Who’s in control? Principles of Rab GTPase activation in endolysosomal membrane trafficking and beyond

The eukaryotic endomembrane system consists of multiple interconnected organelles. Rab GTPases are organelle-specific markers that give identity to these membranes by recruiting transport and trafficking proteins. During transport processes or along organelle maturation, one Rab is replaced by another, a process termed Rab cascade, which requires at its center a Rab-specific guanine nucleotide exchange factor (GEF). The endolysosomal system serves here as a prime example for a Rab cascade. Along with endosomal maturation, the endosomal Rab5 recruits and activates the Rab7-specific GEF Mon1-Ccz1, resulting in Rab7 activation on endosomes and subsequent fusion of endosomes with lysosomes. In this review, we focus on the current idea of Mon1-Ccz1 recruitment and activation in the endolysosomal and autophagic pathway. We compare identified principles to other GTPase cascades on endomembranes, highlight the importance of regulation, and evaluate in this context the strength and relevance of recent developments in in vitro analyses to understand the underlying foundation of organelle biogenesis and maturation.

Membrane identity in the endomembrane systemOne key feature of eukaryotic cells is the presence of membrane-enclosed organelles, which constantly exchange proteins, lipids, or metabolites via vesicular transport or membrane contact sites (MCSs). Along the endomembrane system, vesicular trafficking requires vesicle budding from the donor membrane and directed transport toward and fusion with the acceptor compartment. The resulting trafficking routes form a regulated network that connects not only the internal organelles, but also the interior and exterior of the cell.The specific identity of organelles within the endomembrane system is defined by the lipid and protein composition of their membranes. This includes signaling lipids such as phosphoinositides (PIPs) and small GTPases of the Ras superfamily of small G proteins, namely of the Rab, Arf, and Arl families, which act as binding platforms for accessory proteins involved in multiple membrane trafficking processes (Balla, 2013).Rab GTPases, like other small GTPases, are key regulatory proteins that switch between an inactive GDP-bound (Rab-GDP) and an active GTP-bound (Rab-GTP) state (Barr, 2013; Goody et al., 2017; Hutagalung and Novick, 2011). Rabs are posttranslationally modified by the addition of geranylgeranyl moieties to C-terminal cysteine residues, which allow their reversible membrane association. Within the cytosol, Rab-GDP is kept soluble by binding to the chaperone-like GDP dissociation inhibitor (GDI). At the target membrane, an organelle-specific guanine nucleotide exchange factor (GEF) activates the Rab after its previous release from GDI, a process possibly supported by other factors (Dirac-Svejstrup et al., 1997). GTP binding stabilizes two loops in the Rab GTPase domain, which allows recruitment and binding of various so-called effector proteins to the Rab-GTP on the membrane. Rab GTPases are inefficient enzymes with a low intrinsic GTP hydrolysis rate and thus depend on a GTPase-activating protein (GAP) to hydrolyze bound GTP. GDI then extracts the Rab-GDP and keeps it soluble in the cytosol until the next activation cycle (Barr, 2013; Goody et al., 2017; Hutagalung and Novick, 2011). In addition to their conserved GTPase domain, Rabs contain a hypervariable C-terminal domain (HVD), which supports GEF recognition and therefore correct localization of the Rab (Thomas et al., 2018)Among various other functions, Rab GTPases are critical for the fusion of vesicles with the acceptor membrane by recruiting tethering proteins, which bring the two membranes into close proximity. Tethers, together with Sec1/Munc18 proteins, promote the folding of membrane-bound SNAREs at the vesicle and the target membrane into tetrameric coiled-coil complexes. This process further reduces the distance between the membranes, bypasses the hydration layer on membranes, and results in mixing of lipid bilayers and consequently membrane fusion (Wickner and Rizo, 2017; Ungermann and Kümmel, 2019).Organization and function of the endolysosomal pathwayEndocytosis allows the rapid adaptation of plasma membrane composition in response to changing environmental conditions by the uptake of membrane proteins from the plasma membrane, which are either transported to and finally degraded in the lysosome or sorted back to the plasma membrane, e.g., receptors after releasing their cargo within the endosomal lumen (Sardana and Emr, 2021). A third fate of endocytosed cargo is trafficking to the Golgi (Laidlaw and MacDonald, 2018). In addition, various kinds of endocytosis allow the uptake of very large particles such as bacteria during phagocytosis or fluids during pinocytosis (Huotari and Helenius, 2011; Babst, 2014). The endocytic pathway is also involved in the quality control system of plasma membrane proteins and allows degradation of damaged cell surface proteins as well as the down-regulation of nutrient transporters and receptors (Sardana and Emr, 2021). During endocytosis, membrane proteins marked by ubiquitination are incorporated into endocytic vesicles, which pinch off the plasma membrane and fuse with the tubular-shaped early endosome (EE) in the cell periphery (Fig. 1 A). The EE serves as a sorting station, at which membrane proteins are either sorted into tubular structures and brought to the recycling endosome (RE) or get incorporated into intraluminal vesicles (ILVs) with the help of four endosomal sorting complexes required for transport (ESCRTs; Sardana and Emr, 2021). A prerequisite for the degradation of cargo in the lysosome is the maturation of EEs into late endosomes (LEs) by changing the organelle surface composition, including specific Rab GTPases and PIPs, and organelle shape. The LE is eventually spherically shaped, containing multiple ILVs and a more acidified lumen. Therefore, it is also called Multivesicular Body (MVB). Upon fusion with the lysosome, ILVs and their content are degraded into precursor molecules, which are reused by the cell (Fig. 1 A; Sardana and Emr, 2021; Huotari and Helenius, 2011).Open in a separate windowFigure 1.Rab GTPases in the endolysosomal pathway.(A) Localization of key Rab GTPases along the endolysosomal pathway. Endocytic vesicles containing cargo (blue dot) or receptor proteins (red) are substrates of endocytosis. Endocytic vesicles (EV) fuse with the EE. Rabs are shown by numbers: Rab5 (green) on early EE is replaced by Rab7 (black) on multivesicular bodies (MVBs). GEFs are shown in blue. Positioning of lysosomes (Lys) depends on binding to motor proteins by either Arl8b (orange, 8b) or Rab7. Recycling occurs via REs involving Rab4, Rab11, and Rab14. MTOC, microtubule organizing center; Nuc, nucleus. (B) Spatiotemporal Rab5-to-Rab7 transition during endosomal maturation. Rab5 (green graph) is rapidly recruited to EE and replaced by Rab7. (C) Model of Rab7 GEF recruitment and activation on endosomes. Mon1-Ccz1 (or the trimeric complex additionally containing Rmc1/C18orf8/Bulli, as indicated by the unlabeled hexagon) requires Rab5-GTP for activation to promote Rab7 recruitment. For details, see text.Central functions of Rab5 and Rab7Along the endolysosomal system, several Rabs coordinate sorting and recycling processes at the EE and LE. Early endosomal Rab5 and late endosomal Rab7 are here the key Rabs conserved among species. Their spatiotemporal activation and therefore functions are tightly coordinated on the level of the MVB/LE (Fig. 1 B).In yeast, the Rab5-like GTPases Vps21, Ypt52, Ypt52, and Ypt10 and the Rab7-like Ypt7 structure the endocytic pathway (Singer-Krüger et al., 1994; Wichmann et al., 1992). In mammalian cells, Rab5 (with Rab5a, b, and c isoforms having nonredundant functions in the endocytic network; Chen et al., 2014, 2009) and Rab7 (with Rab7a and b isoforms, of which Rab7a is the main actor in transport processes along the endocytic pathway [Guerra and Bucci, 2016], whereas Rab7b has a role in the transport from endosome to the Golgi [Kjos et al., 2017; Progida et al., 2010]) are present (Wandinger-Ness and Zerial, 2014). While the overall organization of the endocytic pathway into EE and LE is conserved, yeast seems to have a more ancestral minimal endomembrane system, where the trans-Golgi network acts as EE and RE (Day et al., 2018). In mammalian cells, the more complex endolysosomal system depends on additional Rabs. Rab4 is involved in protein sorting at the EE, activation of Rab5, and recycling of cargo back to the plasma membrane (Kälin et al., 2015; Wandinger-Ness and Zerial, 2014; de Renzis et al., 2002), whereas Rab11 and Rab14 function at REs (Fig. 1 A; Linford et al., 2012; Takahashi et al., 2012). Furthermore, Rab9 is required for retrograde transport between LEs and the trans-Golgi network (Lombardi et al., 1993), and Rab32 and Rab38 function in the biogenesis of lysosome-related organelles (Bowman et al., 2019; Gerondopoulos et al., 2012; Wasmeier et al., 2006).During endosomal maturation, Rab5 is exchanged for Rab7 (Rink et al., 2005; Poteryaev et al., 2010). This Rab switch is highly conserved and a prime example of coordinated Rab turnover during organelle maturation. The rapid transition from Rab5 to Rab7 was explained by a so-called cutout switch, where activation of Rab5 fosters at a threshold value activation of Rab7, which in turn suppresses further Rab5 activation (Fig. 1 B; Del Conte-Zerial et al., 2008). Such a principle may apply to most Rab cascades (Barr, 2013).Rab5 has multiple functions on EEs (Wandinger-Ness and Zerial, 2014). It interacts with a number of effectors such as the lipid kinase Vps34, Rabaptin-5, which is found in complex with the Rab5-GEF Rabex5, Rabenosyn-5, and tethers such as the class C core vacuole/endosome tethering (CORVET) complex or EEA1. Therefore, Rab5 is critical for the homotypic fusion of EEs (Gorvel et al., 1991; Ohya et al., 2009; Christoforidis et al., 1999a, b; Perini et al., 2014; Marat and Haucke, 2016). Vps34 was initially identified in yeast (Schu et al., 1993) and exists in two heterotetrametric complexes, which differ by just one subunit (Kihara et al., 2001). Complex I resides on autophagosomes, whereas complex II functions on endosomes (Fig. 2 D). Both complexes generate a local pool of phosphatidylinositol-3-phosphate (PI3P), to which several effectors bind, including the early endosomal tether EEA1 and ESCRTs (Wallroth and Haucke, 2018). Recent structural insights revealed that Rab5 recruits and activates endosomal complex II, whereas Rab1 acts similarly on autophagosomal complex I (Tremel et al., 2021). This explains how Rab5-GTP promotes the formation of a local endosomal PI3P pool (Franke et al., 2019). Interestingly, Caenorhabditis elegans VPS-34 can recruit the Rab5 GAP TBC-2 to endosomal membranes, suggesting a possible link between PI3P generation and Rab5 inactivation (Law et al., 2017).Open in a separate windowFigure 2.Rab7 activation on autophagosomes.