Plant Physiology

doi: N/Apmid: N/A

Yu, Su-May

doi: 10.1104/pp.121.3.687pmid: 10557216

Photosynthesis converts solar energy to chemical energy, which then drives the synthesis of sugars from carbon dioxide and water. Sugars play multiple roles in all aspects of plant life. First, they provide the main respiratory substrates for the generation of energy and metabolic intermediates that are then used for the synthesis of macromolecules and other cell constituents. Second, Rib and deoxy-Rib sugars form part of the structure of DNA and RNA. Third, polysaccharides are the major structural elements of plant cell walls. Fourth, linkage to sugar is required for proper functioning of many proteins and lipids. As a consequence, the abundance and depletion of sugars or their derivatives initiate various responses in plants and have profound effects on plant metabolism, growth, and development. Plants are considered to be carbon autotrophs, but they can be considered as carbon heterotrophs during some part of their life cycle and in some of their non-green organs such roots, stems, and flowers, which are not involved in photosynthesis. Furthermore, carbohydrate depletion can occur and is a fact of life in most plants. For instance, variations in environmental factors such as light, water, or temperature and attacks by pathogens or herbivores may lead to a significant decrease in the efficiency of photosynthesis in source tissues (such as leaves that synthesize and export carbohydrates) and thus reduce the supply of carbohydrates to sink tissues (such as the non-green tissues that import carbohydrates for respiration, growth, and development). Under certain growth conditions, such as during an annual resting season or after leaf shedding, photosynthesis is turned off or operates to a lower degree, and carbohydrate reserves must be utilized and may become limited in nonphotosynthetic tissues. In germinating seeds under unfavorable environmental conditions the mobilization of stores in the cotyledons is delayed, which may result in the depletion of available carbohydrates and a decrease in seedling vigor. Knowledge about the response to sugar starvation and adaptation mechanisms in plants is of both fundamental and agronomic importance. In nature, the cessation of growth of a heterotrophic living organism is often brought about by a poor nutrient environment, a commonly encountered stress. Environmental changes affect various biochemical reactions, often disturbing the balanced distribution of metabolites within cells. In most instances, living cells show a rapid molecular response to overcome adverse environmental conditions. How a living organism survives during periods of environmental stress is an exciting area of research. The most extensive studies have been done with microorganisms. The ability of microorganisms to sense and respond to unscheduled changes in their environment is crucial to their survival. When cells of microorganisms encounter unfavorable nutrient conditions, they ultimately enter a stationary phase. Cells in the stationary phase are physiologically, biochemically, and morphologically different from cells growing exponentially. Studies using Escherichia coliand yeast (Saccharomyces cerevisiae) have indicated that entry into the stationary phase is a complex, highly regulated process that activates a program for long-term survival. The program includes the lack of a requirement for added nutrients and an absence of cell division. The similarities of eukaryotic and prokaryotic microorganisms in their responses to nutrient limitations suggest that such responses are based on evolutionarily conserved genetic mechanisms. Similarly, sugar starvation initiates changes in substantial physiological and biochemical processes with the goal of sustaining respiration and other essential metabolic processes in plants. Sugar starvation also initiates changes in cellular processes to recycle cellular constituents and dramatically changes the pattern of gene expression. However, the underlying mechanisms used by plant cells to cope with sugar starvation are largely unknown, and only recently have these questions been addressed experimentally. This lack of knowledge contrasts with the situation in bacteria and yeast, where the molecular biology and physiology of mutants have yielded extensive information about responses to sugar starvation. This review discusses the recent advances made in our understanding of the molecular events that operate in microorganisms upon sugar starvation, as well as the cellular and genetic responses of plants to sugar starvation. SUGAR STARVATION IN BACTERIA The majority of bacteria spend most of their time in a nutrient-limited starvation phase and as a result have evolved mechanisms that allow them to survive under these conditions and to resume growth once nutrients become available. Some bacteria, e.g.Bacillus spp., undergo major differentiation programs that lead to the formation of highly stress-resistant endospores or cysts. Other bacteria, e.g. E. coli, even without the formation of differentiated cells, enter starvation-induced programs that allow them to survive long periods of non-growth and to restart growth when nutrients become available. These starvation-induced programs often lead to the formation of metabolically less-active cells that are more resistant to a wide range of environmental stresses. This adaptation to starvation conditions is often accompanied by a change in cell size and the induction of genes and stabilization of proteins essential for long-term survival. Evidence suggests that there is a general starvation response among various bacteria species. For example, Glc- or nitrogen-starved cultures of E. coli exhibit resistance to heat or hydrogen peroxide (Jenkins et al., 1988). The nitrogen-fixing bacterium Rhizobium leguminosarum can survive carbon, nitrogen, and phosphorus starvation for at least 2 months with little loss of viability. Upon carbon starvation, R. leguminosarum cells undergo reductive cell division and the levels of protein, DNA, and RNA synthesis decrease to base levels, mRNA stabilizes, and the starved cells are cross-protected against pH, heat, osmotic, and oxidative shock (Thorne and Williams, 1997). In E. coli, nutrient-starved stationary-phase cells have been used as a model system for studying the molecular mechanism that regulates gene expression under nutrient starvation. Stationary-phase cells have a small spherical shape, are resistant to multiple stresses, synthesize glycogen, and survive long-term starvation. Genes expressed during adaptation to starvation conditions involve several classes of starvation genes that code for special stress-resistant proteins. The two major classes of genes induced upon carbon starvation arecst genes, which require cAMP and enhance the cell's metabolic potential, and pex genes, which do not require cAMP but play a more direct protective role against stresses (Matin, 1991). A protective role of stress-resistant proteins is proposed due to their ability to rescue misfolded macromolecules (Matin, 1991). Expression of the stress-resistant proteins depends on an intactrpoS allele (Hengge-Aronis, 1993). However, a common consensus sequence has not been found among various promoters controlled by rpoS, and thus a regulatory cascade that mediates expression of the rpoS-dependent genes has been suggested (Hengge-Aronis, 1993). The protein encoded by rpoSis an alternative ς factor of RNA polymerase and is designated as ςs. Evidence shows that the ςs factors are regulated primarily at the post-transcriptional level by a mechanism that involves a mRNA secondary structure (McCann et al., 1993). In addition, carbon starvation in E. coli might be sensed through the accumulation of homoserine lactone (Huisman and Kolter, 1994). SUGAR STARVATION IN YEAST Stress conditions imposed on yeast can be as diverse as nutrient starvation, suboptimal temperatures or osmolarity, high ethanol concentrations, the presence of heavy metals or oxidation compounds, and desiccation (Ruis and Schüller, 1995). Similarity in response to these stresses has been observed and the previous exposure to one stress generally increases the acquisition of tolerance against challenge by another stress (cross-protection or cross-resistance) (Lewis et al., 1995; Ruis and Schüller, 1995). These observations indicate that cells possess one central molecular mechanism that can be activated by various factors and, upon activation, will protect cells against a number of conditions threatening their survival. Some carbohydrates or proteins induced by various stresses have been suggested to play a protective role against stresses. For example, a close correlation was observed between the content of trehalose, one of the major reserve carbohydrates in yeast, and the stress resistance of the cells. The levels of trehalose and stress resistance increase rapidly upon exhaustion of Glc in the culture medium (Panek and Panek, 1990). The level of trehalose also increases strongly upon starvation of an essential nutrient such as nitrogen, phosphate, or sulfate in a Glc-containing medium (Attfiel et al., 1992). The same is true during sublethal heat, freeze-thaw, and desiccation treatment (Hottiger et al., 1987; Attfiel et al., 1992). Genes that have been demonstrated to contribute significantly to the ability of yeast cells to survive severe stress include CTT1 (encoding the cytosolic catalase T) and HSP104 and HSP70(encoding heat shock proteins) (Ruis and Schüller, 1995). How yeast cells respond to a wide range of stresses through a convergent molecular mechanism(s) remains largely unclear. Specific gene control elements and stress-activated transcription factors binding to them are probably shared by the stress-responsive genes. A common feature at the transcriptional level, the stress-response element (STRE), a cis-acting element with the core consensus CCCCT, has been found to be present in the promoters of genes induced by various stresses (Varela et al., 1995). STRE activity correlates well with the potential to establish stress tolerance (Ruis and Schüller, 1995). Msn2p, a transcription factor that activates STRE-regulated genes in response to stress, has been identified. Mutants defective in Msn2p exhibit pleiotrophic hypersensitivity to stress factors (Schmitt and McEntee, 1996). How stress signals are transmitted to STREs is not clear, and this raises the question of whether the various stress factors create a common pathway or multiple pathways that then transmit signals to the stress-specific STREs. STRE activities have been shown to be controlled by the high osmolarity glycerol pathway and the protein kinase A pathway (signaling nutrient stress), suggesting that different signals are transmitted through different pathways (Ruis and Schüller, 1995). Dramatic morphological changes can be observed in yeast undergoing nutrient starvation. The depletion of nutrients such as carbon, nitrogen, sulfur, or amino acids induces autophagy in yeast (Takeshige et al., 1992). Autophagy is the major route of delivery of cytoplasmic proteins into vacuoles/lysosomes under conditions in which cells require enhanced protein degradation and remodeling of components (Dunn, 1994). A Ser/Thr protein kinase gene, APG1, is essential for both the autophagic process and the maintenance of viability of yeast under starvation conditions (Matsuura et al., 1997). It is therefore hypothesized that autophagy-dependent reconstruction of cellular constituents is required for long-term viability in starvation conditions and that the process involves regulation by protein phosphorylation (Matsuura et al., 1997). PLANT CELL METABOLISM ALTERED BY SUGAR STARVATION Over the past 20 years, carbohydrate starvation has been studied in a number of plant species. Physiological and cellular changes that occur during a plant's transition to sugar starvation are most extensively studied in excised maize root tips (Brouquisse et al., 1991; Dieuaide et al., 1992), cultured sycamore cells (Journet et al., 1986; Aubert et al., 1996), and cultured rice suspension cells (Chen et al., 1994). These studies have shown that sugar starvation generally triggers sequential changes in the following cellular events: (a) arrest of cell growth, (b) rapid consumption of cellular carbohydrate content and decrease in respiration rate, (c) degradation of lipids and proteins, (d) increase in accumulation of Pi, phosphorylcholine, and free amino acids, and (e) decline in glycolytic enzymatic activities. It appears that changes in metabolism are involved in the adaptation response of plant cells to sugar starvation. For example, cells in roots (Brouquisse et al., 1991) and leaves (Peeters and Van Laere, 1992), cultured suspension cells (Journet et al., 1986; Chen et al., 1994), and callus cells (Tassi et al., 1992) modify their metabolism to survive in the absence of sugar. In sugar-starved cultured cells, there is a decrease in enzymatic activities related to sugar metabolism and respiration (Journet et al., 1986; Brouquisse et al., 1991), nitrate reduction and assimilation (Brouquisse et al., 1992), and protein synthesis (Tassi et al., 1992). Decreases in these enzymatic activities presumably protect cells against nutrient stress by switching off biosynthesis (i.e. growth) to conserve energy. At the same time, an increase in enzymatic activities related to catabolism of fatty acids (Dieuaide et al., 1992), amino acids (Brouquisse et al., 1992), and proteins (Tassi et al., 1992) occurs. Such a change can substitute protein and lipid catabolism for sugar catabolism to sustain respiration and metabolic processes (Journet et al., 1986; Brouquisse et al., 1991). Although these metabolic changes appear to enhance the survival of cultured cells under Glc starvation, they finally result in irreversible damages and cell death (Brouquisse et al., 1991; Chen et al., 1994). Similar metabolic changes occur in plant organs or tissues during senescence or in postharvest situations (Noodén, 1988;King et al., 1990). A common mechanism that regulates metabolic processes during sugar starvation and senescence has been suggested (Noodén, 1988). Sugar starvation has also been described as a component of senescence (Dieuaide et al., 1992). VACUOLAR AUTOPHAGY IN PLANT CELLS In Suc-starved sycamore and rice suspension cells, the decline in cellular sugar and starch contents couples with the decline in metabolic activity and the increase in vacuolar autophagic activity (Journet et al., 1986; Chen et al., 1994). Triggering of such autophagic processes presumably involves the regression of cytoplasm, including the organelles, and the recycling of respiratory substrates (Journet et al., 1986; Chen et al., 1994; Aubert et al., 1996). This process is well documented in animal cells (Marzella and Glaumann, 1987) and has been implicated in the nonselective bulk degradation of proteins triggered by nutrient deprivation. Autophagy in plant, animal, and yeast systems is often associated with nutrient starvation. In Suc-provided rice suspension cells, the size of the vacuole is small (Fig. 1a). Vacuolar autophagic activity begins a few hours after Suc starvation, and vacuole size expands either by engulfing neighboring cytoplasm and organelles (except the nucleus) or by vacuoles fusing together (Fig. 1b). After a long period of Suc starvation, the vacuole volume becomes extremely large and the cytoplasm and the leftover organelles (mostly mitochondria) are confined to a narrow area adjacent to cell walls (Fig. 1c). Plant vacuoles are rich in hydrolases, and cytoplasm sequestered by the autophagic vacuoles is eventually degraded by these enzymes. Vacuolar autophagy has also been observed in plants undergoing senescence (Matile and Winkenbach, 1971). Due to the presence of intracellular pools of carbohydrates and the ability to control the autophagic process, plant cells can survive for some time after carbohydrate starvation. Fig. 1. Open in new tabDownload slide Electron micrographs showing morphological changes of cultured rice suspension cells during Suc starvation. Cells were Suc-starved for 0 d (a), 1 d (b), and 2 d (c), and examined under an electron microscope. AMY, Amyloplast; CW, cell wall; M, mitochondria; N, nucleus; S, starch granule; V, vacuole. Bar = 4 μm. Fig. 1. Open in new tabDownload slide Electron micrographs showing morphological changes of cultured rice suspension cells during Suc starvation. Cells were Suc-starved for 0 d (a), 1 d (b), and 2 d (c), and examined under an electron microscope. AMY, Amyloplast; CW, cell wall; M, mitochondria; N, nucleus; S, starch granule; V, vacuole. Bar = 4 μm. PLANT CELL RESPONSE TO SUGAR STARVATION AT THE GENE EXPRESSION LEVEL Sugar plays an important dual role in regulating the expression of various genes in plants. In general, sugar favors the expression of enzymes in connection with biosynthesis, utilization, and storage of reserves (including starch, lipid, and proteins). On the other hand, sugar represses the expression of enzymes involved in photosynthesis and