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Suspension-cultured
Chenopodium album L. cells are capable of continuous,
long-term growth on a boron-deficient medium. Compared with cultures
grown with boron, these cultures contained more enlarged and detached
cells, had increased turbidity due to the rupture of a small number of
cells, and contained cells with an increased cell wall pore size. These
characteristics were reversed by the addition of boric acid (≥7
μm) to the boron-deficient cells. C. album
cells grown in the presence of 100 μm boric acid entered
the stationary phase when they were not subcultured, and remained
viable for at least 3 weeks. The transition from the growth phase to
the stationary phase was accompanied by a decrease in the wall pore
size. Cells grown without boric acid or with 7 μm boric
acid were not able to reduce their wall pore size at the transition to
the stationary phase. These cells could not be kept viable in the
stationary phase, because they continued to expand and died as a result
of wall rupture. The addition of 100 μm boric acid
prevented wall rupture and the wall pore size was reduced to normal
values. We conclude that boron is required to maintain the normal pore
structure of the wall matrix and to mechanically stabilize the wall at
growth termination.The ultrastructure and physical properties of plant cell walls are
known to be affected by boron deficiency (Kouchi and Kumazawa, 1976;
Hirsch and Torrey, 1980; Fischer and Hecht-Buchholz, 1985; Matoh et
al., 1992; Hu and Brown, 1994; Findeklee and Goldbach, 1996). Moreover,
boron is predominantly localized in the cell wall when plants are grown
with suboptimal boron (Loomis and Durst, 1991; Matoh et al., 1992; Hu
and Brown, 1994; Hu et al., 1996). In radish, >80% of the cell wall
boron is present in the pectic polysaccharide RG-II (Matoh et al.,
1993; Kobayashi et al., 1996), which is now known to exist as a dimer
that is cross-linked by a borate ester between two apiosyl residues
(Kobayashi et al., 1996; O''Neill et al., 1996). Dimeric RG-II is
unusually stable at low pH and is present in a large number of plant
species (Ishii and Matsunaga, 1996; Kobayashi et al., 1996, 1997; Matoh
et al., 1996; O''Neill et al., 1996; Pellerin et al., 1996; Kaneko et
al., 1997). The widespread occurrence and conserved structure of RG-II
(Darvill et al., 1978; O''Neill et al., 1990) have led to the
suggestion that borate ester cross-linked RG-II is required for the
development of a normal cell wall (O''Neill et al., 1996; Matoh, 1997).One approach for determining the function of boron in plant cell walls
is to compare the responses to boron deficiency of growing plant cells
that are dividing and synthesizing primary cell walls with those of
growth-limited plant cells in which the synthesis of primary cell walls
is negligible. Suspension-cultured cells are well suited for this
purpose because they may be reversibly transferred from a growth phase
to a stationary phase. Continuous cell growth phase is maintained by
frequent transfer of the cells into new growth medium (King, 1981;
Kandarakov et al., 1994), whereas a stationary cell population
is obtained by feeding the cells with Suc and by not subculturing them.
Cells in the stationary phase are characterized by mechanically
stabilized primary walls and reduced biosynthetic activity. Here we
describe the responses of suspension-cultured Chenopodium
album L. cells in the growth and stationary phases to boron
deficiency. These cells have a high specific-growth rate, no
significant lag phase, and reproducible changes in their wall pore size
during the transition from the growth phase to the stationary phase
(Titel et al., 1997). 相似文献
6.
