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Craig Brodersen Steven Jansen Brendan Choat Christopher Rico Jarmila Pittermann 《Plant physiology》2014,165(2):895-904
Plant water transport occurs through interconnected xylem conduits that are separated by partially digested regions in the cell wall known as pit membranes. These structures have a dual function. Their porous construction facilitates water movement between conduits while limiting the spread of air that may enter the conduits and render them dysfunctional during a drought. Pit membranes have been well studied in woody plants, but very little is known about their function in more ancient lineages such as seedless vascular plants. Here, we examine the relationships between conduit air seeding, pit hydraulic resistance, and pit anatomy in 10 species of ferns (pteridophytes) and two lycophytes. Air seeding pressures ranged from 0.8 ± 0.15 MPa (mean ± sd) in the hydric fern Athyrium filix-femina to 4.9 ± 0.94 MPa in Psilotum nudum, an epiphytic species. Notably, a positive correlation was found between conduit pit area and vulnerability to air seeding, suggesting that the rare-pit hypothesis explains air seeding in early-diverging lineages much as it does in many angiosperms. Pit area resistance was variable but averaged 54.6 MPa s m−1 across all surveyed pteridophytes. End walls contributed 52% to the overall transport resistance, similar to the 56% in angiosperm vessels and 64% in conifer tracheids. Taken together, our data imply that, irrespective of phylogenetic placement, selection acted on transport efficiency in seedless vascular plants and woody plants in equal measure by compensating for shorter conduits in tracheid-bearing plants with more permeable pit membranes.Water transport in plants occurs under tension, which renders the xylem susceptible to air entry. This air seeding may lead to the rupture of water columns (cavitation) such that the air expands within conduits to create air-vapor embolisms that block further transport. (Zimmermann and Tyree, 2002). Excessive embolism such as that which occurs during a drought may jeopardize leaf hydration and lead to stomatal closure, overheating, wilting, and possibly death of the plant (Hubbard et al., 2001; Choat et al., 2012; Schymanski et al., 2013). Consequently, strong selection pressure resulted in compartmentalized and redundant plant vascular networks that are adapted to a species habitat water availability by way of life history strategy (i.e. phenology) or resistance to air seeding (Tyree et al., 1994; Mencuccini et al., 2010; Brodersen et al., 2012). The spread of drought-induced embolism is limited primarily by pit membranes, which are permeable, mesh-like regions in the primary cell wall that connect two adjacent conduits. The construction of the pit membrane is such that water easily moves across the membrane between conduits, but because of the small membrane pore size and the presence of a surface coating on the membrane (Pesacreta et al., 2005; Lee et al., 2012), the spread of air and gas bubbles is restricted up to a certain pressure threshold known as the air-seeding pressure (ASP). When xylem sap tension exceeds the air-seeding threshold, air can be aspirated from an air-filled conduit into a functional water-filled conduit through perhaps a large, preexisting pore or one that is created by tension-induced membrane stress (Rockwell et al., 2014). Air seeding leads to cavitation and embolism formation, with emboli potentially propagating throughout the xylem network (Tyree and Sperry, 1988; Brodersen et al., 2013). So, on the one hand, pit membranes are critical to controlling the spread of air throughout the vascular network, while on the other hand, they must facilitate the efficient flow of water between conduits (Choat et al., 2008; Domec et al., 2008; Pittermann et al., 2010; Schulte, 2012). Much is known about such hydraulic tradeoffs in the pit membranes of woody plants, but comparatively little data exist on seedless vascular plants such as ferns and lycophytes. Given that seedless vascular plants may bridge the evolutionary transition from bryophytes to woody plants, the lack of functional data on pit membrane structure in early-derived tracheophytes is a major gap in our understanding of the evolution of plant water transport.In woody plants, pit membranes fall into one of two categories: the torus-margo type found in most gymnosperms and the homogenous pit membrane characteristic of angiosperms (Choat et al., 2008; Choat and Pittermann, 2009). In conifers, water