(A and B) Atg8-dependent Mon1-Ccz1 recruitment and activation. Atg8 (violet) recruits Mon1-Ccz1 (and likely also the trimeric GEF complex in higher eukaryotes, as indicated by the unlabeled hexagon) and allows fusion with lysosome. (C) Model of spatiotemporal Rab7 activation on autophagosomes. Maturation is prerequisite for successful fusion. (D) Comparison of proteins involved in maturation of LEs and autophagosomes.Rab7 is a key component in the late endocytic pathway (Langemeyer et al., 2018a). It is found on LEs, lysosomes, and autophagosomes and is required for the biogenesis and positioning of LEs and lysosomes, for MCSs of lysosomes with other organelles, and for the fusion of endosomes and autophagosomes with lysosomes (Fig. 1 A; Guerra and Bucci, 2016; McEwan et al., 2015; Ballabio and Bonifacino, 2020; Cabukusta and Neefjes, 2018). Even though both the metazoan Rab7 and yeast Ypt7 are activated by the homologous Mon1-Ccz1 GEF complex and are required for endosomal maturation, their function on LEs and lysosomes is not entirely conserved. In yeast, active Ypt7 directly binds the hexameric homotypic fusion and vacuole protein sorting (HOPS) tethering complex and mediates SNARE-dependent fusion of LEs or autophagosomes with vacuoles as well as homotypic vacuole fusion (Wickner and Rizo, 2017; Gao et al., 2018a, b). In higher eukaryotes, HOPS also promotes fusion between LEs and lysosomes, yet apparently does not directly interact with Rab7, but rather with the GTPases Rab2 and Arl8b (Gillingham et al., 2014; Fujita et al., 2017; Lőrincz et al., 2017; Khatter et al., 2015). How Rab7 contributes to fusion at the lysosome is still unclear. Rab7 interacts with several proteins on lysosomes, including the cholesterol sensor ORPL1 and the dynein-interacting lysosomal RILP (Jordens et al., 2001; Cantalupo et al., 2001; Rocha et al., 2009). Both proteins also bind HOPS (van der Kant et al., 2015, 2013), as does another multivalent adaptor protein, PLEKHM1 (McEwan et al., 2015), which binds both Arl8b and Rab7 (Marwaha et al., 2017). Interestingly, Arl8b in complex with its effector SKIP also binds TBC1D15, a Rab7 GAP, which may displace Rab7 from LEs before their fusion with lysosomes (Jongsma et al., 2020). It is thus possible that fusion of LEs and autophagosomes with lysosomes requires a complex coordination of the three GTPases, Rab7, Arl8b, and Rab2, with the HOPS complex and other effectors. Some of this complexity may be explained by a second function of Rab7 and Arl8b in binding adapters of the kinesin or dynein motor protein family, which connect LEs and lysosomes to the microtubule network. Thereby Rab7 and Arl8b control the positioning of these organelles to the periphery or perinuclear area via the microtubule network, which has functional implications (Fig. 1 A; Cabukusta and Neefjes, 2018; Bonifacino and Neefjes, 2017). Perinuclear lysosomes are the main places for degradation of cargo delivered by endosomes and autophagosomes, whereas peripheral lysosomes are involved in the regulation of mammalian target of rapamycin complex1 (mTORC1), the master regulator switching between cell growth and autophagy (Johnson et al., 2016; Korolchuk et al., 2011). This also may be connected to the role of lysosomes in lipid homeostasis, as Rab7 seems to control cholesterol export via the lysosomal NPC1 (van den Boomen et al., 2020; Shin and Zoncu, 2020; Castellano et al., 2017). How far the acidification state of perinuclear and peripheral lysosomes also affects their Rab7 and Arl8b mediated localization is still under debate (Ponsford et al., 2021). Thus, it is likely that Rab7 coordinates LE and lysosomal transport and fusion activity in coordination with endosomal biogenesis and cellular metabolism.GEF function and regulation in endosomal maturationThe heterodimeric complex Mon1-Ccz1 was identified as the GEF for Ypt7 in yeast and for Rab7 in higher eukaryotes (Nordmann et al., 2010; Gerondopoulos et al., 2012). The Mon1-Ccz1 complex is an effector of Rab5 (Kinchen and Ravichandran, 2010; Langemeyer et al., 2020; Cui et al., 2014; Li et al., 2015; Poteryaev et al., 2010; Singh et al., 2014), suggesting a direct link to endosomal maturation and Rab turnover (Fig. 1 B). Structural analyses uncovered how the two central longin domains in Mon1 and Ccz1 displace the bound nucleotide from Ypt7 (Kiontke et al., 2017). Unlike yeast, the metazoan Mon1-Ccz1 complex contains a third subunit termed RMC1 or C18orf8 in mammals and Bulli in Drosophila (Vaites et al., 2017; Dehnen et al., 2020; van den Boomen et al., 2020). Even though loss of this subunit impairs endosomal and autophagosomal biogenesis, this subunit does not affect GEF activity toward Rab7 in vitro (Dehnen et al., 2020; Langemeyer et al., 2020), indicating that the general GEF mechanism is conserved across species. As Rab7 is required on LEs, autophagosomes, and lysosomes, spatial recruitment and activity of the Rab7 GEF must be tightly regulated.Rab5 activates the Mon1-Ccz1 GEF complexDuring endosomal maturation, the Mon1-Ccz1 complex is recruited to Rab5- and PI3P-positive endosomes and activates Rab7 for subsequent fusion of endosomes with lysosomes (Nordmann et al., 2010; Poteryaev et al., 2010; Cabrera and Ungermann, 2013; Cabrera et al., 2014; Singh et al., 2014; Fig. 1 C). However, it was postulated that (but remained unclear how) Rab5 affects Rab7 GEF activity. The activity of GEFs is in the simplest way determined in solution, where the respective Rab, which has been loaded with a fluorescent- or radioactive-labeled nucleotide, is incubated with the GEF (Schoebel et al., 2009; Bergbrede et al., 2009). GDP or GTP addition then triggers displacement of the bound nucleotide, which results in a decrease of fluorescence or increase of radioactive signal in solution. Such in-solution assays can uncover the Rab specificity of GEFs yet cannot recapitulate the membrane context and potential regulating factors. Recent approaches therefore used liposomes and prenylated Rab:GDI complexes to address the role of membrane lipids and proteins in GEF activation (Thomas and Fromme, 2016; Thomas et al., 2018; Langemeyer et al., 2020, 2018b; Cezanne et al., 2020; Bezeljak et al., 2020). Details of these reconstituted systems are discussed below. In yeast, prenylated, membrane-bound, and GTP-loaded Rab5-like Vps21 was surprisingly inefficient as a single factor to recruit Mon1-Ccz1 to membranes, whereas addition of PIPs together with Vps21 enhanced recruitment (Langemeyer et al., 2020). However, activity of both the yeast and metazoan Rab7 GEF complexes showed a striking dependence on membrane-bound Rab5-GTP in the GEF assay, whereas PIPs alone were not sufficient to drive GEF activation. These observations demonstrate that the Mon1-Ccz1 complex depends on membrane-bound Rab5 for its Rab7 GEF activity, which nicely explains some of the previous in vivo observations on endosomal Rab5-to-Rab7 exchange (Poteryaev et al., 2010; Rink et al., 2005).This Rab exchange, which occurs similarly on phagosomes (Jeschke and Haas, 2016), is in vivo likely regulated in space and time. Time-lapse microscopy studies revealed that levels of fluorescently labeled Rab5 decreased, while fluorescently labeled Rab7 increased on the surface of a tracked endosome (Poteryaev et al., 2010; Yasuda et al., 2016). Analysis of the spatiotemporal Rab5-to-Rab7 transition in mammalian cells revealed that Rab5-positive endosomes can separate from Rab7-positive membranes, suggesting that a stepwise maturation process also occurs in some cells (Skjeldal et al., 2021). However, in all cases, only some insights on Mon1-Ccz1 regulation are presently available. Phosphorylation is one potential regulatory mechanism in GEF regulation (Kulasekaran et al., 2015). Indeed, yeast Mon1-Ccz1 is a substrate of the vacuolar casein kinase 1 Yck3 (Lawrence et al., 2014). When added to the Rab5-dependent GEF assay, Yck3-mediated phosphorylation inhibited Mon1-Ccz1 GEF activity, presumably by blocking the Rab5 interaction (Langemeyer et al., 2020). How the kinase is in turn regulated and whether this is the only mechanism of Mon1-Ccz1 GEF control is currently unknown.Rab7 activation and function in autophagyThe lysosome is also the destination of the autophagic catabolic pathway. During autophagy, portions of the cytosol, specific organelles, aggregates, or pathogens are engulfed into a double-layered membrane, which upon closure fuses with the lysosome for degradation and reuse of its content (Fig. 2 A; Zhao and Zhang, 2019; Nakatogawa, 2020). Autophagy is a versatile pathway required for adaptation of a cell’s organelle repertoire and quality control.Rab7 is found not just on LEs, but also on autophagosomes (Hegedűs et al., 2016; Gao et al., 2018a), although its precise function seems to differ between organisms (Kuchitsu and Fukuda, 2018). In yeast, the Rab7-homologue Ypt7 mediates HOPS-dependent fusion of autophagosomes with vacuoles (Gao et al., 2018a). In metazoan cells, Rab7 and its effectors PLEKHM1 and WDR91 are required for autolysosome/amphisome-lysosome fusion, yet Rab7 does not seem to directly bind HOPS during fusion of autophagosomes with lysosomes (Xing et al., 2021; McEwan et al., 2015; Gutierrez et al., 2004; Kuchitsu and Fukuda, 2018).Given the striking Rab5 dependence on endosomes in Mon1-Ccz1 activation, the question arises, how does Mon1-Ccz1-mediated Rab7 activation happen on autophagosomes? Some data suggest that yeast and metazoan Rab5 is directly involved in the autophagy process such as autophagosome closure (Ravikumar et al., 2008; Bridges et al., 2012; Zhou et al., 2019, 2017), whereas others do not find direct evidence, for instance in Drosophila (Hegedűs et al., 2016). Studies in yeast revealed that the LC3–like Atg8 protein directly binds and recruits Mon1-Ccz1 to the autophagosomal membrane during starvation, which results in Ypt7 activation as a prerequisite of HOPS-dependent fusion with the vacuole (Gao et al., 2018a; Fig. 2 B). Tight regulation of Mon1-Ccz1 GEF-activity is apparently mandatory to avoid fusion of premature autophagosomes with the vacuole (Fig. 2 C). How Mon1-Ccz1 localization to either endosomes or autophagosomes is coordinated (also with regard to similarities in organelle features; Fig. 2 D) and whether Atg8/LC3 also regulates the activity of the GEF complex are not yet known.Of note, an endosomal-like Rab5-to-Rab7 cascade also occurs on the mitochondrial outer membrane during mitophagy in metazoan cells, a selective pathway to degrade damaged mitochondria (Yamano et al., 2018). Here, Rab5 is activated by a mitochondrially localized Rab5 GEF, followed by Mon1-Ccz1 recruitment and Rab7A activation, which then orchestrates the subsequent mitophagy process. How this process is coupled to autophagosome maturation, and whether Rab7 is then again needed on the formed autophagosome, has not been addressed so far.These data nevertheless demonstrate the adjustable recruitment of Mon1-Ccz1 during endosomal maturation and autophagosome formation and even to the mitochondrial surface. Targeting of the Mon1-Ccz1 complex is likely coordinated between all these processes.A role for ER-endosome MCSs in endosome maturationEndosomes form MCSs with the ER. Such contact sites have multiple roles ranging from lipid transport to ion exchange (Scorrano et al., 2019; Reinisch and Prinz, 2021). The endosome-ER contact depends on Rab7 and contributes to transport and positioning of endosomes, supports endosomal fission, and facilitates endocytic cargo transport and cholesterol transfer between LEs and the ER (Rocha et al., 2009; Friedman et al., 2013; Rowland et al., 