Heme and chlorophyll accumulate to
high levels in legume root nodules and in photosynthetic tissues,
respectively, and they are both derived from the universal tetrapyrrole
precursor δ-aminolevulinic acid (ALA). The first committed step in
ALA and tetrapyrrole synthesis is catalyzed by glutamyl-tRNA reductase
(GTR) in plants. A soybean (Glycine max) root-nodule
cDNA encoding GTR was isolated by complementation of an
Escherichia coli GTR-defective mutant for restoration of
ALA prototrophy. Gtr mRNA was very low in uninfected
roots but accumulated to high levels in root nodules. The induction of
Gtr mRNA in developing nodules was subsequent to that of
the gene Enod2 (early nodule)
and coincided with leghemoglobin mRNA accumulation. Genomic analysis
revealed two Gtr genes, Gtr1 and a 3′
portion of Gtr2, which were isolated from the soybean
genome. RNase-protection analysis using probes specific to
Gtr1 and Gtr2 showed that both genes were
expressed, but Gtr1 mRNA accumulated to significantly
higher levels. In addition, the qualitative patterns of expression of
Gtr1 and Gtr2 were similar to each other
and to total Gtr mRNA in leaves and nodules of mature
plants and etiolated plantlets. The data indicate that
Gtr1 is universal for tetrapyrrole synthesis and that a
Gtr gene specific for a tissue or tetrapyrrole is
unlikely. We suggest that ALA synthesis in specialized root nodules
involves an altered spatial expression of genes that are otherwise
induced strongly only in photosynthetic tissues of uninfected plants.Soybean (Glycine max) and numerous other legumes can
establish a symbiosis with rhizobia, resulting in the formation of root
nodules comprising specialized plant and bacterial cells (for review,
see Mylona et al., 1995). Rhizobia reduce atmospheric nitrogen to
ammonia within nodules, which is assimilated by the plant host to
fulfill its nutritional nitrogen requirement. The high energy
requirement for nitrogen fixation necessitates efficient respiration by
the prokaryote within the microaerobic milieu of the nodule. The plant
host synthesizes a nodule-specific hemoglobin (leghemoglobin) that
serves to facilitate oxygen diffusion to the bacterial endosymbiont and
to buffer the free oxygen concentration at a low
tension (for review, see Appleby, 1992). Both of these functions
require that the hemoglobin concentration be high, and, indeed, it
exceeds 1 mm in soybean nodules (Appleby, 1984)
and is the predominant plant protein in that organ. Once thought to be
confined to legume nodules, hemoglobins are found throughout the plant
kingdom, and leghemoglobin likely represents a specialization of a
general plant phenomenon (for review, see Hardison, 1996). A gene
encoding a nonsymbiotic hemoglobin has been identified in soybean and
other legumes (Andersson et al., 1996); therefore, expression in
nodules involves the specific activation of a subset of genes within a
gene family. Leghemoglobin genes may have arisen from gene duplication,
followed by specialization (Andersson et al., 1996).Hemes and chlorophyll are tetrapyrroles synthesized
from common precursors; chlorophyll is quantitatively the major
tetrapyrrole in plants, with heme and other tetrapyrroles being present
in minor amounts. Legume root nodules represent an exception, in which
heme is synthesized in high quantity in the absence of chlorophyll,
thus requiring the activity of enzymes not normally expressed highly in
nonphotosynthetic tissues. Heme is synthesized from the universal
tetrapyrrole precursor ALA by seven successive enzymatic steps;
chlorophyll formation diverges after the synthesis of protoporphyrin,
the immediate heme precursor (for review, see O''Brian, 1996).
Biochemical and genetic evidence shows that soybean heme biosynthesis
genes are strongly induced in root nodules (Sangwan and O''Brian, 1991,
1992, 1993; Madsen et al., 1993; Kaczor et al., 1994; Frustaci et al.,
1995; Santana et al., 1998), and immunohistochemical studies
demonstrate that induction is concentrated in infected nodule cells
(Santana et al., 1998).ALA is synthesized from Glu in plants by a three-step mechanism called
the C5 pathway (Fig.
(Fig.1);1); the latter two steps are committed to
ALA synthesis and are catalyzed by GTR and GSAT, respectively (for
review, see Beale and Weinstein, 1990; Jahn et al., 1991). Plant cDNA
or genes encoding GTR (Gtr, also called HemA) and
GSAT (Gsa) have been identified in several plant species
(Grimm, 1990; Sangwan and O''Brian, 1993; Hofgen et al., 1994; Ilag et
al., 1994; Frustaci et al., 1995; Wenzlau and Berry-Lowe, 1995; Bougri
and Grimm, 1996; Kumar et al., 1996; Tanaka et al., 1996). Two genes
for each enzyme have been described, and some genes are reported to be
specific to a tissue, tetrapyrrole, or light regimen (Bougri and Grimm,
1996; Kumar et al., 1996; Tanaka et al., 1996). However, soybean
Gsa1 is highly expressed in both leaves and nodules and
contains a cis-acting element in its promoter that binds to
a nuclear factor found in both tissues. (Frustaci et al., 1995). In
this study we isolated soybean Gtr1 and characterized the
genetic basis of GTR expression in root nodules.
Figure 1C5 pathway for ALA synthesis. The
committed steps for ALA synthesis catalyzed by GTR and GSAT are boxed.