moves from one tracheid to another through the margo region of the membrane, with the torus sealing the pit aperture should one conduit become embolized. Air seeding occurs when water potential in the functional conduit drops low enough to dislodge the torus from its sealing position, letting air pass through the pit aperture into the water-filled tracheid (Domec et al., 2006; Delzon et al., 2010; Pittermann et al., 2010; Schulte, 2012; but see Jansen et al., 2012). Across north-temperate conifer species, larger pit apertures correlate with lower pit resistance to water flow (rpit; MPa s m−1), but it is the ratio of torus-aperture overlap that sets a species cavitation resistance (Pittermann et al., 2006, 2010; Domec et al., 2008; Hacke and Jansen, 2009). A similar though mechanistically different tradeoff exists in angiosperm pit membranes. Here, air seeding reflects a probabilistic relationship between membrane porosity and the total area of pit membranes present in the vessel walls. Specifically, the likelihood of air aspirating into a functional conduit is determined by the combination of xylem water potential and the diameter of the largest pore and/or the weakest zone in the cellulose matrix in the vessel’s array of pit membranes (Wheeler et al., 2005; Hacke et al., 2006; Christman et al., 2009; Rockwell et al., 2014). As it has come to be known, the rare-pit hypothesis suggests that the infrequent, large-diameter leaky pore giving rise to that rare pit reflects some combination of pit membrane traits such as variation in conduit membrane area (large or small), membrane properties (tight or porous), and hydrogel membrane chemistry (Hargrave et al., 1994; Choat et al., 2003; Wheeler et al., 2005; Hacke et al., 2006; Christman et al., 2009; Lee et al., 2012; Plavcová et al., 2013; Rockwell et al., 2014). The maximum pore size is critical because, per the Young-Laplace law, the larger the radius of curvature, the lower the air-water pressure difference under which the contained meniscus will fail (Jarbeau et al., 1995; Choat et al., 2003; Jansen et al., 2009). Consequently, angiosperms adapted to drier habitats may exhibit thicker, denser, smaller, and less abundant pit membranes than plants occupying regions with higher water availability (Wheeler et al., 2005; Hacke et al., 2007; Jansen et al., 2009; Lens et al., 2011; Scholz et al., 2013). However, despite these qualitative observations, there is no evidence that increased cavitation resistance arrives at the cost of higher rpit. Indeed, the bulk of the data suggest that prevailing pit membrane porosity is decoupled from the presence of the single largest pore that allows air seeding to occur (Choat et al., 2003; Wheeler et al., 2005
Hacke et al., 2006, 2007).As water moves from one conduit to another, pit membranes offer considerable hydraulic resistance throughout the xylem network. On average, rpit contributes 64% and 56% to transport resistance in conifers and angiosperms, respectively (Wheeler et al., 2005; Pittermann et al., 2006; Sperry et al., 2006). In conifers, the average rpit is estimated at 6 ± 1 MPa s m−1, almost 60 times lower than the 336 ± 81 MPa s m−1 computed for angiosperms (Wheeler et al., 2005; Hacke et al., 2006; Sperry et al., 2006). Presumably, the high porosity of conifer pits compensates for the higher transport resistance offered by a vascular system composed of narrow, short, single-celled conduits (Pittermann et al., 2005; Sperry et al., 2006).Transport in seedless vascular plants presents an interesting conundrum because, with the exception of a handful of species, their primary xylem is composed of tracheids, the walls of which are occupied by homogenous pit membranes (Gibson et al., 1985; Carlquist and Schneider, 2001, 2007; but see Morrow and Dute, 1998, for torus-margo membranes in Botrychium spp.). At first pass, this combination of traits appears hydraulically maladaptive, but several studies have shown that ferns can exhibit transport capacities that are on par with more recently evolved plants (Wheeler et al., 2005; Watkins et al., 2010; Pittermann et al., 2011, 2013; Brodersen et al., 2012). Certainly, several taxa possess large-diameter, highly overlapping conduits, some even have vessels such as Pteridium aquilinum and many species have high conduit density, all of which could contribute to increased hydraulic efficiency (Wheeler et al., 2005; Pittermann et al., 2011, 2013). But how do the pit membranes of seedless vascular plants compare? Scanning electron micrographs of fern and lycopod xylem conduits