The nuclease activity of DNA2 promotes exonuclease 1–independent mismatch repair

The DNA mismatch repair (MMR) system is a major DNA repair system that corrects DNA replication errors. In eukaryotes, the MMR system functions via mechanisms both dependent on and independent of exonuclease 1 (EXO1), an enzyme that has multiple roles in DNA metabolism. Although the mechanism of EXO1-dependent MMR is well understood, less is known about EXO1-independent MMR. Here, we provide genetic and biochemical evidence that the DNA2 nuclease/helicase has a role in EXO1-independent MMR. Biochemical reactions reconstituted with purified human proteins demonstrated that the nuclease activity of DNA2 promotes an EXO1-independent MMR reaction via a mismatch excision-independent mechanism that involves DNA polymerase δ. We show that DNA polymerase ε is not able to replace DNA polymerase δ in the DNA2-promoted MMR reaction. Unlike its nuclease activity, the helicase activity of DNA2 is dispensable for the ability of the protein to enhance the MMR reaction. Further examination established that DNA2 acts in the EXO1-independent MMR reaction by increasing the strand-displacement activity of DNA polymerase δ. These data reveal a mechanism for EXO1-independent mismatch repair.

The mismatch repair (MMR) system has been conserved from bacteria to humans (1, 2). It promotes genome stability by suppressing spontaneous and DNA damage-induced mutations (1, 3, 4, 5, 6, 7, 8, 9, 10, 11). The key function of the MMR system is the correction of DNA replication errors that escape the proofreading activities of replicative DNA polymerases (1, 4, 5, 6, 7, 8, 9, 10, 12). In addition, the MMR system removes mismatches formed during strand exchange in homologous recombination, suppresses homeologous recombination, initiates apoptosis in response to irreparable DNA damage caused by several anticancer drugs, and contributes to instability of triplet repeats and alternative DNA structures (1, 4, 5, 7, 8, 9, 10, 11, 13, 14, 15, 16, 17, 18). The principal components of the eukaryotic MMR system are MutSα (MSH2-MSH6 heterodimer), MutLα (MLH1-PMS2 heterodimer in humans and Mlh1-Pms1 heterodimer in yeast), MutSβ (MSH2-MSH3 heterodimer), proliferating cell nuclear antigen (PCNA), replication factor C (RFC), exonuclease 1 (EXO1), RPA, and DNA polymerase δ (Pol δ). Loss-of-function mutations in the MSH2, MLH1, MSH6, and PMS2 genes of the human MMR system cause Lynch and Turcot syndromes, and hypermethylation of the MLH1 promoter is responsible for ∼15% of sporadic cancers in several organs (19, 20). MMR deficiency leads to cancer initiation and progression via a multistage process that involves the inactivation of tumor suppressor genes and action of oncogenes (21).MMR occurs behind the replication fork (22, 23) and is a major determinant of the replication fidelity (24). The correction of DNA replication errors by the MMR system increases the replication fidelity by ∼100 fold (25). Strand breaks in leading and lagging strands as well as ribonucleotides in leading strands serve as signals that direct the eukaryotic MMR system to remove DNA replication errors (26, 27, 28, 29, 30). MMR is more efficient on the lagging than the leading strand (31). The substrates for MMR are all six base–base mismatches and 1 to 13-nt insertion/deletion loops (25, 32, 33, 34). Eukaryotic MMR commences with recognition of the mismatch by MutSα or MutSβ (32, 34, 35, 36). MutSα is the primary mismatch-recognition factor that recognizes both base–base mismatches and small insertion/deletion loops whereas MutSβ recognizes small insertion/deletion loops (32, 34, 35, 36, 37). After recognizing the mismatch, MutSα or MutSβ cooperates with RFC-loaded PCNA to activate MutLα endonuclease (38, 39, 40, 41, 42, 43). The activated MutLα endonuclease incises the discontinuous daughter strand 5′ and 3′ to the mismatch. A 5'' strand break formed by MutLα endonuclease is utilized by EXO1 to enter the DNA and excise a discontinuous strand portion encompassing the mismatch in a 5''→3′ excision reaction stimulated by MutSα/MutSβ (38, 44, 45). The generated gap is filled in by the Pol δ holoenzyme, and the nick is ligated by a DNA ligase (44, 46, 47). DNA polymerase ε (Pol ε) can substitute for Pol δ in the EXO1-dependent MMR reaction, but its activity in this reaction is much lower than that of Pol δ (48). Although MutLα endonuclease is essential for MMR in vivo, 5′ nick-dependent MMR reactions reconstituted in the presence of EXO1 are MutLα-independent (44, 47, 49).EXO1 deficiency in humans does not seem to cause significant cancer predisposition (19). Nevertheless, it is known that Exo1^-/- mice are susceptible to the development of lymphomas (50). Genetic studies in yeast and mice demonstrated that EXO1 inactivation causes only a modest defect in MMR (50, 51, 52, 53). In agreement with these genetic studies, a defined human EXO1-independent MMR reaction that depends on the strand-displacement DNA synthesis activity of Pol δ holoenzyme to remove the mismatch was reconstituted (54). Furthermore, an EXO1-independent MMR reaction that occurred in a mammalian cell extract system without the formation of a gapped excision intermediate was observed (54). Together, these findings implicated the strand-displacement activity of Pol δ holoenzyme in EXO1-independent MMR.In this study, we investigated DNA2 in the context of MMR. DNA2 is an essential multifunctional protein that has nuclease, ATPase, and 5''→3′ helicase activities (55, 56, 57). Previous research ascertained that DNA2 removes long flaps during Okazaki fragment maturation (58, 59, 60), participates in the resection step of double-strand break repair (61, 62, 63), initiates the replication checkpoint (64), and suppresses the expansions of GAA repeats (65). We have found in vivo and in vitro evidence that DNA2 promotes EXO1-independent MMR. Our data have indicated that the nuclease activity of DNA2 enhances the strand-displacement activity of Pol δ holoenzyme in an EXO1-independent MMR reaction.     

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

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

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

CYP86A33-Targeted Gene Silencing in Potato Tuber Alters Suberin Composition,Distorts Suberin Lamellae,and Impairs the Periderm's Water Barrier Function

Suberin is a cell wall lipid polyester found in the cork cells of the periderm offering protection against dehydration and pathogens. Its biosynthesis and assembly, as well as its contribution to the sealing properties of the periderm, are still poorly understood. Here, we report on the isolation of the coding sequence CYP86A33 and the molecular and physiological function of this gene in potato (Solanum tuberosum) tuber periderm. CYP86A33 was down-regulated in potato plants by RNA interference-mediated silencing. Periderm from CYP86A33-silenced plants revealed a 60% decrease in its aliphatic suberin load and greatly reduced levels of C18:1 ω-hydroxyacid (approximately 70%) and α,ω-diacid (approximately 90%) monomers in comparison with wild type. Moreover, the glycerol esterified to suberin was reduced by 60% in the silenced plants. The typical regular ultrastructure of suberin, consisting of dark and light lamellae, disappeared and the thickness of the suberin layer was clearly reduced. In addition, the water permeability of the periderm isolated from CYP86A33-silenced lines was 3.5 times higher than that of the wild type. Thus, our data provide convincing evidence for the involvement of ω-functional fatty acids in establishing suberin structure and function.Periderm, the boundary tissue that replaces the epidermis in the secondary organs of plants, provides efficient protection against dehydration, UV radiation, and pathogens (Esau, 1965). Periderm is regularly found in the bark of woody plants, but herbaceous plants may also form a well-developed periderm in roots, tubers, and the oldest portions of stem. The periderm has been widely studied in potato (Solanum tuberosum) tubers because of the latter''s great agronomic significance (Schmidt and Schönherr, 1982; Vogt et al., 1983; Lulai and Freeman, 2001; Sabba and Lulai, 2002). Shrinkage and flaccidity occur in tubers if the protection afforded by the periderm against water loss is compromised (Lulai et al., 2006). Suberin and waxes embedded into the suberin matrix are considered essential for periderm imperviousness (Franke and Schreiber, 2007). Chemically, suberin is a complex lipid polymer consisting of a fatty acid-derived domain (aliphatic suberin) cross-linked by ester bonds to a polyaromatic lignin-like domain (aromatic suberin; Kolattukudy, 2001; Bernards, 2002; Franke and Schreiber, 2007). Aliphatic suberin has been widely analyzed in potato periderm (Kolattukudy and Agrawal, 1974; Graça and Pereira, 2000; Schreiber et al., 2005). The main monomers released from potato aliphatic suberin are a mixture of ω-hydroxyacids and α,ω-diacids with chain lengths ranging from C16 to C28 (mainly C18), together with glycerol. Small amounts of monofunctional fatty acids, alcohols, and ferulic acid are also released. Waxes are complex mixtures of lipids extractable with chloroform that in potato periderm consist mostly of linear very-long-chain aliphatics up to C32 (Schreiber et al., 2005). Suberin is deposited in the cell wall as a continuous deposit or secondary cell wall that overlays the polysaccharide primary cell wall from the inside (Esau, 1965). These suberin deposits appear under the transmission electron microscope (TEM) as regularly alternating opaque and translucent lamellae (Schmidt and Schönherr, 1982). Although several molecular models for suberin have been proposed (Kolattukudy, 1980; Bernards, 2002; Graça and Santos, 2007), how the suberin and wax components are organized in the lamellated suberin secondary cell wall is a matter of debate (Graça and Santos, 2007). Moreover, to what extent suberin and wax deposition and composition determine sealing properties of periderm still remains unclear (Schreiber et al., 2005). Several studies confirm the importance of waxes for the diffusion barrier (Soliday et al., 1979; Vogt et al., 1983; Schreiber et al., 2005), but the significance of aliphatic suberin has hardly been investigated at all. Interestingly, an Arabidopsis (Arabidopsis thaliana) knockout mutant for the GLYCEROL-3-PHOSPHATE ACYLTRANSFERASE5 gene (GPAT5) with altered suberin showed higher tetrazolium salt permeability in the seed coat (Beisson et al., 2007).ω-Hydroxylation of fatty acids, a reaction carried out in plants by cytochrome P450 monooxygenases, is a crucial step in the biosynthesis of plant biopolymers (Kolattukudy, 1980; Nawrath, 2002). The Arabidopsis mutant lacerata, which shows phenotype defects compatible with a cutin deficiency, is defective in CYP86A8 encoding a fatty acid ω-hydroxylase (Wellesen et al., 2001). The aberrant induction of type three genes1 (att1) mutant, showing an altered cuticle ultrastructure and a higher transpiration rate than wild type, is defective in CYP86A2 and contains reduced amounts of ω-functionalized cutin monomers (Xiao et al., 2004). Moreover, a genome-wide study of cork oak (Quercus suber) bark highlighted a member of the cytochrome P450 of the CYP86A subfamily as a strong candidate gene for aliphatic suberin biosynthesis (Soler et al., 2007); and a role for CYP86A1 in the biosynthesis of suberin has recently been confirmed in the primary root of Arabidopsis knockout mutants (Li et al., 2007; Hofer et al., 2008). However, how the lack of fatty acid ω-hydroxylase activity may affect the structural features and sealing properties of suberin in periderm cell walls has not been documented.To provide experimental evidence of the role of CYP86A genes in periderm fine structure and water transpiration properties, especially quantitative permeability studies so far unexplored in Arabidopsis, we followed a strategy based on the potato (cv Desirée). The potato is a model of choice for such studies because transgenic plants can be produced in reasonable time and sufficient amounts of periderm can easily be obtained from tubers for chemical and physiological studies (Vogt et al., 1983; Schreiber et al., 2005). Based on the CYP86A gene that was highlighted in cork oak periderm as a strong suberin candidate for aliphatic suberin biosynthesis, we isolated the putative ortholog in potato and used a reverse genetic approach to analyze the effects of down-regulation on the chemical and ultrastructural features of suberin and on the physiological properties of the tuber periderm. Our findings indicate that down-regulation of CYP86A33, as this gene was designated, results in a strong decrease of the ω-functionalized monomers in aliphatic suberin, which are necessary for the suberin typical lamellar organization and for the periderm resistance to water loss.     