Glutamyl-tRNA synthetase (GluRS) and glutamyl-tRNAGlu also
participate in protein synthesis. The gene designations in plants are
shown in parentheses ... 相似文献
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SPA2 encodes a yeast protein that is one of the first proteins to localize to sites of polarized growth, such as the shmoo tip and the incipient bud. The dynamics and requirements for Spa2p localization in living cells are examined using Spa2p green fluorescent protein fusions. Spa2p localizes to one edge of unbudded cells and subsequently is observable in the bud tip. Finally, during cytokinesis Spa2p is present as a ring at the mother–daughter bud neck. The bud emergence mutants bem1 and bem2 and mutants defective in the septins do not affect Spa2p localization to the bud tip. Strikingly, a small domain of Spa2p comprised of 150 amino acids is necessary and sufficient for localization to sites of polarized growth. This localization domain and the amino terminus of Spa2p are essential for its function in mating. Searching the yeast genome database revealed a previously uncharacterized protein which we name, Sph1p (Spa2p homolog), with significant homology to the localization domain and amino terminus of Spa2p. This protein also localizes to sites of polarized growth in budding and mating cells. SPH1, which is similar to SPA2, is required for bipolar budding and plays a role in shmoo formation. Overexpression of either Spa2p or Sph1p can block the localization of either protein fused to green fluorescent protein, suggesting that both Spa2p and Sph1p bind to and are localized by the same component. The identification of a 150–amino acid domain necessary and sufficient for localization of Spa2p to sites of polarized growth and the existence of this domain in another yeast protein Sph1p suggest that the early localization of these proteins may be mediated by a receptor that recognizes this small domain.Polarized cell growth and division are essential cellular processes that play a crucial role in the development of eukaryotic organisms. Cell fate can be determined by cell asymmetry during cell division (Horvitz and Herskowitz, 1992; Cohen and Hyman, 1994; Rhyu and Knoblich, 1995). Consequently, the molecules involved in the generation and maintenance of cell asymmetry are important in the process of cell fate determination. Polarized growth can occur in response to external signals such as growth towards a nutrient (Rodriguez-Boulan and Nelson, 1989; Eaton and Simons, 1995) or hormone (Jackson and Hartwell, 1990a
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; Segall, 1993; Keynes and Cook, 1995) and in response to internal signals as in Caenorhabditis elegans (Goldstein et al., 1993; Kimble, 1994; Priess, 1994) and Drosophila melanogaster (St Johnston and Nusslein-Volhard, 1992; Anderson, 1995) early development.
Saccharomyces cerevisiae undergo polarized growth towards an external cue during mating and to an internal cue during budding. Polarization towards a mating partner (shmoo formation) and towards a new bud site requires a number of proteins (Chenevert, 1994; Chant, 1996; Drubin and Nelson, 1996). Many of these proteins are necessary for both processes and are localized to sites of polarized growth, identified by the insertion of new cell wall material (Tkacz and Lampen, 1972; Farkas et al., 1974; Lew and Reed, 1993) to the shmoo tip, bud tip, and mother–daughter bud neck. In yeast, proteins localized to growth sites include cytoskeletal proteins (Adams and Pringle, 1984; Kilmartin and Adams, 1984; Ford, S.K., and J.R. Pringle. 1986. Yeast. 