suggest that they are thin, diaphanous, and susceptible to damage during specimen preparation (Carlquist and Schneider 2001, 2007). Consistent with such observations, two estimates of rpit imply that rpit in ferns may be significantly lower than in angiosperms; Wheeler et al. (2005) calculated rpit in the fern Pteridium aquilinum at 31 MPa s m−1, while Schulte et al. (1987) estimated rpit at 1.99 MPa s m−1 in the basal fern Psilotum nudum. The closest structural analogy to seedless vascular plant tracheids can be found in the secondary xylem of the early-derived vesselless angiosperms, in which tracheids possess homogenous pit membranes with rpit values that at 16 MPa s m−1 are marginally higher than those of conifers (Hacke et al., 2007). Given that xylem in seedless vascular plants is functionally similar to that in vesselless angiosperms, we expected convergent rpit values in these two groups despite their phylogenetic distance. We tested this hypothesis, as well as the intrinsic cavitation resistance of conduits in seedless vascular plants, by scrutinizing the pit membranes of ferns and fern allies using the anatomical and experimental approaches applied previously to woody taxa. In particular, we focused on the relationship between pit membrane traits and cavitation resistance at the level of the individual conduit. 相似文献
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O Maximyuk V Khmyz C-J Lindskog V Vukojevi? T Ivanova I Bazov K F Hauser G Bakalkin O Krishtal 《Cell death & disease》2015,6(3):e1683
Neuropeptides induce signal transduction across the plasma membrane by acting through cell-surface receptors. The dynorphins, endogenous ligands for opioid receptors, are an exception; they also produce non-receptor-mediated effects causing pain and neurodegeneration. To understand non-receptor mechanism(s), we examined interactions of dynorphins with plasma membrane. Using fluorescence correlation spectroscopy and patch-clamp electrophysiology, we demonstrate that dynorphins accumulate in the membrane and induce a continuum of transient increases in ionic conductance. This phenomenon is consistent with stochastic formation of giant (~2.7 nm estimated diameter) unstructured non-ion-selective membrane pores. The potency of dynorphins to porate the plasma membrane correlates with their pathogenic effects in cellular and animal models. Membrane poration by dynorphins may represent a mechanism of pathological signal transduction. Persistent neuronal excitation by this mechanism may lead to profound neuropathological alterations, including neurodegeneration and cell death.Neuropeptides are the largest and most diverse family of neurotransmitters. They are released from axon terminals and dendrites, diffuse to pre- or postsynaptic neuronal structures and activate membrane G-protein-coupled receptors. Prodynorphin (PDYN)-derived opioid peptides including dynorphin A (Dyn A), dynorphin B (Dyn B) and big dynorphin (Big Dyn) consisting of Dyn A and Dyn B are endogenous ligands for the κ-opioid receptor. Acting through this receptor, dynorphins regulate processing of pain and emotions, memory acquisition and modulate reward induced by addictive substances.1, 2, 3, 4 Furthermore, dynorphins may produce robust cellular and behavioral effects that are not mediated through opioid receptors.5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29 As evident from pharmacological, morphological, genetic and human neuropathological studies, these effects are generally pathological, including cell death, neurodegeneration, neurological dysfunctions and chronic pain. Big Dyn is the most active pathogenic peptide, which is about 10- to 100-fold more potent than Dyn A, whereas Dyn B does not produce non-opioid effects.16, 17, 22, 25 Big Dyn enhances activity of acid-sensing ion channel-1a (ASIC1a) and potentiates ASIC1a-mediated cell death in nanomolar concentrations30, 31 and, when administered intrathecally, induces characteristic nociceptive behavior at femtomolar doses.17, 22 Inhibition of endogenous Big Dyn degradation results in pathological pain, whereas prodynorphin (Pdyn) knockout mice do not maintain neuropathic pain.22, 32 Big Dyn differs from its constituents Dyn A and Dyn B in its unique pattern of non-opioid memory-enhancing, locomotor- and anxiolytic-like effects.25Pathological role of dynorphins is emphasized by the identification of PDYN missense mutations that cause profound neurodegeneration in the human brain underlying the SCA23 (spinocerebellar ataxia type 23), a very rare dominantly inherited neurodegenerative