Wettability,Polarity, and Water Absorption of Holm Oak Leaves: Effect of Leaf Side and Age

Plant trichomes play important protective functions and may have a major influence on leaf surface wettability. With the aim of gaining insight into trichome structure, composition, and function in relation to water-plant surface interactions, we analyzed the adaxial and abaxial leaf surface of holm oak (Quercus ilex) as a model. By measuring the leaf water potential 24 h after the deposition of water drops onto abaxial and adaxial surfaces, evidence for water penetration through the upper leaf side was gained in young and mature leaves. The structure and chemical composition of the abaxial (always present) and adaxial (occurring only in young leaves) trichomes were analyzed by various microscopic and analytical procedures. The adaxial surfaces were wettable and had a high degree of water drop adhesion in contrast to the highly unwettable and water-repellent abaxial holm oak leaf sides. The surface free energy and solubility parameter decreased with leaf age, with higher values determined for the adaxial sides. All holm oak leaf trichomes were covered with a cuticle. The abaxial trichomes were composed of 8% soluble waxes, 49% cutin, and 43% polysaccharides. For the adaxial side, it is concluded that trichomes and the scars after trichome shedding contribute to water uptake, while the abaxial leaf side is highly hydrophobic due to its high degree of pubescence and different trichome structure, composition, and density. Results are interpreted in terms of water-plant surface interactions, plant surface physical chemistry, and plant ecophysiology.Plant surfaces have an important protecting function against multiple biotic and abiotic stress factors (Riederer, 2006). They may, for example, limit the attack of insects (Eigenbrode and Jetter, 2002) or pathogenic fungi (Gniwotta et al., 2005; Łaźniewska et al., 2012), avoid damage caused by high intensities of UV and visible radiation (Reicosky and Hanover, 1978; Karabourniotis and Bormann, 1999), help to regulate leaf temperature (Ehleringer and Björkman, 1978; Ripley et al., 1999), and chiefly prevent plant organs from dehydration (Riederer and Schreiber, 2001).The epidermis of plants has been found to have a major degree of physical and chemical variability and may often contain specialized cells such as trichomes or stomata (Roth-Nebelsick et al., 2009; Javelle et al., 2011). Most aerial organs are covered with an extracellular and generally lipid-rich layer named the cuticle, which is typically composed of waxes embedded in (intracuticular waxes) or deposited on (epicuticular waxes) a biopolymer matrix of cutin, forming a network of cross-esterified hydroxy C₁₆ and/or C₁₈ fatty acids, and/or cutan, with variable amounts of polysaccharides and phenolics (Domínguez et al., 2011; Yeats and Rose, 2013). Different nano- and/or microscale levels of plant surface sculpturing have been observed by scanning electron microscopy (SEM), generally in relation to the topography of epicuticular waxes, cuticular folds, and epidermal cells (Koch and Barthlott, 2009). Such surface features together with their chemical composition (Khayet and Fernández, 2012) may lead to a high degree of roughness and hydrophobicity (Koch and Barthlott, 2009; Konrad et al., 2012). The interactions of plant surfaces with water have been addressed in some investigations (Brewer et al., 1991; Brewer and Smith, 1997; Pandey and Nagar, 2003; Hanba et al., 2004; Dietz et al., 2007; Holder, 2007a, 2007b; Fernández et al., 2011, 2014; Roth-Nebelsick et al., 2012; Wen et al., 2012; Urrego-Pereira et al., 2013) and are a topic of growing interest for plant ecophysiology (Helliker and Griffiths, 2007; Aryal and Neuner, 2010; Limm and Dawson, 2010; Kim and Lee, 2011; Berry and Smith, 2012; Berry et al., 2013; Rosado and Holder, 2013; Helliker, 2014). On the other hand, the mechanisms of foliar uptake of water and solutes by plant surfaces are still not fully understood (Fernández and Eichert, 2009; Burkhardt and Hunsche, 2013), but they may play an important ecophysiological role (Limm et al., 2009; Johnstone and Dawson, 2010; Adamec, 2013; Berry et al., 2014).The importance of trichomes and pubescent layers on water drop-plant surface interactions and on the subsequent potential water uptake into the organs has been analyzed in some investigations (Fahn, 1986; Brewer et al., 1991; Grammatikopoulos and Manetas, 1994; Brewer and Smith, 1997; Pierce et al., 2001; Kenzo et al., 2008; Fernández et al., 2011, 2014; Burrows et al., 2013). Trichomes are unicellular or multicellular and glandular or nonglandular appendages, which originate from epidermal cells only and develop outwards on the surface of plant organs (Werker, 2000). Nonglandular trichomes are categorized according to their morphology and exhibit a major variability in size, morphology, and function. On the other hand, glandular trichomes are classified by the secretory materials they excrete, accumulate, or absorb (Johnson, 1975; Werker, 2000; Wagner et al., 2004). Trichomes can be often found in xeromorphic leaves and in young organs (Fahn, 1986; Karabourniotis et al., 1995). The occurrence of protecting leaf trichomes has been also reported for Mediterranean species such as holm oak (Quercus ilex; Karabourniotis et al., 1995, 1998; Morales et al., 2002; Karioti et al., 2011; Camarero et al., 2012). There is limited information about the nature of the surface of trichomes, but they are also covered with a cuticle similarly to other epidermal cell types (Fernández et al., 2011, 2014).In this study and using holm oak as a model, we assessed, for the first time, the leaf surface-water relations of the abaxial (always pubescent) versus the adaxial (only pubescent in developing leaves and for a few months) surface, including their capacity to absorb surface-deposited water drops. Based on membrane science methodologies (Fernández et al., 2011; Khayet and Fernández, 2012) and following a new integrative approach, the chemical, physical, and anatomical properties of holm oak leaf surfaces and trichomes were analyzed, with the aim of addressing the following questions. Are young and mature adaxial and abaxial leaf surfaces capable of absorbing water deposited as drops on to the surfaces? Are young and mature abaxial and adaxial leaf surfaces similar in relation to their wettability, hydrophobicity, polarity, work of adhesion (W_a) for water, solubility parameter (δ), and surface free energy (γ)? What is the physical and chemical nature of the adaxial versus the abaxial trichomes, chiefly in relation to young leaves?     