reserve mobilization (Koch, 1996). The events of cellular responses to sugar starvation is shown in Figure 2. Generally, gene expression repressed by sugar is up-regulated by sugar starvation, whereas that enhanced by sugar is down-regulated. The alteration of gene expression by sugar starvation results in the induction of synthesis of preexisting or new proteins and repression of normally expressed proteins. Fig. 2. Open in new tabDownload slide Events in cellular responses to sugar starvation in plants. Fig. 2. Open in new tabDownload slide Events in cellular responses to sugar starvation in plants. A large and specific set of genes whose expression is induced by sugar starvation has been reported (Koch, 1996). For example, sugar starvation induces the expression of photosynthetic genes in maize mesophyll protoplasts (Sheen, 1990), α-amylase genes in rice suspension cells and germinating embryos (Yu et al., 1991, 1996), Suc synthase (Sh1) gene in maize root tips (Koch et al., 1992), and malate synthase and isocitrate lyse genes in cucumber (Graham et al., 1994). At the beginning of rice seed germination, active metabolism and a rise in the respiration rate cause rapid sugar depletion in the embryo, which then triggers the expression of α-amylase genes and degradation of starch in this tissue (Yu et al., 1996). Sugar depletion is also proposed to be a primary factor in initiating the synthesis of phytohormone GA in the embryo, since sugar reduces the quantity of GA in this tissue (Yu et al., 1996). Most studies on the mechanisms of sugar repression of gene expression in microorganisms and plants have emphasized regulation at the transcriptional level. In plants, while sugar repression of genes involved in photosynthesis (Sheen, 1990) and the glyoxylate cycle (Graham et al., 1994) operates at the transcriptional level, sugar repression of α-amylase gene expression involves control of transcription and mRNA stability (Sheu et al., 1996; Chan et al., 1994,1998; Lu et al., 1998). Search for cis-regulatory elements in the promoters of sugar-regulated genes is important in understanding the mechanism of sugar regulation of gene expression. Although carbohydrate depletion induces expression of a large set of genes essential for various physiological processes, the cis-acting sugar response elements in the promoters of these genes have not been extensively studied. A sugar response complex in the promoter region of a Suc-deprivation-induced rice α-amylase gene, αAmy3, has been identified. This complex contains three essential motifs for a high level of sugar-starvation-induced gene expression in rice cells (Lu et al., 1998). One of the motifs, a TATCCA element, along with its variants, are present at a proximity upstream of the transcription start sites of 18 α-amylase genes isolated from various plant species (Yu, 1999) and several other sugar-repressible genes. The TATCCA element is present in tandem repeat between position −116 to −105 of the transcription start site of αAmy3 (Lu et al., 1998). Nuclear proteins from rice suspension cells that bind to the TATCCA element in a sequence-specific and sugar-dependent manner have also been identified (Lu et al., 1998). A 20-bp sequence upstream of the transcription start site of the maize Suc synthase geneShrunken is sufficient to confer sugar inhibition of downstream reporter gene expression (Maas et al., 1990). There is no homology between the sugar response sequences of the αAmy3and the 20-bp sequence of the Shrunken promoters. However, the TATCCA element is present between position −136 and position −141 of the Shrunken promoter, which could be another control element that exhibits a function similar to the 20-bp sequence (Maas et al., 1990). SUGAR SENSING AND SIGNAL TRANSDUCTION IN PLANT CELLS Information concerning the sugar status of plant cells is of great importance in initiating changes in gene expression and subsequent metabolic and developmental responses. The mechanisms used by plant cells to sense sugars and to process this information are largely unknown. Yeast has been an essential model for studies on the mechanisms of sugar sensing and signal transduction employed in plant cells. In yeast, genes required for growth on carbon sources other than Glc are repressed by the presence of Glc in the medium and can be derepressed when Glc is removed. This is the phenomenon of Glc repression that requires a mechanism for sensing the availability of Glc. Hexokinase, the enzyme that catalyzes the phosphorylation of hexose sugars at the first step of the glycolytic pathway, has been implicated as a Glc sensor in organisms as diverse as yeast (Rose et al., 1991) and mammals (Efrat et al., 1994). Recent studies suggest that hexokinase also acts as a primary sugar sensor in plants (Jang and Sheen, 1997; Smeekens and Rook, 1997). However, the notion that hexokinase is a primary sugar sensor was recently challenged, and multiple sugar-sensing pathways were proposed to exist in plants (Halford et al., 1999). The other sugar-sensing systems proposed to exist in plants are a hexose transporter and/or receptor signaling pathway and a Suc transporter and/or receptor signaling pathway (Smeekens and Rook, 1997; Halford et al., 1999). Knowledge of the downstream components of the Glc-signaling pathway in plants has just begun to emerge. In fungi, the SNF1 protein (Suc non-fermenting 1) is required for derepression of nearly all Glc-repressed genes and is an integral component of the sugar signal transduction pathway (Ronne, 1995). SNF1 is a Ser/Thr protein kinase and the active kinase is a high-molecular-mass complex. The SNF1 complex contains three proteins that are homologs of three subunits of the mammalian AMPK (AMP-activated protein kinase) (Hardie et al., 1998). AMPK is one component of a kinase cascade that is activated in a highly sensitive manner by the elevation of AMP and the depletion of ATP. The AMPK cascade has been shown to be activated by environmental stresses that deplete cellular ATP, for example, in pancreatic β cells by Glc deprivation (Salt et al., 1998). It is therefore suggested that the SNF1 complex in yeast might be activated in a manner similar to AMPK in mammals in response to Glc deprivation, and a change in the ATP level might be the signal that indicates the availability of sugar (Halford et al., 1999). Recently, the requirement for a SNF1-related protein kinase-1 (SnRK1) in Suc-activated expression of a Suc synthase gene was demonstrated in potato by an antisense RNA approach (Purcell et al., 1998). This study indicated that SNF1 in plants may play a role analogous to that of SNF1 in yeast (Halford et al., 1999). However, whether SnRK1 activity is regulated by Glc or another hexose and whether plant SNF1 homologs also play a role in the derepression of sugar-repressible genes remains to be determined. Identification of other functional components in the sugar signal transduction pathway are also important for determining whether the mechanisms through which cells sense sugar availability and respond by changing gene expression are conserved or diverged between yeast and plants throughout evolution. Based on the available information, a model of sugar sensing, signal transduction, and mechanisms of gene regulation in plant cells is shown in Figure3. Fig. 3. Open in new tabDownload slide Hypothetical model of genetic and cellular responses of plant cells to sugar, including a sugar signal transduction pathway and a mechanism of gene regulation. Three elements function as sugar sensors: a hexokinase, a sugar transporter, and a change in the AMP/ATP ratio. Protein phosphatase and protein kinase are involved in the signal transduction pathway. In some cases, the SNF1 complex may be a component of the signal transduction pathway. The expression level of sugar-regulated genes is determined by the control of promoter transcriptional activity and mRNA stability. In the presence of sugar, the expression of sugar-starvation-induced genes is suppressed and there is no change in metabolism or vacuolar autophagic activity. Under sugar starvation, an opposite action of these events likely occurs. Fig. 3. Open in new tabDownload slide Hypothetical model of genetic and cellular responses of plant cells to sugar, including a sugar signal transduction pathway and a mechanism of gene regulation. Three elements function as sugar sensors: a hexokinase, a sugar transporter, and a change in the AMP/ATP ratio. Protein phosphatase and protein kinase are involved in the signal transduction pathway. In some cases, the SNF1 complex may be a component of the signal transduction pathway. The expression level of sugar-regulated genes is determined by the control of promoter transcriptional activity and mRNA stability. In the presence of sugar, the expression of sugar-starvation-induced genes is suppressed and there is no change in metabolism or vacuolar autophagic activity. Under sugar starvation, an opposite action of these events likely occurs. CONCLUSIONS Bacteria and yeast have developed mechanisms to react to depletion of nutrients in their environment and protect themselves against damage caused by nutrient stress and other stresses. Some components of stress signal pathways have been shown to be conserved among yeast, mammal, and plant cells (Ruis and Schüller, 1995; Hardie et al., 1998). Studies on the mechanisms of signal transduction and gene regulation in response to sugar deprivation will determine which strategies nature uses to deal with problems encountered by cells living in an unfavorable environment. However, many questions with respect to the underlying molecular mechanisms employed by plants in the adaptation to sugar deprivation remain to be answered. An understanding of how plants respond to sugar starvation and regulate the mobilization of stored carbohydrates can also help us to design crops with higher stress-tolerant capacity and is thus of biotechnological importance. ACKNOWLEDGMENTS I thank Dr. Maarten J. Chrispeels for critical review of the manuscript, and Lin-Tze Yu and Douglas Platt for help in preparing the manuscript. 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J Biol Chem 266 1991 21131 21137 Google Scholar Crossref Search ADS PubMed WorldCat 49 Yu S-M Lee Y-C Fang S-C Chan M-T Hwa S-F Liu L-F Sugars act as signal molecules and osmotica to regulate the expression of α-amylase genes and metabolic activities in germinating cereal grains. Plant Mol Biol 30 1996 1277 1289 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 Research in the author's laboratory is supported by grants from Academia Sinica and the National Science Council of the Republic of China. * E-mail [email protected]; fax 886–2–2788–2695 or 886–2–2782–6085. Copyright © 1999 American Society of Plant Physiologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Miernyk, Jan A.

doi: 10.1104/pp.121.3.695pmid: 10557217

Whole-genome sequencing projects have drastically changed the landscape of biological research. The lexicon of contemporary biology contains a plethora of new terms, including: genomics, research pertaining to the genome; proteomics, description of the protein complement of an organism; and bioinformatics, the collection and interpretation of biological information (primarily nucleic acid and amino acid sequence data) (Bouchez and Höfte, 1998). However, obtaining sequence information is not an end unto itself. It is essential that the products of these genes be identified and their function and physiological significance discovered (Bork et al., 1998;Saier, 1998). In complex eukaryotes such as flowering plants, as many as 5 × 104 genes can be selectively expressed in individual cells. It is the products of these genes, the proteins, that determine the fate and function of the cells. Protein function is determined by how the protein folds to form a specific three-dimensional structure. The way that a protein folds is determined by the free energy of the constituent amino acid residues (Levitt et al., 1997). As much as 50% of the primary amino acid sequence is necessary just to define the three-dimensional structure of a typical protein (Dobson et al., 1998). PROTEIN FOLDING IN THE CELL IS NOT AUTONOMOUS The classic in vitro studies of Anfinsen, which resulted in his receipt of the 1972 Nobel prize in chemistry, demonstrated that the primary amino acid sequence of a protein can contain all of the information necessary to direct the folding of a polypeptide chain to the correct final structure (for review, see Anfinsen, 1973). However, the conditions of temperature and pH, the salt concentration, and especially the total protein concentration found in vivo tend to promote a plethora of side reactions that compete with the single pathway that will lead to the correct final structure. Unfolded and partially folded proteins tend to aggregate when present at the concentrations found in vivo, which are estimated to be as high as 340 mg mL −1 in Escherichia coli. Molecular chaperones, proteins that prevent inappropriate association or aggregation of exposed hydrophobic surfaces of unfolded or partially folded proteins and direct them into productive folding, transport, or degradation pathways, function to minimize protein aggregation and can promote dissociation of aggregates that have formed (Boston et al., 1996; Miernyk, 1997; Netzer and Hartl, 1998; Sigler et al., 1998). Protein-folding catalysts, conventional enzymes that accelerate the rate-limiting steps in protein folding, allowing folding intermediates to avoid aggregation and non-productive interactions with other proteins, also assist cellular proteins to avoid aggregation by accelerating the rate of correct folding (Schmid, 1993; Boston et al., 1996; Huppa and Ploegh, 1998). The molten globule state is an intermediate stage where a protein is “partly unfolded,” and it is thought that proteins are in the molten globule state when recognized by chaperones or for membrane translocation. MOLECULAR CHAPERONES PREVENT NON-PRODUCTIVE INTERACTIONS Molecular chaperones assist in the assembly/disassembly of proteins but are not themselves components of the final structures. Molecular chaperones do not have an active role in protein folding, do not accelerate the folding reactions, nor do they provide steric information directing protein folding. Rather, they serve to reduce the divergence of folding intermediates into non-productive side reactions. Molecular chaperones reversibly bind to and shield unfolded segments of polypeptides that would otherwise act as loci for aggregation. For in vitro analysis, it is assumed the protein folding and unfolding are equivalent. Molecular chaperones are often characterized by their ability to prevent aggregation of proteins unfolded by mild acid or heat treatment (Fig. 1). Under stress conditions such as heat shock, it is the synthesis of molecular chaperones that allows cellular proteins to avoid and/or recover from aggregation. A brief description of each of the major classes of stress proteins/molecular chaperones from the large to small subunit size is presented. Fig. 1. Open in new tabDownload slide The Stress70 chaperone machine prevents thermal aggregation of the model protein malate dehydrogenase. Malate dehydrogenase (300 nm) was incubated at 45°C with no chaperone (○), with recombinant Arabidopsis Stress70 (▿), Stress70 plus the chaperone activating protein AtJ2 (▵), or Stress70 plus AtJ2 plus the nucleotide exchange factor AtE1 (⋄). Light scattering, measured at 320 nm, increased as the enzyme was unfolded during heat treatment. Data are the means ± se of three measurements. Fig. 1. Open in new tabDownload slide The Stress70 chaperone machine prevents thermal aggregation of the model protein malate dehydrogenase. Malate dehydrogenase (300 nm) was incubated at 45°C with no chaperone (○), with recombinant Arabidopsis Stress70 (▿), Stress70 plus the chaperone activating protein AtJ2 (▵), or Stress70 plus AtJ2 plus the nucleotide exchange factor AtE1 (⋄). Light scattering, measured at 320 nm, increased as the enzyme was unfolded during heat treatment. Data are the means ± se of three measurements. THERE ARE SIX MAJOR FAMILIES OF CHAPERONES, EACH WITH UNIQUE HOMOLOGS IN DIFFERENT SUBCELLULAR COMPARTMENTS Stress100/Clp The 100-kD stress protein is found in all organisms, with the actual size ranging from 84 to 104 kD. There are two major subclasses: class 1 proteins (A, B, C, and D) have two ATP-binding sites, and class 2 proteins (M, N, X, and Y) have a single ATP-binding site (Schirmer et al., 1996). Class 1 Stress100/Clp proteins have a coiled-coil secondary structure separating the ATP-binding domains (Nieto-Sotelo et al., 1999). Transmission electron microscopy of Stress100 reveals ring-shaped particles with a 6-fold rotational symmetry. Side views show two rings stacked together (a dodecamer). The HSP100 proteins have been extensively studied in E. colias subunits of an ATP-dependent protease (caseino-lytic protease [Clp]) (Hoskins et al., 1998). Clp consists of two distinct subunits: ClpP is the actual protease, while the ClpA/B/C/X