disorder.27, 33 Most PDYN mutations are located in the Big Dyn domain, demonstrating its critical role in neurodegeneration. PDYN mutations result in marked elevation in dynorphin levels and increase in its pathogenic non-opioid activity.27, 34 Dominant-negative pathogenic effects of dynorphins are not produced through opioid receptors.ASIC1a, glutamate NMDA (N-methyl-d-aspartate) and AMPA (α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid)/kainate ion channels, and melanocortin and bradykinin B2 receptors have all been implicated as non-opioid dynorphin targets.5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 30, 31, 35, 36 Multiplicity of these targets and their association with the cellular membrane suggest that their activation is a secondary event triggered by a primary interaction of dynorphins with the membrane. Dynorphins are among the most basic neuropeptides.37, 38 The basic nature is also a general property of anti-microbial peptides (AMPs) and amyloid peptides that act by inducing membrane perturbations, altering membrane curvature and causing pore formation that disrupts membrane-associated processes including ion fluxes across the membrane.39 The similarity between dynorphins and these two peptide groups in overall charge and size suggests a similar mode of their interactions with membranes.In this study, we dissect the interactions of dynorphins with the cell membrane, the primary event in their non-receptor actions. Using fluorescence imaging, correlation spectroscopy and patch-clamp techniques, we demonstrate that dynorphin peptides accumulate in the plasma membrane in live cells and cause a profound transient increase in cell membrane conductance. Membrane poration by endogenous neuropeptides may represent a novel mechanism of signal transduction in the brain. This mechanism may underlie effects of dynorphins under pathological conditions including chronic pain and tissue injury. 相似文献
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Carla Minguet-Parramona Yizhou Wang Adrian Hills Silvere Vialet-Chabrand Howard Griffiths Simon Rogers Tracy Lawson Virgilio L. Lew Michael R. Blatt 《Plant physiology》2016,170(1):33-42
Oscillations in cytosolic-free Ca2+ concentration ([Ca2+]i) have been proposed to encode information that controls stomatal closure. [Ca2+]i oscillations with a period near 10 min were previously shown to be optimal for stomatal closure in Arabidopsis (Arabidopsis thaliana), but the studies offered no insight into their origins or mechanisms of encoding to validate a role in signaling. We have used a proven systems modeling platform to investigate these [Ca2+]i oscillations and analyze their origins in guard cell homeostasis and membrane transport. The model faithfully reproduced differences in stomatal closure as a function of oscillation frequency with an optimum period near 10 min under standard conditions. Analysis showed that this optimum was one of a range of frequencies that accelerated closure, each arising from a balance of transport and the prevailing ion gradients across the plasma membrane and tonoplast. These interactions emerge from the experimentally derived kinetics encoded in the model for each of the relevant transporters, without the need of any additional signaling component. The resulting frequencies are of sufficient duration to permit substantial changes in [Ca2+]i and, with the accompanying oscillations in voltage, drive the K+ and anion efflux for stomatal closure. Thus, the frequency optima arise from emergent interactions of transport across the membrane system of the guard cell. Rather than encoding information for ion flux, these oscillations are a by-product of the transport activities that determine stomatal aperture.Stomata in the leaf epidermis are the main pathway both for CO2 entry for photosynthesis and for foliar water loss by transpiration. Guard cells surround the stomatal pore and regulate the aperture, balancing the often conflicting demands for CO2 and water conservation. Guard cells open and close the pore by expanding and contracting through the uptake and loss, respectively, of osmotic solutes, notably of K+, Cl−, and malate2− (Mal2−; Pandey et al., 2007; Kim et al., 2010; Roelfsema and Hedrich, 2010; Lawson and Blatt, 2014). These transport processes comprise the final effectors of a regulatory network that coordinates transport across the plasma membrane and tonoplast, and maintains the homeostasis of the guard cell. A number of well-defined signals—including light, CO2, drought and the water stress hormone