Effect of Rubisco Activase Deficiency on the Temperature Response of CO2 Assimilation Rate and Rubisco Activation State: Insights from Transgenic Tobacco with Reduced Amounts of Rubisco Activase

The activation of Rubisco in vivo requires the presence of the regulatory protein Rubisco activase. To elucidate its role in maintaining CO₂ assimilation rate at high temperature, we examined the temperature response of CO₂ assimilation rate at 380 μL L⁻¹ CO₂ concentration (A₃₈₀) and Rubisco activation state in wild-type and transgenic tobacco (Nicotiana tabacum) with reduced Rubisco activase content grown at either 20°C or 30°C. Analyses of gas exchange and chlorophyll fluorescence showed that in the wild type, A₃₈₀ was limited by ribulose 1,5-bisphosphate regeneration at lower temperatures, whereas at higher temperatures, A₃₈₀ was limited by ribulose 1,5-bisphosphate carboxylation irrespective of growth temperatures. Growth temperature induced modest differences in Rubisco activation state that declined with measuring temperature, from mean values of 76% at 15°C to 63% at 40°C in wild-type plants. At measuring temperatures of 25°C and below, an 80% reduction in Rubisco activase content was required before Rubisco activation state was decreased. Above 35°C, Rubisco activation state decreased slightly with more modest decreases in Rubisco activase content, but the extent of the reductions in Rubisco activation state were small, such that a 55% reduction in Rubisco activase content did not alter the temperature sensitivity of Rubisco activation and had no effect on in vivo catalytic turnover rates of Rubisco. There was a strong correlation between Rubisco activase content and Rubisco activation state once Rubisco activase content was less that 20% of wild type at all measuring temperatures. We conclude that reduction in Rubisco activase content does not lead to an increase in the temperature sensitivity of Rubisco activation state in tobacco.The catalytic sites of Rubisco must be activated for CO₂ fixation to take place. This requires the carbamylation of a Lys residue at the catalytic sites to allow the binding of Mg²⁺ and ribulose 1,5-bisphosphate (RuBP; Andrews and Lorimer, 1987). Rubisco activase facilitates carbamylation and the maintenance of Rubisco activity by removing inhibitors such as tight-binding sugar phosphates from Rubisco catalytic sites in an ATP-dependent manner (Andrews, 1996; Spreitzer and Salvucci, 2002; Portis, 2003; Parry et al., 2008). The activity of Rubisco activase is regulated by the ATP/ADP ratio and redox state in the chloroplast (Zhang and Portis, 1999; Zhang et al., 2002; Portis, 2003).In many plant species, Rubisco activation state decreases at high temperature in vivo (Crafts-Brandner and Salvucci, 2000; Salvucci and Crafts-Brandner, 2004b; Cen and Sage, 2005; Yamori et al., 2006b; Makino and Sage, 2007). However, it is unclear what the primary mechanisms underlying the inhibition of Rubisco activation are and whether Rubisco deactivation limits CO₂ assimilation rate at high temperature. It has been proposed that Rubisco activation state decreases at high temperature, because the activity of Rubisco activase is insufficient to keep pace with the faster rates of Rubisco inactivation at high temperatures (Crafts-Brandner and Salvucci, 2000; Salvucci and Crafts-Brandner, 2004a, 2004c; Kim and Portis, 2006). In in vitro assays using purified Rubisco and Rubisco activase, the activity of Rubisco activase was sufficient for the activation of Rubisco at the optimum temperature but not at high temperatures (Crafts-Brandner and Salvucci, 2000; Salvucci and Crafts-Brandner, 2004a, 2004c). ATP hydrolysis activity of Rubisco activase in vitro has varying temperature optima among species (e.g. 25°C in Antarctic hairgrass [Deschampsia antarctica] and spinach [Spinacia oleracea] but 35°C in tobacco [Nicotiana tabacum] and cotton [Gossypium hirsutum]), and Rubisco activase more readily dissociates into inactive forms at high temperature, causing a loss of Rubisco activase capacity (Crafts-Brandner and Law, 2000; Salvucci and Crafts-Brandner, 2004b). Moreover, the rates of inhibitor formation by misprotonation of RuBP during catalysis increased at higher temperatures (Salvucci and Crafts-Brandner, 2004c; Kim and Portis, 2006). CO₂ assimilation rates and plant growth were improved under heat stress in transgenic Arabidopsis expressing thermotolerant Rubisco activase isoforms generated by either gene-shuffling technology (Kurek et al., 2007) or chimeric Rubisco activase constructs (Kumar et al., 2009). These results support the view that the reduction of Rubisco activase activity limits the Rubisco activation and, therefore, the CO₂ assimilation rates at high temperatures.It has also been suggested that the decrease in CO₂ assimilation rate at high temperatures is caused by a limitation of RuBP regeneration capacity (e.g. electron transport capacity) rather than by Rubisco deactivation per se (Schrader et al., 2004; Wise et al., 2004; Cen and Sage, 2005; Makino and Sage, 2007; Kubien and Sage, 2008). These groups suggest that Rubisco deactivation at high temperature may be a regulatory response to the limitation of one of the processes contributing to electron transport capacities. For example, at high temperature, protons can leak through the thylakoid membrane, impairing the coupling of ATP synthesis to electron transport (Pastenes and Horton, 1996; Bukhov et al., 1999, 2000). As the electron transport capacity becomes limiting, ATP/ADP ratios and the redox potential of the chloroplast decline, causing a loss of Rubisco activase activity and, in turn, a reduction in the Rubisco activation state (Zhang and Portis, 1999; Zhang et al., 2002; Sage and Kubien, 2007). Based on this understanding, the decline in the Rubisco activation state at high temperature may be a regulated response to a limitation in electron transport capacity rather than a consequence of a direct effect of heat on the integrity of Rubisco activase.Temperature dependence of CO₂ assimilation rate shows a considerable variation with growth temperature (Berry and Björkman, 1980; Hikosaka et al., 2006; Sage and Kubien, 2007). Plants grown at low temperature generally exhibit higher CO₂ assimilation rates at low temperatures compared with plants grown at high temperature, but they exhibit lower rates at high temperature. Furthermore, both the temperature response of Rubisco activation state and the limiting step of CO₂ assimilation rate (a Rubisco versus RuBP regeneration limitation) have been shown to differ depending on growth temperature (Hikosaka et al., 1999; Onoda et al., 2005; Yamori et al., 2005, 2006a, 2006b, 2008). This suggests that the regulation of Rubisco activation state could also differ in plants grown at different growth temperatures. Here, we analyzed the effects of Rubisco activase content on Rubisco activation state and CO₂ assimilation rate at leaf temperatures ranging from 15°C to 40°C in tobacco grown under two different temperature regimes (day/night temperatures of 20°C/15°C or 30°C/25°C). We used wild-type and transgenic tobacco with a range of reductions in Rubisco activase content to examine the dependence of Rubisco activation on Rubisco activase content over the range of leaf temperatures (Mate et al., 1993, 1996).     

Lambda Red Recombineering in Escherichia coli Occurs Through a Fully Single-Stranded Intermediate