2014; Raiborg et al., 2015; Jongsma et al., 2016). Rab7 activation via the Mon1-Ccz1 complex is required for cholesterol export from the lysosome, likely in the context of MCSs. Rab7 binds to the NPC1 cholesterol transporter and may thus promote cholesterol export only at MCSs with the ER or other organelles (van den Boomen et al., 2020). The ER is also involved in endosome maturation, which requires an MCS between Reticulon-3L on the ER and endosomal Rab9. In fact, Rab9 is recruited shortly before the Rab5-to-Rab7 transition (Wu and Voeltz, 2021; Kucera et al., 2016). How Rab9 activation and MCS formation are coordinated with endosomal maturation has not yet been revealed. It is likely that the spatial positioning of endosomes (Fig. 1 A), their acidification, and TORC1 activity also contribute to this process (Bonifacino and Neefjes, 2017; Johnson et al., 2016).Retromer opposes Rab7 activationRetromer is a conserved heteropentameric complex that mediates the formation of vesicular carriers at the endosome and thus allows the transport of receptors back to the Golgi or plasma membrane. The complex consists of a trimeric core (Vps35, Vps26, and Vps29), which binds either a SNX1-SNX4 heterodimer or a SNX3 monomer (Simonetti and Cullen, 2018; Leneva et al., 2021; Kovtun et al., 2018). Retromer is an effector of Rab7, but also recruits the Rab7 GAP TBC1D5 in metazoan cells (Rojas et al., 2008; Kvainickas et al., 2019; Jimenez-Orgaz et al., 2018; Distefano et al., 2018; Seaman et al., 2009). This dual function of retromer may facilitate the formation of endosomal tubules after the Rab5-to-Rab7 transition, and these tubules eventually lose Rab7 once scission has occurred (Jongsma et al., 2020).It is not yet clear how conserved the Rab7-retromer-GAP connection is. Yeast retromer is also an effector of the Rab7-like Ypt7 and coordinates protein recycling at the endosome (Liu et al., 2012; Balderhaar et al., 2010), yet a role of a Rab7 GAP has not been described. However, yeast retromer also binds to the Rab5 GEFs Vps9 and Muk1 (Bean et al., 2015), which suggests that both Rab5 and Rab7 function contribute to efficient tubule formation at the endosome. Whether and how the Rab7 GEF Mon1-Ccz1 is functionally coordinated with retromer will be a topic of future studies.GEF regulation along the endomembrane systemIn the previous section, we focused mainly on the role of the Rab7 GEF in the context of endosome and autophagosome maturation. However, the timing of GEF activation and the subsequent recruitment of their target Rabs is critical for all membrane trafficking processes along the endomembrane system to guarantee maintenance of intracellular organelle organization. Rabs in turn interact with effectors, and effectors such as the lysosomal HOPS complex not only bind SNAREs but also catalyze their assembly and thus drive membrane fusion (Fig. 3 A). The spatiotemporal regulation of GEF activation is therefore at the heart of organelle biogenesis and maturation, and thus membrane trafficking. Within this section, we will now broaden our view by comparing different regulatory principles of GEFs.Open in a separate windowFigure 3.Regulatory mechanisms influence the activity of GEFs.(A) Hierarchical cascade of factors controlling membrane fusion. GEFs integrate various signals and initiate a cascade of protein activities, finally leading to membrane fusion. Signaling lipids, the presence of cargo proteins, upstream GTPases, and kinases influence the activity of GEFs and therefore determine Rab GTPase activation. Consequently, effector proteins such as tethering factors are recruited. This ultimately leads to SNARE-mediated lipid bilayer mixing and membrane fusion. (B) A Rab cascade in yeast exocytosis. Active Ypt32 and PI4P (yellow) on late Golgi compartments and secretory vesicles recruit the GEF Sec2, which in turn promotes activation and stable membrane insertion of the Rab Sec4. (C) Mon1-Ccz1 regulation by phosphorylation. Mon1-Ccz1 is recruited to and activated on LEs by coincidence detection of membrane-associated Rab5 and PI3P (red, Fig. 1 C) and promotes stable membrane insertion of Rab7. This process is terminated by Mon1-Ccz1 phosphorylation by the type I casein kinase Yck3 in yeast (orange). (D) A positive feedback loop of GEF activation on endocytic vesicles and EEs. The Rab5 GEF Rabex-5 binds ubiquitinated cargo on endocytic vesicles and is autoinhibited. Rab5 recruits Rabaptin-5, which binds Rabex-5 and releases the GEF from autoinhibition, generating a positive feedback loop. (E) Membrane factors determine GEF activity of TRAPPII at the trans-Golgi. TRAPPII activity for the Rab Ypt32 requires membrane-associated Arf1 and PI4P. (F) The length of the hypervariable domain of Golgi Rabs defines the substrate specificity for TRAPP complexes. The yeast Rab GTPases Ypt1 and Ypt32 differ in the length of their C-terminal HVD (box). TRAPPII and TRAPPIII complexes have the same active site, which is positioned away from the membrane, and thus discriminate Rab accessibility. (G) Phosphorylation as a mechanism to promote GEF activity. DENND1 GEF activity is autoinhibited, which is released by Akt-mediated phosphorylation. For details, see text.A Rab cascade in exocytosisAnother well-characterized Rab cascade is involved in the exocytic transport of secretory vesicles from the trans-Golgi network to the plasma membrane. At the trans-Golgi, the GEF transport protein particle II (TRAPPII) activates the Rab GTPase Ypt32, which then recruits the GEF Sec2 to secretory vesicles. Sec2 in turn activates the Rab Sec4, which binds the Sec15 subunit of the Exocyst tethering complex and allows vesicles to dock and fuse with the plasma membrane (Fig. 3 B; Walch-Solimena et al., 1997; Ortiz et al., 2002; Dong et al., 2007; Itzen et al., 2007). This cascade is conserved in humans. During ciliogenesis at the plasma membrane, the Ypt32 homologue Rab11 recruits the GEF Rabin 8, which in turn activates the human Sec4 homologue Rab8, a process regulated by phosphorylation (Hattula et al., 2002; Wang et al., 2015; Knödler et al., 2010). Interestingly, yeast Sec2 not only is a GEF, but also interacts with the Sec4 effector Sec15 (Medkova et al., 2006), a principle also observed in the endocytic Rab5 activation cycle, where the GEF Rabex5 interacts with the Rab5 effector Rabaptin-5. This dual role may also apply to Mon1-Ccz1, as the Mon1 homologue in C. elegans, SAND1, and yeast Mon1-Ccz1 can bind the HOPS tethering complex (Poteryaev et al., 2010; Nordmann et al., 2010).At the Golgi, phosphatidylinositol-4-phosphate (PI4P) contributes to directionality and spatiotemporal regulation of the exocytic Rab cascade. Sec2 binds both Ypt32 and PI4P on secretory vesicles via two binding sites, a process called coincidence detection. However, PI4P binding inhibits the interaction of Sec2 with Sec15. As vesicles reach the cell periphery, PI4P levels drop by the activity of Osh4, a lipid transporter, which allows Sec2 to bind the Exocyst subunit rather than Ypt32 (Ling et al., 2014; Mizuno-Yamasaki et al., 2010). In addition, Sec2 is phosphorylated by the plasma membrane–localized casein kinases Yck1 and Yck2 (Stalder et al., 2013; Stalder and Novick, 2016), resulting in effector recruitment rather than further Rab activation.Such a regulation may also apply to yeast Mon1-Ccz1. Anionic phospholipids and PI3P support Mon1-Ccz1 recruitment to liposomes and vacuoles (Langemeyer et al., 2020; Cabrera et al., 2014; Lawrence et al., 2014), whereas phosphorylation of the complex by the casein kinase Yck3 inhibits the binding of Mon1-Ccz1 to the Rab5-like Ypt10 and consequently reduces its GEF activity toward Rab7 (Fig. 3 C; Langemeyer et al., 2020). These observations suggest that the phosphorylation of GEFs by kinases may be a general regulatory principle in Rab cascades.Autoinhibition controls the Rab5 GEFAnother widely used regulatory mechanism is the autoinhibition of GEFs to control their activity. This has been analyzed in detail for the early endosomal Rab5-specific GEF Rabex-5, which interacts with the Rab5-effector Rabaptin-5 (Horiuchi et al., 1997). One factor for Rabex-5 recruitment to endocytic vesicles are ubiquitinated cargo proteins at the plasma membrane (Fig. 3 D; Mattera et al., 2006; Lee et al., 2006). Yet, isolated Rabex-5 has only low GEF activity in vitro (Delprato and Lambright, 2007). Structural analysis revealed that binding of Rabaptin-5 to Rabex-5 causes a rearrangement in the Rabex-5 C-terminus, which releases the GEF from autoinhibition and therefore facilitates nucleotide exchange of Rab5 (Delprato and Lambright, 2007; Zhang et al., 2014). On endosomes, increasing amounts of Rab5-GTP further promotes recruitment of the Rabex-5–Rabaptin-5 complex, resulting in a positive feedback loop of Rab5 activation and GEF recruitment (Lippé et al., 2001). Overall, Rabex-5 GEF activity is regulated by autoinhibition, a feedback loop with the Rab5 effector protein Rabaptin-5, and ubiquitinated cargo, which guarantees precise timing in establishing a Rab5-positive endosome. Of note, the Mon1 subunit of the Rab7 GEF can displace Rabex-5 from endosomal membranes (Poteryaev et al., 2010), which suggests a negative feedback loop of the Rab5 activation cascade once the next GEF is present.Regulation of Arf1 GEFs at different Golgi subcompartmentsThese key principles of GEF regulation in GTPase cascades are also found for Arf GTPases. Arf GTPases are soluble in their GDP-bound state by shielding their N-terminal myristate anchor in a hydrophobic pocket. Like Rabs, Arf GTPases are activated by specific GEFs, and their inactivation requires a specific GAP (Sztul et al., 2019). However, this review only highlights some key findings in the regulation of Rab GEFs and does not address regulation of the corresponding GAPs. Once activated, Arfs insert their lipid anchor and an adjacent amphipathic helix into membranes and are then able to bind effector proteins (Sztul et al., 2019). One of the best-studied Arf-GEFs is Sec7, which activates Arf1, an Arf GTPase involved in intra-Golgi trafficking (Achstetter et al., 1988). Studies on yeast Sec7 revealed that the protein is autoinhibited in solution and depends on three small GTPases—Arf1, the Rab Ypt1, and the Arf-like Arl1—for recruitment to the Golgi, a process supported by anionic lipids found in the late Golgi compartment. Importantly, the late Golgi Rabs Ypt31/32 strongly stimulate GEF activity (McDonold and Fromme, 2014; Richardson et al., 2012, 2016), indicating allosteric activation, as observed for Rab5-dependent Mon1-Ccz1 activation (Langemeyer et al., 2020). In this process, Sec7 dimerizes and promotes Arf1 recruitment and thus establishes a positive feedback loop. Interestingly, membrane binding of two additional Arf1 GEFs of the early Golgi, Gea1/2, depends on Rab1/Ypt1 and neutral membranes. Under these conditions, Gea1/2 is released from autoinhibition, although no positive feedback loop was observed (Gustafson and Fromme, 2017). Thus, Arf GEF regulation and Arf activation are tightly linked to multiple small GTPases and the membrane environment to establish Golgi compartments.Regulation and specificity of TRAPP complexes at the GolgiArf1 activation is also linked to the activation of Golgi-specific Rabs. Arf1-GTP binds to the highly conserved TRAPP GEF complexes at the Golgi (Fig. 3 E). Yeast and mammalian