abscisic acid (ABA)—act on this network, altering transport, solute content, turgor and cell volume, and ultimately stomatal aperture.Much research has focused on stomatal closure, underscoring both Ca2+-independent and Ca2+-dependent signaling. Of the latter, elevated cytosolic-free Ca2+ concentration ([Ca2+]i) inactivates inward-rectifying K+ channels (IK,in) to prevent K+ uptake and activates Cl− (anion) channels (ICl) at the plasma membrane to depolarize the membrane and engage K+ efflux through outward-rectifying K+ channels (IK,out; Keller et al., 1989; Blatt et al., 1990; Thiel et al., 1992; Lemtiri-Chlieh and MacRobbie, 1994). ABA, and most likely CO2 (Kim et al., 2010), elevate [Ca2+]i by facilitating Ca2+ entry at the plasma membrane to trigger Ca2+ release from endomembrane stores, a process often described as Ca2+-induced Ca2+ release (Grabov and Blatt, 1998, 1999). The hormone promotes Ca2+ influx by activating Ca2+ channels (ICa) at the plasma membrane, even in isolated membrane patches (Hamilton et al., 2000, 2001), which is linked to reactive oxygen species (Kwak et al., 2003; Wang et al., 2013). In parallel, cADP-ribose and nitric oxide promote endomembrane Ca2+ release and [Ca2+]i elevation (Leckie et al., 1998; Neill et al., 2002; Garcia-Mata et al., 2003; Blatt et al., 2007). Best estimates indicate that endomembrane release accounts for more than 95% of the Ca2+ entering the cytosol to raise [Ca2+]i (Chen et al., 2012; Wang et al., 2012).One feature of stomatal response to ABA, and indeed to a range of stimuli both hormonal as well as external, is its capacity for oscillations both in membrane voltage and [Ca2+]i. Guard cell [Ca2+]i at rest is typically around 100 to 200 nm, as it is in virtually all living cells. In response to ABA, [Ca2+]i can rise above 1 μm—and locally, most likely above 10 μm—often in cyclic transients of tens of seconds to several minutes’ duration in association with oscillations in voltage and stomatal closure (Gradmann et al., 1993; McAinsh et al., 1995; Webb et al., 1996; Grabov and Blatt, 1998, 1999; Staxen et al., 1999; Allen et al., 2001). In principle, cycling in voltage and [Ca2+]i arises as closure is accelerated with a controlled release of K+, Cl−, and Mal2− from the guard cell and is subject to extracellular ion concentrations (Gradmann et al., 1993; Chen et al., 2012). However, it has been proposed that these, and similar oscillations in a variety of plant cell models, serve as physiological signals in their own right (McAinsh et al., 1995; Ehrhardt et al., 1996; Taylor et al., 1996). In support of such a signaling role, experiments designed to impose [Ca2+]i (and voltage) oscillations in guard cells have yielded an optimal frequency for closure with a period near 10 min (Allen et al., 2001). Nonetheless, the studies offer no mechanistic explanation for this optimum that could validate a causal role in signaling, and none has been forthcoming since. Here we address questions of how such optimal frequencies in [Ca2+]i oscillation arise and their relevance for stomatal closure, using quantitative systems analysis of guard cell transport and homeostasis. Our findings indicate that oscillations in voltage and [Ca2+]i, and their optima associated with stomatal closure, are most simply explained as emerging from the interactions between ion transporters that drive stomatal closure. Thus, we conclude that these oscillations do not control, but are a by-product of the transport that determines stomatal aperture. 相似文献
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Jose Salvador Rubio Asensio Shimon Rachmilevitch Arnold J. Bloom 《Plant physiology》2015,168(1):156-163
A major contributor to the global carbon cycle is plant respiration. Elevated atmospheric CO2 concentrations may either accelerate or decelerate plant respiration for reasons that have been uncertain. We recently established that elevated CO2 during the daytime decreases plant mitochondrial respiration in the light and protein concentration because CO2 slows the daytime conversion of nitrate (NO3−) into protein. This derives in part from the inhibitory effect of CO2 on photorespiration and the dependence of shoot NO3− assimilation on photorespiration. Elevated CO2 also inhibits the translocation of nitrite into the chloroplast, a response that influences shoot NO3− assimilation during both day and night. Here, we exposed Arabidopsis (Arabidopsis thaliana) and wheat (Triticum aestivum) plants to daytime or nighttime elevated CO2 and supplied them with NO3− or ammonium as a sole nitrogen (N) source. Six