chaperone subunits designate target specificity. The ClpP sequence is unrelated to ClpA/B/C/X, and ClpP is not a chaperone. Either as subunits of the homomeric ring structure or as subunits of the protease, HSP100/Clp employs ATP hydrolysis to promote changes in protein folding and assembly. Thus, the HSP100/Clp proteins constitute a class of molecular chaperones (Boston et al., 1996; Hoskins et al., 1998). In plant cells HSP100/Clp is both cytoplasmic and organellar (within plastids and/or mitochondria). HSP100/Clp is expressed in developmental and organ-specific patterns and is up-regulated by a variety of environmental stress conditions (heat, cold, high salt, and heavy metals) (Schirmer et al., 1994). Stress90 The 90-kD stress proteins, which actually range in size from 82 to 96 kD, are highly conserved and abundant in the cytoplasm of both eukaryotic and prokaryotic cells (Csermely et al., 1998). Despite the high levels of expression under non-stress conditions, the term HSP90 is widely employed. Although considered controversial for many years, the question of ATP binding to Stress90 was answered unequivocally when the x-ray structure was solved (Fig. 2A). Thus, HSP90 is an ATP-dependent molecular chaperone that binds transiently to late, probably highly structured folding intermediates, preventing aggregation. While the majority of cellular HSP90 is in the cytoplasm, there are distinct organellar forms found in the rough endoplasmic reticulum (ER), plastids, and mitochondria (Boston et al., 1996; Møgelsvang and Simpson, 1998). Fig. 2. Open in new tabDownload slide Structures of selected molecular chaperones. A, Ribbon diagram derived from the 1.8-Å x-ray structure of the tetragonal form of the N-terminal domain of Saccharomyces cerevisiae Stress90 complexed with the specific inhibitor geldanamycin. This inhibitor binds to the ATP-binding site. B, Ribbon diagram derived from the 1.7-Å x-ray structure of the 44-kD N-terminal ATP-binding domain of bovine Stress70. C, Ribbon diagram derived from the NMR structure of the J-domain of the human DnaJ homolog HdJ1. D through F, Structures of E. coli GroE reconstructed from cryo-electron microscopy. D, GroEL; E, GroEL plus 30 mm ATP; F, GroEL-ATP-GroES (http://bioc09.uthscsa.edu/ approximately seale/chap/em1.html). Fig. 2. Open in new tabDownload slide Structures of selected molecular chaperones. A, Ribbon diagram derived from the 1.8-Å x-ray structure of the tetragonal form of the N-terminal domain of Saccharomyces cerevisiae Stress90 complexed with the specific inhibitor geldanamycin. This inhibitor binds to the ATP-binding site. B, Ribbon diagram derived from the 1.7-Å x-ray structure of the 44-kD N-terminal ATP-binding domain of bovine Stress70. C, Ribbon diagram derived from the NMR structure of the J-domain of the human DnaJ homolog HdJ1. D through F, Structures of E. coli GroE reconstructed from cryo-electron microscopy. D, GroEL; E, GroEL plus 30 mm ATP; F, GroEL-ATP-GroES (http://bioc09.uthscsa.edu/ approximately seale/chap/em1.html). HSP90 can function independently as a chaperone; however, it also acts in concert with a group of other proteins that together comprise the foldosome, or cytoplasmic chaperone heterocomplex (CCH). The CCH has been studied most extensively in mammalian cells, where it plays an important role in signal transduction via interaction with steroid hormone receptors and protein kinases including the Src and Raf components of the MAP kinase system (Buchner, 1999). HSP90 is associated with at least six partner proteins complexed with the hormone-free receptor. Formation of this complex is essential for subsequent hormone binding. After binding of the ligand and dissociation of the CCH, the activated hormone-bound receptor functions as a transcription factor. In the absence of ligands, the receptors interact with HSP70 to start a new cycle. Two important factors have led to the characterization of CCH function: (a) the CCH can be assembled in vitro from isolated components, and (b) this assembly can be prevented by geldanamycin, a specific inhibitor of Stress90 function (Buchner, 1999). HSP90 partner proteins in the CCH include: Stress70, Hip (HSP70 interacting protein; p48), Hop (HSP70/HSP90 organizing protein; p60 or Sti1p), a DnaJ homolog, the folding catalyst prolyl-isomerase (PPI), and p23 (Sba1p in yeast). An additional component, p50 (Cdc37), has only been detected in complexes of the CCH with protein kinases. Although the CCH has been best characterized in mammalian systems, a complex that is very similar in both composition and function can also be found in both yeast and filamentous fungi (e.g. Brunt et al., 1998). Plant cells also contain a CCH capable of activating the mammalian glucocorticoid receptor in vitro. HSP70, HSP90, and an FKBP-type prolyl-isomerase have been identified as components of the wheat CCH (Reddy et al., 1998). The maize CCH also contains a DnaJ homolog and a cyclophilin-type prolyl-isomerase (J.A. Miernyk, unpublished data). Plant and yeast homologs of the mammalian steroid hormone receptors have not yet been reported, and the native targets for CCH function in these cells are at this time unknown. Stress70 The 70-kD stress proteins comprise a ubiquitous set of highly conserved molecular chaperones that range in actual size from 68 to 110 kD (Vierling, 1991; Boston et al., 1996; Miernyk, 1997). Some family members are constitutively expressed and are often referred to as HSC70 (70-kD heat shock cognate). Other family members are expressed only when the organism is challenged by environmental stresses such as temperature extremes, anoxia, heavy metals, and predation. These family members are generally referred to as HSP70 (70-kD heat shock protein), even, for example, when they are induced by cold shock. No differences in actual chaperone function have been described between the constitutive and stress-induced proteins, and hereafter they will be discussed collectively as Stress70. Several excellent recent reviews cover the molecular details of stress protein induction in plants (e.g.Schöffl et al., 1998), which are outside the scope of this review. Specific species of the Stress70 proteins are found in all subcellular compartments (Boston et al., 1996; Miernyk, 1997). While plant cytoplasmic Stress70 proteins are more closely related to their mammalian isologs, the mitochondrial and plastidic proteins are more similar to their prokaryotic counterparts. The rough ER luminal resident form of Stress70 is variously referred to as BiP (from early studies on its function as the IgG-binding protein of mammalian cells), GRP78 (78-kD Glc-regulated protein), and in yeast, KAR2 (karyogamy). The prokaryotic and organellar Stress70 proteins do not function as chaperones by themselves, but rather in concert with two accessory or co-chaperone proteins. The functional association of these components is often referred to as the Stress70 chaperone machine (Miernyk, 1997;Bukau and Horwich, 1998) (Fig. 3). DnaK (Stress70) is the central component of the machine, and functions as a chaperone in association with DnaJ, a chaperone-activating protein (CAP), and GrpE, a nucleotide exchange factor (NEF). Eukaryotic counterparts of DnaJ are widespread and are referred to variously as DnaJ homologs, Hsp40, or Stress40. Thus far the occurrence of eukaryotic cytoplasmic nucleotide exchange proteins has been controversial, although structurally related GrpE homologs have been found in mitochondria and plastids. Fig. 3. Open in new tabDownload slide Schematic presentation of the Stress70 chaperone machine showing the interactions among the central ATP-dependent chaperone, the chaperone-activating protein/DnaJ, and the nucleotide exchange factor/GrpE. The 44-kD N-terminal ATPase domain of Stress70 is indicated as a gray rectangle; the peptide-binding domain is indicated in black. A newly synthesized polypeptide can be recognized and bound by the ATP-ligated form of Stress70 while still nascent or immediately after release from the ribosome. This binary complex is recognized by the chaperone-activating protein (blue trapezoid) that binds and stimulates ATP hydrolysis. The ADP-ligated form of Stress70 has a lower affinity for the polypeptide, which is then released from the machine. If the polypeptide folds incorrectly, it is prone to aggregation. Similarly, a mature correctly folded protein will aggregate when unfolded/denatured. The ADP-ligated form of Stress70 is recognized by the nucleotide exchange factor (orange triangle) that promotes the exchange of ADP for ATP, allowing another cycle by the machine. A typical protein would require several cycles of binding and release before reaching the final correctly folded conformation. The polypeptide chain is not folded by the machine, but rather “held” in a conformation that allows expression of the folding information present in the primary sequence. Fig. 3. Open in new tabDownload slide Schematic presentation of the Stress70 chaperone machine showing the interactions among the central ATP-dependent chaperone, the chaperone-activating protein/DnaJ, and the nucleotide exchange factor/GrpE. The 44-kD N-terminal ATPase domain of Stress70 is indicated as a gray rectangle; the peptide-binding domain is indicated in black. A newly synthesized polypeptide can be recognized and bound by the ATP-ligated form of Stress70 while still nascent or immediately after release from the ribosome. This binary complex is recognized by the chaperone-activating protein (blue trapezoid) that binds and stimulates ATP hydrolysis. The ADP-ligated form of Stress70 has a lower affinity for the polypeptide, which is then released from the machine. If the polypeptide folds incorrectly, it is prone to aggregation. Similarly, a mature correctly folded protein will aggregate when unfolded/denatured. The ADP-ligated form of Stress70 is recognized by the nucleotide exchange factor (orange triangle) that promotes the exchange of ADP for ATP, allowing another cycle by the machine. A typical protein would require several cycles of binding and release before reaching the final correctly folded conformation. The polypeptide chain is not folded by the machine, but rather “held” in a conformation that allows expression of the folding information present in the primary sequence. While both the primary sequence and structural organization of the Stress70 proteins are highly conserved, results from recent studies suggest that the Stress70 reaction cycle might be significantly different in prokaryotes and the eukaryotic cytoplasm. The eubacterial paradigm begins with DnaJ binding to an unfolded peptide and this binary complex subsequently interacts with ATP-ligated DnaK. When ATP occupies the nucleotide site of DnaK, the chaperone is in the open state and can effectively bind the extended peptide (Fig. 2B). DnaJ promotes the hydrolysis of ATP by DnaK, and the concomitant conformational change drives the release of a more structurally organized portion of the polypeptide chain. GrpE subsequently facilitates the exchange of ADP for ATP. This cycle repeats until a stable structure is achieved. In the mammalian Stress70 cycle, the unfolded peptide reacts directly with HSP70 (Fig. 3). Next, the DnaJ homolog binds to the polypeptide/HSP70 complex and stimulates the ATPase activity. The HSP70 complex then adopts the more stable ADP-ligated state. This state is further stabilized by association with Hip, the HSP70 interacting protein. In the absence of a nucleotide exchange factor, the cycle ends with slow dissociation of ADP, acquisition of the next ATP molecule, and release of the polypeptide from the ATP-bound, open form of Stress70. In addition to Hip and Hop, which are mentioned in the section on HSP90, there are at least two other proteins that bind to the mammalian Stress70 complex and can potentially control chaperone activity. In keeping with the Hip/Hop vernacular, these have been termed Hap and Hup. Hap, the HSP70-associating protein, also known as BAG-1 or Rap46, is a negative modulator of Stress70 chaperone activity. Hup, the HSP70-unbinding protein, also known as p16 or Nm23, is ostensibly a nucleoside diphosphate kinase. Hup is also a stress protein and modulates HSP70 chaperone activity by promoting dissociation (unbinding) of the chaperone-target complex. The elaborate, but until now comprehensible, model of the mammalian Stress70 chaperone machine became a bit more difficult to fathom with a recent report that Hop interacts with TriC and modulates the protein folding activity of the chaperonin (molecular chaperone proteins that are related to GroE either by primary sequence homology or overall structure) by affecting nucleotide exchange (Gebauer et al., 1998). This would represent an unprecedented economy of function, with Hop regulating two structurally unrelated chaperone complexes. Less is known about the Stress70 chaperone machine in the cytoplasm of plant cells. The HSP70 proteins and DnaJ homologs of plant cells are structurally similar to their microbial or mammalian counterparts (Boston et al., 1996; Miernyk, 1997). While GrpE is a well-defined nucleotide exchange factor for the prokaryotic Stress70 chaperone machine, it is currently believed that the mammalian cytoplasmic machine functions without such a component. A small Arabidopsis protein (GB U64825) that stimulates ATP/ADP exchange by a plant Stress70/DnaJ-homolog complex has recently been characterized (B. Kroczyńska and J.A. Miernyk, unpublished observations). This protein (which should possibly be named the HSP70-exchange protein) has no obvious sequence relationship with GrpE, Hip, Hop, Hap, or Hup. To date, only one other plant component, a soybean Hop homolog (accession no. GBX79770), has been cloned or studied. Thus, it appears that the plant cytoplasmic Stress70 chaperone machine has some features in common with both bacterial and mammalian complexes, and perhaps unique features as well. The Chaperonins The chaperonins comprise the best-studied family of molecular chaperones (Boston et al., 1996; Netzer and Hartl, 1998; Sigler et al., 1998). They are double-ring, oligomeric structures that provide a closed compartment that shields folding proteins from the cellular environment (Fig. 2, D–F). Based upon evolutionary relationship, there are two distinct groups of chaperonins. The archetype group I chaperonin is GroEL, which occurs in the cytoplasm of eubacteria and within the mitochondria and plastids of eukaryotic cells, where it is referred to as cpn60 (Boston et al., 1996). Group II chaperonins include the archaeal thermosome and the eukaryotic cytoplasmic chaperonin complex, variously referred to as C-cpn, TCP-1 (T-complex protein-1), CCT (chaperonin containing TCP-1), or TriC (TCP-1 ring complex). Members of both groups require energy derived from ATP hydrolysis to drive protein folding, and share the same basic mode of action. However, details at the molecular level differ considerably (Sigler et al., 1998). Group I chaperonins depend upon a partner protein, chaperonin 10 (cpn10, or GroES in bacteria), while a distinct cpn10 protein is not part of the group II system. In group I chaperonin-mediated protein folding, an asymmetrical double-ring structure, in which GroES/cpn10 is bound to only one end, acts as the polypeptide acceptor (Fig. 2F). Binding occurs exclusively to the GroEL/cpn60 ring not occupied by GroES/cpn10. After polypeptide binding, a round of ATP hydrolysis in the occupied GroEL/cpn60 ring induces release of GroES/cpn10 (Fig. 2D). After release, GroES/cpn10 binds to the GroEL/cpn60 ring, where the polypeptide is bound. In this form the polypeptide is sequestered under a GroES/cpn10 “lid” in an enlarged central cavity. ATP then binds to the unoccupied ring, initiating polypeptide folding (Fig. 1E). ATP hydrolysis drives GroES/cpn10 release and allows the polypeptide to exit the GroEL/cpn60 central cavity. This cycle is repeated until the polypeptide reaches the final native structure (Bakau and Horwich, 1998; Sigler et al., 1998). Nuclear-encoded group I chaperonins are found within the plastids and mitochondria of plant cells, and it was based upon studies of the chloroplast cpn60 (or Rubisco large-subunit-binding protein, as it was previously known) that the molecular chaperone concept was revived and extended (Hemmingsen et al., 1988). The structures and reaction cycles of the plant mitochondrial chaperonins are highly homologous to those of GroEL/S. The reaction cycle of the plastid chaperonins is also homologous to that of the prokaryotic counterparts. There are, however, some intriguing structural differences. Plastids contain two distinct sets of GroEL homologs, cpn60α (61 kD) and cpn60β (60 kD). It remains unclear if there are distinct α- and β-ring structures in vivo or if there are structures composed of varying proportions of α- and β-subunits. The existence of distinct subunits implies distinct structures and distinct protein target specificity, however, this has not yet been experimentally demonstrated. Initial analyses of chloroplast cpn10 revealed that the protein was unusually large at 