The phage lambda-derived Red recombination system is a powerful tool for making targeted genetic changes in Escherichia coli, providing a simple and versatile method for generating insertion, deletion, and point mutations on chromosomal, plasmid, or BAC targets. However, despite the common use of this system, the detailed mechanism by which lambda Red mediates double-stranded DNA recombination remains uncertain. Current mechanisms posit a recombination intermediate in which both 5′ ends of double-stranded DNA are recessed by λ exonuclease, leaving behind 3′ overhangs. Here, we propose an alternative in which lambda exonuclease entirely degrades one strand, while leaving the other strand intact as single-stranded DNA. This single-stranded intermediate then recombines via beta recombinase-catalyzed annealing at the replication fork. We support this by showing that single-stranded gene insertion cassettes are recombinogenic and that these cassettes preferentially target the lagging strand during DNA replication. Furthermore, a double-stranded DNA cassette containing multiple internal mismatches shows strand-specific mutations cosegregating roughly 80% of the time. These observations are more consistent with our model than with previously proposed models. Finally, by using phosphorothioate linkages to protect the lagging-targeting strand of a double-stranded DNA cassette, we illustrate how our new mechanistic knowledge can be used to enhance lambda Red recombination frequency. The mechanistic insights revealed by this work may facilitate further improvements to the versatility of lambda Red recombination.OVER the past decade, lambda Red recombination (“recombineering”) has been used as a powerful technique for making precisely defined insertions, deletions, and point mutations in Escherichia coli, requiring as few as 35 bp of homology on each side of the desired alteration (Thomason et al. 2007a; Sharan et al. 2009). With this system, single-stranded DNA (ssDNA) oligonucleotides have been used to efficiently modify E. coli chromosomal targets (Ellis et al. 2001; Costantino and Court 2003), BACs (Swaminathan et al. 2001), and plasmids (Thomason et al. 2007b), as well as to rapidly optimize a metabolic pathway coding for the production of lycopene (Wang et al. 2009). Furthermore, linear double-stranded DNA (dsDNA) recombineering has been used to replace chromosomal genes (Murphy 1998; Murphy et al. 2000), to disrupt gene function (Datsenko and Wanner 2000), and to develop novel cloning methods (Lee et al. 2001; Li and Elledge 2005). Large-scale dsDNA recombineering projects include creating a library of single-gene knockout E. coli strains (Baba et al. 2006) and removing 15% of the genomic material from a single E. coli strain (Posfai et al. 2006). Linear dsDNA recombineering has also been used to insert heterologous genes and entire pathways into the E. coli chromosome (Zhang et al. 1998; Wang and Pfeifer 2008) and BACs (Lee et al. 2001; Warming et al. 2005), including those used for downstream applications in eukaryotes (Chaveroche et al. 2000; Bouvier and Cheng 2009). However, despite the broad use of this method, the mechanism of lambda Red recombination has not achieved scientific consensus, particularly in the case of dsDNA recombination. A clearer understanding of the mechanism underlying this process could suggest ways to improve the functionality, ease, and versatility of lambda Red recombination.Three phage-derived lambda Red proteins are necessary for carrying out dsDNA recombination: Gam, Exo, and Beta. Gam prevents the degradation of linear dsDNA by the E. coli RecBCD and SbcCD nucleases; lambda exonuclease (Exo) degrades dsDNA in a 5′ to 3′ manner, leaving single-stranded DNA in the recessed regions; and Beta binds to the single-stranded regions produced by Exo and facilitates recombination by promoting annealing to the homologous genomic target site (Sawitzke et al. 2007). Current mechanisms claim that Exo binds to both 5′ ends of the dsDNA and degrades in both directions simultaneously to produce a double-stranded region flanked on both sides by 3′ overhangs (Sharan et al. 2009; Szczepanska 2009). However, a comprehensive explanation of how this construct ultimately recombines with the chromosome has not yet been advanced.Initially, it was proposed that this recombination occurs via strand invasion (Thaler et al. 1987). However, it has more recently been shown that strand invasion is unlikely to be the dominant mechanism in the absence of long regions of homology, as recombination remains highly proficient in a recA^- background (Yu et al. 2000). Furthermore, a detailed analysis of lambda Red recombination products showed characteristics consistent with strand annealing rather than a strand invasion model (Stahl et al. 1997). Finally, lambda Red dsDNA recombination has been shown to preferentially target the lagging strand during DNA replication, which suggests strand annealing rather than strand invasion (Lim et al. 2008; Poteete 2008).To explain these results, Court et al. (2002) proposed a strand-annealing model for insertional dsDNA recombination (Figure 1A), in which one single-stranded 3′ end anneals to its homologous target at the replication fork. The replication fork then stalls, due to the presence of a large dsDNA nonhomology (i.e., the insertion cassette). The stalled replication fork is ultimately rescued by the other replication fork traveling in the opposite direction around the circular bacterial chromosome. The other 3′ end of the recombinogenic DNA anneals to the homology region exposed by the second replication fork, forming a crossover structure, which is then resolved by unspecified E. coli enzymes (Court et al. 2002).Open in a separate windowFigure 1.—Previously proposed lambda Red-mediated dsDNA recombination mechanisms. Heterologous dsDNA is shown in green; Exo is an orange oval, and Beta is a yellow oval. In both mechanisms the recombination intermediate is proposed to be a dsDNA core flanked on either side by 3′ ssDNA overhangs. (A) The Court mechanism posits that (1) Beta facilitates annealing of one 3′ overhang to the lagging strand of the replication fork. (2) This replication fork then stalls and backtracks so that the leading strand can template switch onto the synthetic dsDNA. The heterologous dsDNA blocks further replication from this fork. (3) Once the second replication fork reaches the stalled fork, the other 3′ end of the integration cassette is annealed to the lagging strand in the same manner as prior. Finally, the crossover junctions must be resolved by unspecified E. coli enzymes (Court et al. 2002). (B) The Poteete mechanism suggests that (1) Beta facilitates 3′ overhang annealing to the lagging strand of the replication fork and (2) positions the invading strand to serve as the new template for leading-strand synthesis. This structure is resolved by an unspecified host endonuclease (red triangle), and (3) the synthetic dsDNA becomes template for both lagging and leading-strand synthesis. A second template switch must then occur at the other end of the synthetic dsDNA (Poteete 2008). The figure was adapted from the references cited.The Court mechanism was challenged by Poteete (2008), who showed that the dsDNA recombination of a linear lambda phage chromosome occurs readily onto a unidirectionally replicating plasmid, which does not have the second replication fork required by the Court mechanism (Court et al. 2002). Thus, Poteete proposed an alternate mechanism (Poteete 2008), termed “replisome invasion” (Figure 1B), in which a 3′ overhang of the Exo-processed dsDNA first anneals to its complementary sequence on the lagging strand of the recombination target. Subsequently, this overhang displaces the leading strand, thereby serving as the new template for leading-strand synthesis. The resulting structure is resolved by an unspecified endonuclease, after which the recombinogenic DNA becomes the template for the synthesis of both new strands. In the context of recombineering using a linear dsDNA cassette, the author indicates that a second strand-switching event must occur at the other end of the incoming dsDNA.While Poteete''s mechanism addresses some of the weaknesses of the Court mechanism, it remains largely speculative. This mechanism does not identify the endonuclease responsible for resolving the structure after the first template switching event, nor does it explain how the recombinogenic DNA and replication machinery form a new replication fork. Additionally, this template-switching mechanism would have to operate two times in a well-controlled manner, which may not be consistent with the high-recombination frequencies often observed (Murphy et al. 2000) for lambda Red-mediated dsDNA insertion. Finally, little experimental evidence has been advanced to directly support this hypothesis.To address the deficiencies in these mechanisms, we propose that lambda Red dsDNA recombination proceeds via a ssDNA intermediate rather than a dsDNA core flanked by 3′ overhangs (Figure 2). In this mechanism, Exo binds to one of the two dsDNA strands and degrades that strand completely, leaving behind full-length ssDNA. This ssDNA then anneals to its homology target at the lagging strand of the replication fork and is incorporated as part of the newly synthesized strand as if it were an Okazaki fragment. This process is analogous to the accepted mechanism for the lambda Red-mediated recombination of ssDNA oligonucleotides (Court et al. 2002) and, therefore, unifies the mechanisms for ssDNA and dsDNA recombination. Notably, our mechanism uses one replication fork for the incorporation of a full-length heterologous cassette, thereby addressing Poteete''s criticism of the Court mechanism.Open in a separate windowFigure 2.—Lambda Red mediated dsDNA recombination proceeds via a ssDNA intermediate. Instead of a recombination intermediate involving dsDNA flanked by 3′-ssDNA overhangs, we propose that one strand of linear dsDNA is entirely degraded by Exo (orange oval). Beta (yellow oval) then facilitates annealing to the lagging strand of the replication fork in place of an Okazaki fragment. The heterologous region does not anneal to the genomic sequence. This mechanism could account for gene replacement (as shown) or for insertions in which no genomic DNA is removed.The degradation of an entire strand by lambda Exo is feasible, given the highly processive nature of the enzyme (Subramanian et al. 2003). Whereas previously proposed mechanisms assume that both dsDNA ends are degraded approximately simultaneously, our hypothesis implies that some dsDNA molecules will be entirely degraded to ssDNA before a second Exo can bind to the other end. In this article, we demonstrate that single-stranded DNA is a viable recombinogenic intermediate with lagging-strand bias. Furthermore, we show that genetic information from one strand of a recombinogenic dsDNA cassette cosegregates during lambda Red-mediated recombination. These results provide strong support of our proposed mechanism.     

Alpha-Synuclein affects neurite morphology,autophagy, vesicle transport and axonal degeneration in CNS neurons

Many neuropathological and experimental studies suggest that the degeneration of dopaminergic terminals and axons precedes the demise of dopaminergic neurons in the substantia nigra, which finally results in the clinical symptoms of Parkinson disease (PD). The mechanisms underlying this early axonal degeneration are, however, still poorly understood. Here, we examined the effects of overexpression of human wildtype alpha-synuclein (αSyn-WT), a protein associated with PD, and its mutant variants αSyn-A30P and -A53T on neurite morphology and functional parameters in rat primary midbrain neurons (PMN). Moreover, axonal degeneration after overexpression of αSyn-WT and -A30P was analyzed by live imaging in the rat optic nerve in vivo. We found that overexpression of αSyn-WT and of its mutants A30P and A53T impaired neurite outgrowth of PMN and affected neurite branching assessed by Sholl analysis in a variant-dependent manner. Surprisingly, the number of primary neurites per neuron was increased in neurons transfected with αSyn. Axonal vesicle transport was examined by live imaging of PMN co-transfected with EGFP-labeled synaptophysin. Overexpression of all αSyn variants significantly decreased the number of motile vesicles and decelerated vesicle transport compared with control. Macroautophagic flux in PMN was enhanced by αSyn-WT and -A53T but not by αSyn-A30P. Correspondingly, colocalization of αSyn and the autophagy marker LC3 was reduced for αSyn-A30P compared with the other αSyn variants. The number of mitochondria colocalizing with LC3 as a marker for mitophagy did not differ among the groups. In the rat optic nerve, both αSyn-WT and -A30P accelerated kinetics of acute axonal degeneration following crush lesion as analyzed by in vivo live imaging. We conclude that αSyn overexpression impairs neurite outgrowth and augments axonal degeneration, whereas axonal vesicle transport and autophagy are severely altered.Growing evidence suggests that Parkinson''s disease (PD) pathology starts at the presynaptic terminals and the distal axons and is then propagated back to the soma in a ''dying back'' pattern.^{1, 2} Accordingly, at the time of clinical onset, there is only a 30% loss of total substantia nigra pars compacta neurons but a far more severe loss of striatal dopaminergic markers (70–80%), suggesting that axonal terminals of the nigrostriatal pathway are affected earlier.¹ It is thus essential to understand the pathomechanisms specifically affecting the axon in PD in order to interfere with early disease progression.Neurodegeneration in PD is accompanied by the appearance of intraneuronal protein aggregates, denoted Lewy bodies (LBs).³ Interestingly, also LB pathology is initially found in the distal axons before becoming evident in the neuronal somata, and dystrophic neurites, so called ''Lewy neurites'', outnumber LBs in the early stages of PD.^{2, 4, 5} A main component of LBs is the protein alpha-synuclein (αSyn) that is not only widely used as a histopathological marker for PD but is also believed to have a major role in PD pathogenesis.^{6, 7} The importance of αSyn is further underlined by the discovery of αSyn point mutations (e.g. Ala53Thr (A53T), Ala30Pro (A30P)) and multiplications of the αSyn gene, all of which cause autosomal dominant forms of PD.^{8, 9, 10} However, neither the physiological functions nor the pathogenetic mechanisms of αSyn are well understood.⁷The biological effects of αSyn expression strongly depend on the model system. Wild-type (WT) human αSyn does not lead to major clinical or histological abnormalities when expressed in transgenic mice,^{11, 12} but its overexpression mediated by adeno-associated viral vectors (AAV) results in severe neurodegeneration, suggesting a dose-dependent toxic effect.^{13, 14} Different human αSyn-A30P and -A53T transgenic mouse lines develop severe motor impairments, partly resembling symptoms of human PD, accompanied by a degeneration of the nigrostriatal neuronal system and LB-like pathology.^{11, 12, 15} In line with the pathological findings in human PD, the axonal compartment is affected early and most prominently in these animal models.Different putative pathomechanisms of αSyn toxicity have been explored. For example, the cytoskeleton is an important molecular target of αSyn. Multimeric forms of αSyn were shown to impair the polymerization of tubulin and microtubule formation.^{16, 17} Overexpression of αSyn increased actin instability and induced actin bundling in cultured hippocampal neurons.¹⁸ There are, however, divergent data on the resulting effects of αSyn overexpression on neurite outgrowth and integrity in different model systems.^{19, 20, 21, 22}Moreover, a dysregulation of autophagy has been implicated in PD pathology. Aberrant αSyn is normally degraded by autophagy and only to a negligible degree by the proteasome.²³ Several studies have shown that the inhibition of autophagy results in an accumulation and increased toxicity of αSyn, whereas the activation of autophagy has therapeutic effects in PD models.^{23, 24, 25, 26} However, the direct effects of αSyn and its mutants on autophagy seem to rely strongly on the model system and the published data are highly controversial.^{24, 26, 27, 28, 29, 30, 31, 32}Given the central role of axonal degeneration in PD, it is likely that disturbances of axonal transport are involved.³³ In support of this proposition, the motor protein kinesin was shown to be decreased early and stage-dependently in PD patients, preceding the loss of substantia nigra neurons.³⁴ αSyn itself is actively transported along the axons, mainly by the slow component of axonal transport, but the role of αSyn in axonal vesicle transport is unclear.³⁵Here, we present a comprehensive analysis of the effects of αSyn on neurite morphology and examine important pathomechanisms.     