independent measures (plant biomass, shoot NO3−, shoot organic N, 15N isotope fractionation, 15NO3− assimilation, and the ratio of shoot CO2 evolution to O2 consumption) indicated that elevated CO2 at night slowed NO3− assimilation and thus decreased dark respiration in the plants reliant on NO3−. These results provide a straightforward explanation for the diverse responses of plants to elevated CO2 at night and suggest that soil N source will have an increasing influence on the capacity of plants to mitigate human greenhouse gas emissions.The CO2 concentration in Earth’s atmosphere has increased from about 270 to 400 µmol mol–1 since 1800, and may double before the end of the century (Intergovernmental Panel on Climate Change, 2013). Plant responses to such increases are highly variable, but plant nitrogen (N) concentrations generally decline under elevated CO2 (Cotrufo et al., 1998; Long et al., 2004). One explanation for this decline is that CO2 inhibits nitrate (NO3−) assimilation into protein in the shoots of C3 plants during the daytime (Bloom et al., 2002, 2010, 2012, 2014; Cheng et al., 2012; Pleijel and Uddling, 2012; Myers et al., 2014; Easlon et al., 2015; Pleijel and Högy, 2015). This derives in part from the inhibitory effect of CO2 on photorespiration (Foyer et al., 2009) and the dependence of shoot NO3− assimilation on photorespiration (Rachmilevitch et al., 2004; Bloom, 2015).A key factor in global carbon budgets is plant respiration at night (Amthor, 1991; Farrar and Williams, 1991; Drake et al., 1999; Leakey et al., 2009). Nighttime elevated CO2 may inhibit, have a negligible effect on, or stimulate dark respiration, depending on the plant species (Bunce, 2001, 2003; Wang and Curtis, 2002), plant development stage (Wang et al., 2001; Li et al., 2013), experimental approach (Griffin et al., 1999; Baker et al., 2000; Hamilton et al., 2001; Bruhn et al., 2002; Jahnke and Krewitt, 2002; Bunce, 2004), and total N supply (Markelz et al., 2014). The current study is, to our knowledge, the first to examine the influence of N source, NO3− versus ammonium (NH4+), on plant dark respiration at elevated CO2 during the night.Plant organic N compounds account for less than 5% of the total dry weight of a plant, but conversion of NO3− into organic N expends about 25% of the total energy in shoots (Bloom et al., 1989) and roots (Bloom et al., 1992). During the day, photorespiration supplies a portion of the energy (Rachmilevitch et al., 2004; Foyer et al., 2009), but at night, this energetic cost is borne entirely by the respiration of C substrates (Amthor, 1995) and may divert a substantial amount of reductant from the mitochondrial electron transport chain (Cousins and Bloom, 2004). The relative importance of NO3− assimilation at night versus the day, however, is still a matter of intense debate (Nunes-Nesi et al., 2010). Here, we estimated NO3− assimilation using several independent methods and show in Arabidopsis (Arabidopsis thaliana) and wheat (Triticum aestivum), two diverse C3 plants, that NO3− assimilation at night can be substantial, and that elevated CO2 at night inhibits this process. 相似文献
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Lyudmila Y. Kadyrova Basanta K. Dahal Vaibhavi Gujar James M. Daley Patrick Sung Farid A. Kadyrov 《The Journal of biological chemistry》2022,298(4)
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. 相似文献
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Victoria Fernández Domingo Sancho-Knapik Paula Guzmán José Javier Peguero-Pina Luis Gil George Karabourniotis Mohamed Khayet Costas Fasseas José Alejandro Heredia-Guerrero Antonio Heredia Eustaquio Gil-Pelegrín 《Plant physiology》2014,166(1):168-180
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 (Wa) 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? 相似文献
11.
Girdhar K. Pandey Poonam Kanwar Amarjeet Singh Leonie Steinhorst Amita Pandey Akhlilesh K. Yadav Indu Tokas Sibaji K. Sanyal Beom-Gi Kim Sung-Chul Lee Yong-Hwa Cheong J?rg Kudla Sheng Luan 《Plant physiology》2015,169(1):780-792
The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca2+) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca2+ changes are primarily perceived by several Ca2+ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca2+-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca2+-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca2+ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca2+ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca2+ sensors CBL2 and CBL3 under salt stress condition. 相似文献
12.