21 kD. The spinach cpn10 cDNA encodes an open reading frame that consists of two cpn10 sequences fused head to tail, explaining this apparent anomaly (Boston et al., 1996). It has been suggested that the two halves of the protein might play subtly different roles in vivo. The large fused-dimer form of cpn10 is found only in photosynthetic eukaryotes (Boston et al., 1996). It is noteworthy that cyanobacterial cpn10 is of the usual size, suggesting that the apparent gene-fusion event took place after the presumptive endosymbiosis that gave rise to chloroplasts. In mammalian cells, TriC forms a double-ring structure that is very similar in appearance to GroEL. The TriC is hetero-oligomeric; nine different subunits have been characterized (α–λ). TriC has been best characterized as a chaperone for actin and tubulin in mammalian cells (Boston et al., 1996; Sigler et al., 1998). In contrast to the organellar chaperonins, relatively little is known about TriC in plant cells. The larger subunit (e.g. α or 1) has been cloned from Arabidopsis and oat, and the deduced amino acid sequences are similar to those of their mammalian and yeast counterparts. In plant cells the TriC subunits are of relatively low abundance, and there have not yet been any reports of specific chaperone activity. The Small Stress Proteins The low-molecular-mass (small) heat shock proteins (smHSPs) (15–30 kD) are ubiquitous among eukaryotes, and represent a particularly important class of molecular chaperones in plant cells (Vierling, 1991). The smHSPs share a conserved C-terminal domain with the mammalian α-crystallin proteins. Biochemical analyses indicate that in vivo the smHSPs are found not as monomer/dimers but, rather, in high-molecular-mass complexes of 200 to 400 kD. Despite the occurrence of multiple species of smHSPs, they appear to exist as separate high-molecular-mass homo-complexes rather than as mixed-subunit hetero-complexes. This suggests subtly different functions for the different smHSPs, even when present in the same cellular compartment. In vitro, the smHSPs can both facilitate the reactivation of chemically denatured proteins and prevent heat-induced protein aggregation (Boston et al., 1996). Based on these observations, it is likely that the smHSPs can also act in vivo as a type of molecular chaperone. In marked contrast to members of the other classes of molecular chaperones, the activity of the smHSPs is ATP independent. In higher plants, six nuclear gene families encoding smHSPs have been described. Each family encodes proteins localized within distinct cellular compartments, including the cytoplasm, plastids, rough ER, and mitochondria (Waters et al., 1996). Additionally, specific smHSPs are expressed during various phases of plant development (Boston et al., 1996). Calnexin and Calreticulin Calnexin (CNX) is a rough ER-localized chaperone that is also a low-affinity, high-capacity calcium-binding protein (Helenius et al., 1997; Crofts and Denecke, 1998). CNX is a non-glycosylated type I integral membrane protein with a relatively short cytoplasmic domain and a large rough ER lumenal domain. Adjacent to the CNX transmembrane region is the P domain, a Pro-rich sequence that participates in glycan recognition and chaperone function. CNX binds ATP, although no ATPase activity has been reported, and ATP binding promotes a monomer to oligomer shift (Chapman et al., 1997). CNX interacts with a wide range of newly synthesized proteins, then dissociates from these proteins upon folding prior to export or further assembly (Helenius et al., 1997). Thus, CNX is a component of the rough ER quality control system that allows only proteins that have acquired a final native conformation to move on through the secretory pathway (Boston et al., 1996). Calreticulin (CRT) is a soluble lumenal protein of the rough ER. Like CNX, CRT is a low-affinity, high-capacity calcium-binding protein (Borisjuk et al., 1998). CRT has been found associated with other proteins in the rough ER lumen, and has similar behavior and target specificity to that of CNX. Possibly, CNX and CRT cooperate in the rough ER as chaperones in the folding of secretory proteins (Helenius et al., 1997; Crofts and Denecke, 1998). Because of the environment with the lumen of the rough ER (Huppa and Ploegh, 1998), protein folding is a particularly complex process involving multiple molecular chaperones and folding catalysts. Many of the interactions must occur sequentially in order for proteins to achieve the correct native structure. While CNX binds to all folding intermediates, CRT associates preferentially with the earliest oxidative species either immediately prior to or coincident with disulfide bond formation (Helenius et al., 1997). Thus, recognition and binding by CRT seems to precede interaction with the rough ER-localized folding catalysts. Overall, CNX and CRT promote correct protein folding in the rough ER by inhibiting aggregation, preventing premature oxidation and oligomerization, and suppressing degradation of incompletely or incorrectly folded intermediates. FOLDING CATALYSTS ACCELERATE THE RATE OF FOLDING In contrast to molecular chaperones, the folding catalysts are conventional enzymes. Protein-folding catalysts accelerate the slow chemical reactions in protein folding that might otherwise be rate limiting. Without the acceleration of the rate-limiting reactions, cellular proteins would be trapped in intermediate states, and most folding intermediates are vulnerable to aggregation and to non-productive interactions with other proteins (Dobson et al., 1998). The two best-studied protein-folding catalysts are protein disulfide isomerase (EC 5.3.4.1) and peptidyl-prolyl cis-trans-isomerase (EC5.2.1.8). Protein Disulfide Isomerase Protein disulfide isomerase (PDI) is a member of the thioredoxin superfamily of proteins, and contains two copies of the characteristic active-site motif CXXC. Protein disulfide isomerase facilitates folding through its ability to reduce or oxidize disulfide bridges in the presence of an oxidizing or reducing agent such as glutathione (Huppa and Ploegh, 1998). To accomplish this, PDI expedites disulfide interchange by shuffling the disulfide bonds to quickly find the most thermodynamically stable pairing (Laboissière et al., 1995). In eukaryotic cells, PDI is localized exclusively within the lumen of the rough ER. This is a unique environment for protein folding in terms of pH, redox conditions, and high calcium ion concentrations (Huppa and Ploegh, 1998; Møgelsvang and Simpson, 1998). Peptidyl Prolyl Cis-Trans Isomerases Attempts to follow the literature on PPI are complicated by the diverse terminology employed by various researchers. In addition to the formal enzyme name, PPIs are variously referred to as: cyclophilins, immunophilins, rotamase, FKBPs, and parvulins (Dolinski and Heitman, 1997). The term immunophilin comes from the ability of PPI to bind immunosuppressive drugs. The cyclophilins are immunophilins that bind the cyclic undecapeptide cyclosporin A, while FKBPs are immunophilins that bind the macrolide drug FK506. Whatever the term used, PPI participates in protein folding by accelerating the cis-trans isomerization of prolyl peptide bonds. Pro isomerization can be the rate-limiting step in overall protein folding, and this reaction is accelerated 300-fold by PPI (Schmid, 1993). After isomerization, the correct form of Pro is stabilized by the polypeptide secondary or tertiary structure. While most PPI proteins are localized in the cytoplasm/nucleus, there are unique forms present within the rough ER/nucleus, the plastid stroma, and the mitochondrial matrix (Boston et al., 1996; Kurek et al., 1999). What's in a Name? The distinctions between chaperones and folding catalysts have in some instances become blurred. The folding catalysts bind to their target proteins, and in some in vitro experiments it has been observed that this binding can prevent aggregation. Can the folding catalysts also therefore be chaperones? Furthermore, site-directed mutagenesis has been used to modify catalytically essential residues in PDI or PPI, and the resultant mutant proteins still had chaperone activity in vitro. However, such results must be interpreted with care. Recently it was shown that a mutant form of PPI that had no activity in vitro with an artificial substrate, remained fully active when assayed with an unfolded polypeptide as the substrate (Scholz et al., 1997). The distinction between chaperones and folding catalysts remains that the latter accelerate the rate of target polypeptide folding. PROTEIN FOLDING IN THE CYTOPLASM The majority of nuclear-encoded proteins reside in the cytoplasm throughout their lifetime. It is important to identify which molecular chaperones and folding catalysts assist the newly synthesized cytoplasmic proteins to their final native state. This is a particularly interesting question in plant cells that are uniquely complex in their array of cytoplasmic chaperones. Some proteins, especially those that are small, single-domain, and monomeric, are able to fold spontaneously in the cytoplasm without any external assistance, or perhaps only with the assistance of a folding catalyst. Other proteins are recognized by the Stress70 chaperone machine and bound while still nascent. There are three potential fates for this complex: the polypeptide might fold to the correct final structure and be released from the chaperone, it might be maintained in an unfolded translocation-competent conformation until transferred to the protein machinery associated with an organellar outer membrane, or the polypeptide might be transferred to a different chaperone for further folding, assembly, or oligomerization. Among the other known cytoplasmic chaperones, the chaperonins, the CCH, and the smHSPs are the best understood. For the most part, the actual native target proteins for each of the chaperone systems remain unidentified. The inner chamber of the chaperonin structure, or “Anfinsen cage” as it is sometimes referred to, is limited to polypeptides no larger than 55 kD (Netzer and Hartl, 1998; Sigler et al., 1998). A precise structural model of the CCH has not yet been defined; however, in principle, the HSP70 and HSP90 components are capable of assisting the folding of proteins considerably larger than 55 kD. In mammalian cells the only CCH target polypeptides identified to date are various receptors and some protein kinases (Pratt and Toft, 1997). The oligomeric small HSPs are thought to bind target polypeptides on the external surfaces and should be capable of assisting in the folding of any size protein (Vierling, 1991; Boston et al., 1996). For the most part, HSP100/Clp are thought to mediate the unfolding/refolding of damaged proteins (Schirmer et al., 1996; Hoskins et al., 1998), but a potential role in the initial stages of protein folding cannot be excluded. Protein refolding by HSP100/Clp is typically assisted by the Stress70 machine and/or HSP90. The complexity of the various interactions has not yet been fully defined. While chaperone-mediated protein folding is generally considered to be unidirectional, there is some in vitro evidence for retrograde transfer between chaperones (Netzer and Hartl, 1998). Additionally, there are some instances in which more than two of the chaperone systems seem to be involved in the folding of a single target protein (i.e. the prion protein has been observed during its “normal” lifetime to be associated with HSP70, the chaperonins, and HSP90). FROM THE CYTOPLASM INTO THE ER Molecular chaperones have a dual interaction with proteins destined to reside within the ER or any of other compartment of the endo-membrane system. These proteins are first inserted into a channel in the ER membrane that is aligned with the ribosome (Vitale and Denecke, 1999). Translocation must occur in a luminal direction. The directionality and driving force for polypeptide translocation into the ER lumen are provided by the resident Stress70 chaperone, BiP, which acts as a “molecular ratchet” (Matlack et al., 1999). Multiple BiP molecules interact with the translocating polypeptide to minimize any retrograde movement. The BiP:target polypeptide complex interacts with the J-domain of an integral membrane protein (Sec63p in yeast), which activates the ATPase function of the chaperone (Miernyk, 1997). Once within the ER lumen, polypeptide folding begins immediately. As in the cytoplasm, non-productive folding is forestalled by interaction with the Stress70 chaperone machine (Møgelsvang and Simpson, 1998). BiP is absolutely essential for protein translocation and folding in the ER (Li et al., 1993). In addition to variously interacting with the specific ER homologs of Stress70 (BiP), DnaJ, PDI and PPI, the unique environment found within the rough ER lumen (Huppa and Ploegh, 1998) has given rise to the CNX and CRT chaperones and the PDI-folding catalyst (Chapman et al., 1997; Borisjuk et al., 1998; Crofts and Denecke, 1998; Møgelsvang and Simpson, 1998). While all of the Stress70 proteins are subject to multiple posttranslational modifications (phosphorylation,N-methylation) (e.g. Miernyk et al., 1992), BiP is unique in that it is additionally ADP-ribosylated (Boston et al., 1996). Specific roles have not been defined for any of the modifications, however, it has been shown that the extent of phosphorylation and ADP-ribosylation varies with physiological state or in some mutants, suggesting regulatory functions (Fontes et al., 1991). PROTEIN FOLDING IN OTHER SUBCELLULAR COMPARTMENTS Most organellar proteins are nuclear encoded and synthesized as precursors in the cytoplasm. As such, they will interact with various cytoplasmic chaperones prior to membrane translocation. After membrane translocation, protein folding within other subcellular organelles of a plant cell likely follows pathways similar to those in the cytoplasm or ER, and specific forms of HSP70, HSP90, the chaperonins, and the small HSPs have been found in plastids, mitochondria, and peroxisomes (Boston et al., 1996; Waters et al., 1996; Corpas and Trelease, 1997; Miernyk, 1997). PROSPECTUS: WHERE DO WE GO FROM HERE? I have attempted to provide an overview of the roles played by several of the better-understood chaperones in protein folding. Such a presentation is by definition limited in two ways: by space and by knowledge. SecB is a general chaperone for proteins that will be secreted by bacteria. MSF is a mammalian ATP-dependent protein that specifically chaperones mitochondrial precursors. Atx1p from yeast and UreE from bacteria are metal ion chaperones for copper and nickel, respectively. PapD and CssC chaperone pilus formation in prokaryotes. This list could go on, but the unifying feature of these otherwise diverse chaperones is that no homolog has yet been identified in plants. Thus, it seems likely that many different molecular chaperones remain, awaiting discovery. Necessarily, most of the early studies on molecular chaperones and folding catalysts have been conducted in vitro, based upon mammalian and microbial paradigms, and used model target proteins. It is crucially important that we bridge the gap between these studies and actual in vivo activities. Only then can we be said to have truly “cracked the second half of the genetic code” (Ellis, 1991). A promising step in this direction has recently been provided byForreiter et al. (1997). 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van der Luit, Arnold H.; Olivari, Claudio; Haley, Ann; Knight, Marc R.; Trewavas, Anthony J.

doi: 10.1104/pp.121.3.705pmid: 10557218