Naturally Occurring Disseminated Group B Streptococcus Infections in Postnatal Rats

Group B Streptococcus (Streptococcus agalactiae, GBS) is a gram-positive commensal and occasional opportunistic pathogen of the human vaginal, respiratory, and intestinal tracts that can cause sepsis, pneumonia, or meningitis in human neonates, infants, and immunosuppressed persons. We report here on a spontaneous outbreak of postnatal GBS-associated disease in rats. Ten of 26 (38.5%) 21- to 24-d-old rat pups died or were euthanized due to a moribund state in a colony of rats transgenic for the human diphtheria toxin receptor on a Munich–Wistar–Frömter genetic background. Four pups had intralesional coccoid bacteria in various organs without accompanying inflammation. GBS was isolated from the liver of 2 of these pups and from skin abscesses in 3 littermates. A connection with the transgene could not be established. A treatment protocol was evaluated in the remaining breeding female rats. GBS is a potentially clinically significant spontaneous infection in various populations of research rats, with some features that resemble late-onset postnatal GBS infection in human infants.Abbreviations: GBS, Group B Streptococcus; MWF, Munich Wistar Frömter; hDTR, human diphtheria toxin receptorStreptococci are gram-positive, coccoid bacteria that typically are classified according to their hemolytic capacity. α-hemolytic streptococci produce a zone of partial hemolysis that appears greenish on blood agar, whereas β-hemolytic streptococci produce a zone of complete hemolysis, and γ-hemolytic organisms produce no hemolysis on blood agar.²⁴ The β-hemolytic streptococci are further subdivided into Lancefield groups (A through G), according to cell-wall carbohydrate antigens.^24,29,39 The group B β-hemolytic Streptococcus (GBS) have been speciated as Streptococcus agalactiae.^28,39 It was first isolated as a causative agent of mastitis in cattle.²⁹ This organism has since been recognized as a cause of severe infection in human neonates.^28,39 In humans, GBS is harbored asymptomatically in the maternal genitourinary tract.^24,28 Infants can be infected and present with serious systemic disease in the first week of life (early-onset GBS) or from 1 wk to 3 mo of age (late-onset GBS).³⁹ In laboratory animals, rats have been used experimentally as models for neonatal^{1,6,7,20,37,38,43,44,47,50,51} or adult⁴⁵ GBS infection, but to our knowledge, GBS has not been associated with spontaneous disease in rats.     

A Host RNA Helicase-Like Protein,AtRH8, Interacts with the Potyviral Genome-Linked Protein,VPg, Associates with the Virus Accumulation Complex,and Is Essential for Infection

The viral genome-linked protein, VPg, of potyviruses is a multifunctional protein involved in viral genome translation and replication. Previous studies have shown that both eukaryotic translation initiation factor 4E (eIF4E) and eIF4G or their respective isoforms from the eIF4F complex, which modulates the initiation of protein translation, selectively interact with VPg and are required for potyvirus infection. Here, we report the identification of two DEAD-box RNA helicase-like proteins, PpDDXL and AtRH8 from peach (Prunus persica) and Arabidopsis (Arabidopsis thaliana), respectively, both interacting with VPg. We show that AtRH8 is dispensable for plant growth and development but necessary for potyvirus infection. In potyvirus-infected Nicotiana benthamiana leaf tissues, AtRH8 colocalizes with the chloroplast-bound virus accumulation vesicles, suggesting a possible role of AtRH8 in viral genome translation and replication. Deletion analyses of AtRH8 have identified the VPg-binding region. Comparison of this region and the corresponding region of PpDDXL suggests that they are highly conserved and share the same secondary structure. Moreover, overexpression of the VPg-binding region from either AtRH8 or PpDDXL suppresses potyvirus accumulation in infected N. benthamiana leaf tissues. Taken together, these data demonstrate that AtRH8, interacting with VPg, is a host factor required for the potyvirus infection process and that both AtRH8 and PpDDXL may be manipulated for the development of genetic resistance against potyvirus infections.Plant viruses are obligate intracellular parasites that infect many agriculturally important crops and cause severe losses each year. One of the common characteristics of plant viruses is their relatively small genome that encodes a limited number of viral proteins, making them dependent on host factors to fulfill their infection cycles (Maule et al., 2002; Whitham and Wang, 2004; Nelson and Citovsky, 2005; Decroocq et al., 2006). In order to establish a successful infection, the invading virus must recruit an array of host proteins (host factors) to translate and replicate its genome and to move locally from cell to cell via the plasmodesmata and systemically via the vascular system. It has been suggested that down-regulation or mutation of some of the required host factors may result in recessively inherited resistance to viruses (Kang et al., 2005b).Potyviruses, belonging to the genus Potyvirus in the family Potyviradae, constitute the largest group of plant viruses (Rajamäki et al., 2004). Potyviruses have a single positive-strand RNA genome approximately 10 kb in length, with a viral genome-linked protein (VPg) covalently attached to the 5′ end and a poly(A) tail at the 3′ end (Urcuqui-Inchima et al., 2001; Rajamäki et al., 2004). The viral genome contains a single open reading frame (ORF) that translates into a polypeptide with a molecular mass of approximately 350 kD, which is cleaved into 10 mature proteins by viral proteases (Urcuqui-Inchima et al., 2001). Recently, a novel viral protein resulting from a frameshift in the P3 cistron has been reported (Chung et al., 2008). Of the 11 viral proteins, VPg is a multifunctional protein and the only other viral protein present in the viral particles (virions) besides the coat protein and the cylindrical inclusion protein (CI; Oruetxebarria et al., 2001; Puustinen et al., 2002; Gabrenaite-Verkhovskaya et al., 2008). The nonstructural protein is linked to the viral RNA by a phosphodiester bond between the 5′ terminal uridine residue of the RNA and the O⁴-hydroxyl group of amino acid Tyr (Murphy et al., 1996; Oruetxebarria et al., 2001; Puustinen et al., 2002). Mutation of the Tyr residue that links VPg to the viral RNA abolishes virus infectivity completely (Murphy et al., 1996). In infected cells, VPg and its precursor NIa are present in the nucleus and in the membrane-associated virus replication vesicles in the cytoplasm (Carrington et al., 1993; Rajamäki and Valkonen, 2003; Cotton et al., 2009). As a component of the replication complex, VPg may serve as a primer for viral RNA replication (Puustinen and Mäkinen, 2004) and as an analog of the m⁷G cap of mRNAs for the viral genome to recruit the translation complex for translation (Michon et al., 2006; Beauchemin et al., 2007; Khan et al., 2008). Furthermore, VPg has been suggested to be an avirulence factor for recessive resistance genes in diverse plant species (Moury et al., 2004; Kang et al., 2005b; Bruun-Rasmussen et al., 2007). Thus, VPg plays a pivotal role in the virus infection process. The molecular identification of VPg-interacting host proteins and the subsequent functional characterization of such interactions may advance knowledge of the intricate virus replication mechanisms and help develop novel antiviral strategies.Previous studies have shown that VPg and its precursor NIa interact with several host proteins, including three essential components of the host protein translation apparatus (Thivierge et al., 2008). The first protein is the cellular translation initiation factor eIF4E or its isoform eIF(iso)4E, identified through a yeast two-hybrid screen using VPg as a bait (Wittmann et al., 1997; Schaad et al., 2000). The protein complex of VPg and eIF4E is an essential component for virus infectivity (Robaglia and Caranta, 2006). Mutations and knockout of eIF4E or eIF(iso)4E confer resistance to infection (Lellis et al., 2002; Ruffel et al., 2002; Nicaise et al., 2003; Gao et al., 2004; Kang et al., 2005a; Ruffel et al., 2005; Decroocq et al., 2006; Bruun-Rasmussen et al., 2007). It is well known that potyviruses recruit selectively one of the eIF4E isoforms, depending on specific virus-host combinations (German-Retana et al., 2008). For instance, in Arabidopsis (Arabidopsis thaliana), eIF(iso)4E is required for infection by Turnip mosaic virus (TuMV), Plum pox virus (PPV), and Lettuce mosaic virus, while eIF4E is indispensable for infection by Clover yellow vein virus (Duprat et al., 2002; Lellis et al., 2002; Sato et al., 2005; Decroocq et al., 2006). The second cellular protein interacting with VPg is another translation initiation factor, eIF4G. Analysis of Arabidopsis knockout mutants for eIF4G or its isomers eIF(iso)4G1 and eIF(iso)4G2 has yielded results supporting the idea that the recruitment of eIF4G for potyvirus infection is also isoform dependent (Nicaise et al., 2007). Recently, poly(A)-binding protein (PABP), the translation initiation factor that bridges the 5′ and 3′ termini of the mRNA into proximity, has been proposed to be essential for efficient multiplication of TuMV (Dufresne et al., 2008). PABP was previously documented to interact with NIa, a VPg precursor containing both VPg and the proteinase NIa-Pro (Léonard et al., 2004). As the translation factors eIF(iso)4E and PABP have been found to be internalized in virus-induced vesicles, it has been suggested that the interactions between VPg and these translation factors are crucial for viral RNA translation and/or replication (Beauchemin and Laliberté, 2007; Beauchemin et al., 2007; Cotton et al., 2009). Besides these three translation factors, a Cys-rich plant protein, potyvirus VPg-interaction protein, was also found to associate with VPg (Dunoyer et al., 2004). This plant-specific VPg-interacting host protein contains a PHD finger domain and acts as an ancillary factor to support potyvirus infection and movement (Dunoyer et al., 2004).In this study, we describe the identification of an Arabidopsis DEAD-box RNA helicase (DDX), AtRH8, and a peach (Prunus persica) DDX-like protein, PpDDXL, both interacting with the potyviral VPg protein. Using the atrh8 mutant, we demonstrate that AtRH8 is not required for plant growth and development in Arabidopsis but is necessary for infection by two plant potyviruses, PPV and TuMV. Furthermore, we present evidence that AtRH8 colocalizes with the virus accumulation complex in potyvirus-infected leaf tissues, which reveals a possible role of AtRH8 in virus infection. Finally, we have identified the VPg-binding region (VPg-BR) of AtRH8 and PpDDX and show that overexpression of the VPg-BR either from AtRH8 or PpDDXL suppresses virus accumulation.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