Nikolaus Pfanner Martin van der Laan Paolo Amati Roderick A. Capaldi Amy A. Caudy Agnieszka Chacinska Manjula Darshi Markus Deckers Suzanne Hoppins Tateo Icho Stefan Jakobs Jianguo Ji Vera Kozjak-Pavlovic Chris Meisinger Paul R. Odgren Sang Ki Park Peter Rehling Andreas S. Reichert M. Saeed Sheikh Susan S. Taylor Nobuo Tsuchida Alexander M. van der Bliek Ida J. van der Klei Jonathan S. Weissman Benedikt Westermann Jiping Zha Walter Neupert Jodi Nunnari 《The Journal of cell biology》2014,204(7):1083-1086
The mitochondrial inner membrane contains a large protein complex that functions in inner membrane organization and formation of membrane contact sites. The complex was variably named the mitochondrial contact site complex, mitochondrial inner membrane organizing system, mitochondrial organizing structure, or Mitofilin/Fcj1 complex. To facilitate future studies, we propose to unify the nomenclature and term the complex “mitochondrial contact site and cristae organizing system” and its subunits Mic10 to Mic60.Mitochondria possess two membranes of different architecture and function (Palade, 1952; Hackenbrock, 1968). Both membranes work together for essential shared functions, such as protein import (Schatz, 1996; Neupert and Herrmann, 2007; Chacinska et al., 2009). The outer membrane harbors machinery that controls the shape of the organelle and is crucial for the communication of mitochondria with the rest of the cell. The inner membrane harbors the complexes of the respiratory chain, the F1Fo-ATP synthase, numerous metabolite carriers, and enzymes of mitochondrial metabolism. It consists of two domains: the inner boundary membrane, which is adjacent to the outer membrane, and invaginations of different shape, termed cristae (Werner and Neupert, 1972; Frey and Mannella, 2000; Hoppins et al., 2007; Pellegrini and Scorrano, 2007; Zick et al., 2009; Davies et al., 2011). Tubular openings, termed crista junctions (Perkins et al., 1997), connect inner boundary membrane and cristae membranes (Fig. 1, A and B). Respiratory chain complexes and the F1Fo-ATP synthase are preferentially located in the cristae membranes, whereas preprotein translocases are enriched in the inner boundary membrane (Vogel et al., 2006; Wurm and Jakobs, 2006; Davies et al., 2011). Contact sites between outer membrane and inner boundary membrane promote import of preproteins, metabolite channeling, lipid transport, and membrane dynamics (Frey and Mannella, 2000; Sesaki and Jensen, 2004; Hoppins et al., 2007, 2011; Neupert and Herrmann, 2007; Chacinska et al., 2009; Connerth et al., 2012; van der Laan et al., 2012).Open in a separate windowFigure 1.MICOS complex. (A) The MICOS complex (hypothetical model), previously also termed MINOS, MitOS, or Mitofilin/Fcj1 complex, is required for maintenance of the characteristic architecture of the mitochondrial inner membrane (IM) and forms contact sites with the outer membrane (OM). In budding yeast, six subunits of MICOS have been identified. All subunits are exposed to the intermembrane space (IMS), five are integral inner membrane proteins (Mic10, Mic12, Mic26, Mic27, and Mic60), and one is a peripheral inner membrane protein (Mic19). Mic26 is related to Mic27; however, mic26Δ yeast cells show considerably less severe defects of mitochondrial inner membrane architecture than mic27Δ cells (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011). The MICOS complex of metazoa additionally contains Mic25, which is related to Mic19, yet subunits corresponding to Mic12 and Mic26 have not been identified so far. MICOS subunits that have been conserved in most organisms analyzed are indicated by bold boundary lines. (B, top) Wild-type architecture of the mitochondrial inner membrane with crista junctions and cristae. (bottom) This architecture is considerably altered in micos mutant mitochondria: most cristae membranes are detached from the inner boundary membrane and form internal membrane stacks. In some micos mutants (deficiency of mammalian Mic19 or Mic25), a loss of cristae membranes was observed (Darshi et al., 