Abstract Cold shock and wind stimuli initiate Ca2+ transients in transgenic tobacco (Nicotiana plumbaginifolia) seedlings (named MAQ 2.4) containing cytoplasmic aequorin. To investigate whether these stimuli initiate Ca2+ pathways that are spatially distinct, stress-induced nuclear and cytoplasmic Ca2+ transients and the expression of a stress-induced calmodulin gene were compared. Tobacco seedlings were transformed with a construct that encodes a fusion protein between nucleoplasmin (a major oocyte nuclear protein) and aequorin. Immunocytochemical evidence indicated targeting of the fusion protein to the nucleus in these plants, which were named MAQ 7.11. Comparison between MAQ 7.11 and MAQ 2.4 seedlings confirmed that wind stimuli and cold shock invoke separate Ca2+ signaling pathways. Partial cDNAs encoding two tobacco calmodulin genes, NpCaM-1 andNpCaM-2, were identified and shown to have distinct nucleotide sequences that encode identical polypeptides. Expression ofNpCaM-1, but not NpCaM-2, responded to wind and cold shock stimulation. Comparison of the Ca2+dynamics with NpCaM-1 expression after stimulation suggested that wind-induced NpCaM-1 expression is regulated by a Ca2+ signaling pathway operational predominantly in the nucleus. In contrast, expression ofNpCaM-1 in response to cold shock is regulated by a pathway operational predominantly in the cytoplasm. Calmodulin is highly conserved in eukaryotes and is considered to be a multifunctional protein because of its ability to interact and regulate the activity of a number of other proteins (Hepler and Wayne, 1985; Gilroy et al., 1993; Poovaiah and Reddy, 1993; Trewavas and Knight, 1994). In plant cells, calmodulin is considered to be the primary sensor for changes in cellular free Ca2+levels ([Ca2+]i) (Roberts and Harmon, 1992). As [Ca2+]i rises transiently after signaling, the combination of Ca2+ with calmodulin leads to the activation of numerous target proteins initiating the physiological response. Calmodulin has been purified and characterized from a number of plant species. Genomic and/or cDNA clones encoding calmodulin have been isolated and characterized from Arabidopsis (Ling et al., 1991; Perera and Zielinski, 1992), potato (Takezawa et al., 1995), and wheat (Yang et al., 1996). In all multicellular organisms in which it has been examined, genes encoding the different calmodulin isoforms are under the control of different promoters that exhibit distinct temporal and spatial expression (Ling et al., 1991; Gannon and McEwen, 1994; Shimoda et al., 1995; Solà et al., 1996). In plant cells, stimuli such as touch, wind, or temperature shocks induce the rapid accumulation of mRNA levels encoding calmodulin and calmodulin-related proteins (Jena et al., 1989; Braam, 1992; Perera and Zielinski, 1992; Watillon et al., 1992; Takezawa et al., 1995). Since many of these signals also elevate [Ca2+]i (Knight et al., 1991, 1992, 1997), and artificial elevation of [Ca2+]i in cultured cells increases calmodulin mRNA accumulation (Braam, 1992), it has been suggested that the transduction of environmental signals regulating calmodulin gene expression are in part regulated by [Ca2+]i levels (Braam and Davis, 1990; Braam, 1992). Calmodulin has been detected in several plant cell compartments (Biro et al., 1984; Collinge and Trewavas, 1989). In particular, a substantial amount of calmodulin has been found in both plant and animal nuclei and in combination with nuclear Ca2+ signals, gene expression is thought to be regulated via Ca2+/calmodulin interaction with transcription factors or via specific protein kinases (Bachs et al., 1992; Gilchrist et al., 1994; Kocsis et al., 1994; Zimprich et al., 1995; Szymanski et al., 1996). Plants transformed with a cDNA encoding the Ca2+-sensitive luminescent protein aequorin provides a simple, non-invasive means of measuring [Ca2+]i in whole plants. Many new signals initiating rapid changes in [Ca2+]i have subsequently been detected with this technology, including the mechanical signals of touch and wind, salt/drought, heat shock, and osmotic pressure (Trewavas and Knight, 1994; Haley et al., 1995; Knight et al., 1996,1997; Takahashi et al., 1997; Gong et al., 1998). Furthermore, aequorin targeted to chloroplasts (Johnson et al., 1995) and the vacuole membrane (Knight et al., 1996) clearly indicated that the [Ca2+]i signal is strictly compartmentalized within the cell. In a previous paper (Knight et al., 1992), wind and cold shock stimulation were investigated in tobacco (Nicotiana plumbaginifolia) seedlings. Mechanical stimulation induced by puffs of air blown over the seedling resulted in a slight movement of the seedling around the hypocotyl/root junction lasting about 0.02 to 0.3 s. Temperature shocks can be induced by irrigating the plant briefly with cold water at 0°C to 5°C. Both signals induce [Ca2+]i spikes in aequorin transgenic tobacco seedlings. However, careful titration with different inhibitors suggested specific spatial organization of the Ca2+ signal depending on the type of stimulation. Ruthenium red at low concentrations specifically blocked the transient induced by wind or touch and did not affect the cold shock [Ca2+]i transient; lanthanum and gadolinium chlorides, which are Ca2+-channel blockers, specifically blocked the cold shock signal without influencing the wind-induced [Ca2+]i transient. It was concluded that the two signals were mobilizing separate pools of [Ca2+]i. At present, no direct evidence is available to indicate whether compartmentalization of the Ca2+ signal is significant for other downstream responses such as calmodulin gene expression. To address this question, we created a fusion protein between nucleoplasmin (a major oocyte nuclear protein) and aequorin, which was then used to transform tobacco seedlings. Transfection of animal cells with this construct allowed measurement of nuclear Ca2+ concentrations and indicated the presence of compartmentalized regulation of Ca2+ signaling pathways (Badminton et al., 1995, 1996, 1998). The use of the same construct in plant cells could help to clarify the Ca2+ signaling pathways involved in the control of calmodulin gene expression by wind stimuli and cold shock. MATERIALS AND METHODS All enzymes used for recombinant DNA manipulation were purchased from Promega Biotech (Southampton, UK). Plasmid DNA isolation kits were obtained from Qiagen (Dorking, UK), agar was from Difco Laboratories (Detroit), and all plant tissue culture reagents and other chemicals were from Sigma (Dorset, UK). Exceptions were 1,2-bis(o-aminophenoxy)ethane-N;N;N;N-tetraacetic acid-acetoxymethyl ester (BAPTA-AM) from Calbiochem (Nottingham, UK) and ruthenium red from LC Laboratories (Woburn, MA). Macerozyme and cellulase used for the production of protoplasts were from Yakult Honsha (Tokyo). Native coelenterazine and cp-coelenterazine were purchased from Molecular Probes (Leiden, The Netherlands). Oligonucleotide primers were prepared by Genosys (Cambridge, UK). Plant Materials and Growth Conditions MAQ 2.4, the transgenic tobacco (Nicotiana plumbaginifolia) line that expresses cytosolic aequorin (Knight et al., 1991), was used to measure changes in cytosolic free Ca2+ concentrations ([Ca2+]cyt). All seedlings used for experiments were grown on one-half-strength Murashige and Skoog medium (Murashige and Skoog, 1962) and 0.8% (w/v) agar either in luminometer cuvettes or on plates at 25°C with a 16-h photoperiod, and used when 7 to 10 d old. Design and Expression of Nuclear-Targeted Chimeric Aequorin To target aequorin to plant nuclei, a chimeric construct in which the nucleoplasmin coding region was placed in frame with the coding region of apoaequorin (Badminton et al., 1995; kindly provided by Dr. M. Badminton, University of Wales, UK) was cloned into pDH51 (Pietrzak et al., 1986) as a SmaI-SalI fragment. The entire construct, including the 35S promoter and terminator, was cloned into the Agrobacterium tumefaciens binary vector pBIN19.Escherichia coli JM101 and XL-1 Blue were used as hosts for all recombinant DNA manipulations (Sambrook et al., 1989). A. tumefaciens LBA4404 and N. plumbaginifolia were used for plant genetic transformation (Draper et al., 1988). Immunolocalization of Aequorin For immunolocalization using fluorescein isothiocyanate (FITC)-labeled secondary antibodies, protoplasts were washed and pelleted in 0.5% (w/v) MES (pH 5.8), 80 mmCaCl2, 300 mm mannitol, and fixed for 15 min on poly-l-Lys treated slides using 4% (w/v) paraformaldehyde. Cells were permeabilized for 40 min with 0.5% (v/v) Triton X-100 in 50 mm PIPES (pH 6.9), 5 mmMgSO4, 5 mm EGTA, and 300 mm mannitol. Samples were incubated for 5 min with 1% (w/v) BSA followed by a 1.5-h incubation at 37°C with mouse anti-aequorin (1:1,000) obtained as previously described (Knight et al., 1991), and then with FITC-labeled goat anti-mouse IgG from Sigma (Dorset, UK) (1:30) in PBS (pH 5.8), 1% (w/v) BSA, 20 mmNaN3 for 45 min at 37°C. Cells were stained with DAPI, mounted in Citifluor (Citifluor Products, Kent, UK), and photographed with an epifluorescence microscope (Polyvar, Reichert-Jung, Vienna, Austria) using Ektachrome T film (ASA 64, Eastman-Kodak, Rochester, NJ). For immunoelectron microscopy using gold-labeled secondary antibodies, protoplasts were prepared as described above and fixed for 15 min using PBS-buffered one-fourth-strength Karnovsky's fixative at pH 5.8 (Karnovsky, 1965). The fixed tissue was dehydrated by consecutive 10-min incubations in 30%, 50%, 70%, and 90% (v/v) ethanol, and for 20 min in three changes of dehydrated absolute ethanol followed by propylene oxide (twice for 15 min). The embedding of fixed and dehydrated tissue was carried out using resin from Agar Scientific (Essex, UK). Thin sections (80–90 nm) placed on gold grids were incubated in 1% (w/v) BSA in PBS for 5 min at room temperature. Sections were incubated with mouse anti-aequorin (1:200) obtained as previously described (Knight et al., 1991) for 2 h at room temperature or for 18 to 24 h at about 4°C in a moist chamber. The antisera or immunosorbent-purified antibodies were diluted in 1% (w/v) BSA-PBS (pH 7.4). The grids were placed on drops of a 20-fold dilution of the 1 nM gold-labeled goat anti-mouse IgG solution from British Biocell International (Cardiff, UK) for 1 h at room temperature in a moist chamber. The sections were stained with 5% (v/v) uranyl acetate (5–7 min), and washed thoroughly with distilled water and PBS and a subsequent Reynold's lead citrate solution (2–5 min). Gold particles were stained with a silver enhancement kit from Sigma prior to examination using an electron microscope (model 100S, JEOL, Hartfordshire, UK). In Vitro and in Vivo Aequorin Reconstitution, Wind and Cold Shock Stimulation, and Ca2+ Measurements For in vitro reconstitution of aequorin, seedlings were homogenized in 50 mm Tris-Cl (pH 7.4), 500 mmNaCl, 5 mm β-mercaptoethanol, 10 mm EGTA, and 0.1% (w/v) BSA, incubated with 2 μm coelenterazine for at least 4 h in the dark, discharged by adding an equal volume of 100 mm CaCl2 (Knight et al., 1991), and the total amount of luminescence produced was measured. Luminescence was measured using a digital chemiluminometer consisting of an photomultiplier (model 9829A, EMI, Middlesex, UK) with a cooling system (FACT50, EMI) (Badminton et al., 1995). For in vivo reconstitution of aequorin, seedlings were germinated as described above. Aequorin was reconstituted in vivo by placing a 3-μL droplet of 2 μm coelenterazine between the cotyledons and incubating at least 4 h in the dark. For experiments with inhibitors, seedlings were submerged and incubated for 4 h. A long period is needed to allow the compounds to penetrate into the seedling. Following this treatment, the liquid was drained and a 3-μL droplet of 2 μm coelenterazine with the relevant inhibitor was placed between the cotyledons and left for at least another 4 h in the dark, after which time the liquid was removed and the luminescence measurement carried out. Wind stimulation was simulated by instantly injecting 5 mL of air into the sample housing of the luminometer. Cold shock was simulated by slowly injecting 1 mL of ice-cold water into the sample housing of the luminometer. The light emitted by the seedling is a measure of the change in the [Ca2+]i and was recorded every 0.2 s using a cooled photomultiplier tube. For measurement of changes in cytosolic Ca2+native coelenterazine, we used the luminophore used in our previous experiments (Knight et al., 1991, 1992). Initial investigations showed nuclear Ca2+ changes to be smaller than those in the cytoplasm in response to cold shock. To reduce errors in the measurement of emitted light, the more sensitivecp-coelenterazine was used, which enabled approximate equality in light emission measurements between the cytosolic and nuclear compartments (Shimomura et al., 1993). Reconstitutedcp-aequorin shows improved light emission in the lower Ca2+ ranges and is thus useful for detecting smaller changes in [Ca2+]i (Shimomura et al., 1993). Calibration constants for cp-coelenterazine (and many other coelenterazines) in comparison with native coelenterazine have been published previously (Shimomura et al., 1993). The luminescent light was calibrated into Ca2+concentrations by a method based on the calibration curve of Allen et al. (1977): L/L max = ([1 +K R × {Ca2+}]/[1 +K TR +K R × {Ca2+}])3, whereL is the amount of light per second,L max is the total amount of light present in the entire sample over the course of the experiment, [Ca2+] is the calculated Ca2+ concentration,K R is the dissociation constant for the first Ca2+ ion to bind, andK TR is the binding constant of the second Ca2+ ion to bind to aequorin;K R = 26 × 106 m −1 andK TR = 57m −1 forcp-coelenterazine (Shimomura et al., 1993) andK R = 2 ×106 m −1 andK TR = 55m −1 for native coelenterazine. Total RNA Extraction, RACE (3′ RACE), and Northern-Blot Analysis Total RNA from seedlings was extracted according to the method ofLópez-Gómez and Gómez-Lim (1992), a method designed to obtain RNA free of polysaccharide contamination. Seedlings for RNA extraction were 7 to 10 d old and grown under the same conditions as seedlings for luminometry. Inhibitors were applied for a 4-h period, after which time the solution was drained and the seedlings were allowed to recover overnight. For 3′-RACE, total RNA was extracted from unstimulated seedlings (T0), 1 h after wind stimulation (T1W), and 2 h after cold shock stimulation (T2CS). cDNA was synthesized from 5 μg of total RNA in a buffer consisting of 50 mm Tris-Cl (pH 8.3), 3 mm MgCl2, 75 mm KCl, 10 mm DTT, and 0.5 μm of each dNTP, 10 units of RNasin (Promega Biotech), 100 ng μL−1 of dT17-adapter primer, QT (5′- CCAGTGAGCAGAGTGACGAGGACTCGAGCTCAAGC- TT17VN-3′, with V = G, C, A and N = G, C, T, A), 10 units of SuperScript II RNase H− reverse transcriptase from Life Technologies (Paisley, UK) in a total volume of 20 μL. The mixture was incubated for 5 min at room temperature, for 1 h at 42°C, for 10 min at 50°C, and for 15 min at 70°C. The RNA was then removed with 0.2 unit of RNase H from Life Technologies and the whole reaction was diluted with 1 mL of TE buffer (10 mm Tris-Cl [pH 7.6] and 1 mm EDTA) to produce the cDNA pool for amplification. For amplification, a PCR cocktail was prepared consisting of: 5 μL of 10× PCR buffer (670 mm Tris-Cl, pH 8.8, 67 mmMgCl2, 17 mg mL−1 BSA, 166 mm[NH4]2SO4), 5 μL of DMSO, 5 μL of 10× dNTPs (10 mm each), and 30 μL of distilled water, 1 μL of adapter primer, Qi (ACGAGGACTCGAGCTCAAGC, 25 pmol μL−1), 1 μL of a calmodulin-specific primer, E086 (GCATCACGACTAAGGAGCTT, 25 pmol μL−1), and 1 μL of cDNA pool. The cDNA was denatured 5 min at 95°C and cooled to 72°C. Then 2.5 units of Taq polymerase and 30 μL of mineral oil were added. Primers were annealed and extended at 52°C or 56°C for 5 min and at 72°C for 40 min to ensure correct replication, respectively, followed by a 20- to 35-times cycle: 95°C for 40 s, 52°C or 56°C for 1 min, 72°C for 3 min, and ended by a 15-min incubation at 72°C to complete the reaction.NpCaM-1 (accession no. AJ005039) and NpCaM-2(accession no. AJ005040) were cloned using the pCR-Script Amp SK(+) cloning kit from Stratagene (Cambridge, UK) and used for sequence analysis. Sequence analysis was carried out on both the DNA strands in quadruple. For each strand, sequencing reactions were performed using a dye-terminator cycle sequencing ready reaction kit (PRISM, ABI, Perkin-Elmer, Cheshire, UK) and sequenced using an automatic sequencer (Perkin-Elmer). For northern-blot analysis, 5 to 15 μg of total RNA was size-fractionated on a 1.3% (w/v) denaturing-formaldehyde agarose gel (Sambrook et al., 1989). To ensure that an equal amount of RNA was loaded, a picture of the ethidium bromide-stained gel was taken, scanned, and quantified with Imagequant software (Molecular Dynamics, 's-Hertogenbosch, The Netherlands). The 3′-untranslated regions (UTRs) of NpCaM-1 andNpCaM-2 were used as DNA hybridization probes and were labeled with [32P]dCTP by random-primed labeling from Amersham (Buckinghamshire, UK), and hybridized in 4× SSC, 1% (w/v) SDS, 200 mm Tris-Cl (pH 7.6), 10% (w/v) dextran sulfate, 100 μg mL−1herring-sperm DNA, and 2× Denhardt's solution, at 65°C overnight, and washed for 20 min in 2× SSC at 65°C, followed by a brief wash in 2× SSC, 1% (w/v) SDS at room temperature. Filters were either exposed to HyperfilmMP from Amersham or a phosphor plate, and imaged with a phosphor imager from Molecular Dynamics. Intensities of hybridizing bands were quantified using Imagequant software. RESULTS Transformation of Tobacco with a Nucleoplasmin Aequorin Construct and Localization of the Expressed Fusion Protein Wind and cold shock stimulation initiate specific Ca2+ signaling pathways (Knight et al., 1992). The use of different inhibitors suggested the specific organization of the Ca2+ signal depending on the type of stimulation. To investigate the organization of the Ca2+ signal in more detail, we transformed tobacco with a nucleoplasmin/aequorin construct. This construct was used previously (Badminton et al., 1995, 1996, 1998) to investigate the putative independence of the regulation of nuclear ([Ca2+]nuc) and cytoplasmic ([Ca2+]cyt) Ca2+ in transfected mammalian cell lines. Nucleoplasmin is an abundant nuclear protein in Xenopus laevis oocytes (Philpott and Leno, 1992). After leaf disc transformation, 7-d-old F1seedlings of >20 individual transformants were homogenized in 50 mm Tris-Cl (pH 7.4), 500 mm NaCl, 5 mm β-mercaptoethanol, 10 mm EGTA, and 0.1% (w/v) BSA, aequorin was reconstituted with added