Gee et al., 2013; Gupta et al., 2013), ARHGAP24 (Akilesh et al., 2011), INF2 (Brown et al., 2010), MYO1E (Krendel et al., 2009; Mele et al., 2011), and ANLN (Gbadegesin et al., 2014), which encode for known regulators of the actin cytoskeleton, have all been implicated in the etiology of proteinuric diseases.The cytoskeletal dynamics and structural plasticity of podocytes are also regulated by calcium signaling, lipid–protein interactions at the SD, and endocytosis. In podocytes, the ion channels Trpc5 and Trpc6 mediate distinctive calcium influx in response to angiotensin II, eliciting the reorganization of the actin cytoskeleton via modulation of the Rho GTPases Rac1 and RhoA (Tian et al., 2010). Gain-of-function mutations of TRPC6 are known to cause proteinuria in humans, whereas genetic loss of Trpc5 or Trpc6 prevents podocyte injury (Reiser et al., 2005; Winn et al., 2005; Schaldecker et al., 2013). These findings suggest that unbalanced elevation of intracellular calcium mitigates podocyte dysfunction and provide an explanation as to the protective benefits of blockade of angiotensin signaling in the progression of proteinuric renal diseases such as glomerulosclerosis and diabetic nephropathy. Interestingly, Trpc6 interacts with podocin, which suggests how intimately calcium signaling is coupled to the SD complex (Huber et al., 2007; Schurek et al., 2014). Lipid-dependent autocrine signaling in podocytes involving sFlt1 is required in the regulation of actin dynamics and the proper formation of FPs and the SDs (Jin et al., 2012). Specifically, sFlt1 secreted by podocytes binds to the glycosphingolipid GM3 at the podocyte surface, promoting cell adhesion, nephrin phosphorylation, and consequent remodeling of the cytoskeleton. Defective endocytosis in podocytes has also been shown to impair renal filtration. FP effacement and proteinuria ensues in the absence of endocytosis-related lipid-binding proteins, specifically dynamins, endophilins, and synaptojanin-1 (Soda et al., 2012). It has been postulated that clathrin-mediated endocytosis could dynamically sculpt FPs by regulating the turnover of SD components (Soda and Ishibe, 2013).Podocytes also play multiple important functions in maintaining the GFB independent of the formation of the SDs. Podocytes have a vital role in promoting the proliferation, survival, and development of endothelial cells. The pro-angiogenic factors VEGFA, Angpt1, and SDF1 are secreted by podocytes and are essential for the normal development of the glomerular endothelium (Simon et al., 1998; Yuan et al., 1999; Satchell et al., 2002; Takabatake et al., 2009; Haege et al., 2012). Podocytes together with the glomerular endothelium also collaborate in building the GBM (Byron et al., 2014). Whereas α1α1α2 (IV) collagen is produced jointly by endothelial cells and podocytes, the α3α4α5 (IV) collagen network is derived primarily from podocytes, as seen in vivo (Abrahamson et al., 2009). Macromolecules and proteins that traverse the GBM can be sequestered by podocytes via endocytosis, a mechanism that likely prevents the GFB from clogging (Eyre et al., 2007; Akilesh et al., 2008). Megalin and cubilin, which form a multifunctional endocytic receptor complex commonly found in absorptive epithelia, are coexpressed in podocytes and could mediate the retrieval of urinary albumin by podocytes (Yamazaki et al., 2004; Prabakaran et al., 2012). Overwhelming genetic evidence undeniably underscores the fact that the podocyte is an essential component of the GFB.