2:S114; Drubin et al., 1988; Snyder, 1989; Snyder et al., 1991; Amatruda and Cooper, 1992; Lew and Reed, 1993; Waddle et al., 1996), neck filament components (septins) (Byers and Goetsch, 1976; Kim et al., 1991; Ford and Pringle, 1991; Haarer and Pringle, 1987; Longtine et al., 1996), motor proteins (Lillie and Brown, 1994), G-proteins (Ziman, 1993; Yamochi et al., 1994; Qadota et al., 1996), and two membrane proteins (Halme et al., 1996; Roemer et al., 1996; Qadota et al., 1996). Septins, actin, and actin-associated proteins localize early in the cell cycle, before a bud or shmoo tip is recognizable. How this group of proteins is localized to and maintained at sites of cell growth remains unclear.Spa2p is one of the first proteins involved in bud formation to localize to the incipient bud site before a bud is recognizable (Snyder, 1989; Snyder et al., 1991; Chant, 1996). Spa2p has been localized to where a new bud will form at approximately the same time as actin patches concentrate at this region (Snyder et al., 1991). An understanding of how Spa2p localizes to incipient bud sites will shed light on the very early stages of cell polarization. Later in the cell cycle, Spa2p is also found at the mother–daughter bud neck in cells undergoing cytokinesis. Spa2p, a nonessential protein, has been shown to be involved in bud site selection (Snyder, 1989; Zahner et al., 1996), shmoo formation (Gehrung and Snyder, 1990), and mating (Gehrung and Snyder, 1990; Chenevert et al., 1994; Yorihuzi and Ohsumi, 1994; Dorer et al., 1995). Genetic studies also suggest that Spa2p has a role in cytokinesis (Flescher et al., 1993), yet little is known about how this protein is localized to sites of polarized growth.We have used Spa2p green fluorescent protein (GFP)1 fusions to investigate the early localization of Spa2p to sites of polarized growth in living cells. Our results demonstrate that a small domain of ∼150 amino acids of this large 1,466-residue protein is sufficient for targeting to sites of polarized growth and is necessary for Spa2p function. Furthermore, we have identified and characterized a novel yeast protein, Sph1p, which has homology to both the Spa2p amino terminus and the Spa2p localization domain. Sph1p localizes to similar regions of polarized growth and sph1 mutants have similar phenotypes as spa2 mutants. 相似文献
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NAD-isocitrate
dehydrogenase (NAD-IDH) from the eukaryotic microalga
Chlamydomonas reinhardtii was purified to
electrophoretic homogeneity by successive chromatography steps on
Phenyl-Sepharose, Blue-Sepharose, diethylaminoethyl-Sephacel, and
Sephacryl S-300 (all Pharmacia Biotech). The 320-kD enzyme was found to
be an octamer composed of 45-kD subunits. The presence of isocitrate
plus Mn2+ protected the enzyme against thermal inactivation
or inhibition by specific reagents for arginine or lysine. NADH was a
competitive inhibitor (Ki, 0.14
mm) and NADPH was a noncompetitive inhibitor
(Ki, 0.42 mm) with respect to
NAD+. Citrate and adenine nucleotides at concentrations
less than 1 mm had no effect on the activity, but 10
mm citrate, ATP, or ADP had an inhibitory effect. In
addition, NAD-IDH was inhibited by inorganic monovalent anions, but
l-amino acids and intermediates of glycolysis and the
tricarboxylic acid cycle had no significant effect. These data support
the idea that NAD-IDH from photosynthetic organisms may be a key
regulatory enzyme within the tricarboxylic acid cycle.IDH catalyzes the oxidative decarboxylation of isocitrate to
produce 2-oxoglutarate. According to the specificity for the electron
acceptor, two enzymes with IDH activity are known, NAD-IDH (EC
1.1.1.41) and NADP-IDH (EC 1.1.1.42) (Chen and Gadal, 1990a).In photosynthetic organisms NADP-IDH has been detected in the cytosol,
chloroplasts, mitochondria, and peroxisomes. Cytosolic NADP-IDH has
been purified from higher plants (Chen et al., 1988) and eukaryotic
algae (Martínez-Rivas et al., 1996), and its cDNA has been
cloned from alfalfa (Shorrosh and Dixon, 1992), soybean (Udvardi et
al., 1993), potato (Fieuw et al., 1995), and tobacco (Gálvez et
al., 1996). This 80-kD isoenzyme is a dimer, and it is likely to be
involved in the synthesis of NADPH for biosynthetic purposes in the
cytosol (Chen et al., 1988), in the synthesis of 2-oxoglutarate for
ammonium assimilation (Chen and Gadal, 1990b), and in the cycling,
redistribution, and export of amino acids (Fieuw et al., 1995).