coelenterazine overnight as described previously (Knight et al., 1991, 1993, 1996), and aequorin levels were measured by light emission. Homogenates of all transformants were separated on SDS gels and the relative amounts of apoaequorin confirmed using western blotting and mouse anti-apoaequorin as described previously (Knight et al., 1991). The transformant containing the highest levels of expression was designated MAQ 7.11. The cellular distribution of the nucleoplasmin/aequorin fusion protein was examined using immunocytochemistry with anti-apoaequorin and either FITC or gold-labeled secondary antibodies. Protoplasts were isolated from mature leaves of untransformed tobacco and from MAQ 2.4 and MAQ 7.11 containing the nucleoplasmin/aequorin construct, and stained for apoaequorin distribution. Figure 1, A to C, shows protoplasts stained first with 4′,6-diamidino-2-phenylindole dihydrochloride or DAPI (to highlight DNA) and then stained with anti-apoaequorin followed by fluorescent secondary antibody (Fig. 1, D–F). The distribution of staining between the MAQ 2.4 and the MAQ 7.11 construct is clearly very different. The aequorin is distributed throughout the cytoplasm of the highly vacuolated protoplasts of MAQ 2.4 (Fig. 1, B and E), while the nucleoplasmin/aequorin construct is predominantly concentrated in the nuclear region of the protoplasts for MAQ 7.11 (Fig. 1, C and F). Fig. 1. Open in new tabDownload slide Targeting of aequorin to tobacco cell nuclei. Protoplasts of wild-type tobacco, MAQ 2.4, and MAQ 7.11 stained with DAPI are shown in A, B, and C, respectively. The same protoplasts treated with anti-apoaequorin and FITC-labeled secondary antibody are shown in D, E, and F. G shows a MAQ 7.11 protoplast treated with anti-apoaequorin and gold-labeled secondary antibody. Bar = 1 μm. C, Cytoplasm; N, nucleus. Fig. 1. Open in new tabDownload slide Targeting of aequorin to tobacco cell nuclei. Protoplasts of wild-type tobacco, MAQ 2.4, and MAQ 7.11 stained with DAPI are shown in A, B, and C, respectively. The same protoplasts treated with anti-apoaequorin and FITC-labeled secondary antibody are shown in D, E, and F. G shows a MAQ 7.11 protoplast treated with anti-apoaequorin and gold-labeled secondary antibody. Bar = 1 μm. C, Cytoplasm; N, nucleus. Confirmation of this distribution was obtained using gold-labeled secondary antibody. Figure 1G shows a nucleus and two associated areas of chloroplast/cytoplasm of a MAQ 7.11 protoplast. The intact nucleolus and nuclear envelope are clearly visible. Staining with gold-labeled secondary antibody revealed a gold particle distribution that was much more highly concentrated over the nuclear regions than the neighboring cytoplasm and strongly localized in dense chromatin. We quantified the gold particle distribution on a large number of sections and observed that 86% was localized in nuclei. Of the remainder, 9% was found in the chloroplasts and 5% in the cytoplasm. The distribution of nucleoplasmin/aequorin between the nucleus and other cytoplasmic compartments was similar to that reported for the distribution of nucleoplasmin in HeLa cells, 90% to 92% nuclear localization (Greber and Gerace, 1995), with a slightly higher proportion outside the nucleus. Isolation of Wind- and Cold-Shock-Induced and Non-Induced Tobacco Calmodulin Genes In all plants examined so far, calmodulin is represented by multigene families, and the individual calmodulin members exhibit both tissue-specific and developmental-stage-specific expression (Ling et al., 1991; Takezawa et al., 1995). As the length and the sequence of 3′-UTRs of calmodulin isoforms were reported to be different (Takezawa et al., 1995), 3′-RACE was carried out in tobacco to identify differentially expressed calmodulin genes. Using this technique, several potential calmodulin transcripts were identified. One of these putative calmodulin transcripts, designated NpCaM-1, appeared to be induced by wind and cold shock, while another,NpCaM-2, was not (data not shown). These two cDNAs were cloned and sequenced. In Figure 2, the partial sequence of two calmodulin isoforms is shown starting from the first Ca2+-binding site of calmodulin. The partial sequences of NpCaM-1 and NpCaM-2 are different in nucleotide sequence; however, they encode polypeptides with the same amino acid sequence. The 3′-UTRs were subcloned and used as DNA hybridization probes to study the expression kinetics ofNpCaM-1 and NpCaM-2 using northern-blot analysis. As shown in Figure 3, this type of analysis indicated that NpCaM-1 mRNA accumulates after wind and cold shock signaling, whereas NpCaM-2 does not. Fig. 2. Open in new tabDownload slide Partial cDNA sequence of NpCaM-1and NpCaM-2 showing nucleotide and predicted amino acid identities. A, Nucleotide sequence; B, amino acid sequence. Primers used for 3′-RACE and subsequent PCR are indicated in lowercase; stop codons are underlined. Homology is indicated with bars. Fig. 2. Open in new tabDownload slide Partial cDNA sequence of NpCaM-1and NpCaM-2 showing nucleotide and predicted amino acid identities. A, Nucleotide sequence; B, amino acid sequence. Primers used for 3′-RACE and subsequent PCR are indicated in lowercase; stop codons are underlined. Homology is indicated with bars. Fig. 3. Open in new tabDownload slide Expression kinetics of NpCaM-1 andNpCaM-2 determined by northern-blot analysis after stimulation by a single wind signal or a single cold shock. The 3′-UTRs of NpCaM-1 and NpCaM-2 were used as DNA hybridization probes to study the expression kinetics ofNpCaM-1 and NpCaM-2. Water of room temperature was used as a control. Fig. 3. Open in new tabDownload slide Expression kinetics of NpCaM-1 andNpCaM-2 determined by northern-blot analysis after stimulation by a single wind signal or a single cold shock. The 3′-UTRs of NpCaM-1 and NpCaM-2 were used as DNA hybridization probes to study the expression kinetics ofNpCaM-1 and NpCaM-2. Water of room temperature was used as a control. Wind- and Cold-Shock-Induced Ca2+ Changes in MAQ 2.4 and MAQ 7.11 and NpCaM-1 mRNA Accumulation Changes in [Ca2+]cytin tobacco seedlings in response to wind and cold shock have been previously reported (Knight et al., 1991, 1992). By constructing plants in which the distribution of aequorin is clearly different from cytoplasmic aequorin, we were able to examine the spatial organization of the Ca2+ signal in response to wind and cold shock. Prior to [Ca2+]imeasurements, coelenterazine was placed between the cotyledons to allow the reconstitution of aequorin. Wind and cold shock stimulation were achieved respectively by injecting air instantly or ice-cold water gently from above into the sampling housing of the luminometer. Conversions of emitted luminescence at each time point to free Ca2+ levels were performed as described in “Materials and Methods.” Figure 4 shows the effects of wind and cold shock signaling in MAQ 2.4 and MAQ 7.11. Because individual seedlings varied slightly in their absolute response, we have indicated only the ses of the peak values. For wind response the mean peak Ca2+ increase in the MAQ 2.4 and MAQ 7.11 were, respectively, 1.08 μm (n = 7) and 0.79 μm (n = 8) (Fig. 4A) and 1.25 μm (n = 7) and 0.55 μm (n = 8) for the cold shock response (Fig. 4B). Fig. 4. Open in new tabDownload slide Wind- and cold-shock-induced changes in the cytosolic and nuclear free Ca2+ concentrations and the expression levels of NpCaM-1 and NpCaM-2. A, Ca2+ changes in cytoplasm (cyt) and nucleoplasm (nuc) after stimulation with 5 mL of air at t = 10 s. B, Ca2+ changes in cytoplasm (cyt) and nucleoplasm (nuc) after stimulation with 1 mL of ice-cold water at t = 10 s. C, Wind- (■ and ▪) and cold-shock (○ and ●)-induced changes in mRNA levels of NpCaM-1 andNpCaM-2 are indicated and are averages of three experiments. Data are shown as hybridization relative to non-induced mRNA levels (given a value of 1) and is plotted against time in minutes. ○, NpCaM-1; ●, NpCaM-2; ■,NpCaM-1; ▪, NpCaM-2. Fig. 4. Open in new tabDownload slide Wind- and cold-shock-induced changes in the cytosolic and nuclear free Ca2+ concentrations and the expression levels of NpCaM-1 and NpCaM-2. A, Ca2+ changes in cytoplasm (cyt) and nucleoplasm (nuc) after stimulation with 5 mL of air at t = 10 s. B, Ca2+ changes in cytoplasm (cyt) and nucleoplasm (nuc) after stimulation with 1 mL of ice-cold water at t = 10 s. C, Wind- (■ and ▪) and cold-shock (○ and ●)-induced changes in mRNA levels of NpCaM-1 andNpCaM-2 are indicated and are averages of three experiments. Data are shown as hybridization relative to non-induced mRNA levels (given a value of 1) and is plotted against time in minutes. ○, NpCaM-1; ●, NpCaM-2; ■,NpCaM-1; ▪, NpCaM-2. The kinetics of the signals in the nucleus and cytoplasm differ in response to both stimuli. For wind stimulation, the average rise time (the time required to reach the peak) for MAQ 2.4 (cytoplasm) was 0.31 ± 0.04 and 0.60 ± 0.04 s for MAQ 7.11. For cold shock the average rise times for MAQ 2.4 and MAQ 7.11 were, respectively, 4.8 ± 0.3 and 9.0 ± 1.1 s. The MAQ 7.11 signals always peaked later than those in the cytoplasm, and the peak value was always lower. In more recent unpublished studies of ours using heat shock, signal-induced elevations of MAQ 2.4 and MAQ 7.11 were separated by minutes (M. Gong, A.H. van der Luit, and A.J. Trewavas, unpublished observations). This response, a much later and lower peak value found in MAQ 7.11 (compared with cytoplasmic MAQ 2.4), was similar to that recorded for [Ca2+]nuc in mammalian cells. The average length of the Ca2+ transient was similar in both compartments for wind stimulation, but was about 6 s longer in cold-shocked MAQ 7.11 compared with the cytoplasm. In separate experiments, tobacco seedlings were given wind signals (one treatment of 5 mL of air) or cold shock signals (one treatment of 1 mL of ice-cold water) similar to those used for Figure 4, A and B. RNA was extracted and the levels of NpCaM-1 and NpCaM-2mRNAs estimated from northern blots. These data (n = 3) are shown in Figure 4C. The total increase of NpCaM-1 mRNA after wind stimulation was about 5-fold after 60 to 90 min, whereas after cold shock it was about 10-fold after 90 to 120 min.NpCaM-2 exhibited only a slight increase throughout the experimental period. Use of Inhibitors on MAQ 2.4 and MAQ 7.11 Emphasize That Spatially Separate Ca2+ Pathways Can Regulate Calmodulin Gene Expression To try to deduce which Ca2+ compartment is used to regulate NpCaM-1 RNA concentrations, we treated seedlings with several inhibitors that modify [Ca2+]i kinetics. To establish suitable concentrations for use, we titrated the concentrations of these inhibitors to obtain an inhibition of about 50% or less in the Ca2+ signal. We then quantified the inhibitor-induced alterations in the [Ca2+]i kinetics and the alterations, if any, in NpCaM-1 andNpCaM-2 accumulation. MAQ 2.4 and MAQ 7.11 seedlings treated with thapsigargin, ruthenium red, or BAPTA-acetoxymethyl ester (AM) were subjected to wind signals (Fig. 5). There was a clear correlation between the behavior of the [Ca2+]i signals in the MAQ 7.11 compartment and NpCaM-1 RNA accumulation. Thapsigargin increased the MAQ 7.11 Ca2+ signal and subsequent calmodulin RNA accumulation, ruthenium red had no effect on the Ca2+ signal in MAQ 7.11 or the subsequent accumulation of calmodulin RNA, whereas BAPTA-AM decreased both. Ruthenium red did decrease the MAQ 2.4 signal but with no effect on NpCaM-1 accumulation. BAPTA-AM led to a slight decrease in the mean Ca2+ peak height in the MAQ 2.4 seedlings, but the difference was not significant, falling within these of the experiment. Treatment with the inhibitors alone had no detectable effect on either mRNA levels or cytosolic or nuclear Ca2+ (data not shown). Fig. 5. Open in new tabDownload slide The effect of Ca2+ modulators on wind-induced changes in cytosolic and nuclear Ca2+ and NpCaM-1 andNpCaM-2 mRNA accumulation. Wind stimulation was applied by 5 mL of air at t = 10 s. The se for the peak values from eight experiments is indicated for the mean peak. CON, Control; THAP, thapsigargin; RR, ruthenium red; BA, BAPTA-AM. A, The effect of 200 μm thapsigargin. ○, CONNpCaM-1; ●, CON NpCaM-2; □, THAPNpCaM-1; ▪, THAP NpCaM-2. B, 50 μm Ruthenium red. ○, CON NpCaM-1; ●, CON NpCaM-2; □, RR NpCaM-1; ▪, RRNpCaM-2.; C, 1 mm BAPTA-AM; solvents were used as control. ○, CON NpCaM-1; ●, CONNpCaM-2; □, BA NpCaM-1; ▪, BANpCaM-2. Fig. 5. Open in new tabDownload slide The effect of Ca2+ modulators on wind-induced changes in cytosolic and nuclear Ca2+ and NpCaM-1 andNpCaM-2 mRNA accumulation. Wind stimulation was applied by 5 mL of air at t = 10 s. The se for the peak values from eight experiments is indicated for the mean peak. CON, Control; THAP, thapsigargin; RR, ruthenium red; BA, BAPTA-AM. A, The effect of 200 μm thapsigargin. ○, CONNpCaM-1; ●, CON NpCaM-2; □, THAPNpCaM-1; ▪, THAP NpCaM-2. B, 50 μm Ruthenium red. ○, CON NpCaM-1; ●, CON NpCaM-2; □, RR NpCaM-1; ▪, RRNpCaM-2.; C, 1 mm BAPTA-AM; solvents were used as control. ○, CON NpCaM-1; ●, CONNpCaM-2; □, BA NpCaM-1; ▪, BANpCaM-2. MAQ 2.4 and MAQ 7.11 seedlings were treated with lanthanum and gadolinium chlorides and subjected to cold shock (Fig.6). With both inhibitors there was a substantial decrease in the MAQ 2.4 signal, which was associated with a severe inhibition of subsequent NpCaM-1 accumulation. The different behavior of the MAQ 7.11 seedlings, in which a slight increase in Ca2+ response was observed when the lanthanides were present, emphasizes a correlation between cold-shock-induced [Ca2+]i kinetics in MAQ 2.4 and NpCaM-1 expression. Treatment with the inhibitors alone had no detectable effect on mRNA levels or on cytosolic or nuclear Ca2+ (data not shown). Fig. 6. Open in new tabDownload slide The effect of Ca2+ modulators on cold shock-induced changes in cytosolic and nuclear Ca2+ andNpCaM-1 and NpCaM-2 mRNA accumulation. Cold shock stimulation was applied by a 1-mL injection of ice-cold water at t = 10 s. The se for the peak values from eight experiments is indicated for the mean peak. A, The effect of 10 mm LaCl3 (LA). ○, CONNpCaM-1; ●, CON NpCaM-2; ■, LANpCaM-1; ▪, LA NpCaM-2; B, 20 mm GdCl3 (GD); MgCl2 concentrations of identical ionic strength were used as a control (CON). ○, CONNpCaM-1; ●, CON NpCaM-2; ■, GDNpCaM-1; ▪, GD NpCaM-2. Fig. 6. Open in new tabDownload slide The effect of Ca2+ modulators on cold shock-induced changes in cytosolic and nuclear Ca2+ andNpCaM-1 and NpCaM-2 mRNA accumulation. Cold shock stimulation was applied by a 1-mL injection of ice-cold water at t = 10 s. The se for the peak values from eight experiments is indicated for the mean peak. A, The effect of 10 mm LaCl3 (LA). ○, CONNpCaM-1; ●, CON NpCaM-2; ■, LANpCaM-1; ▪, LA NpCaM-2; B, 20 mm GdCl3 (GD); MgCl2 concentrations of identical ionic strength were used as a control (CON). ○, CONNpCaM-1; ●, CON NpCaM-2; ■, GDNpCaM-1; ▪, GD NpCaM-2. DISCUSSION MAQ 7.11 Reports Changes in Nuclear Ca2+ We transformed tobacco seedlings with a nucleoplasmin aequorin construct to investigate further the apparent compartmentalization of wind and cold shock Ca2+ signals. For a variety of reasons, we believe that MAQ 7.11 seedlings report [Ca2+]nuc in response to wind and cold shock stimulation. Immunolocalization techniques indicated that 86% of the nucleoplasmin/aequorin fusion protein was found located in the nucleus of MAQ 7.11 cells. To be nuclear targeted, this oocyte polypeptide must be recognized by the nuclear import machinery of plants. Increasing evidence suggests that the mechanism of nuclear protein translocation is highly conserved among higher eukaryotes. About 9% of the aequorin in MAQ 7.11 was associated with chloroplasts. We have previously targeted aequorin to chloroplasts in tobacco (designated MAQ 6.3,Johnson et al., 1995). No changes in the chloroplastic Ca2+ levels were detected during mechanical and cold shock treatment of these seedlings (A.H. van der Luit, A. Haley, and A.J. Trewavas, unpublished observation). The low level of aequorin in the chloroplast therefore did not contribute to the measurements described here. Another 5% of the nucleoplasmin aequorin was found in the cytoplasm. The nucleoplasmin/aequorin construct is synthesized in the cytoplasm and then partitions to the nucleus. However, this residual cytoplasmic aequorin does not contribute significantly to the luminescence signal of MAQ 7.11. The Ca2+kinetics of the MAQ 7.11 are different from those of MAQ 2.4. Furthermore, there was no evidence of MAQ 7.11 kinetics of two components or two peaks, or even a broadening of the MAQ 7.11 peak, which might have resulted from a contaminating cytoplasmic signal. There was a clear difference in the kinetics of the Ca2+ response to cold shock between MAQ 2.4 and MAQ 7.11. This difference in kinetics was not due to fusion to nucleoplasmin, as aequorin in the cytoplasm and nucleoplasm reported identical Ca2+ values (Badminton et al., 1998). Wind signals induced [Ca2+]cyt (MAQ 2.4) to peak at 0.3 s, while the MAQ 7.11 peaked later at 0.6 s. The quick response of the nuclear and