The importance of cell adhesion to the GBM

Biophysical studies demonstrate that GBM compression reduces permeability to albumin and the polysaccharide Ficoll (Robinson and Walton, 1989; Fissell et al., 2009). It is therefore tempting to speculate based on this that GECs and podocytes physically constrain and mitigate compression of the GBM. Consistent with this supposition are genetic studies showing that loss of Itga3 (Kreidberg et al., 1996), Itgb1 (Pozzi et al., 2008), Cd151 (Karamatic Crew et al., 2004; Sachs et al., 2006), Ddr1 (Gross et al., 2004), Ilk (Dai et al., 2006; El-Aouni et al., 2006), Tln1 (Tian et al., 2014), and Rap1a/b (Potla et al., 2014), genes encoding for proteins implicated in the adhesion of podocytes to GBM components such as collagens and laminins, results in impairment of the GFB. Intrinsic structural reorganization of the GBM such as in Alport and Pierson syndrome may also perturb the normal anchorage of podocytes and endothelial cells, cumulatively altering the compressibility of the GBM. This is not at all far-fetched, as ECM stiffness, by way of mechanotransduction via adhesion molecules, is known to influence a diverse range of cellular behavior including restructuring of the actin cytoskeleton, contractility, motility, gene expression, proliferation, and overall differentiation (DuFort et al., 2011).Effaced podocyte FPs could very well be indicative of not just the remodeling of cell–cell junctions or SDs but also of a maladaptive response to reestablish weakening focal contacts on the GBM. In fact, unfastening of podocytes and denuded GBM are common in the progression of diabetic nephropathy and chronic kidney disease (Toyoda et al., 2007; Weil et al., 2012; Kriz and Lemley, 2015), whereas endothelial detachment has been observed in Vegfa-null mutant mice (Eremina et al., 2003). Interestingly, comparison of the elastic properties of purified glomeruli by atomic force microscopy reveals that glomerular rigidity is reduced by as much as 30% in mouse models of Alport syndrome (Col4a3 knockout) and HIV-induced nephropathy (HIVAN) before the onset of overt pathological histology (Wyss et al., 2011). Increased glomerular deformability correlating with increased permeability of the GFB is likely symptomatic of an aberrant interaction between the GBM and the cells attached to it.

Streaming potential and charge selectivity in renal filtration

An attractive hypothesis based on electrokinetic principles has been proposed to account for the charge selectivity in renal filtration. Micropuncture measurements on salamander (Necturus maculosus) glomeruli demonstrate that filtration pressure establishes a distinctive streaming potential or charge difference across the GFB, with the Bowman’s space being more negative than the endothelial lumen (Fig. 4, A and B; Hausmann et al., 2010). The phenomenon of streaming potentials arises when electrolytes are forced by a pressure gradient across porous media or a channel carrying a permanent charge. In essence, the electrokinetic model posits that when small cations traverse the GFB, they bind and counterbalance the negatively charged surfaces within the GFB, reaching a threshold at which net ionic movement of small cations lags behind that of small anions (Hausmann et al., 2012; Moeller and Tenten, 2013). This differential advance of oppositely charged small ions thereby establishes a net charge separation and a measurable electrical field that polarizes the GFB. The streaming potential hypothesis therefore predicts that larger anions such as native albumin would encounter a retrograde electrophoretic field running opposite to the direction of hydraulic flux (Fig. 4 C). Given the minor importance of negative charges within the GBM, it can be inferred that the streaming potential is largely initiated from the highly charged ESL. The finding that neutral albumin traverses the GFB independent of the glomerular filtration rate whereas native anionic albumin passage becomes increasingly more restricted with increasing glomerular pressure is congruent with an electrokinetic model of the GFB (Lund et al., 2003). Similarly, albumin readily diffuses and equilibrates across the GFB once plasma flow is halted (Ryan and Karnovsky, 1976).Open in a separate windowFigure 4.Electrokinetic model of renal filtration. (A) Experimental setup used to demonstrate the existence of a flow-dependent electrical potential (streaming potential) across the GFB in salamander (N. maculosus) glomeruli (Hausmann et al., 2010). P, potential electrode; R, reference electrode. Small ions, due to differential interaction with the negatively charged GFB, create a net gradient of charges measurable as a streaming potential (blue arrows), making the endothelial lumen (EL) more positive than the Bowman’s space (BS). (B) Filtration pressure dependence of glomerular streaming potential. (C) Retrograde electrophoretic field created by streaming potentials. Due to streaming potentials, macromolecules encounter a dynamic electrophoretic field (green arrow) that is opposite to that of diffusive and convective fluxes (purple arrow). Albumin, a negatively charged macromolecule, would not only encounter size-dependent exclusion by the GFB but would effectively be electrophoresed away from the GFB during the course of active filtration.The hypothesis also predicts that podocyte FP effacement can be detrimental to the generation of a streaming potential as more rapidly advancing small anions bounce back upon encountering the broadened FPs, causing the electrical field across the GFB to be short-circuited (Hausmann et al., 2010, 2012). Hence, normal podocytes with their elaborate and regular network of FPs and SDs guarantees that streaming potential and filtration occur uniformly across the GFB. Further proof regarding this prediction is needed and should ideally be based on recording glomerular streaming potentials in the context of proteinuric disease models in mice. Nevertheless, in salamander glomeruli, streaming potentials are reversibly blocked by protamine, a polycationic protein that neutralizes the negative charge of the GFB and is well-known to induce proteinuria (Hausmann et al., 2010).