Differential Sodium and Potassium Transport Selectivities of the Rice OsHKT2;1 and OsHKT2;2 Transporters in Plant Cells

Na⁺ and K⁺ homeostasis are crucial for plant growth and development. Two HKT transporter/channel classes have been characterized that mediate either Na⁺ transport or Na⁺ and K⁺ transport when expressed in Xenopus laevis oocytes and yeast. However, the Na⁺/K⁺ selectivities of the K⁺-permeable HKT transporters have not yet been studied in plant cells. One study expressing 5′ untranslated region-modified HKT constructs in yeast has questioned the relevance of cation selectivities found in heterologous systems for selectivity predictions in plant cells. Therefore, here we analyze two highly homologous rice (Oryza sativa) HKT transporters in plant cells, OsHKT2;1 and OsHKT2;2, that show differential K⁺ permeabilities in heterologous systems. Upon stable expression in cultured tobacco (Nicotiana tabacum) Bright-Yellow 2 cells, OsHKT2;1 mediated Na⁺ uptake, but little Rb⁺ uptake, consistent with earlier studies and new findings presented here in oocytes. In contrast, OsHKT2;2 mediated Na⁺-K⁺ cotransport in plant cells such that extracellular K⁺ stimulated OsHKT2;2-mediated Na⁺ influx and vice versa. Furthermore, at millimolar Na⁺ concentrations, OsHKT2;2 mediated Na⁺ influx into plant cells without adding extracellular K⁺. This study shows that the Na⁺/K⁺ selectivities of these HKT transporters in plant cells coincide closely with the selectivities in oocytes and yeast. In addition, the presence of external K⁺ and Ca²⁺ down-regulated OsHKT2;1-mediated Na⁺ influx in two plant systems, Bright-Yellow 2 cells and intact rice roots, and also in Xenopus oocytes. Moreover, OsHKT transporter selectivities in plant cells are shown to depend on the imposed cationic conditions, supporting the model that HKT transporters are multi-ion pores.Intracellular Na⁺ and K⁺ homeostasis play vital roles in growth and development of higher plants (Clarkson and Hanson, 1980). Low cytosolic Na⁺ and high K⁺/Na⁺ ratios aid in maintaining an osmotic and biochemical equilibrium in plant cells. Na⁺ and K⁺ influx and efflux across membranes require the function of transmembrane Na⁺ and K⁺ transporters/channels. Several Na⁺-permeable transporters have been characterized in plants (Zhu, 2001; Horie and Schroeder, 2004; Apse and Blumwald, 2007). Na⁺/H⁺ antiporters mediate sequestration of Na⁺ into vacuoles under salt stress conditions in plants (Blumwald and Poole, 1985, 1987; Sze et al., 1999). Na⁺ (cation)/H⁺ antiporters are encoded by six AtNHX genes in Arabidopsis (Arabidopsis thaliana; Apse et al., 1999; Gaxiola et al., 1999; Yokoi et al., 2002; Aharon et al., 2003). A distinct Na⁺/H⁺ antiporter, Salt Overly Sensitive1, mediates Na⁺/H⁺ exchange at the plasma membrane and mediates cellular Na⁺ extrusion (Shi et al., 2000, 2002; Zhu, 2001; Ward et al., 2003). Electrophysiological analyses reveal that voltage-independent channels, also named nonselective cation channels, mediate Na⁺ influx into roots under high external Na⁺ concentrations (Amtmann et al., 1997; Tyerman et al., 1997; Buschmann et al., 2000; Davenport and Tester, 2000); however, the underlying genes remain unknown.Potassium is the most abundant cation in plants and an essential nutrient for plant growth. The Arabidopsis genome includes 13 genes encoding KUP/HAK/KT transporters (Quintero and Blatt, 1997; Santa-María et al., 1997; Fu and Luan, 1998; Kim et al., 1998), and 17 genes have been identified encoding this family of transporters in rice (Oryza sativa ‘Nipponbare’; Bañuelos et al., 2002). Several KUP/HAK/KT transporters have been characterized as mediating K⁺ uptake across the plasma membrane of plant cells (Rigas et al., 2001; Bañuelos et al., 2002; Gierth et al., 2005).Ionic balance, especially the Na⁺/K⁺ ratio, is a key factor of salt tolerance in plants (Niu et al., 1995; Maathuis and Amtmann, 1999; Shabala, 2000; Mäser et al., 2002a; Tester and Davenport, 2003; Horie et al., 2006; Apse and Blumwald, 2007; Chen et al., 2007; Gierth and Mäser, 2007). Salinity stress is a major problem for agricultural productivity of crops worldwide (Greenway and Munns, 1980; Zhu, 2001). The Arabidopsis AtHKT1;1 transporter plays a key role in salt tolerance of plants by mediating Na⁺ exclusion from leaves (Mäser et al., 2002a; Berthomieu et al., 2003; Gong et al., 2004; Sunarpi et al., 2005; Rus et al., 2006; Davenport et al., 2007; Horie et al., 2009). athkt1;1 mutations cause leaf chlorosis and elevated Na⁺ accumulation in leaves under salt stress conditions in Arabidopsis (Mäser et al., 2002a; Berthomieu et al., 2003; Gong et al., 2004; Sunarpi et al., 2005). AtHKT1;1 and its homolog in rice, OsHKT1;5 (SKC1), mediate leaf Na⁺ exclusion by removing Na⁺ from the xylem sap to protect plants from salinity stress (Ren et al., 2005; Sunarpi et al., 2005; Horie et al., 2006, 2009; Davenport et al., 2007).The land plant HKT gene family is divided into two classes based on their nucleic acid sequences and protein structures (Mäser et al., 2002b; Platten et al., 2006). Class 1 HKT transporters have a Ser residue at a selectivity filter position in the first pore loop, which is replaced by a Gly in all but one known class 2 HKT transporter (Horie et al., 2001; Mäser et al., 2002b; Garciadeblás et al., 2003). While the Arabidopsis genome includes only one HKT gene, AtHKT1;1 (Uozumi et al., 2000), seven full-length OsHKT genes were found in the japonica rice cv Nipponbare genome (Garciadeblás et al., 2003). Members of class 1 HKT transporters, AtHKT1;1 and SKC1/OsHKT1;5, have a relatively higher Na⁺-to-K⁺ selectivity in Xenopus laevis oocytes and yeast than class 2 HKT transporters (Uozumi et al., 2000; Horie et al., 2001; Mäser et al., 2002b; Ren et al., 2005). The first identified plant HKT transporter, TaHKT2;1 from wheat (Triticum aestivum), is a class 2 HKT transporter (Schachtman and Schroeder, 1994). TaHKT2;1 was found to mediate Na⁺-K⁺ cotransport and Na⁺ influx at high Na⁺ concentrations in heterologous expression systems (Rubio et al., 1995, 1999; Gassmann et al., 1996; Mäser et al., 2002b). Thus, class 1 HKT transporters have been characterized as Na⁺-preferring transporters with a smaller K⁺ permeability (Fairbairn et al., 2000; Uozumi et al., 2000; Su et al., 2003; Jabnoune et al., 2009), whereas class 2 HKT transporters function as Na⁺-K⁺ cotransporters or channels (Gassmann et al., 1996; Corratgé et al., 2007). In addition, at millimolar Na⁺ concentrations, class 2 HKT transporters were found to mediate Na⁺ influx, without adding external K⁺ in Xenopus oocytes and yeast (Rubio et al., 1995, 1999; Gassmann et al., 1996; Horie et al., 2001). However, the differential cation transport selectivities of the two types of HKT transporters have not yet been analyzed and compared in plant cells.A study of the barley (Hordeum vulgare) and wheat class 2 transporters has suggested that the transport properties of HvHKT2;1 and TaHKT2;1 expressed in yeast are variable, depending on the constructs from which the transporter is expressed, and have led to questioning of the K⁺ transport activity of HKT transporters characterized in Xenopus oocytes and yeast (Haro et al., 2005). It was further proposed that the 5′ translation initiation of HKT proteins in yeast at nonconventional (non-ATG) sites affects the transporter selectivities of HKT transporters (Haro et al., 2005), although direct evidence for this has not yet been presented. However, recent research has shown a K⁺ permeability of OsHKT2;1 but not of OsHKT1;1 and OsHKT1;3 in Xenopus oocytes. These three OsHKT transporters show overlapping and also distinctive expression patterns in rice (Jabnoune et al., 2009).The report of Haro et al. (2005) has opened a central question addressed in this study: are the Na⁺/K⁺ transport selectivities of plant HKT transporters characterized in heterologous systems of physiological relevance in plant cells, or do they exhibit strong differences in the cation transport selectivities in these nonplant versus plant systems? To address this question, we analyzed the Na⁺/K⁺ transport selectivities of the OsHKT2;1 and OsHKT2;2 transporters expressed in cultured tobacco (Nicotiana tabacum ‘Bright-Yellow 2’ [BY2]) cells. OsHKT2;1 and OsHKT2;2 are two highly homologous HKT transporters from indica rice cv Pokkali, sharing 91% amino acid and 93% cDNA sequence identity (Horie et al., 2001). OsHKT2;1 mediates mainly Na⁺ uptake, which correlates with the presence of a Ser residue in the first pore loop of OsHKT2;1 (Horie et al., 2001, 2007; Mäser et al., 2002b; Garciadeblás et al., 2003). In contrast, OsHKT2;2 mediates Na⁺-K⁺ cotransport in Xenopus oocytes and yeast (Horie et al., 2001). Furthermore, at millimolar Na⁺ concentrations, OsHKT2;2 mediates Na⁺ influx in the absence of added K⁺ (Horie et al., 2001). Recent research on oshkt2;1 loss-of-function mutant alleles has revealed that OsHKT2;1 from japonica rice mediates a large Na⁺ influx component into K⁺-starved roots, thus compensating for lack of K⁺ availability (Horie et al., 2007). But the detailed Na⁺/K⁺ selectivities of Gly-containing, predicted K⁺-transporting class 2 HKT transporters have not yet been analyzed in plant cells.Here, we have generated stable OsHKT2;1- and OsHKT2;2-expressing tobacco BY2 cell lines and characterized the cell lines by ion content measurements and tracer influx studies to directly analyze unidirectional fluxes (Epstein et al., 1963). These analyses showed that OsHKT2;1 exhibits Na⁺ uptake activity in plant BY2 cells in the absence of added K⁺, but little K⁺ (Rb⁺), influx activity. In contrast, OsHKT2;2 was found to function as a Na⁺-K⁺ cotransporter/channel in plant BY2 cells, showing K⁺-stimulated Na⁺ influx and Na⁺-stimulated K⁺ (Rb⁺) influx. The differential K⁺ selectivities of the two OsHKT2 transporters were consistently reproduced by voltage clamp experiments using Xenopus oocytes here, as reported previously (Horie et al., 2001). OsHKT2;2 was also found to mediate K⁺-independent Na⁺ influx at millimolar external Na⁺ concentrations. These findings demonstrate that the cation selectivities of OsHKT2;1 and OsHKT2;2 in plant cells are consistent with past findings obtained from heterologous expression analyses under similar ionic conditions (Horie et al., 2001; Garciadeblás et al., 2003; Tholema et al., 2005). Furthermore, the shift in OsHKT2;2 Na⁺-K⁺ selectivity depending on ionic editions is consistent with the model that HKT transporters/channels are multi-ion pores (Gassmann et al., 1996; Corratgé et al., 2007). Classical studies of ion channels have shown that ion channels, in which multiple ions can occupy the pore at the same time, can change their relative selectivities depending on the ionic conditions (Hille, 2001). Moreover, the presence of external K⁺ and Ca²⁺ was found here to down-regulate OsHKT2;1-mediated Na⁺ influx both in tobacco BY2 cells and in rice roots. The inhibitory effect of external K⁺ on OsHKT2;1-mediated Na⁺ influx into intact rice roots, however, showed a distinct difference in comparison with that of BY2 cells, which indicates a possible posttranslational regulation of OsHKT2;1 in K⁺-starved rice roots.     