cytoplasmic signals to wind stimulation probably resulted in part from the speed with which the mechanical signal is perceived. Wind induced a slight movement of the seedling around the hypocotyl/root junction that lasted 0.02 to 0.3 s. In animal cells [Ca2+]nuc usually peaks later than [Ca2+]cyt, and the peak height is lower (Badminton et al., 1995, 1996, 1998). With cold shock stimulation, in which seedlings were irrigated with ice-cold water, MAQ 2.4 peaked at 4 to 5 s but MAQ 7.11 peaked at 9 s. Furthermore, MAQ 7.11 Ca2+ transients peaked at a substantially lower [Ca2+] than MAQ 2.4 (Fig. 4) in both cases. Unpublished evidence using MAQ 2.4 and MAQ 7.11 supports the apparent independence of the Ca2+ response in the different compartments. Heat shock treatments induce [Ca2+]cyt and [Ca2+]nuc transients, which are separated by minutes (M. Gong, A.H. van der Luit, and A.J. Trewavas, unpublished observations). While we could detect circadian variations in [Ca2+]cytin MAQ 2.4, we could not detect them in MAQ 7.11 (N.T. Wood, A. Haley, M. Moussaid, A.H. van der Luit, and A.J. Trewavas, unpublished data). There is therefore some autonomy in nuclear Ca2+signaling in plant cells, much as there seems to be in animal cells. There is an ongoing debate as to the extent to which the nucleus regulates [Ca2+]nuc(Carafoli et al., 1997; Malviya and Rogue, 1998). A common view is that alterations in [Ca2+]cytare the basic element in Ca2+ signaling and that they pass through the nuclear membrane, albeit in an attenuated and later form; in this case the nucleus is not thought to independently regulate [Ca2+]nuc. The alternative view regards the nuclear envelope and associated endoplasmic reticulum as an intracellular store of Ca2+ able to respond to signals independently of cytoplasmic changes. This latter view does not preclude parallel changes in [Ca2+]nuc and [Ca2+]cyt. Meyer et al. (1995) suggested that if Ca2+ signals in the cytoplasm and nucleus differ from each other in kinetics by at least 1 s, then the nuclear membrane is a substantial barrier to Ca2+ movement from the cytoplasm, greatly increasing the likelihood of separate regulation of nuclear Ca2+. In the case of cold shock at least, the nuclear membrane may act as a significant barrier to Ca2+ movement, because there is a 4-s difference between the peak values of MAQ 2.4 and MAQ 7.11. Distinct Ca2+ Signaling Pathways Regulate Calmodulin Gene Expression in Tobacco There is definite evidence that the flow of Ca2+ resulting from activation of different receptors regulates different pathways of gene expression (Bading et al., 1993; Finkbeiner and Greenberg, 1997), presumably through spatial separation of the pathways themselves. Hardingham et al. (1997)microinjected dextran-linked BAPTA into nuclei and concluded that some signals require a pathway through [Ca2+]cyt, while others involve [Ca2+]nuc. This technology is not yet currently feasible with cells in tobacco seedlings. In the experiments described in this paper for wind signals, some component of the signaling pathways controlling NpCaM-1expression could clearly be through [Ca2+]nuc. Prior treatment with BAPTA-AM inhibited the nuclear Ca2+ signal, leaving the cytosolic Ca2+ signal unaffected, ruthenium red greatly reduced the cytoplasmic signal without influencing that in the nucleus, while treatment with thapsigargin increased the subsequent nuclear signal without influencing the subsequent cytoplasmic signal. Variations in the accumulation of NpCaM-1 mRNA as a result of inhibitor treatments were correlated with [Ca2+]nuc but not with [Ca2+]cyt. Selective inhibition of the cold-shock-induced cytosolic Ca2+ signal by lanthanum and gadolinium chlorides, indicative of a cytosolic pathway for regulation ofNpCaM-1 calmodulin gene expression, helps confirm the spatial separation of signaling pathways between wind and cold shock stimuli. This apparent spatial distribution of signaling pathways may be further complicated by clear evidence that different downstream events are switched on at different stages of the Ca2+transient (Dolmetsch et al., 1997) and by a requirement that cytoplasmic signaling must take place near the plasma membrane. This latter observation of Finkbeiner and Greenberg (1997) might explain why reductions of about 40% in the cold-shock-dependent cytoplasmic Ca2+ signal nevertheless completely blocksNpCaM-1 mRNA accumulation. Based on previously reported effects of neomycin, we suspect that only part of the cold-shock-induced cytosolic signal originates with increased Ca2+ flux through the plasma membrane, with the remainder being released from internal stores by InsP3 (Knight et al., 1996). The reduction of 40% might then disguise a quantitatively greater inhibition of Ca2+ flux through the plasma membrane by the lanthanides, the cellular region critical perhaps to switching on the cytosolic pathway leading to NpCaM-1 transcription. By generating artificial Ca2+ transients,Dolmetsch et al. (1997) implicated early events in the rise time, peak value, and duration of the decay back to resting levels as controlling different transduction processes, including changes in gene expression. It is for this reason that we included measurements of rise time, peak Ca2+ values, decay times, and resting values where relevant for the data in Figures 4 to 6. However, in tobacco seedlings the kinetics of the Ca2+ transient seemed to be directly determined by the nature of the original signal. A wind signal induced a transient lasting some 20 s but reaching a peak within less than 0.5 s. Cold shock induced a transient lasting some 40 s and reaching a peak within 5 to 9 s. Both signals induced NpCaM-1 mRNA accumulation, although cold shock accumulations were higher than those of wind induction. Even when inhibitors are used, there is little alteration to the overall kinetics except in the peak height. There is a slight lengthening of about 5 s of the transient with thapsigargin; only more detailed studies directly modifying Ca2+ transients will determine whether this is a significant change. Certainly at present for theNpCaM-1 gene, peak height seems to be the more critical factor determining final mRNA accumulation. At present, two possible ways can be proposed in which [Ca2+]nuc exerts transcriptional regulation. The first may operate through Ca2+ or Ca2+-sensitive protein kinases located in the nucleus. As reported many years ago (Trewavas, 1979; Melanson and Trewavas, 1981), plant nuclei contain protein kinase activity and changes in specific phosphorylation of discrete nuclear proteins during cell development or cell division could be detected using two-dimensional electrophoretic separations. Clearly, plant nuclei have the potential for the regulation of transcription through phosphorylation, although whether there are Ca2+ or Ca2+/calmodulin-sensitive protein kinases in the plant nucleus remains to be established. The second possibility is that there is direct interaction of Ca2+/calmodulin with promoters or particular transcription factors. This mechanism is supported by recent work byCorneliussen et al. (1994), who reported binding of calmodulin to the basic helix-loop-helix domains of several mice basic helix-loop-helix transcription factors that inhibit their DNA binding in vitro, and with those of Szymanski et al. (1996), who reported that calmodulin isoforms enhance the binding of TGA3 to the Arabidopsis CaM-3promoter. The mechanism whereby wind signals can apparently selectively modify nuclear Ca2+ requires further investigation. Nuclei are surrounded by a basket of microfilaments. Distortion of these microfilamentous structures is thought to be one of the major means by which plant cells sense mechanical signals (Trewavas and Knight, 1994). In addition, Ca2+ channels localized to nuclei of amphibian epithelial cells (Prat and Cantiello, 1996) have been shown to be associated with actin filaments. Equivalent channels in tobacco cells might regulate nuclear Ca2+ levels in plant cells after wind stimulation. ACKNOWLEDGMENTS We would like to thank Dr. M. 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Akamatsu, Taku; Hanzawa, Yoshie; Ohtake, Yuhko; Takahashi, Taku; Nishitani, Kazuhiko; Komeda, Yoshibumi

doi: 10.1104/pp.121.3.715pmid: 10557219

Abstract A mutant of Arabidopsis with reduced internodal cell length, acaulis5 (acl5), has recently been shown to have reduced transcript levels of a gene for endoxyloglucan transferase, EXGT-A1 (Y. Hanzawa, T. Takahashi, Y. Komeda [1997] Plant J 12: 863–874). In the present study, we cloned genomic fragments of five members of theEXGT gene family, EXGT-A1,EXGT-A3, EXGT-A4, XTR2, and XTR3, and examined their expression in the wild type and in a series of acl mutants. In wild-type plants, theEXGT-A3 gene showed higher expression in lower internodes (internodes between nodes bearing axillary shoots) than in upper and young internodes, in which EXGT-A1 was highly expressed. EXGT-A4 was preferentially expressed in roots and XTR3 in siliques. The XTR2 gene was constitutively expressed. In acl1, acl3, and acl4 mutants, which have a severe defect in leaf expansion as well as in internode elongation, theEXGT-A1 gene showed reduced levels of expression before bolting of plants. In contrast, XTR3 was increased in these mutant seedlings. Reduction of EXGT-A1 expression was also detected after bolting of all acl mutants except acl2, whose growth defect is restricted to lower internodes. These results suggest the involvement of each EXGT in different aspects of organ development. The growth of plant cells depends on the balance between the turgor pressure and the extensibility of the cell wall. While the turgor pressure, which provides the driving force for cell extension, is influenced by the availability of water, the wall extensibility is to a large extent regulated by enzymes involved in the cleavage or formation of cross-links between cell wall polymers and in the turnover of certain wall components. In dicots, xyloglucan is a major structural polysaccharide of primary cell walls and is hydrogen-bonded to cellulose microfibrils to form cross-links between them (for review, see Hayashi, 1989; Carpita and Gibeaut, 1993). The cleavage and molecular grafting of xyloglucan polymers are catalyzed by endoxyloglucan transferase (EXGT) enzymes (also called xyloglucan endo-transglycosylase; XET). Therefore, EXGT has been suggested as one of the most likely agents responsible for wall loosening (for review, see Fry, 1995; Nishitani, 1995, 1997). Cloning of EXGT genes from several plant species has led us to realize that plants possess a large gene family of EXGTs. They have been classified into three subfamilies based on their sequence similarities (Nishitani, 1995, 1997; Xu et al., 1996). Subfamily I includes EXGT-V1 from azuki bean epicotyls, the first enzyme proved to mediate a transglycosylation reaction between xyloglucans (Nishitani and Tominaga, 1992). Subfamily II includes Arabidopsis meristem-expressed Meri5 (Medford et al., 1991) and mechanostimulus-inducible TCH4 (Xu et al., 1995), soybean brassinosteroid-inducible BRU1 (Zurek and Clouse, 1994), maize flooding-responsive WUSL1005 (Saab and Sachs, 1996), and tomato-fruit-expressed XET-B1 (Arrowsmith and de Silva, 1995). Germinating seed-specific NXG1 of nasturtium (de Silva et al., 1993) belongs to subfamily III and was originally identified as a hydrolase (xyloglucanase) (Edwards et al., 1986). Expression patterns of these genes are in good agreement with their proposed roles in cell wall modification during cell elongation (Nishitani, 1997), fruit ripening (Redgwell and Fry, 1993), vascular differentiation (Oh et al., 1998), and adaptive growth to physical stimuli (Antosiewicz et al., 1997). Dwarf phenotypes of the Arabidopsis brassinosteroid-responsive mutants have been shown to correlate with a reduced expression ofTCH4 (Kauschmann et al., 1996). It is still not known whether each member of the EXGT gene family within a single plant species plays a distinct and vital role in cell morphogenesis. We have isolated and studied Arabidopsis mutants with reduced internodal cell length, acaulis (acl), to determine the molecular basis of cell elongation in stem internodes. In rosette plants, including Arabidopsis, initiation of the internode elongation (bolting) follows flower bud formation. This process is probably mediated by phytohormones, but how their effects are exerted is not clear. Our previous study revealed that the acl5mutant, whose defect is sharply restricted to internodal growth, shows a reduced expression of the EXGT-A1 gene after flowering (Hanzawa et al., 1997). We report the cloning of genomic fragments of five members of the EXGT gene family and their expression patterns in the wild type and in a series of acl mutants, to which acl3 and acl4 have recently been added. MATERIALS AND METHODS Plant Material Arabidopsis ecotype Columbia was used in all experiments. Plants were grown on rock-wool bricks watered with Murashige and Skoog solution under continuous fluorescent light at 22°C. For RNA preparation from root tissue, seeds were surface-sterilized and sown on solidified Murashige and Skoog medium with 3% (w/v) Suc in Petri dishes. Petri dishes were kept under continuous fluorescent light at 22°C. The acl3-1 and acl4-1 mutants were selected in a screen for mutants with short internodes from M2 plants derived from fast-neutron-mutagenized seeds homozygous for gl1 (Lehle Seeds, Tucson, AZ). These were backcrossed five times into the wild-type Columbia (Col-0). Mapping was performed using molecular markers polymorphic between Columbia and Landsberg erecta(Konieczny and Ausubel, 1993; Bell and Ecker, 1994). Mutant alleles ofACL1 and ACL2 used in this study wereacl1-2 and acl2-1, respectively (Tsukaya et al., 1993). The acl5-1 allele in the Landsberg erectabackground (Hanzawa et al., 1997) was backcrossed at least five times into the Columbia background. Isolation of Genomic Clones Encoding EXGT Four cDNA clones with homology to EXGT-A1 (Okazawa et al., 1993) were previously isolated from an Arabidopsis cDNA library by screening at low stringency with the EXGT-A1 cDNA fragment, and were designated EXGT-A2, EXGT-A3,EXGT-A4, and EXGT-A5 (Nishitani, 1997; S. Okamoto and K. Nishitani, unpublished data). The nucleotide sequences ofEXGT-A2 and EXGT-A5 were found to be identical to those isolated and named XTR2 and XTR3, respectively, by Xu et al. (1996). An Arabidopsis genomic library constructed in λGEM12 was generously provided by J. Mulligan and R.W. Davis (Stanford University, Stanford, CA). The library was screened by plaque hybridization using a mixture of cDNA fragments as the probes. Subclones were prepared in pBluescript SK+ (Stratagene, La Jolla, CA) and sequenced using a Taq dye terminator cycle sequencing kit and a DNA sequencer (model 373A, Applied Biosystems, Foster City, CA). RNA Gel-Blot Analysis Total RNA was isolated from different tissues as described byTakahashi et al. (1992), separated by agarose/formaldehyde gel electrophoresis, and blotted onto nylon membranes (GeneScreen, New England Nuclear, Boston). Hybridization was performed at 42°C in 50% (w/v) formamide, 10% (w/v) dextran sulfate, 1% (w/v) SDS, 1m NaCl, 0.25 mg mL−1 salmon-sperm DNA, and the labeled gene-specific probe (see below). The filters were washed twice for 15 min at 65°C in 2× SSC, 1% (w/v) SDS and once at room temperature in 0.1× SSC. For all blots, equal loading was confirmed by ethidium bromide staining of ribosomal RNAs (25S, 18S). Probe Preparation To specifically detect each of the EXGT transcripts in the RNA gel-blot hybridization, 3′-end-specific probes were synthesized by PCR using cDNA clones as templates. The PCR primers were A1F (5′-GGCGGTTTAGAGAAGACCAA-3′), A1R (5′-GTAACTTATGCGTCTCTGTC-3′), A2F (5′-AAGCGTCTCAGGGTCTATGA-3′), A2R (5′-GTTCAT- AAAATGGAGGAAATC-3′), A3F (5′-CAGTTTCCGAGGT-GCG ATGA-3′), A3R (5′-GGCCAAATCTCACCCATACT3′), A4F (5′-TTGCACTGA CCGCGTCCG-3′), A4R (5′-CCAAACTTTTCTAGATTAAATTG-3′), A5F (5′-TAGCTAC-GAGAATTAATGTG-3′), and A5R (5′-AACCAACATAA-CT-CACGCCC-3′). The specificity of each probe was confirmed by DNA gel-blot analysis. No cross-hybridization was observed (data not shown). The PCR products were agarose gel purified and labeled by the random-primer protocol (BcaBest Labeling Kit, Takara, Kyoto). RESULTS Genomic Structure of EXGT Genes Five EXGT cDNA clones (Okazawa et al., 1993; Nishitani, 1997; S. Okamoto and K. Nishitani, unpublished data) were used as probes to screen an Arabidopsis genomic library in λGEM12. Sequence analysis of subcloned genomic DNA fragments revealed the presence of two or three introns whose placement within each of theEXGT coding regions is conserved (Fig.1A). The phylogenetic tree for these genes and those identified from other plant species is shown in Figure1B. Genomic DNA-blot analysis indicated that 3′-end-specific probes prepared from these EXGT genes (see “Materials and Methods”) hybridized to a single-copy gene at high-stringency conditions (data not shown). Fig. 1. Open in new tabDownload slide Comparison of EXGT genes. A, Genomic structure