Chloroplastic NADP-IDH has been studied in higher plants (Gálvez
et al., 1994) and eukaryotic algae (Martínez-Rivas and Vega,
1994). It is a 154-kD dimer that has been proposed to be involved in
the supply of NADPH for biosynthetic reactions in the chloroplast when
photosynthetic NADPH production is low (Gálvez et al., 1994). The
mitochondrial NADP-IDH of higher plants may have a physiological role
in the production of NADPH, which can be converted to NADH by a
transhydrogenase or used to reduce glutathione in the mitochondrial
matrix (Rasmusson and Møller, 1990). NADP-IDH activity has also been
detected in peroxisomes from spinach leaves (Yamazaki and Tolbert,
1970).NAD-IDH is localized exclusively in the mitochondria in association
with the TCA cycle. This enzyme has been purified from several
nonphotosynthetic eukaryotes such as fungi (Keys and McAlister-Henn,
1990; Alvarez-Villafañe et al., 1996) and animals (Giorgio et
al., 1970), in which it appears to be a 300-kD octamer. Its key
regulatory role in the TCA cycle is well documented. The NAD-IDH from
yeast is activated by AMP and citrate (Hathaway and Atkinson, 1963),
whereas the animal enzyme is activated by ADP and citrate (Cohen and
Colman, 1972). In addition, the NAD-IDH cDNAs have been cloned from
yeast (Cupp and McAlister-Henn, 1991, 1992) and animals (Nichols et
al., 1995; Zeng et al., 1995). In these organisms, the enzyme is
composed of two (yeast) or more (animals) different subunits encoded by
different genes.To our knowledge, no NAD-IDH from photosynthetic organisms has yet been
purified to homogeneity, mainly because of the low stability of the
enzyme (Oliver and McIntosh, 1995). However, partial purifications have
been reported from pea (Cox and Davies, 1967; Cox, 1969; McIntosh
and Oliver, 1992), potato (Laties, 1983), spruce (Cornu et al., 1996),
and the eukaryotic microalga Chlamydomonas reinhardtii
(Martínez-Rivas and Vega, 1994). Matrix and membrane forms of
the enzyme have been detected in potato (Tezuka and Laties, 1983) and
pea (McIntosh, 1997). Although it is an allosteric enzyme that exhibits
sigmoidal kinetics with respect to isocitrate (Cox and Davies, 1967;
McIntosh and Oliver, 1992) and is activated in vitro by ABA (Tezuka et
al., 1990), the regulatory importance of NAD-IDH in photosynthetic
organisms is still under debate.To elucidate the regulatory significance of NAD-IDH in photosynthetic
organisms and its apparent contribution to the 2-oxoglutarate
supply for ammonium assimilation, we have purified and characterized
the NAD-IDH from C. reinhardtii. 相似文献
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Lidia Osuna Jean-N?el Pierre María-Cruz González Rosario Alvarez Francisco J. Cejudo Cristina Echevarría Jean Vidal 《Plant physiology》1999,119(2):511-520
Phosphoenolpyruvate
carboxylase (PEPC) activity was detected in aleurone-endosperm extracts
of barley (Hordeum vulgare) seeds during germination,
and specific anti-sorghum (Sorghum bicolor)
C4 PEPC polyclonal antibodies immunodecorated constitutive
103-kD and inducible 108-kD PEPC polypeptides in western analysis. The
103- and 108-kD polypeptides were radiolabeled in situ after imbibition
for up to 1.5 d in 32P-labeled inorganic phosphate. In
vitro phosphorylation by a Ca2+-independent PEPC protein
kinase (PK) in crude extracts enhanced the enzyme''s velocity and
decreased its sensitivity to l-malate at suboptimal pH and
[PEP]. Isolated aleurone cell protoplasts contained both
phosphorylated PEPC and a Ca2+-independent PEPC-PK that was
partially purified by affinity chromatography on blue dextran-agarose.
This PK activity was present in dry seeds, and PEPC phosphorylation in
situ during imbibition was not affected by the cytosolic
protein-synthesis inhibitor cycloheximide, by weak acids, or by various
pharmacological reagents that had proven to be effective blockers of
the light signal transduction chain and PEPC phosphorylation in
C4 mesophyll protoplasts. These collective data support the
hypothesis that this Ca2+-independent PEPC-PK was formed
during maturation of barley seeds and that its presumed underlying
signaling elements were no longer operative during germination.Higher-plant PEPC (EC 4.1.1.31) is subject to in vivo
phosphorylation of a regulatory Ser located in the N-terminal domain of
the protein. In vitro phosphorylation by a
Ca2+-independent, low-molecular-mass (30–39 kD)
PEPC-PK modulates PEPC regulation interactively by opposing metabolite
effectors (e.g. allosteric activation by Glc-6-P and feedback
inhibition by l-malate; Andreo et al., 1987), decreasing
significantly the extent of malate inhibition of the leaf enzyme
(Carter et al., 1991; Chollet et al., 1996; Vidal et al., 1996; Vidal
and Chollet, 1997). These metabolites control the rate of
phosphorylation of PEPC via an indirect target-protein effect (Wang and
Chollet, 1993; Echevarría et al., 1994; Vidal and Chollet,
1997).Several lines of evidence support the view that this protein-Ser/Thr
kinase is the physiologically relevant PEPC-PK (Li and Chollet, 1993;
Chollet et al., 1996; Vidal et al., 1996; Vidal and Chollet, 1997). The
presence and inducible nature of leaf PEPC-PK have been established
further in various C3, C4,
and CAM plant species (Chollet et al., 1996). In all cases, CHX proved
to be a potent inhibitor of this up-regulation process so that apparent
changes in the turnover rate of PEPC-PK itself or another, as yet
unknown, protein factor were invoked to account for this observation
(Carter et al., 1991; Jiao et al., 1991; Chollet et al., 1996).