cells contain two TRAPP complexes. In yeast, both complexes share seven core components. TRAPPIII in addition contains Trs85, while accessory TRAPPII subunits are instead Trs130, Trs120, Trs65, and Tca17. Metazoan TRAPP complexes contain additional subunits (Lipatova and Segev, 2019).Interestingly, both complexes share the same catalytic site for Rab1/Ypt1 and Rab11/Ypt32. However, TRAPPIII provides GEF activity toward Rab1/Ypt1. Initially, it was proposed that TRAPPII can activate both Rab1/Ypt1 and Rab11/Ypt32 (Thomas et al., 2019, 2018; Thomas and Fromme, 2016; Riedel et al., 2018); however, it was recently shown that the TRAPPII complex is specific for Rab11/Ypt32 (Riedel et al., 2018; Thomas et al., 2019). Reconstitution of GEF activity on liposomes helped here to unravel TRAPP complex substrate specificity, since in solution assays are not adequate to address some of the features important for specific interactions: Rab11/Ypt32 has a longer HVD between the prenyl anchor and the GTPase domain compared with Rab1/Ypt1 (Fig. 3 F, box). The HVD not only binds TRAPPII but also stretches a longer distance from the membrane (Fig. 3 F). Thereby it allows Rab11/Ypt32, but not Rab1/Ypt1, to reach the active site of membrane-bound TRAPPII. Thus, substrate specificity is controlled by the distance of the GTPase domain from the membrane surface, since the active site seems to be located on the opposing site of the complex from the site of membrane interaction (Fig. 3 F; Thomas et al., 2019). The smaller TRAPPIII has its active site closer to the membrane, binds Ypt1 via its shorter HVD, and facilitates its activation, while Ypt32 with its longer HVD may be positioned too far away from the active site. In addition, both complexes require their respective membrane environment for optimal activity, indicating how Arf and Rab GEFs cooperate in Golgi biogenesis.The GEF DENND1 requires Arf5 for Rab35 activationRecently, another example of Arf-mediated Rab activation was reported (Kulasekaran et al., 2021). Rab35, an endocytic Rab found at the plasma membrane and REs (Sato et al., 2008; Kouranti et al., 2006), is involved in cell adhesion and cell migration by controlling the trafficking of β1-integrin and the EGF receptor (Klinkert and Echard, 2016; Allaire et al., 2013). Arf5 binds the Rab35 GEF DENND1 and stimulates its GEF activity, with dysregulation of this cascade linked to glioblastoma growth (Kulasekaran et al., 2021). DENND1 GEF activity is initially autoinhibited and relieved by phosphorylation via the central Akt kinase (Fig. 3 G; Kulasekaran et al., 2015). Similarly, another DENN-domain containing GEF, DENND3, is phosphorylated by the autophagy-specific ULK kinase and then activates Rab12, a small GTPase involved in autophagosome trafficking (Xu et al., 2015). Thus, it seems that Rab GEF activation is more generally linked to other trafficking proteins, such as Arfs, and controlled by kinases and likely also phosphatases.Lessons from reconstitutionOrganelle biogenesis and maintenance in the endomembrane system are tightly linked to the correct spatial and temporal activation of Rab GTPases. A small yeast cell gets by with 11 Rabs, while human cells encode >60 (Hutagalung and Novick, 2011). Rab activation, and therefore membrane identity, of each organelle depends on the cognate GEF. This puts GEFs into the driver’s seat of any Rab-directed function at cellular membranes. It seems that GEFs integrate, by several regulatory loops, incoming signals from various sources such as membrane composition, cargo proteins, upstream GTPases, or kinases/phosphatases (Fig. 3 A). Yet our insights on the specific membrane targeting and regulation of GEFs remain incomplete for want of available experimental approaches. We briefly discuss here how recent advances on the reconstitution of GEF-mediated Rab activation at model membranes have advanced our understanding of organelle maturation and biogenesis.Reconstitution of any reaction to uncover the essential constituents is limited by the available tools. GEFs, Rabs, Sec18/Munc1 proteins, tethering factors, and SNAREs are for instance required for membrane fusion (Fig. 3 A). Initial assays focused on SNAREs and revealed their important but rather inefficient fusogenicity (Weber et al., 1998). Further analyses uncovered critical activation steps for SNAREs (Malsam et al., 2012; Pobbati et al., 2006; Südhof and Rothman, 2009; Jahn and Scheller, 2006), yet fusion at physiological SNARE concentrations in various in vitro systems does not occur, unless assisted by chaperoning Sec1/Munc18 proteins and tethering factors (Bharat et al., 2014; Lai et al., 2017; Mima and Wickner, 2009; Ohya et al., 2009; Wickner and Rizo, 2017). Most tethers again depend on Rabs for their localization, and Rab localization to membranes requires a GEF (Cabrera and Ungermann, 2013), whose activity can be a limiting factor for fusion (Langemeyer et al., 2020, 2018b). The long avenue of understanding the mechanism and regulation of membrane fusion exemplifies the challenges in dissecting the complexity of a cellular reaction, but also demonstrates the powerful insights obtained from reconstitution of these processes.GEFs determine the localization of the corresponding Rab, and consequently, Rabs follow their GEF if they are mistargeted (Gerondopoulos et al., 2012; Blümer et al., 2013; Cabrera and Ungermann, 2013). However, these anchor-away approaches completely bypass the tight cellular regulation of GEF activation by the mistargeting and additional overexpression of the GEF protein and may allow only statements about GEF/substrate specificity. The spatiotemporal activation of each GEF at the right organelle is vital for the timing of all downstream reactions. GEFs are recruited to membranes by coincidence detection, which includes membrane lipids such as PIPs, membrane packaging defects, and peripheral membrane proteins such as upstream Rabs or other small GTPases. This recruitment is often accompanied by the release from autoinhibition, which may be triggered or inhibited by other regulatory processes such as phosphorylation. It comes as no surprise that pathogens such as Legionella and Salmonella take advantage of the central function of GEFs to establish and nourish their intracellular organellar niche by manipulating small GTPase activity (Spanò and Galán, 2018).To understand the specificity of Rab GEFs (and GAPs), mostly very simplified systems were used. Most GEF assays analyze soluble Rabs loaded with fluorescent 2′-O-(N-methylanthraniloyl) (MANT)-nucleotide or radioactively labeled GTP/GDP and soluble GEF in a test tube, where nucleotide exchange activity is observed upon addition of unlabeled nucleotide (Fig. 4 A). This strategy allows the identification of substrate (Rab) specificity of GEFs, but could also lead to misleading results, as pointed out earlier on the example of the TRAPP complexes and Rab1/Ypt1 or Rab11/Ypt32. In addition, GEF-Rab pairs negatively regulated by one of the above principles could easily be missed.Open in a separate windowFigure 4.Approaches to determine GEF activity in vitro. Methods to determine GEF activity for Mon1-Ccz1. In all approaches, Rab7 is preloaded with fluorescent MANT-GDP. Fluorescence decreases upon GEF-mediated nucleotide exchange. (A) GEF assays. (Ai) In-solution Rab GEF assay. Mon1-Ccz1 (blue, Bulli/Rmc1/C18orf8 subunit, indicated by unlabeled hexagon) and Rab7 (gray) are freely diffusible in the test tube, which results in random collision and Rab activation. (Aii) GEF-mediated activation of artificially recruited Rab7 on liposomes. Rab7 with a C-terminal 6xHis-tag is permanently immobilized on membranes containing the cationic lipid DOGS-NTA. Mon1-Ccz1 unspecifically binds to this membrane surface and activates Rab7. Diffusion is limited to the membrane surface, thus increasing chances of interactions. (Aiii) Reconstitution of Rab5-mediated Rab7 activation by Mon1-Ccz1 on liposomes. Chemically activated, prenylated Rab5 (green), delivered to the membrane by the Rab Escort Protein (REP), allows Mon1-Ccz1 recruitment and Rab7 activation from the GDI complex (see text for further details). (B) Summary of Ai–Aiii. pren., prenylation.As Rabs and GEFs function on membranes, we and others adopted strategies for measuring Rab activation by GEFs on membranes (Fig. 4 B). In a first approach, Rab and other small GTPases (Sot et al., 2013; Schmitt et al., 1994) were immobilized with C-terminal hexahistidine tags on liposomes containing the polycationic lipid 1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl] (DOGS-NTA) and observed higher activity of the added GEF (Cabrera et al., 2014; Thomas and Fromme, 2016). A drawback of this technique is the artificial membrane composition. To avoid potential artifacts of unnaturally charged membranes and permanently membrane-bound Rab, recent studies relied on prenylated Rabs in complex with GDI. Reflecting the natural source of the cytoplasmic Rab pool, this complex was used as a GEF substrate in the presence of liposomes mimicking the natural membrane composition (Cezanne et al., 2020; Bezeljak et al., 2020; Langemeyer et al., 2020, 2018b; Thomas et al., 2018, 2019; Thomas and Fromme, 2016).Even though these observations are recent, the outcome and the understanding of GEF regulation is encouraging. For the Rab5 GEF complex consisting of Rabex5 and Rabaptin5, GEF-dependent Rab5 recruitment to membranes revealed a self-organizing system, nonlinear Rab5 patterning, and collective switching of the Rab5 population (Bezeljak et al., 2020; Cezanne et al., 2020). This is in agreement with mathematical modeling and predictions on bistability and ultrasensitivity of Rab networks (Del Conte-Zerial et al., 2008; Barr, 2013). For the Golgi-resident TRAPPII and TRAPPIII complexes, the membrane composition, the length of the Rab HVD, and the presence of membrane-bound Arf1 determined the GEF specificity for their Rabs (Fig. 3 F; Thomas et al., 2019, 2018; Thomas and Fromme, 2016; Riedel et al., 2018), which is nicely supported by recent structural analyses of yeast and metazoan TRAPPIII (Galindo et al., 2021; Joiner et al., 2021)Our own data uncovered that the yeast and metazoan Mon1-Ccz1(-RMC1) complex required membrane-bound Rab5-GTP to activate Rab7 out of the GDI complex (Langemeyer et al., 2020). Surprisingly, Rab5-GTP not only determined membrane binding of Mon1-Ccz1, but also activated the GEF on membranes by a yet-unknown mechanism (Fig. 1 C). Phosphorylation of yeast Mon1-Ccz1 by the casein kinase Yck3 inhibited this activation, demonstrating possible regulation of GEF activity (Fig. 3 C). Importantly, this finding agrees with the observed Rab5-to-Rab7 switch in vivo (Poteryaev et al., 2010; Rink et al., 2005).Taken together, the available tools open exciting avenues for our understanding of organelle maturation. Reconstitution will allow the investigation of an entire Rab cascade and its regulation by kinases or membrane lipids. It will be possible to determine the cross-talk with lipid kinases and observe possible regulatory loops between Rabs and PI kinases (Tremel et al., 2021). We are confident that such analyses, complemented by in vivo analyses of Rabs or other small GTPases and their GEFs, will clarify the underlying mechanism of organelle maturation and biogenesis along the endomembrane system of eukaryotic cells.     