Class XI Myosins Move Specific Organelles in Pollen Tubes and Are Required for Normal Fertility and Pollen Tube Growth in Arabidopsis

Pollen tube growth is an essential aspect of plant reproduction because it is the mechanism through which nonmotile sperm cells are delivered to ovules, thus allowing fertilization to occur. A pollen tube is a single cell that only grows at the tip, and this tip growth has been shown to depend on actin filaments. It is generally assumed that myosin-driven movements along these actin filaments are required to sustain the high growth rates of pollen tubes. We tested this conjecture by examining seed set, pollen fitness, and pollen tube growth for knockout mutants of five of the six myosin XI genes expressed in pollen of Arabidopsis (Arabidopsis thaliana). Single mutants had little or no reduction in overall fertility, whereas double mutants of highly similar pollen myosins had greater defects in pollen tube growth. In particular, myo11c1 myo11c2 pollen tubes grew more slowly than wild-type pollen tubes, which resulted in reduced fitness compared with the wild type and a drastic reduction in seed set. Golgi stack and peroxisome movements were also significantly reduced, and actin filaments were less organized in myo11c1 myo11c2 pollen tubes. Interestingly, the movement of yellow fluorescent protein-RabA4d-labeled vesicles and their accumulation at pollen tube tips were not affected in the myo11c1 myo11c2 double mutant, demonstrating functional specialization among myosin isoforms. We conclude that class XI myosins are required for organelle motility, actin organization, and optimal growth of pollen tubes.Pollen tubes play a crucial role in flowering plant reproduction. A pollen tube is the vegetative cell of the male gametophyte. It undergoes rapid polarized growth in order to transport the two nonmotile sperm cells to an ovule. This rapid growth is supported by the constant delivery of secretory vesicles to the pollen tube tip, where they fuse with the plasma membrane to enlarge the cell (Bove et al., 2008; Bou Daher and Geitmann, 2011; Chebli et al., 2013). This vesicle delivery is assumed to be driven by the rapid movement of organelles and cytosol throughout the cell, a process that is commonly referred to as cytoplasmic streaming (Shimmen, 2007). Cytoplasmic streaming in angiosperm pollen tubes forms a reverse fountain: organelles moving toward the tip travel along the cell membrane, while organelles moving away from the tip travel through the center of the tube (Heslop-Harrison and Heslop-Harrison, 1990; Derksen et al., 2002). Drug treatments revealed that pollen tube cytoplasmic streaming and tip growth depend on actin filaments (Franke et al., 1972; Mascarenhas and Lafountain, 1972; Heslop-Harrison and Heslop-Harrison, 1989; Parton et al., 2001; Vidali et al., 2001). Curiously, very low concentrations of actin polymerization inhibitors can prevent growth without completely stopping cytoplasmic streaming, indicating that cytoplasmic streaming is not sufficient for pollen tube growth (Vidali et al., 2001). At the same time, however, drug treatments have not been able to specifically inhibit cytoplasmic streaming; thus, it is unknown whether cytoplasmic streaming is necessary for pollen tube growth.Myosins are actin-based motor proteins that actively transport organelles throughout the cell and are responsible for cytoplasmic streaming in plants (Shimmen, 2007; Sparkes, 2011; Madison and Nebenführ, 2013). Myosins can be grouped into at least 30 different classes based on amino acid sequence similarity of the motor domain, of which only class VIII and class XI myosins are found in plants (Odronitz and Kollmar, 2007; Sebé-Pedrós et al., 2014). Class VIII and class XI myosins have similar domain architecture. The N-terminal motor domain binds actin and hydrolyzes ATP (Tominaga et al., 2003) and is often preceded by an SH3-like (for sarcoma homology3) domain of unknown function. The neck domain, containing IQ (Ile-Gln) motifs, acts as a lever arm and is bound by calmodulin-like proteins that mediate calcium regulation of motor activity (Kinkema and Schiefelbein, 1994; Yokota et al., 1999; Tominaga et al., 2012). The coiled-coil domain facilitates dimerization (Li and Nebenführ, 2008), and the globular tail functions as the cargo-binding domain (Li and Nebenführ, 2007). Class VIII myosins also contain an N-terminal extension, MyTH8 (for myosin tail homology8; Mühlhausen and Kollmar, 2013), and class XI myosins contain a dilute domain in the C-terminal globular tail (Kinkema and Schiefelbein, 1994; Odronitz and Kollmar, 2007; Sebé-Pedrós et al., 2014). Recently, Mühlhausen and Kollmar (2013) proposed a new nomenclature for plant myosins based on a comprehensive phylogenetic analysis of all known plant myosins that clearly identifies paralogs and makes interspecies comparisons easier (Madison and Nebenführ, 2013).The localization of class VIII myosins, as determined by immunolocalization and the expression of fluorescently labeled full-length or tail constructs, has implicated these myosins in cell-to-cell communication, cell division, and endocytosis in angiosperms and moss (Reichelt et al., 1999; Van Damme et al., 2004; Avisar et al., 2008; Golomb et al., 2008; Sattarzadeh et al., 2008; Yuan et al., 2011; Haraguchi et al., 2014; Wu and Bezanilla, 2014). On the other hand, class XI myosin mutants have been studied extensively in Arabidopsis (Arabidopsis thaliana), which revealed roles for class XI myosins in cell expansion and organelle motility (Ojangu et al., 2007, 2012; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Park and Nebenführ, 2013). Very few studies have examined the reproductive tissues of class XI myosin mutants. In rice (Oryza sativa), one myosin XI was shown to be required for normal pollen development under short-day conditions (Jiang et al., 2007). In Arabidopsis, class XI myosins are required for stigmatic papillae elongation, which is necessary for normal fertility (Ojangu et al., 2012). Even though pollen tubes of myosin XI mutants have not been examined, the tip growth of another tip-growing plant cell has been thoroughly examined in myosin mutants. Root hairs are tubular outgrowths of root epidermal cells that function to increase the surface area of the root for water and nutrient uptake. Two myosin XI mutants have shorter root hairs, of which the myo11e1 (xik; myosin XI K) mutation has been shown to be associated with a slower root hair growth rate and reduced actin dynamics compared with the wild type (Ojangu et al., 2007; Peremyslov et al., 2008; Park and Nebenführ, 2013). Higher order mutants have a further reduction in root hair growth and have altered actin organization (Prokhnevsky et al., 2008; Peremyslov et al., 2010). Disruption of actin organization was also observed in myosin XI mutants of the moss Physcomitrella patens (Vidali et al., 2010), where these motors appear to coordinate the formation of actin filaments in the apical dome of the tip-growing protonemal cells (Furt et al., 2013). Interestingly, organelle movements in P. patens are much slower than in angiosperms and do not seem to depend on myosin motors (Furt et al., 2012).The function of myosins in pollen tubes is currently not known, although it is generally assumed that they are responsible for the prominent cytoplasmic streaming observed in these cells by associating with organelle surfaces (Kohno and Shimmen, 1988; Shimmen, 2007). Myosin from lily (Lilium longiflorum) pollen tubes was isolated biochemically and shown to move actin filaments with a speed of about 8 µm s⁻¹ (Yokota and Shimmen, 1994) in a calcium-dependent manner (Yokota et al., 1999). Antibodies against this myosin labeled small structures in both the tip region and along the shank (Yokota et al., 1995), consistent with the proposed role of this motor in moving secretory vesicles to the apex.In Arabidopsis, six of 13 myosin XI genes are highly expressed in pollen: Myo11A1 (XIA), Myo11A2 (XID), Myo11B1 (XIB), Myo11C1 (XIC), Myo11C2 (XIE), and Myo11D (XIJ; Peremyslov et al., 2011; Sparkes, 2011). The original gene names (Reddy and Day, 2001) are given in parentheses. Myo11D is the only short-tailed myosin XI in Arabidopsis (Mühlhausen and Kollmar, 2013) and lacks the typical myosin XI globular tail involved in cargo binding (Li and Nebenführ, 2007). The remaining genes have the same domain architecture as the conventional class XI myosins that have been shown to be involved in the elongation of trichomes, stigmatic papillae, and root hairs (Ojangu et al., 2007, 2012; Peremyslov et al., 2008, 2010; Prokhnevsky et al., 2008; Park and Nebenführ, 2013). Therefore, we predicted that these five pollen-expressed, conventional class XI myosins are required for the rapid elongation of pollen tubes. In this study, we examined transfer DNA (T-DNA) insertion mutants of Myo11A1, Myo11A2, Myo11B1, Myo11C1, and Myo11C2 for defects in fertility and pollen tube growth. Organelle motility and actin organization were also examined in myo11c1 myo11c2 pollen tubes.     