Conclusions

The robustness of the GFB depends on the plasticity and the dynamic signaling between its distinctive layers. Vigorous investigations on this subject over the years have shown that targeted damage to any one layer can lead to collapse of the GFB, and that in many cases compromising one layer has inevitable deleterious repercussions for the other layers. These highlight an emerging theme that the GFB, despite being multilayered, consists of components with dynamically intertwined roles, no single one of which is more important than the others, that harmonize together into one functionally elegant ensemble. Continued efforts to refine our understanding of the mechanism of renal filtration and the biology of the GFB are invaluable for the development of better therapeutic strategies to alleviate the burden of proteinuric diseases.     

Functional Characterization of the Plastidial 3-Phosphoglycerate Dehydrogenase Family in Arabidopsis

This work contributes to unraveling the role of the phosphorylated pathway of serine (Ser) biosynthesis in Arabidopsis (Arabidopsis thaliana) by functionally characterizing genes coding for the first enzyme of this pathway, 3-phosphoglycerate dehydrogenase (PGDH). We identified two Arabidopsis plastid-localized PGDH genes (3-PGDH and EMBRYO SAC DEVELOPMENT ARREST9 [EDA9]) with a high percentage of amino acid identity with a previously identified PGDH. All three genes displayed a different expression pattern indicating that they are not functionally redundant. pgdh and 3-pgdh mutants presented no drastic visual phenotypes, but eda9 displayed delayed embryo development, leading to aborted embryos that could be classified as early curled cotyledons. The embryo-lethal phenotype of eda9 was complemented with an EDA9 complementary DNA under the control of a 35S promoter (Pro-35S:EDA9). However, this construct, which is poorly expressed in the anther tapetum, did not complement mutant fertility. Microspore development in eda9.1eda9.1 Pro-35S:EDA9 was arrested at the polarized stage. Pollen from these lines lacked tryphine in the interstices of the exine layer, displayed shrunken and collapsed forms, and were unable to germinate when cultured in vitro. A metabolomic analysis of PGDH mutant and overexpressing plants revealed that all three PGDH family genes can regulate Ser homeostasis, with PGDH being quantitatively the most important in the process of Ser biosynthesis at the whole-plant level. By contrast, the essential role of EDA9 could be related to its expression in very specific cell types. We demonstrate the crucial role of EDA9 in embryo and pollen development, suggesting that the phosphorylated pathway of Ser biosynthesis is an important link connecting primary metabolism with development.Plant primary metabolism is a complex process where many interacting pathways must be finely coordinated and integrated in order to achieve proper plant development and acclimation to the environment. An example of such complexity is the biosynthesis of the amino acid l-Ser, which takes place in at least two different organelles and by different pathways. This amino acid is essential for the synthesis of proteins and other biomolecules needed for cell proliferation, including nucleotides and Ser-derived lipids, such as phosphatidylserine and sphingolipids. Additionally, d-Ser has been attributed a signaling function in male gametophyte-pistil communication (Michard et al., 2011).Despite the important role played by Ser in plants, the biological significance of the coexistence of several Ser biosynthetic pathways and how they interact to maintain amino acid homeostasis in cells is not yet understood. Three different Ser biosynthesis pathways have been described in plants (Kleczkowski and Givan, 1988; Ros et al., 2013; Fig. 1). One is the glycolate pathway, which takes place in mitochondria and is associated with photorespiration (Tolbert, 1980, 1997; Douce et al., 2001; Bauwe et al., 2010; Maurino and Peterhansel, 2010). In this