ATG8ylation of proteins: A way to cope with cell stress?

The ATG8 family of proteins regulates autophagy in a variety of ways. Recently, ATG8s were demonstrated to conjugate directly to cellular proteins in a process termed “ATG8ylation,” which is amplified by mitochondrial damage and antagonized by ATG4 proteases. ATG8s may have an emerging role as small protein modifiers.

ATG8 proteins directly conjugate to cellular proteinsAutophagy describes the capture of intracellular material by autophagosomes and their delivery to lysosomes for destruction (Kaur and Debnath, 2015). This process homeostatically remodels the intracellular environment and is necessary for an organism to overcome starvation (Kaur and Debnath, 2015). The autophagy pathway is coordinated by autophagy-related (ATG) proteins that are controlled by diverse post-translational modifications (e.g., phosphorylation, acetylation, ubiquitination, and lipidation; Ichimura et al., 2000; McEwan and Dikic, 2011). Recently, a previously uncharacterized post-translational modification termed “ATG8ylation” was uncovered (Agrotis et al., 2019; Nguyen et al., 2021). ATG8ylation is the direct covalent attachment of the small ubiquitin-like family of ATG8 proteins to cellular proteins (Agrotis et al., 2019; Nguyen et al., 2021). Until now, the only known instances of ATG8 conjugation to proteins were of a transient nature, as E1- and E2-like intermediates with ATG7 and ATG3, respectively, as a way of ligating ATG8 to the lipid phosphatidylethanolamine during autophagy (Ichimura et al., 2000). Therefore, ATG8ylation may represent an underappreciated regulatory mechanism for many cellular proteins that coordinate pathways such as mitophagy.ATG8s play many roles in the autophagy pathwayDuring canonical autophagy, the ATG8 family (comprising LC3A, -B, and -C and GABARAP, -L1, and -L2) undergoes molecular processing that concludes with their attachment to phosphatidylethanolamine, enabling proper construction of autophagosomes and subsequent autophagosome–lysosome fusion (Nguyen et al., 2016). The ATG4 family of cysteine proteases (ATG4A, -B, -C, and -D) cleaves ATG8 proteins immediately after a conserved glycine residue in their C terminus in a process dubbed “priming,” which leads to the formation of ATG8-I (Skytte Rasmussen et al., 2017; Tanida et al., 2004). ATG7 then attaches to the exposed glycine residue of ATG8-I via a thioester linkage to form an E1 ubiquitin-like complex that transfers ATG8-I to ATG3 in a similar way to generate an E2-like complex (Ichimura et al., 2000). The ATG5–ATG12–ATG16L1 complex then catalyzes the E3-like transfer of ATG8-I from ATG3 to phosphatidylethanolamine to form ATG8-II, which is the lipidated species that is incorporated into double membrane–bound compartments such as autophagosomes (Hanada et al., 2007). The lipidation of ATG8s and their recruitment to the phagophore are not essential for the formation of autophagosomes but are important for phagophore expansion, the selective capture of autophagic substrates, and autophagosome–lysosome fusion (Kirkin and Rogov, 2019; Nguyen et al., 2016). Intriguingly, ATG8 lipidation is multifaceted, as ATG8s can be alternatively lipidated with phosphatidylserine (instead of phosphatidylethanolamine) to enable their recruitment to single membrane–bound compartments during LC3-associated phagocytosis, influenza infection, and lysosomal dysfunction (Durgan et al., 2021).The discovery of ATG8ylationKey insights into ATG8ylation came from the observation that various ATG8s form high-molecular-weight species in cells following the expression of their primed forms that have their C-terminal glycine exposed (for example, LC3B-G), bypassing the need for cleavage by ATG4 (Agrotis et al., 2019; Nguyen et al., 2021). Indeed, on an immunoblot, ATG8⁺ “smears” resemble that of ubiquitinated proteins (Agrotis et al., 2019; Nguyen et al., 2021). Traditionally, in the autophagy field, ATG8⁺ smears were thought to arise from poor antibody specificity. However, in light of recent findings, this widely accepted interpretation has been challenged, given that ATG8⁺ smears are enriched following ATG8 overexpression and disappear in the absence of ATG8s (Agrotis et al., 2019; Nguyen et al., 2021). Smearing has also been detected after immunoprecipitation of epitope-tagged ATG8s from cell extracts under denaturing conditions, ruling out noncovalent interactions accounting for this upshift (Agrotis et al., 2019; Nguyen et al., 2021). Further, smearing is not abolished by deubiquitinase treatment, arguing strongly against ATG8 ubiquitination as the cause (Nguyen et al., 2021). Everything considered, the most plausible explanation is that ATG8 itself undergoes covalent linkage to cellular proteins, akin to ubiquitin and NEDD8 modifiers, which are structurally similar to ATG8s. Remarkably, the protease ATG4 antagonizes the ATG8ylation state of many proteins (Agrotis et al., 2019; Nguyen et al., 2021).ATG4 displays isoform-specific proteolytic cleavage of ATG8ATG4 is required for the formation of autophagosomes, but its protease activity is not (Nguyen et al., 2021). The protease activity of ATG4 is, however, required for ATG8 processing, such as priming ahead of lipidation and de-lipidation, which removes excess ATG8 from autophagosomes and other membranes (Nguyen et al., 2021; Tanida et al., 2004; Fig. 1 A). Apart from these functions, ATG4 regulates the deubiquitinase-like removal of ATG8 from cellular proteins (de-ATG8ylation; Agrotis et al., 2019; Nguyen et al., 2021; Fig. 1 A). Consistent with this role, deletion of all four ATG4 isoforms (A, B, C, and D) increases the abundance of ATG8ylated proteins (Nguyen et al., 2021). In contrast, overexpression of ATG4B has the opposite effect, but only if its protease activity is intact (Agrotis et al., 2019). As such, ATG4 inhibits the ATG8ylation state of many proteins, which is likely to modulate their downstream functions.Open in a separate windowFigure 1.The many roles of ATG4 in ATG8 processing. (A) Molecular processing of ATG8 proteins by ATG4 illustrating its roles in priming, de-lipidation, and de-ATG8ylation. The structure of LC3B (Protein Data Bank accession no. 1V49) was used to denote ATG8 (G, glycine; PE, phosphatidylethanolamine). (B) Heatmap summarizing relationships between ATG4 isoforms and ATG8 family members. Data were summarized for qualitative interpretation (Agrotis et al., 2019; Li et al., 2011; Nguyen et al., 2021). Int., intermediate; N.d., not determined. (C) Graphical summary of questions moving forward with ATG8ylation (P, phosphorylation).ATG4 is an important “gatekeeper” for ATG8 conjugation events. ATG4 primes ATG8s to expose their C-terminal glycine, which is required for conjugation to proteins or lipids; however, ATG4 also catalyzes de-ATG8ylation and de-lipidation events, respectively (Agrotis et al., 2019; Nguyen et al., 2021; Tanida et al., 2004). Because the C-terminal glycine of a single ATG8 is occupied when conjugated to a protein or lipid, it is unlikely that ATG8ylated proteins directly engage with phagophore membranes in the same way as ATG8-II. Indeed, protease protection assays with recombinant ATG4B reveal that de-ATG8ylation of cell lysates remains unchanged with or without organellar membrane disruption, suggesting that ATG8ylated proteins are largely cytoplasmic facing rather than intraluminal (Agrotis et al., 2019). Paradoxically, however, ATG8ylation is enhanced by lysosomal V-type ATPase inhibition, which blocks the degradation of lysosomal contents, indicating that ATG8ylated substrates may undergo lysosome-dependent turnover (Agrotis et al., 2019; Nguyen et al., 2021). One explanation for these differences may be that the process of ATG8ylation is itself sensitive to lysosomal dysfunction.Functional relationships between ATG4s and ATG8sIsoforms of ATG4 show clear preferences for proteolytically processing ATG8 subfamilies (i.e., LC3s and GABARAPs) for de-ATG8ylation and priming upstream of phosphatidylethanolamine ligation (Agrotis et al., 2019; Li et al., 2011; Nguyen et al., 2021; Fig. 1 B). ATG4A strongly reduces the abundance of proteins that have been ATG8ylated with the GABARAP family while promoting ligation of GABARAPs to phosphatidylethanolamine (Agrotis et al., 2019; Nguyen et al., 2021; Fig. 1 B). In contrast, ATG4B strongly reduces the abundance of proteins that have been ATG8ylated with LC3 proteins while promoting ligation of LC3s to phosphatidylethanolamine (Agrotis et al., 2019; Nguyen et al., 2021; Fig. 1 B). In comparison, ATG4C and -D lack obvious de-ATG8ylation activity, although the latter weakly promotes phosphatidylethanolamine ligation to GABARAPL1 only (Nguyen et al., 2021). These functional similarities between ATG4 isoforms are consistent with both their sequence and structural homology (i.e., ATG4A and -B are most similar; Maruyama and Noda, 2018; Satoo et al., 2009). Structurally, ATG4B adopts an auto-inhibited conformation with its regulatory loop and N-terminal tail blocking substrate entry to its proteolytic core (Maruyama and Noda, 2018). LC3B induces conformational rearrangements in ATG4B that involve displacement of its regulatory loop and its N-terminal tail, with the latter achieved by an interaction between the ATG8-interacting region in its N-terminal tail with a second copy of LC3B that functions allosterically (Maruyama and Noda, 2018; Satoo et al., 2009). These rearrangements permit entry of LC3B into the proteolytic core of ATG4B, where cleavage of LC3B following its C-terminal glycine occurs (Li et al., 2011; Maruyama and Noda, 2018). ATG4B^L232 is directly involved in LC3B binding and its selectivity for LC3s (Satoo et al., 2009). This residue corresponds to ATG4A^I233 and, when substituted for leucine, gives ATG4A^I233L the ability to efficiently process LC3 proteins, whereas without this mutation it preferentially processes GABARAPs (Satoo et al., 2009). Moreover, the ATG8–ATG4 interaction is necessary for the de-ATG8ylation of cellular proteins, as an LC3B-G^Q116P mutant that cannot bind to ATG4 leads to widespread ATG8ylation (Agrotis et al., 2019). Altogether, these observations hint toward a common mechanism of ATG8 cleavage that regulates priming, de-lipidation, and de-ATG8ylation.Mitochondrial damage promotes ATG8ylationATG8ylation of cellular proteins appears to be enhanced by mitochondrial depolarization and inhibition of the lysosomal V-type ATPase (Agrotis et al., 2019; Nguyen et al., 2021). This may be the consequence of acute ATG4A and -B inhibition, given that cells lacking all ATG4 isoforms display an increased abundance of ATG8ylated proteins and are insensitive to further increase by mitochondrial depolarization or lysosomal V-type ATPase inhibition (Agrotis et al., 2019; Nguyen et al., 2021). Indeed, mitochondrial depolarization leads to activation of ULK1, which phosphorylates ATG4B^S316 to inhibit its protease activity (Pengo et al., 2017). Similarly, mitochondrial depolarization stimulates TBK1 activation, which prevents de-lipidation of ATG8s by blocking the ATG8–ATG4 interaction through phosphorylation of LC3C^S93/S96 and GABARAP-L2^S87/S88 (Herhaus et al., 2020; Richter et al., 2016). As such, ATG8 phosphorylation may render ATG8ylated substrates more resistant to de-ATG8ylation by ATG4s. This may be analogous to how chains of phosphorylated ubiquitin^S65 are more resistant to hydrolysis by deubiquitinating enzymes than unphosphorylated ones (Wauer et al., 2015). Moreover, ATG8ylation is insensitive to nutrient deprivation and pharmacological inhibition of mTOR, which rules out a functional contribution of this process to starvation-induced autophagy (Agrotis et al., 2019). Therefore, ATG8ylation may be a unique aspect of mitophagy (and perhaps also other forms of selective autophagy) given that depolarization potently activates Parkin-dependent mitophagy (Agrotis et al., 2019; Nguyen et al., 2021).Substrates of ATG8ylationBased on ATG8⁺ smearing, ATG4 regulates the de-ATG8ylation of numerous proteins (Agrotis et al., 2019; Nguyen et al., 2021). For the majority, their identity, induced structural and functional changes, and the cellular contexts during which these modifications occur await exploration. Considering that the ATG8 interactome is well characterized, it is likely that at least some ATG8ylated proteins have been mistaken for ATG8-binding partners (Behrends et al., 2010). Given their E2- and E3-like roles in ATG8 lipidation, it is remarkable that ATG3 and ATG16L1 are themselves modified by ATG8ylation (Agrotis et al., 2019; Hanada et al., 2007; Ichimura et al., 2000; Nguyen et al., 2021). Lysine mutagenesis indicates that ATG3^K243 is the “acceptor” site for ATG8ylation (Agrotis et al., 2019). ATG3^K243 is essential for its conjugation to either LC3B or ATG12 and is required for autophagosomes to form around damaged mitochondria (Agrotis et al., 2019; Radoshevich et al., 2010). This also raises the possibility that key functions originally attributed to ATG3–ATG12 conjugation may be, at least in part, due to ATG3–ATG8 conjugation. Because multiple high-molecular-weight species of ATG3 are enriched following immunoprecipitation of primed LC3B-G from cells lacking ATG4B, it is likely that ATG3 is either mono-ATG8ylated at several sites or poly-ATG8ylated (Agrotis et al., 2019). ATG8ylation of ATG3 may also reflect the stabilization of its E2-like intermediate (Ichimura et al., 2000). ATG8ylation of ATG16L1 may regulate whether canonical or noncanonical autophagy pathways are activated (Durgan et al., 2021; Nguyen et al., 2021). In line with this possibility, the WD40 domain mutant of ATG16L1^K490A prevents lipidation of ATG8s with phosphatidylserine (i.e., during noncanonical autophagy pathways) but not phosphatidylethanolamine (i.e., during canonical autophagy; Durgan et al., 2021). Moreover, given that ATG8ylation of protein targets correlates with the activation of mitophagy, it is tempting to speculate that it may stimulate the E2-/E3-like activity of the ATG8 conjugation machinery to amplify mitochondrial capture and destruction.Concluding remarksThe finding that numerous cellular proteins are modified by ATG8ylation poses several questions about how signaling networks are coordinated during selective autophagy (i.e., mitophagy). Whether ATG8ylation is augmented by mitochondrial injury per se or is the consequence of mitophagy activation is yet to be determined, as is whether this phenomenon occurs during other types of selective autophagy (e.g., ER-phagy, ribophagy, and lysophagy; Kirkin and Rogov, 2019; Fig. 1 C). While the in vivo relevance of ATG8ylation is not yet understood, it is plausible that this process could be altered in diseases with defective mitophagy (e.g., Parkinson’s disease and atherosclerosis). Exploring the mechanistic aspects of ATG8ylation (e.g., ATG8 ligases and regulatory proteins, linkage types, acceptor sites, etc.) and de-ATG8ylation by ATG4 will improve our understanding about how this modifier alters the structure and biological function of cellular proteins (Fig. 1 C). By identifying ATG8ylated substrates, or the ATG8ylome, insights into whether ATG8ylation is a ubiquitous epiphenomenon or a post-translational modification that is selective to proteins of distinct biological function(s) will become clearer (Fig. 1 C). Considering the similarity of ATG8s with bona fide modifier proteins (e.g., ubiquitin and ubiquitin-like proteins) and the diversity of their substrates (e.g., lipid species and proteins), only now are we beginning to understand the functional complexities of the ATG8 protein family.     