2011; An et al., 2012). Figure by M. Bohnert (Institute of Biochemistry and Molecular Biology, University of Freiburg, Freiburg, Germany).To understand the complex architecture of mitochondria, it will be crucial to identify the molecular machineries that control the interaction between mitochondrial outer and inner membranes and the characteristic organization of the inner membrane. A convergence of independent studies led to the identification of a large heterooligomeric protein complex of the mitochondrial inner membrane conserved from yeast to humans that plays crucial roles in the maintenance of crista junctions, inner membrane architecture, and formation of contact sites to the outer membrane (Fig. 1 A). Several names were used by different research groups to describe the complex, including mitochondrial contact site (MICOS) complex, mitochondrial inner membrane organizing system (MINOS), mitochondrial organizing structure (MitOS), Mitofilin complex, or Fcj1 (formation of crista junction protein 1) complex (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012). Mitofilin, also termed Fcj1, was the first component identified (Icho et al., 1994; Odgren et al., 1996; Gieffers et al., 1997; John et al., 2005) and was observed enriched at crista junctions (Rabl et al., 2009). Mutants of Mitofilin/Fcj1 as well as of other MICOS/MINOS/MitOS subunits show a strikingly altered inner membrane architecture. They lose crista junctions and contain large internal membrane stacks, the respiratory activity is reduced, and mitochondrial DNA nucleoids are altered (Fig. 1 B; John et al., 2005; Hess et al., 2009; Rabl et al., 2009; Mun et al., 2010; Harner et al., 2011; Head et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012; Itoh et al., 2013). It has been reported that the complex interacts with a variety of outer membrane proteins, such as channel proteins and components of the protein translocases and mitochondrial fusion machines, and defects impair the biogenesis of mitochondrial proteins (Xie et al., 2007; Darshi et al., 2011; Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012; An et al., 2012; Bohnert et al., 2012; Körner et al., 2012; Ott et al., 2012; Zerbes et al., 2012; Jans et al., 2013; Weber et al., 2013). The MICOS/MINOS/MitOS/Mitofilin/Fcj1 complex thus plays crucial roles in mitochondrial architecture, dynamics, and biogenesis. However, communication of results in this rapidly developing field has been complicated by several different nomenclatures used for the complex as well as for its subunits (Standard name Former names Yeast ORF References Complex MICOS MINOS, MitOS, MIB, Mitofilin complex, and Fcj1 complex Xie et al., 2007; Rabl et al., 2009; Darshi et al., 2011; Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012; An et al., 2012; Bohnert et al., 2012; Ott et al., 2012; Jans et al., 2013; Weber et al., 2013 Subunits Mic10 Mcs10, Mio10, Mos1, and MINOS1 YCL057C-A Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Alkhaja et al., 2012; Itoh et al., 2013; Jans et al., 2013; Varabyova et al., 2013 Mic12 Aim5, Fmp51, and Mcs12 YBR262C Hess et al., 2009; Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Varabyova et al., 2013 Mic19 Aim13, Mcs19, CHCH-3, CHCHD3, and MINOS3 YFR011C Xie et al., 2007; Hess et al., 2009; Darshi et al., 2011; Head et al., 2011; Alkhaja et al., 2012; Ott et al., 2012; Jans et al., 2013; Varabyova et al., 2013 Mic25 (metazoan Mic19 homologue) CHCHD6 and CHCM1 Xie et al., 2007; An et al., 2012 Mic26 Mcs29, Mio27, and Mos2 YGR235C Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011 Mic27 Aim37, Mcs27, APOOL, and MOMA-1 YNL100W Hess et al., 2009; Harner et al., 2011; Head et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011; Weber et al., 2013 Mic60 Fcj1, Aim28, Fmp13, Mitofilin, HMP, IMMT, and MINOS2 YKR016W Icho et al., 1994; Odgren et al., 1996; Gieffers et al., 1997; John et al., 2005; Wang et al., 2008; Rabl et al., 2009; Rossi et al., 2009; Mun et al., 2010; Park et al., 2010; Körner et al., 2012; Zerbes et al., 2012; Itoh et al., 2013; Varabyova et al., 2013