pathway, two molecules of Gly are converted to one molecule of Ser in a reaction catalyzed by the Gly decarboxylase complex and Ser hydroxymethyltransferase (Fig. 1). Ser synthesis through the glycolate pathway is obtained in green tissues during daylight hours (Tolbert, 1980, 1985; Douce et al., 2001), suggesting that alternative Ser biosynthesis pathways may be required in the dark and/or in nonphotosynthetic organs. In this respect, Ser can be synthesized through two nonphotorespiratory pathways (Kleczkowski and Givan, 1988), the plastidial phosphorylated pathway (Ho et al., 1998, 1999a, 1999b; Ho and Saito, 2001) and the so-called glycerate pathway, which synthesizes Ser by the dephosphorylation of 3-phosphoglycerate (3-PGA; Kleczkowski and Givan, 1988; Fig. 1). This latter pathway includes the reverse sequence of the section of the oxidative photosynthetic carbon cycle linking 3-PGA to Ser (3-PGA-glycerate-hydroxypyruvate-Ser), these reactions being catalyzed by putative enzymes such as 3-PGA phosphatase, glycerate dehydrogenase, Ala-hydroxypyruvate aminotransferase, and Gly hydroxypyruvate aminotransferase. Although the existence of enzymatic activities of this pathway has been demonstrated (Kleczkowski and Givan, 1988), its functional significance is unknown and genes coding for the specific enzymes of the pathway have not been characterized to date.Open in a separate windowFigure 1.Schematic representation of Ser biosynthesis in plants. The enzymes participating in each Ser biosynthetic pathway are listed separately. Photorespiratory pathway (glycolate pathway): GDC, Gly decarboxylase; SHMT, Ser hydroxymethyltransferase. Glycerate pathway: PGAP, 3-phosphoglycerate phosphatase; GDH, glycerate dehydrogenase; AH-AT, Ala-hydroxypyruvate aminotransferase. Phosphorylated pathway: PSAT, 3-phosphoserine aminotransferase; PSP, 3-phosphoserine phosphatase. Abbreviations used for metabolites are as follows: 3-PHP, 3-phosphohydroxypyruvate; 3-PS, 3-phosphoserine; THF, tetrahydrofolate; 5,10-CH₂-THF, 5,10-methylene-tetrahydrofolate. This figure is adapted from Cascales-Miñana et al. (2013).The plastidial phosphorylated pathway of serine biosynthesis (PPSB; Fig. 1), which is conserved in mammals and plants, synthesizes Ser via 3-phosphoserine utilizing 3-PGA as a precursor (Kleczkowski and Givan, 1988). Evidence for the existence of this pathway in plants stems from the isolation and characterization of its enzyme activities (Handford and Davies, 1958; Slaughter and Davies, 1968; Larsson and Albertsson, 1979; Walton and Woolhouse, 1986). The PPSB involves three enzymes catalyzing sequential reactions: 3-phosphoglycerate dehydrogenase (PGDH), 3-phosphoserine aminotransferase, and 3-phosphoserine phosphatase (PSP; Fig. 1). Genes coding for some isoforms of these enzymes have been cloned and biochemically characterized in Arabidopsis (Arabidopsis thaliana; Ho et al., 1998, 1999a, 1999b; Ho and Saito, 2001).In humans, the PPSB plays a crucial role in cell proliferation control and oncogenesis (Bachelor et al., 2011; Locasale et al., 2011; Pollari et al., 2011; Possemato et al., 2011). The functional significance of the PPSB in plants has recently been unraveled by providing evidence for the crucial role of PSP1, the last enzyme of the pathway in embryo, pollen, and root development (Cascales-Miñana et al., 2013). However, the PPSB still requires further characterization. In order to gain a complete understanding of the PPSB function in plants, precise molecular, metabolic, and genetic knowledge of all the enzymes and genes of the pathway is needed. In this work, we follow a gain- and loss-of-function approach in Arabidopsis to characterize a family of genes coding for putative isoforms of PGDH, the first enzyme of the PPSB. Here, we identify the essential gene of this family and provide evidence for its crucial function in embryo and pollen development.     

Pharmacological targeting of guanosine monophosphate synthase suppresses melanoma cell invasion and tumorigenicity