Consistent with this proposal are recent findings about PEPC-PK from
leaves of C3, C4, and CAM
plants that determined activity levels of the enzyme to depend on
changes in the level of the corresponding translatable mRNA (Hartwell
et al., 1996).Using a cellular approach we previously showed in
sorghum (Sorghum bicolor) and hairy crabgrass
(Digitaria sanguinalis) that PEPC-PK is
up-regulated in C4 mesophyll cell protoplasts
following illumination in the presence of a weak base
(NH4Cl or methylamine; Pierre et al., 1992;
Giglioli-Guivarc''h et al., 1996), with a time course (1–2 h) similar
to that of the intact, illuminated sorghum (Bakrim et al., 1992) or
maize leaf (Echevarría et al., 1990). This light- and
weak-base-dependent process via a complex transduction chain is likely
to involve sequentially an increase in pHc, inositol
trisphosphate-gated Ca2+ channels of the
tonoplast, an increase in cytosolic Ca2+, a
Ca2+-dependent PK, and PEPC-PK.Considerably less is known about the up-regulation of PEPC-PK and
PEPC phosphorylation in nongreen tissues. A sorghum root PEPC-PK
purified on BDA was shown to phosphorylate in vitro both recombinant
C4 PEPC and the root
C3-like isoform, thereby decreasing the enzyme''s
malate sensitivity (Pacquit et al., 1993). PEPC from soybean root
nodules was phosphorylated in vitro and in vivo by an endogenous PK
(Schuller and Werner, 1993; Zhang et al., 1995; Zhang and Chollet,
1997). A Ca2+-independent nodule PEPC-PK
containing two active polypeptides (32–37 kD) catalyzed the
incorporation of phosphate on a Ser residue of the target enzyme and
was modulated by photosynthate transported from the shoots (Zhang and
Chollet, 1997). Regulatory seryl phosphorylation of a heterotetrameric
(α2β2) banana fruit
PEPC by a copurifying, Ca2+-independent PEPC-PK
was shown to occur in vitro (Law and Plaxton, 1997). Although
phosphorylation was also detected in vivo and found to concern
primarily the α-subunit, PEPC exists mainly in the dephosphorylated
form in preclimacteric, climacteric, and postclimacteric fruit.In a previous study we showed that PEPC undergoes regulatory
phosphorylation in aleurone-endosperm tissue during germination of
wheat seeds (Osuna et al., 1996). Here we report on PEPC and the
requisite PEPC-PK in germinating barley (Hordeum vulgare)
seeds. PEPC was highly phosphorylated by a
Ca2+-independent Ser/Thr PEPC-PK similar to that
found in other plant systems studied previously (Chollet et al., 1996);
however, the PK was already present in the dry seed and its activity
did not require protein synthesis during imbibition. 相似文献
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Adokiye Berepiki Alexander Lichius Jun-Ya Shoji Jens Tilsner Nick D. Read 《Eukaryotic cell》2010,9(4):547-557
This study demonstrates the utility of Lifeact for the investigation of actin dynamics in Neurospora crassa and also represents the first report of simultaneous live-cell imaging of the actin and microtubule cytoskeletons in filamentous fungi. Lifeact is a 17-amino-acid peptide derived from the nonessential Saccharomyces cerevisiae actin-binding protein Abp140p. Fused to green fluorescent protein (GFP) or red fluorescent protein (TagRFP), Lifeact allowed live-cell imaging of actin patches, cables, and rings in N. crassa without interfering with cellular functions. Actin cables and patches localized to sites of active growth during the establishment and maintenance of cell polarity in germ tubes and conidial anastomosis tubes (CATs). Recurrent phases of formation and retrograde movement of complex arrays of actin cables were observed at growing tips of germ tubes and CATs. Two populations of actin patches exhibiting slow and fast movement were distinguished, and rapid (1.2 μm/s) saltatory transport of patches along cables was observed. Actin cables accumulated and subsequently condensed into actin rings associated with septum formation. F-actin organization was markedly different in the tip regions of mature hyphae and in germ tubes. Only mature hyphae displayed a subapical collar of actin patches and a concentration of F-actin within the core of the Spitzenkörper. Coexpression