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.     

Maturation Stress Generation in Poplar Tension Wood Studied by Synchrotron Radiation Microdiffraction

Tension wood is widespread in the organs of woody plants. During its formation, it generates a large tensile mechanical stress, called maturation stress. Maturation stress performs essential biomechanical functions such as optimizing the mechanical resistance of the stem, performing adaptive movements, and ensuring long-term stability of growing plants. Although various hypotheses have recently been proposed, the mechanism generating maturation stress is not yet fully understood. In order to discriminate between these hypotheses, we investigated structural changes in cellulose microfibrils along sequences of xylem cell differentiation in tension and normal wood of poplar (Populus deltoides × Populus trichocarpa ‘I45-51’). Synchrotron radiation microdiffraction was used to measure the evolution of the angle and lattice spacing of crystalline cellulose associated with the deposition of successive cell wall layers. Profiles of normal and tension wood were very similar in early development stages corresponding to the formation of the S1 and the outer part of the S2 layer. The microfibril angle in the S2 layer was found to be lower in its inner part than in its outer part, especially in tension wood. In tension wood only, this decrease occurred together with an increase in cellulose lattice spacing, and this happened before the G-layer was visible. The relative increase in lattice spacing was found close to the usual value of maturation strains, strongly suggesting that microfibrils of this layer are put into tension and contribute to the generation of maturation stress.Wood cells are produced in the cambium at the periphery of the stem. The formation of the secondary wall occurs at the end of cell elongation by the deposition of successive layers made of cellulose microfibrils bounded by an amorphous polymeric matrix. Each layer has a specific chemical composition and is characterized by a particular orientation of the microfibrils relative to the cell axis (Mellerowicz and Sundberg, 2008). Microfibrils are made of crystalline cellulose and are by far the stiffest constituent of the cell wall. The microfibril angle (MFA) in each layer is determinant for cell wall architecture and wood mechanical properties.During the formation of wood cells, a mechanical stress of a large magnitude, known as “maturation stress” or “growth stress” (Archer, 1986; Fournier et al., 1991), occurs in the cell walls. This stress fulfills essential biomechanical functions for the tree. It compensates for the comparatively low compressive strength of wood and thus improves the stem resistance against bending loads. It also provides the tree with a motor system (Moulia et al., 2006), necessary to maintain the stem at a constant angle during growth (Alméras and Fournier, 2009) or to achieve adaptive reorientations. In angiosperms, a large tensile maturation stress is generated by a specialized tissue called “tension wood.” In poplar (Populus deltoides × Populus trichocarpa), as in most temperate tree species, tension wood fibers are characterized by the presence of a specific layer, called the G-layer (Jourez et al., 2001; Fang et al., 2008), where the matrix is almost devoid of lignin (Pilate et al., 2004) and the microfibrils are oriented parallel to the fiber axis (Fujita et al., 1974). This type of reaction cell is common in plant organs whose function involves the bending or contraction of axes, such as tendrils, twining vines (Bowling and Vaughn, 2009), or roots (Fisher, 2008).The mechanism at the origin of tensile maturation stress has been the subject of a lot of controversy and is still not fully understood. However, several recent publications have greatly improved our knowledge about the ultrastructure, chemical composition, molecular activity, mechanical state, and behavior of tension wood. Different models have been proposed and discussed to explain the origin of maturation stress (Boyd, 1972; Bamber, 1987, 2001; Okuyama et al., 1994, 1995; Yamamoto, 1998, 2004; Alméras et al., 2005, 2006; Bowling and Vaughn, 2008; Goswami et al., 2008; Mellerowicz et al., 2008). The specific organization of the G-layer suggests a tensile force induced in the microfibrils during the maturation process. Different hypotheses have been proposed to explain this mechanism, such as the contraction of amorphous zones within the cellulose microfibrils (Yamamoto, 2004), the action of xyloglucans during the formation of microfibril aggregates (Nishikubo et al., 2007; Mellerowicz et al., 2008), and the effect of changes in moisture content stimulated by pectin-like substances (Bowling and Vaughn, 2008). A recent work (Goswami et al., 2008) argued an alternative model, initially proposed by Münch (1938), which proposed that the maturation stress originates in the swelling of the G-layer during cell maturation and is transmitted to the adjacent secondary layers, where the larger MFAs allow an efficient conversion of lateral stress into axial tensile stress. Although the proposed mechanism is not consistent with the known hygroscopic behavior of tension wood, which shrinks when it dries and not when it takes up water (Clair and Thibaut, 2001; Fang et al., 2007; Clair et al., 2008), this hypothesis focused attention on the possible role of cell wall layers other than the G-layer. As a matter of fact, many types of wood fibers lacking a G-layer are known to produce axial tensile stress, such as normal wood of angiosperms and conifers (Archer, 1986) and the tension wood of many tropical species (Onaka, 1949; Clair et al., 2006b; Ruelle et al., 2007), so that mechanisms strictly based on an action of the G-layer cannot provide a general explanation for the origin of tensile maturation stress in wood.In order to further understanding, direct observations of the mechanical state of the different cell wall layers and their evolution during the formation of the tension wood fibers are needed. X-ray diffraction can be used to investigate the orientation of microfibrils (Cave, 1966, 1997a, 1997b; Peura et al., 2007, 2008a, 2008b) and the lattice spacing of crystalline cellulose. The axial lattice spacing d₀₀₄ is the distance between successive monomers along a cellulose microfibril and reflects its state of mechanical stress (Clair et al., 2006a; Peura et al., 2007). If cellulose microfibrils indeed support a tensile stress, they should be found in an extended state of deformation. Under this assumption, the progressive development of maturation stress during the cell wall formation should be accompanied by an increase in cellulose lattice spacing. Synchrotron radiation allows a reduction in the size of the x-ray beam to some micrometers while retaining a strong signal, whereby diffraction analysis can be performed at a very local scale (Riekel, 2000). This technique has been used to study sequences of wood cell development (Hori et al., 2000; Müller et al., 2002). In this study, we report an experiment where a microbeam was used to analyze the structural changes of cellulose in the cell wall layers of tension wood and normal wood fibers along the sequence of xylem cell differentiation extending from the cambium to mature wood (Fig. 1). The experiment was designed to make this measurement in planta, in order to minimize sources of mechanical disturbance and be as close as possible to the native mechanical state (Clair et al., 2006a). The 200 and 004 diffraction patterns of cellulose were analyzed to investigate the process of maturation stress generation in tension wood.Open in a separate windowFigure 1.Schematic of the experimental setup, showing the x-ray beam passing perpendicular to the longitudinal-radial plane of wood and the contribution of the 004 and 200 crystal planes to the diffraction pattern recorded by the camera. [See online article for color version of this figure.]     