Malignant melanoma possesses one of the highest metastatic potentials among human cancers. Acquisition of invasive phenotypes is a prerequisite for melanoma metastases. Elucidation of the molecular mechanisms underlying melanoma invasion will greatly enhance the design of novel agents for melanoma therapeutic intervention. Here, we report that guanosine monophosphate synthase (GMPS), an enzyme required for the de novo biosynthesis of GMP, has a major role in invasion and tumorigenicity of cells derived from either BRAF^V600E or NRAS^Q61R human metastatic melanomas. Moreover, GMPS levels are increased in metastatic human melanoma specimens compared with primary melanomas arguing that GMPS is an attractive candidate for anti-melanoma therapy. Accordingly, for the first time we demonstrate that angustmycin A, a nucleoside-analog inhibitor of GMPS produced by Streptomyces hygroscopius efficiently suppresses melanoma cell invasion in vitro and tumorigenicity in immunocompromised mice. Our data identify GMPS as a powerful driver of melanoma cell invasion and warrant further investigation of angustmycin A as a novel anti-melanoma agent.Malignant melanoma is one of the most aggressive types of human cancers. Its ability to metastasize in combination with resistance to conventional anticancer chemotherapy makes melanoma extremely difficult to cure, and the median survival of patients with metastatic melanoma is 8.5 months.^{1, 2, 3} A better understanding of the biology behind melanoma aggressiveness is imperative to facilitate the development of novel anti-melanoma strategies.Melanoma and other cancers cells have been shown to strongly rely on de novo nucleotide biosynthesis^{4, 5} and often overexpress several biosynthetic enzymes involved in these pathways.^{6, 7, 8, 9} Recently, we have identified a fundamental connection between melanoma invasion and biosynthesis of guanylates,⁸ suggesting that distortion of the guanylate metabolism facilitates melanoma progression.Guanosine monophosphate reductase (GMPR) reduces GMP to one of its precursors, inosine monophosphate (IMP), and depletes intracellular GTP pools (Figure 1a). We have recently demonstrated that GMPR suppresses melanoma cell invasion and growth of human melanoma cell xenografts. These findings tightly linked guanylate production to the invasive potential of melanoma cells.⁸Open in a separate windowFigure 1GMPS contributes to the invasive capability of melanoma cells. (a) Simplified schematic of the metabolic pathway for guanylates production. (b) SK-Mel-103 and SK-Mel-28 cells were transduced with a control vector or two independent shRNAs to GMPS and tested for invasion through Matrigel (left panel). Where indicated, cells were incubated with 100 μM guanosine for 24 h before the assay and guanosine supplementation was maintained throughout the experimental procedure. The data represent the average ± S.E.M. of at least two independent experiments. GMPS suppression was verified by immunoblotting (right panel). (c) Cells transduced as in (a) were plated on coverslips coated with FITC-conjugated gelatin. After 16 h cells were fixed with 4% PFA and stained for actin (rhodamine-conjugated phallodin) and nuclei (Hoechst). Where indicated, cells were incubated with 100 μM guanosine for 24 h before the assay and guanosine supplementation was maintained throughout the experimental procedure. At least 25 cells/sample were imaged to assess the number of cells with gelatin degradation. The data represent the average ± S.E.M. of two independent experiments. *P<0.05, **P<0.001 compared with control; ^#P<0.05, ^##P<0.001 compared with untreated cells. Statistics performed with Student''s t-Test. See also Supplementary Figure S1Of the several enzymes involved in guanylate biosynthesis, inositol monophosphate dehydrogenases 1 and 2 (IMPDH-1, -2), functional antagonists of GMPR (Figure 1a), have been targeted clinically with several drugs including the most specific one, mycophenolic acid (MPA) and its salt, mycophenolate mofetil (MMF).^{10, 11, 12, 13} Nonetheless, prior studies demonstrated that MPA possesses poor anti-tumor activity,^{14, 15} and it is primarily used as an immunosuppressing agent in organ transplantation.^{10, 11, 12}GMP synthase (GMPS) is the other functional antagonist of GMPR. GMPS catalyzes the amination of xanitol monophosphate (XMP) to GMP to promote GTP synthesis (Figure 1a).^{16, 17} Most of the studies on GMPS have been performed in bacteria, yeast, and insects, where GMPS has been shown to have a key role in sporulation, pathogenicity, and axon guidance, respectively.^{18, 19, 20} Mammalian GMPS has been the subject of several studies addressing its unconventional (GMP-unrelated) roles in the regulation of activity of ubiquitin-specific protease 7 (USP7).^{21, 22, 23, 24} However, because of the newly revealed importance of guanylate metabolism in control of melanoma cell invasion and tumorigenicity,⁸ GMPS emerges as an attractive target for anti-cancer therapy.In the late 1950s, a specific inhibitor of bacterial GMPS, angustmycin A (also known as decoyinine), has been isolated from Streptomyces hygroscopius as a potential antibiotic with sporulation-inducing activity in Bacillus subtilis.^{25, 26, 27, 28, 29} Its anti-tumor activity has never been experimentally explored. In the current study, we investigated the role of GMPS in regulation of melanoma invasion and tumorigenicity, and explored the possibility of targeting GMPS by angustmycin A as a novel anti-melanoma strategy.