of Lifeact-TagRFP and β-tubulin–GFP revealed distinct but interrelated localization patterns of F-actin and microtubules during the initiation and maintenance of tip growth.Actins are highly conserved proteins found in all eukaryotes and have an enormous variety of cellular roles. The monomeric form (globular actin, or G-actin) can self-assemble, with the aid of numerous actin-binding proteins (ABPs), into microfilaments (filamentous actin, or F-actin), which, together with microtubules, form the two major components of the fungal cytoskeleton. Numerous pharmacological and genetic studies of fungi have demonstrated crucial roles for F-actin in cell polarity, exocytosis, endocytosis, cytokinesis, and organelle movement (6, 7, 20, 34, 35, 51, 52, 59). Phalloidin staining, immunofluorescent labeling, and fluorescent-protein (FP)-based live-cell imaging have revealed three distinct subpopulations of F-actin-containing structures in fungi: patches, cables, and rings (1, 14, 28, 34, 60, 63, 64). Actin patches are associated with the plasma membrane and represent an accumulation of F-actin around endocytic vesicles (3, 26, 57). Actin cables are bundles of actin filaments stabilized with cross-linking proteins, such as tropomyosins and fimbrin, and are assembled by formins at sites of active growth, where they form tracks for myosin V-dependent polarized secretion and organelle transport (10, 16, 17, 27, 38, 47, 48). Cables, unlike patches, are absolutely required for polarized growth in the budding yeast Saccharomyces cerevisiae (34, 38). Contractile actomyosin rings are essential for cytokinesis in budding yeast, whereas in filamentous fungi, actin rings are less well studied but are known to be involved in septum formation (20, 28, 34, 39, 40).Actin cables and patches have been particularly well studied in budding yeast. However, there are likely to be important differences between F-actin architecture and dynamics in budding yeast and those in filamentous fungi, as budding yeasts display only a short period of polarized growth during bud formation, which is followed by isotropic growth over the bud surface (10). Sustained polarized growth during hyphal morphogenesis is a defining feature of filamentous fungi (21), making them attractive models for studying the roles of the actin cytoskeleton in cell polarization, tip growth, and organelle transport.In Neurospora crassa and other filamentous fungi, disruption of the actin cytoskeleton leads to rapid tip swelling, which indicates perturbation of polarized tip growth, demonstrating a critical role for F-actin in targeted secretion to particular sites on the plasma membrane (7, 22, 29, 56). Immunofluorescence studies of N. crassa have shown that F-actin localizes to hyphal tips as “clouds” and “plaques” (7, 54, 59). However, immunolabeling has failed to reveal actin cables in N. crassa and offers limited insights into F-actin dynamics. Live-cell imaging of F-actin architecture and dynamics has not been accomplished in N. crassa, yet it is expected to yield key insights into cell polarization, tip growth, and intracellular transport.We took advantage of a recently developed live-cell imaging probe for F-actin called Lifeact (43). Lifeact is a 17-amino-acid peptide derived from the N terminus of the budding yeast actin-binding protein Abp140 (5, 63) and has recently been demonstrated to be a universal live-cell imaging marker for F-actin in eukaryotes (43). Here, we report the successful application of fluorescent Lifeact fusion constructs for live-cell imaging of F-actin in N. crassa. We constructed two synthetic genes consisting of Lifeact fused to “synthetic” green fluorescent protein (sGFP) (S65T) (henceforth termed GFP) (12) or red fluorescent protein (TagRFP) (33) and expressed these constructs in various N. crassa strains. In all strain backgrounds, fluorescent Lifeact constructs clearly labeled actin patches, cables, and rings and revealed a direct association of F-actin structures with sites of cell polarization and active tip growth. Our results demonstrate the efficacy of Lifeact as a nontoxic live-cell imaging probe in N. crassa. 相似文献