Mitofusins: ubiquitylation promotes fusion

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

AUXIN RESPONSE FACTOR17 Is Essential for Pollen Wall Pattern Formation in Arabidopsis

In angiosperms, pollen wall pattern formation is determined by primexine deposition on the microspores. Here, we show that AUXIN RESPONSE FACTOR17 (ARF17) is essential for primexine formation and pollen development in Arabidopsis (Arabidopsis thaliana). The arf17 mutant exhibited a male-sterile phenotype with normal vegetative growth. ARF17 was expressed in microsporocytes and microgametophytes from meiosis to the bicellular microspore stage. Transmission electron microscopy analysis showed that primexine was absent in the arf17 mutant, which leads to pollen wall-patterning defects and pollen degradation. Callose deposition was also significantly reduced in the arf17 mutant, and the expression of CALLOSE SYNTHASE5 (CalS5), the major gene for callose biosynthesis, was approximately 10% that of the wild type. Chromatin immunoprecipitation and electrophoretic mobility shift assays showed that ARF17 can directly bind to the CalS5 promoter. As indicated by the expression of DR5-driven green fluorescent protein, which is an synthetic auxin response reporter, auxin signaling appeared to be specifically impaired in arf17 anthers. Taken together, our results suggest that ARF17 is essential for pollen wall patterning in Arabidopsis by modulating primexine formation at least partially through direct regulation of CalS5 gene expression.In angiosperms, the pollen wall is the most complex plant cell wall. It consists of the inner wall, the intine, and the outer wall, the exine. The exine is further divided into sexine and nexine layers. The sculptured sexine includes three major parts: baculum, tectum, and tryphine (Heslop-Harrison, 1971; Piffanelli et al., 1998; Ariizumi and Toriyama, 2011; Fig. 1A). Production of a functional pollen wall requires the precise spatial and temporal cooperation of gametophytic and sporophytic tissues and metabolic events (Blackmore et al., 2007). The intine layer is controlled gametophytically, while the exine is regulated sporophytically. The sporophytic tapetum cells provide material for pollen wall formation, while primexine determines pollen wall patterning (Heslop-Harrison, 1968).Open in a separate windowFigure 1.Schematic representation of the pollen wall and primexine development. A, The innermost layer adjacent to the plasma membrane is the intine. The bacula (Ba), tectum (Te), and tryphine (T) make up the sexine layer. The nexine is located between the intine and the sexine layers. The exine includes the nexine and sexine layers. B, Primexine (Pr) appears between callose (Cl) and plasma membrane (Pm) at the early tetrad stage (left panel). Subsequently, the plasma membrane becomes undulated (middle panel) and sporopollenin deposits on the peak of the undulated plasma membrane to form bacula and tectum (right panel).After meiosis, four microspores were encased in callose to form a tetrad. Subsequently, the primexine develops between the callose layer and the microspore membrane (Fig. 1B), and the microspore plasma membrane becomes undulated (Fig. 1B; Fitzgerald and Knox, 1995; Southworth and Jernstedt, 1995). Sporopollenin precursors then accumulate on the peak of the undulated microspore membrane to form the bacula and tectum (Fig. 1B; Fitzgerald and Knox, 1995). After callose degradation, individual microspores are released from the tetrad, and the bacula and tectum continue to grow into exine with further sporopollenin deposition (Fitzgerald and Knox, 1995; Blackmore et al., 2007).The callose has been reported to affect primexine deposition and pollen wall pattern formation. The peripheral callose layer, secreted by the microsporocyte, acts as the mold for primexine (Waterkeyn and Bienfait, 1970; Heslop-Harrison, 1971). CALLOSE SYNTHASE5 (CalS5) is the major enzyme responsible for the biosynthesis of the callose peripheral of the tetrad (Dong et al., 2005; Nishikawa et al., 2005). Mutation of Cals5 and abnormal CalS5 pre-mRNA splicing resulted in defective peripheral callose deposition and primexine formation (Dong et al., 2005; Nishikawa et al., 2005; Huang et al., 2013). Besides CalS5, four membrane-associated proteins have also been reported to be involved in primexine formation: DEFECTIVE EXINE FORMATION1 (DEX1; Paxson-Sowders et al., 1997, 2001), NO EXINE FORMATION1 (NEF1; Ariizumi et al., 2004), RUPTURED POLLEN GRAIN1 (RPG1; Guan et al., 2008; Sun et al., 2013), and NO PRIMEXINE AND PLASMA MEMBRANE UNDULATION (NPU; Chang et al., 2012). Mutation of DEX1 results in delayed primexine formation (Paxson-Sowders et al., 2001). The primexine in nef1 is coarse compared with the wild type (Ariizumi et al., 2004). The loss-of-function rpg1 shows reduced primexine deposition (Guan et al., 2008; Sun et al., 2013), while the npu mutant does not deposit any primexine (Chang et al., 2012). Recently, it was reported that Arabidopsis (Arabidopsis thaliana) CYCLIN-DEPENDENT KINASE G1 (CDKG1) associates with the spliceosome to regulate the CalS5 pre-mRNA splicing for pollen wall formation (Huang et al., 2013). Clearly, disrupted primexine deposition leads to aberrant pollen wall patterning and ruptured pollen grains in these mutants.The plant hormone auxin has multiple roles in plant reproductive development (Aloni et al., 2006; Sundberg and Østergaard, 2009). Knocking out the two auxin biosynthesis genes, YUC2 and YUC6, caused an essentially sterile phenotype in Arabidopsis (Cheng et al., 2006). Auxin transport is essential for anther development; defects in auxin flow in anther filaments resulted in abnormal pollen mitosis and pollen development (Feng et al., 2006). Ding et al. (2012) showed that the endoplasmic reticulum-localized auxin transporter PIN8 regulates auxin homeostasis and male gametophyte development in Arabidopsis. Evidence for the localization, biosynthesis, and transport of auxin indicates that auxin regulates anther dehiscence, pollen maturation, and filament elongation during late anther development (Cecchetti et al., 2004, 2008). The role of auxin in pollen wall development has not been reported.The auxin signaling pathway requires the auxin response factor (ARF) family proteins (Quint and Gray, 2006; Guilfoyle and Hagen, 2007; Mockaitis and Estelle, 2008; Vanneste and Friml, 2009). ARF proteins can either activate or repress the expression of target genes by directly binding to auxin response elements (AuxRE; TGTCTC/GAGACA) in the promoters (Ulmasov et al., 1999; Tiwari et al., 2003). The Arabidopsis ARF family contains 23 members. A subgroup in the ARF family, ARF10, ARF16, and ARF17, are targets of miRNA160 (Okushima et al., 2005b; Wang et al., 2005). Plants expressing miR160-resistant ARF17 exhibited pleiotropic developmental defects, including abnormal stamen structure and reduced fertility (Mallory et al., 2005). This indicates a potential role for ARF17 in plant fertility, although the detailed function remains unknown. In addition, ARF17 was also proposed to negatively regulate adventitious root formation (Sorin et al., 2005; Gutierrez et al., 2009), although an ARF17 knockout mutant was not reported and its phenotype is unknown.In this work, we isolated and characterized a loss-of-function mutant of ARF17. Results from cytological observations suggest that ARF17 controls callose biosynthesis and primexine deposition. Consistent with this, the ARF17 protein is highly abundant in microsporocytes and tetrads. Furthermore, we demonstrate that the ARF17 protein is able to bind the promoter region of CalS5. Our results suggest that ARF17 regulates pollen wall pattern formation in Arabidopsis.     

Variability and Constancy in Cellular Growth of Arabidopsis Sepals

Growth of tissues is highly reproducible; yet, growth of individual cells in a tissue is highly variable, and neighboring cells can grow at different rates. We analyzed the growth of epidermal cell lineages in the Arabidopsis (Arabidopsis thaliana) sepal to determine how the growth curves of individual cell lineages relate to one another in a developing tissue. To identify underlying growth trends, we developed a continuous displacement field to predict spatially averaged growth rates. We showed that this displacement field accurately describes the growth of sepal cell lineages and reveals underlying trends within the variability of in vivo cellular growth. We found that the tissue, individual cell lineages, and cell walls all exhibit growth rates that are initially low, accelerate to a maximum, and decrease again. Accordingly, these growth curves can be represented by sigmoid functions. We examined the relationships among the cell lineage growth curves and surprisingly found that all lineages reach the same maximum growth rate relative to their size. However, the cell lineages are not synchronized; each cell lineage reaches this same maximum relative growth rate but at different times. The heterogeneity in observed growth results from shifting the same underlying sigmoid curve in time and scaling by size. Thus, despite the variability in growth observed in our study and others, individual cell lineages in the developing sepal follow similarly shaped growth curves.Cells undergo multiple rounds of growth and division to create reproducible tissues. In some plant tissues, such as expanding cotyledons, reproducibility can occur on a cellular level during specific intervals of development, where cotyledon cells exhibit uniform cellular growth (Zhang et al., 2011). However, several studies on cell division and growth in other developing plant tissues have demonstrated that plant cells exhibit considerable cell-to-cell variability during development (Meyer and Roeder, 2014). For example, in both the Arabidopsis (Arabidopsis thaliana) meristem and leaf epidermis, cells show spatiotemporal variation in individual cell growth rates (GRs; Asl et al., 2011; Elsner et al., 2012; Kierzkowski et al., 2012; Uyttewaal et al., 2012). Furthermore, cell divisions have been observed with marked randomness in their timing and orientation (Roeder et al., 2010; Besson and Dumais, 2011; Roeder, 2012). In this study, we identify a hidden, underlying pattern in the seemingly random GR (Box 1) of cells during the formation of sepals in Arabidopsis.Open in a separate windowBox 1.Definitions of GR terms. (For details on the calculations, see “Materials and Methods.”)Plant cell growth is defined as an increase in cell size due to an irreversible expansion of the cell wall. Neighboring cells physically accommodate one another during plant growth because their cell walls are glued together with a pectin-rich middle lamella, which prevents cell mobility. The cell wall is a thin, stiff layer composed of a polymer matrix including cellulose, hemicellulose, and pectin (Somerville et al., 2004; Cosgrove, 2005). Plant cells change their size and shape by modifying their turgor pressure and/or the mechanical properties of their walls, such as elasticity, plasticity, and extensibility. Growing plant cells exert forces on their neighbors through their walls, and cell wall stresses created by these forces feed back to alter the growth anisotropy (Hamant et al., 2008; Sampathkumar et al., 2014). Although these feedbacks can coordinate growth, they may also amplify differences in growth between neighboring cells (Uyttewaal et al., 2012).Two competing computational models have proposed explanations of the cellular heterogeneity observed in growing tissues by making different assumptions about how cells grow. In the first, it is assumed that relative growth rates (RGRs) of all cells are uniform in space and time, whereas variation in the timing of division causes the heterogeneity of cell sizes (Roeder et al., 2010). This model suggests that cell divisions cut the sepal into semiindependent cells, which grow uniformly within the expanding organ (Kaplan and Hagemann, 1991). The second model postulates the reverse process: timing of cell division is uniform, but cellular growth is variable and depends on the size of the cell (Asl et al., 2011). This model suggests that cells are autonomous. Currently, there is biological evidence for both models. Variability in cell division timing is observed in sepals and meristems, whereas variability in cellular GRs has been observed in leaves and meristem cells (Reddy et al., 2004; Roeder et al., 2010; Asl et al., 2011; Elsner et al., 2012; Kierzkowski et al., 2012; Uyttewaal et al., 2012). Thus, the debate on how the growth of individual cells within an organ relates to one another remains unresolved.The identification of underlying patterns in noisy cellular growth processes is challenging. Technical difficulties include the capability for cellular-resolution imaging of the tissue at sufficiently small time intervals. Previous studies (Zhang et al., 2011; Elsner et al., 2012; Kierzkowski et al., 2012) did not image and track individual cells, or they had a coarse time resolution, with 11- to 48-h intervals between images, which may have hidden important temporal dynamics. We studied growing cells in the Arabidopsis sepal, which allows for live imaging with cellular resolution at 6-h intervals (Roeder et al., 2010). The sepal is the leaf-like outermost floral organ of Arabidopsis (Fig. 1) with four sepals of stereotypical size produced per flower. Its accessibility for live imaging makes the sepal an excellent system for studying organogenesis (Roeder et al., 2010, 2011, 2012; Qu et al., 2014). Sepals exhibit high cellular variability in the timing of division and endoreduplication, an alternative cell cycle in which a cell replicates its DNA but fails to divide (Roeder et al., 2010). Furthermore, quantifying cell growth in sepals may shed light on growth mechanisms of other plant organs, such as leaves (Poethig and Sussex, 1985; Roeder et al., 2010).Open in a separate windowFigure 1.Diverse sizes of Arabidopsis sepal cells. A, Four sepals (s) are the outermost green leaf-like floral organs in Arabidopsis. B and C, Scanning electron micrographs of a mature Arabidopsis sepal show that the outer epidermal cells have a wide range of sizes. Asterisks mark some of the largest cells (giant cells) that can span 1/4 the length of the sepal. Scale = 100 µm.Another key challenge in analyzing cellular growth is the identification of trends in noisy data. Inaccuracies in data acquisition, such as segmentation errors, and noisy growth of individual cells can hide meaningful spatiotemporal trends in growth. GRs measured over longer time intervals will have reduced noise, but they may also obscure important temporal dynamics. Alternatively, previous studies have examined growth of the whole organ or its subregions to avoid cellular noise (De Veylder et al., 2001; Mündermann et al., 2005; Rolland-Lagan et al., 2005, 2014; Kuchen et al., 2012; Remmler and Rolland-Lagan, 2012). However, precise cellular patterns are not resolved. In our study, we use cellular resolution data to define spatially averaged kinematics while keeping the full temporal resolution to identify course-grained spatial trends in the dynamics of cellular growth (Box 1).We analyze the relationships among the growth of individual cell lineages in a developing Arabidopsis sepal by live imaging and computational analyses. We have developed continuous low-order displacement fields to represent the spatially averaged kinematics of the sepal (Box 1). We find that the growth of the tissue surface area, cell lineage area, and wall length follows S curves, suggesting that their GRs vary over time. Additionally, we find that there is a linear correlation between the maximum GR (i.e. size increase per hour) and the size of the cell. We furthermore find that each sepal cell lineage reaches the same maximum RGR (i.e. GR divided by size). However, each cell reaches the maximum RGR at a different time during its development, generating the observed heterogeneity. Thus, we find underlying similarities in the growth curves of sepal cells.     