Exploring the Ultrastructural Localization and Biosynthesis of β(1,4)-Galactan in Pinus radiata Compression Wood

Softwood species such as pines react to gravitropic stimuli by producing compression wood, which unlike normal wood contains significant amounts of β(1,4)-galactan. Currently, little is known regarding the biosynthesis or physiological function of this polymer or the regulation of its deposition. The subcellular location of β(1,4)-galactan in developing tracheids was investigated in Pinus radiata D. Don using anti-β(1,4)-galactan antibodies to gain insight into its possible physiological role in compression wood. β(1,4)-Galactan was prominent and evenly distributed throughout the S2 layer of developing tracheid cell walls in P. radiata compression wood. In contrast, β(1,4)-galactan was not detected in normal wood. Greatly reduced antibody labeling was observed in fully lignified compression wood tracheids, implying that lignification results in masking of the epitope. To begin to understand the biosynthesis of galactan and its regulation, an assay was developed to monitor the enzyme that elongates the β(1,4)-galactan backbone in pine. A β(1,4)-galactosyltransferase (GalT) activity capable of extending 2-aminopyridine-labeled galacto-oligosaccharides was found to be associated with microsomes. Digestion of the enzymatic products using a β(1,4)-specific endogalactanase confirmed the production of β(1,4)-galactan by this enzyme. This GalT activity was substantially higher in compression wood relative to normal wood. Characterization of the identified pine GalT enzyme activity revealed pH and temperature optima of 7.0 and 20°C, respectively. The β(1,4)-galactan produced by the pine GalT had a higher degree of polymerization than most pectic galactans found in angiosperms. This observation is consistent with the high degree of polymerization of the naturally occurring β(1,4)-galactan in pine.The ability to respond to gravitropic stimuli is important for the survival of most terrestrial plants. Arborescent angiosperm and gymnosperm species generate wood with modified properties, called reaction wood, in response to gravitropic stimuli (Timell, 1969, 1986; Du and Yamamoto, 2007). The formation of reaction wood enables the return of bent stems to a vertical orientation. Interestingly, the location and type of the reaction wood deposited in woody gymnosperms and angiosperms generally differs significantly. Gymnosperms respond to gravitropic stimuli by compression wood formation on the underside of leaning stems (Timell, 1986), and arboreal angiosperms generate reaction wood primarily in the form of tension wood on the upper side of inclined stems (Timell, 1969).Compression wood in conifers differs significantly from normal wood in its anatomical, chemical, and physical properties. Typical anatomical features of severe compression wood are short, rounded, and thick-walled tracheids with a prominent band of lignin in the outer S2 layer of the cell wall as well as spiral checks and the absence of an S3 layer (Timell, 1986). Biochemically, compression wood is characterized by high levels of lignin, rich in condensed p-hydroxyphenyl units, as well as reduced cellulose and galactoglucomannan relative to normal wood (Timell, 1986; Nanayakkara et al., 2005; Yeh et al., 2006). Most striking, though, is that β(1,4)-galactan can constitute more than 10% (w/w) of the cell wall material in severe compression wood but is virtually absent in normal wood (Nanayakkara et al., 2005; Yeh et al., 2006). Recent work suggests that β(1,4)-galactan biosynthesis represents an early step in compression wood formation and confirms that its presence is diagnostic for this wood type (Altaner et al., 2007). However, the molecular signal cascades in conifers that lead to the deposition of β(1,4)-galactan are currently not well understood.Immunological studies in conifer species using the monoclonal anti-β(1,4)-galactan LM5 antibody (Jones et al., 1997) indicate that β(1,4)-galactan in compression wood is located in the S1 and outer S2 layers of mature tracheids but is virtually absent from the primary cell walls (Schmitt et al., 2006; Altaner et al., 2007; Möller and Singh, 2007). Instead of β(1,4)-galactan, most conifers contain small amounts of arabinogalactan, a polysaccharide characterized by a highly branched β(1,3)-galactan backbone (Vikkula et al., 1997; Willför et al., 2002; Laine et al., 2004) in their primary cell walls. The ultrastructural distribution of β(1,4)-galactan in compression wood appears to be largely consistent with highly lignified cell wall layers (Möller and Singh, 2007), which might explain the involvement of β(1,4)-galactan in the formation of lignin-carbohydrate complexes (Mukoyoshi et al., 1981; Minor, 1982; Timell, 1986; Laine et al., 2004).The investigation of β(1,4)-galactan structure in preparations from Pinus sylvestris (Laine et al., 2004) and Pinus radiata (Nanayakkara 2007) revealed a linear polymer. In Pinus densiflora Siebold & Zucc., β(1,4)-galactan was found to be slightly branched at positions C2, C3, and C6 (Mukoyoshi et al., 1981). β(1,4)-Galactan in conifers display a high degree of polymerization (DP), which was originally estimated to be in the range of 200 to 300 (Timell, 1986). More recent studies with P. radiata compression wood found the native polysaccharide to have a DP of approximately 380 (Nanayakkara 2007).β(1,4)-Galactan is a very good biochemical marker for compression wood (Altaner et al., 2007), but its physiological role is currently not well understood. Various functions for β(1,4)-galactan in compression wood have been proposed, such as strengthening of the secondary cell wall, absorption of mechanical stresses, and generation of compressive forces (Möller and Singh, 2007). Furthermore, β(1,4)-galactan is also found in tension wood, with a proposed role in cross-linking cellulose microfibrils (Arend, 2008). However, all of those hypotheses on the molecular function of β(1,4)-galactan in reaction wood await experimental verification.Despite substantial efforts to characterize the biosynthesis of this polymer, β(1,4)-galactan biosynthetic enzymes and their corresponding genes are currently unknown (Peugnet et al., 2001; Geshi et al., 2002, 2004; Abdel-Massih et al., 2003; Kato et al., 2003; Ishii et al., 2004; Konishi et al., 2004, 2007; Gorshkova and Morvan, 2006). However, based on other cell wall polysaccharide biosynthetic enzymes, it is likely that the enzymes involved in the biosynthesis of β(1,4)-galactan are either Golgi localized or pass through the Golgi in transit to the apoplastic space (Reyes and Orellana, 2008).To better understand β(1,4)-galactan synthesis in compression wood formation, we sampled both normal wood and severe compression wood from two 6-year-old P. radiata trees, which displayed stark differences in lignin and carbohydrate content and composition. Using these wood samples, new insights into the subcellular localization of β(1,4)-galactan in pine were generated using confocal laser fluorescence microscopy and transmission electron microscopy. An enzyme assay was developed, based on 2-aminopyridine (2AP)-labeled galacto-oligosaccharides as acceptor molecules, which we used to identify and partially purify a robust, microsome-associated, UDP-Gal-dependent β(1,4)-galactosyltransferase (GalT) activity in compression wood that was virtually undetectable in normal wood. Assays of the partially purified GalT revealed that this enzyme has some properties similar to those of previously characterized pectic GalTs, but a marked difference was observed in the size distribution of the enzymatic products.