Ovine Model for Critical-Size Tibial Segmental Defects

A segmental tibial defect model in a large animal can provide a basis for testing materials and techniques for use in nonunions and severe trauma. This study reports the rationale behind establishing such a model and its design and conclusions. After ethics approval of the study, aged ewes (older than 5 y; n = 12) were enrolled. A 5-cm mid diaphyseal osteoperiosteal defect was made in the left tibia and was stabilized by using an 8-mm stainless-steel cross-locked intramedullary nail. Sheep were euthanized at 12 wk after surgery and evaluated by using radiography, microCT, and soft-tissue histology techniques. Radiology confirmed a lack of hard tissue callus bridging across the defect. Volumetric analysis based on microCT showed bone growth across the 16.5-cm³ defect of 1.82 ± 0.94 cm³. Histologic sections of the bridging tissues revealed callus originating from both the periosteal and endosteal surfaces, with fibrous tissue completing the bridging in all instances. Immunohistochemistry was used to evaluate the quality of the healing response. Clinical, radiographic, and histologic union was not achieved by 12 wk. This model may be effective for the investigation of surgical techniques and healing adjuncts for nonunion cases, where severe traumatic injury has led to significant bone loss.Abbreviations: BMP2, bone morphogenic protein 2; CATK, cathepsin K; VEGF, vascular endothelial growth factorThe human tibia is the most frequently broken long bone, often with significant bone loss.⁴ Segmental tibial defects can occur as a result of large tumor removal, trauma such as motor vehicle accidents, and more recently, blast injuries as seen with the escalating number of global conflicts. Treatment of these large bone and surrounding soft tissue defects is an ongoing, costly, and challenging clinical problem; no surgical technique has currently achieved preeminence.⁴ The general consensus on factors that affect healing include concomitant disease, age, and degree of trauma.⁵ When the first 2 factors, which are patient-related, are removed from the equation, healing is influenced by the size, anatomic location, and soft-tissue coverage of the defect. The ability to study these situations in a well-controlled, robust, and reproducible preclinical model would be advantageous to help establish effective surgical techniques and evaluate implants and materials.A literature review revealed that many ovine models for bone defects have been used, but all have limitations^{6,12,14,15,20,21,24,25,27,31,37,39,40} (Figure 1). Variations in protocols, such as age of the animals, size of the defect, and the bone and stabilization techniques used, limit meaningful comparison between studies.^33,34 Although some studies have investigated material performance in the healing of defects, they did not rigorously quantify control defects,^17,20 and others used no controls at all.³⁹ There is often no explanation regarding the use of a particular defect size, leading to the question of whether the defect size was critical.²⁴ The choice of bone used has been also varied; the femur,¹⁵ tibia,³⁷ and metatarsus⁴⁰ have all been studied. A noncritical-size defect implies that healing would eventually occur without the presence of any graft materials. One study,¹² for example, used a 3-cm defect at an average of 1.8 times the diameter of the tibias in question and found that empty controls achieved as much as 26% of the stiffness of an intact tibia after 12 wk. Stabilization methods include plating,^21,40 external fixtures,²⁰ intramedullary nails,^6,16 and a combination of intramedullary nails and plating.³⁷Open in a separate windowFigure 1.A limited summary of the many studies where a segmental tibial has been used with their references.The criteria used in the present study for a critical-size segmental tibial defect model were based on the following factors. The ovine tibia closely resembles that of the human tibia in terms of size, shape, and physical properties and is commonly used when studying human orthopedic diseases.^26,34 Intramedullary nailing has become the most commonly used method of tibial fracture fixation in human orthopedic surgery.^8,22 An 8-mm intramedullary nail is commonly used in the treatment of human fractures, further confirming the size similarity between the ovine and human tibiae.¹⁹The aim of this study was to establish and characterize a preclinical ovine 5-cm osteoperiosteal critical-size tibial segmental defect model in mature sheep. The endpoints included those commonly used clinically, such as radiography and microCT. Histology to investigate the degree of healing and immunohistochemistry to characterize the healing process were included to complete the evaluation process.     

Endemic Viruses of Squirrel Monkeys (Saimiri spp.)

Nonhuman primates are the experimental animals of choice for the study of many human diseases. As such, it is important to understand that endemic viruses of primates can potentially affect the design, methods, and results of biomedical studies designed to model human disease. Here we review the viruses known to be endemic in squirrel monkeys (Saimiri spp.). The pathogenic potential of these viruses in squirrel monkeys that undergo experimental manipulation remains largely unexplored but may have implications regarding the use of squirrel monkeys in biomedical research.Abbreviations: HTLV1, human T-cell leukemia virus type 1; HVS, herpesvirus saimiri; IPF, idiopathic pulmonary fibrosis; SaHV, Saimiriine herpesvirus; SFV, simian foamy virus; SM-CMV, squirrel monkey cytomegalovirus; SMPyV, squirrel monkey polyomavirus; SMRV, squirrel monkey retrovirusThe similarity between the nonhuman primate and human immune systems is a key advantage in the use of nonhuman primates compared with other mammalian models of human disease.^{13,71,88,94,103,113,125} In addition, the diversity of environmental and infectious disease agents encountered by primates is similar to that of humans, providing nonhuman primates a comparable level of biologic complexity.¹ Old World primates, such as macaques and baboons, and New World primates, including squirrel monkeys and marmosets, are commonly used in biomedical research. Squirrel monkeys (Saimiri spp.) are neotropical primates native to the forests of Central and South America. Of the 7 species of squirrel monkey, 3 (S. oerstedii, S. vanzolinii, and S. ustus) are classified as endangered, vulnerable to extinction in the wild, or near threatened, whereas the remaining 4 (S. boliviensis, S. cassiquiarensis, S. macrodon, and S. sciureus) are not endangered, although the S. cassiquiarensis albigena subspecies is near threatened^52,81 (Figure 1). In South America, where squirrel monkeys are indigenous, breeding colonies of S. sciureus have been maintained at the Pasteur Institute in French Guiana and at the Oswaldo Cruz Foundation in Brazil.^7,12 In the United States, the Squirrel Monkey Breeding and Research Resource, an NIH-sponsored national research resource, maintains breeding colonies for S. boliviensis boliviensis, S. sciureus sciureus, and S. boliviensis peruviensis.Open in a separate windowFigure 1.Taxonomy of Saimiri species with associated IUCN designations.^52,81Squirrel monkeys adapt easily to laboratory housing and can be housed in smaller spaces than can Old World primates.¹ Unlike when working with Old World primates, particularly macaques, no additional personnel protective equipment is necessary when working with squirrel monkeys beyond that recommended for working with other New World primates.⁹² Their small size, combined with the reduced need for personnel protective equipment during handling, make squirrel monkeys attractive species for model development and for studies of viral pathogenesis, which cost approximately 30% to 40% less than comparable studies in macaques.¹ The likelihood of zoonotic transmission of infectious pathogens is considerably less than that associated with macaques and the risk of Macacine herpesvirus 1 (B virus) is nonexistent, given that neotropical primates do not harbor this lethal virus.¹ These factors are increasingly important in the current climate of limited grant funding for biomedical research and emphasis on safety for laboratory personnel. The limited availability of immunologic reagents with specificity for neotropical primates has hindered broader use of squirrel monkeys in biomedical research, compared with that of the more commonly used Old World primates. In addition, the small size of neotropical primates limits the volume of blood that can be collected at any one time. To abrogate these limitations, the NIH Nonhuman Primate Reagent Resource (www.nhpreagents.org) provides an increasing repertoire of agents that have been characterized for immunologic studies of neotropical primates.⁸⁹Squirrel monkeys are used in numerous aspects of biomedical research, including studies of viral persistence, neuroendocrinology, infectious diseases, cancer treatments, vaccine development, gene expression, and reproductive physiology.¹¹⁷ The similarity between the squirrel monkey immune system and that of humans means that, as with macaques, there is a high likelihood that research outcomes will recapitulate what occurs in human diseases.^13,71,87,94 This is particularly true for the study of several notable infectious diseases, including malaria, Creutzfeldt–Jakob disease, and human T-cell leukemia virus type 1 (HTLV1) infection.^19,56,128 For these diseases, squirrel monkeys are the model system of choice for studying pathogenesis, experimental treatments, and strategies for prevention.Squirrel monkeys are recognized as some of the most susceptible nonhuman primate species for the experimental transmission of Creutzfeldt–Jakob disease and other transmissible spongiform encephalopathies that cause chronic wasting disease.^11,72,98,130 The experimental infection of squirrel monkeys with HTLV1 has led to their use in vaccine development and chemotherapy research directed against HTLV1.^44,57,58,82 In addition, squirrel monkeys are an important model for studying the immunology of malaria and for testing vaccines against several Plasmodium species.^19,20,68,114 Furthermore, squirrel monkeys have been used in pharmacologic research to raise HDL levels to prevent atherosclerosis and reduce the risk of coronary heart disease.⁶ As the use of squirrel monkeys increases, especially for infectious disease research, accurate information about the endemic viral infections of squirrel monkeys is needed because of the potential for zoonotic transfer of these viruses to humans (and vice versa) and to understand the potential influence these agents may have on research involving other infectious pathogens diseases and immunosuppressive drugs.