of the EXGT genes cloned in this study. Protein coding regions are shown by black boxes with the number of amino acid residues encoded by each exon. Numbers in parentheses indicate the number of nucleotides for intron. Intron splice sites in genomic sequences were deduced by comparison with their corresponding cDNA sequences, EXGT-A1 (Okazawa et al., 1993; accession no. D16454), XTR2 (Xu et al., 1996; accession no.U43487), EXGT-A3 (Nishitani, 1997; accession no.D63509), EXGT-A4 (Nishitani, 1997; accession no.AB026486), and XTR3 (Xu et al., 1996; accession no.U43485). The accession numbers for genomic sequences determined in this study are AF163819 (EXGT-A1), AF163820(XTR2), AF163821 (EXGT-A3), AF163822(EXGT-A4), and AF163823 (XTR3), respectively. B, Phylogenetic relationship between the Arabidopsis and other EXGT-related protein sequences. The entire deduced amino acid sequences were compared using the malign program of DNA Data Bank of Japan (Nishitani, 1997). References: a, Arrowsmith and de Silva (1995); b, Xu et al. (1995); c, Xu et al. (1996); d, Medford et al. (1991); e,Saab and Sachs (1995); f, Zurek and Clouse (1994); g, Nishitani (1997); h, Okazawa et al. (1993); i, Rose et al. (1996); and j, de Silva et al. (1993). Fig. 1. Open in new tabDownload slide Comparison of EXGT genes. A, Genomic structure of the EXGT genes cloned in this study. Protein coding regions are shown by black boxes with the number of amino acid residues encoded by each exon. Numbers in parentheses indicate the number of nucleotides for intron. Intron splice sites in genomic sequences were deduced by comparison with their corresponding cDNA sequences, EXGT-A1 (Okazawa et al., 1993; accession no. D16454), XTR2 (Xu et al., 1996; accession no.U43487), EXGT-A3 (Nishitani, 1997; accession no.D63509), EXGT-A4 (Nishitani, 1997; accession no.AB026486), and XTR3 (Xu et al., 1996; accession no.U43485). The accession numbers for genomic sequences determined in this study are AF163819 (EXGT-A1), AF163820(XTR2), AF163821 (EXGT-A3), AF163822(EXGT-A4), and AF163823 (XTR3), respectively. B, Phylogenetic relationship between the Arabidopsis and other EXGT-related protein sequences. The entire deduced amino acid sequences were compared using the malign program of DNA Data Bank of Japan (Nishitani, 1997). References: a, Arrowsmith and de Silva (1995); b, Xu et al. (1995); c, Xu et al. (1996); d, Medford et al. (1991); e,Saab and Sachs (1995); f, Zurek and Clouse (1994); g, Nishitani (1997); h, Okazawa et al. (1993); i, Rose et al. (1996); and j, de Silva et al. (1993). Developmental Regulation of EXGT Gene Expression Steady-state levels of EXGT transcripts were measured in different organs of adult flowering plants and in young seedlings before bolting. The results of RNA-blot hybridization using 3′-end-specific probes are shown in Figure2. The EXGT-A1 gene was highly expressed in 7-d-old seedlings and in the roots, upper internodes (internodes between nodes bearing flowers), flower buds, and green siliques of 30-d-old flowering plants. Transcript levels in fully expanded leaves and lower internodes (internodes between nodes bearing axillary shoots) were reduced, indicating the preferential expression of the EXGT-A1 gene in young, developing tissues. On the other hand, XTR2 showed a constitutive expression.EXGT-A3 showed a pattern similar to that of XTR2, but was higher in lower internodes. The EXGT-A4 gene was mainly expressed in roots. XTR3 was restricted to siliques and only weakly expressed in mature leaves. We further examined the expression of EXGT-A1 and EXGT-A3 genes during the internode elongation. RNA samples were prepared from upper and lower internodes at 5, 10, and 15 d after bolting, respectively. Our results revealed that, while the EXGT-A3 expression was increased as the day proceeded, the EXGT-A1 expression, especially in lower internodes, was drastically decreased (Fig.3). Fig. 2. Open in new tabDownload slide Analysis of the expression of EXGTgenes in different organs. Total RNA (10 μg per lane) was prepared from 7-d-old seedlings (lane 1), roots (lane 2), rosette leaves (lane 3), internodes between nodes bearing axillary shoots (lane 4), internodes between nodes bearing flowers (lane 5), flower buds (lane 6), and siliques (lane 7). Fig. 2. Open in new tabDownload slide Analysis of the expression of EXGTgenes in different organs. Total RNA (10 μg per lane) was prepared from 7-d-old seedlings (lane 1), roots (lane 2), rosette leaves (lane 3), internodes between nodes bearing axillary shoots (lane 4), internodes between nodes bearing flowers (lane 5), flower buds (lane 6), and siliques (lane 7). Fig. 3. Open in new tabDownload slide Analysis of the expression of EXGTgenes during internode elongation. Total RNA (10 μg per lane) was prepared from internodes between nodes bearing flowers (lanes 1, 3, and 5) and internodes between nodes bearing axillary shoots (lanes 2, 4, and 6). Tissues were harvested at 5 d (lanes 1 and 2), 10 d (lanes 3 and 4), and 15 d (lanes 5 and 6) after bolting. Fig. 3. Open in new tabDownload slide Analysis of the expression of EXGTgenes during internode elongation. Total RNA (10 μg per lane) was prepared from internodes between nodes bearing flowers (lanes 1, 3, and 5) and internodes between nodes bearing axillary shoots (lanes 2, 4, and 6). Tissues were harvested at 5 d (lanes 1 and 2), 10 d (lanes 3 and 4), and 15 d (lanes 5 and 6) after bolting. Identification of New acl Loci The acl mutants have been characterized by a defect in elongation growth of stem internodes after flowering, from which the name “acaulis” originates. In addition to the previously described mutants acl1, acl2, and acl5, two mutants derived from fast-neutron-mutagenized plants were found to represent new recessive loci by complementation tests and defined asacl3 and acl4, respectively (Fig.4). Mapping experiments revealed thatacl3 is tightly linked to the marker GL1 (Konieczny and Ausubel, 1993) on chromosome III and that acl4 is tightly linked to the marker SC5 on the lower arm of chromosome IV (data not shown). These two mutants have a severe defect in rosette leaf expansion before flowering and are phenotypically indistinguishable from the allele of acl1-2 (Fig.5A). Fig. 4. Open in new tabDownload slide Morphology of adult flowering plants withacl mutations. Plants were grown at 22°C under continuous light for 40 d. A, acl1-2; B,acl2-1; C, acl3-1; D,acl4-1; E, acl5-1. Scale bars = 1 cm. Fig. 4. Open in new tabDownload slide Morphology of adult flowering plants withacl mutations. Plants were grown at 22°C under continuous light for 40 d. A, acl1-2; B,acl2-1; C, acl3-1; D,acl4-1; E, acl5-1. Scale bars = 1 cm. Fig. 5. Open in new tabDownload slide Morphology of 10-d-old wild-type andacl seedlings. Plants were grown under continuous light at 22°C (A) or 28°C (B). Fig. 5. Open in new tabDownload slide Morphology of 10-d-old wild-type andacl seedlings. Plants were grown under continuous light at 22°C (A) or 28°C (B). We found that, like the phenotype of acl1 (Tsukaya et al., 1993), the phenotype of acl3 and acl4 could not be rescued by the exogenous addition of phytohormones, but was drastically supressed by elevated growth temperature (28°C; Fig. 5B). On the other hand, acl2 and acl5 mutants were nearly wild-type in appearance before bolting and their defect was only detected in the growth of stem internodes (Fig. 4). In contrast toacl1, acl3, and acl4 mutants, whose internodal growth was markedly restored at 28°C, acl2 andacl5 mutants showed no restoration of the internodal growth at 28°C. The reduction in leaf expansion and/or stem elongation in all of these acl mutants is primarily due to the reduction in cell size (Tsukaya et al., 1993; Hanzawa et al., 1997; data not shown). EXGT Gene Expression in acl Mutants The effect of acl mutations on the expression ofEXGT genes was examined by RNA blots. Figure6A shows that the EXGT-A1expression was reduced in aerial portions of 7-d-old seedlings ofacl1, acl3, and acl4 mutants with the leaf phenotype. Interestingly, these three mutant seedlings exhibited elevated levels of the XTR3 transcript. Reduced expression of EXGT-A1 was also observed in acl5 mutants after flowering, as well as in acl1, acl3, andacl4 flowering plants (Fig. 6B). The transcript levels ofXTR3 in 30-d-old flowering plants, which seems mainly attributable to the expression in siliques (Fig. 2), and those in rosette leaves of flowering plants were unaffected by theseacl mutations (Fig. 6B; data not shown). There were no obvious influences of acl mutations on the transcript levels of XTR2 and EXGT-A3 in aerial tissues (Fig. 6, A and B) or those of EXGT-A1 and EXGT-A4 in roots (Fig. 6C). Fig. 6. Open in new tabDownload slide Analysis of the expression of EXGTgenes in acl mutants. Total RNA (10 μg per lane) was prepared from aerial tissues of 7-d-old seedlings (A and D) and 30-d-old flowering plants (B) and from root tissues of 7-d-old seedlings (C). Plants were grown at 22°C (A–C) or at the indicated temperature (D). Lanes W, Wild type; lanes 1, acl1-2; lanes 2, acl2-1; lanes 3, acl3-1; lanes 4, acl4-1; lanes 5, acl5-1. Fig. 6. Open in new tabDownload slide Analysis of the expression of EXGTgenes in acl mutants. Total RNA (10 μg per lane) was prepared from aerial tissues of 7-d-old seedlings (A and D) and 30-d-old flowering plants (B) and from root tissues of 7-d-old seedlings (C). Plants were grown at 22°C (A–C) or at the indicated temperature (D). Lanes W, Wild type; lanes 1, acl1-2; lanes 2, acl2-1; lanes 3, acl3-1; lanes 4, acl4-1; lanes 5, acl5-1. We further found that the transcript levels of EXGT-A1 inacl1, acl3, and acl4 seedlings grown at 22°C were restored by the growth at 28°C, in parallel with their morphological phenotypes (Fig. 6D). An elevated level ofEXGT-A1 expression was also seen in wild-type seedlings grown at 28°C, in which leaf expansion and petiole elongation were also enhanced (Fig. 5). DISCUSSION One of our major interests was to identify actual molecules involved in the rapid cell growth of stem internodes in Arabidopsis. Previously, we observed that the acaulis5 (acl5) mutant showed a marked reduction of the EXGT-A1 gene expression after flowering, as well as a severely reduced length of stem internodes (Hanzawa et al., 1997). To evaluate the relationship between the expression of EXGT genes and plant cell growth, we extended our analysis to the expression of other members of theEXGT gene family in the wild type and in a series ofacl mutants. This study revealed that the members of the EXGT gene family are under the differential control of expression during development of wild-type plants. Expression of EXGT-A3 appeared to be high in lower (old) internodes, in contrast to that of EXGT-A1 in upper (young) internodes. According to the phylogenic tree established from related protein sequences (Fig. 1B; Nishitani, 1997), EXGT-A1 and root-expressed EXGT-A4 belong to subfamily I, while XTR2 and EXGT-A3 belong to subfamily III. In nasturtium, NXG1 (subfamily III) and XET1 (subfamily I) exhibit mutually exclusive patterns of gene expression and possess different substrate specificities (Rose et al., 1996). NXG1 has been suggested to act predominantly as a hydrolytic enzyme in the mobilization of xyloglucan seed storage reserves in germinating seed cotyledons (Edwards et al., 1986). If hydrolytic action toward xyloglucans is a major role of members of subfamily III, then EXGT-A3, together with XTR2, could be required for the regulated degradation of xyloglucan networks for the maturation and/or maintenance of the fine structure of cell walls, which follows the elongation growth. It will be necessary to determine whether these EXGTs possess different enzyme activities against different xyloglucan substrates and whether they exhibit cell-type-specific patterns of expression. The significance of EXGT-A1 in cell elongation was strengthened by our analysis of the expression in acl mutants. Two loci,acl3 and acl4, were newly identified in this study. The phenotypes of these two mutants could not be restored by exogenously applied phytohormones (data not shown), suggesting that neither of these mutations represent genes involved in hormone biosynthesis. Based on their phenotypes, which are almost identical to the acl1 phenotype, we suggest that these three gene products act in a common regulatory pathway of cell elongation. Our results showed that the defects of acl1,acl3, and acl4 in leaf expansion and in stem elongation are accompanied by the reduced expression of theEXGT-A1 gene. When grown at 28°C, these mutants restore both the phenotype and the transcript level of EXGT-A1. The high temperature also enhances both petiole elongation andEXGT-A1 expression in wild-type seedlings. Xu et al. (1996)have reported that the EXGT-A1 gene (referred to asEXT) is up-regulated in response to touch, auxin, and darkness, all of which can facilitate elongation growth. It is possible that EXGT-A1 functions in the process of cell elongation in young leaves and stem internodes. Further genetic approaches, including the isolation of knockout mutants of this gene and the creation of transgenic plants with altered levels of expression are required to define the exact role. Moreover, it remains to be clarified whether the reduction in cell length, which can be caused by mutations in a vast variety of genes, is generally associated with reduced expression ofEXGT-A1. The possibility cannot be ruled out thatEXGT-A1 expression is changed as a consequence of altered cell morphology. There were no detectable alterations in EXGT-A1 expression in semidominant acl2 mutants. This can be explained by the limited defect of acl2 within the internode elongation between nodes bearing axillary shoots (Tsukaya et al., 1995), which might be accompanied by a temporal and slight reduction inEXGT-A1 expression. However, it is also likely that theacl2 mutation has a negative effect on other molecules involved in cell elongation, while having no influence on EXGT-A1. Preferential expression of the XTR3 gene in wild-type siliques is consistent with the fact that the corresponding cDNAs have been identified as expressed-sequence-tag clones derived from dry seeds by Xu et al. (1996). XTR3, as well as stress-responsive Meri5 and TCH4 (Xu et al., 1996), belongs to subfamily II. We found that, in contrast to EXGT-A1, the XTR3 transcript levels were elevated in acl1, acl3, and acl4seedlings. Such opposite effects on EXGT genes may reflect the complexity of environmental and hormonal regulation of theEXGT gene expression (Xu et al., 1996). Cloning of theACL genes is currently in progress and will help to answer the question of how acl mutations affect regulatory pathways of EXGT gene expression. In summary, our data on the expression of EXGT genes (especially on their responsiveness to environmental stimuli), which are supported by data reported by others, support the possibility that many kinds of mutations can affect the regulatory pathways ofEXGT gene expression, resulting in altered cell morphology. The molecular processes underlying the cell wall architecture consist of various biochemical steps, indicating the involvement of many enzymes other than EXGT. It should be noted that there is increasing evidence suggesting the importance of expansins (Cosgrove, 1998) and endo-1,4-β-glucanases (Shani et al., 1997; Nicol et al., 1998) in plant cell growth. Expansins have been identified as a catalyst for acid growth and have been shown to induce the extension of isolated cell walls (McQueen-Mason and Cosgrove, 1995). ACKNOWLEDGMENTS We are grateful to Drs. John Mulligan and Ronald W. Davis (Stanford University, Stanford, CA) for the gift of the Arabidopsis genomic library. We also thank Dr. Shigehisa Okamoto (Kagoshima University, Kagoshima, Japan) for help with the cloning ofEXGT genes. LITERATURE CITED 1 Antosiewicz DM Purugganan MM Polisensky DH Braam J Cellular localization of Arabidopsis xyloglucan endotransglycosylase-related proteins during development and after wind stimulation. Plant Physiol 115 1997 1319 1328 Google Scholar Crossref Search ADS PubMed WorldCat 2 Arrowsmith DA de Silva J Characterisation of two tomato fruit-expressed cDNAs encoding xyloglucan endo-transglycosylase. Plant Mol Biol 28 1995 391 403 Google Scholar Crossref Search ADS PubMed WorldCat 3 Bell CJ Ecker JR Assignment of 30 microsatellite loci to the linkage map of Arabidopsis. Genomics 19 1994 137 144 Google Scholar Crossref Search ADS PubMed WorldCat 4 Carpita NC Gibeaut DM Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. 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Plant J 9 1996 879 889 Google Scholar Crossref Search ADS PubMed WorldCat 29 Xu W Purugganan MM Polisensky DH Antosiewicz DM Fry SC Braam J Arabidopsis TCH4, regulated by hormones and the environment, encodes a xyloglucan endotransglycosylase. Plant Cell 7 1995 1555 1567 Google Scholar PubMed OpenURL Placeholder Text WorldCat 30 Zurek DM Clouse SD Molecular cloning and characterization of a brassinosteroid-regulated gene from elongationg soybean epicotyls. Plant Physiol 104 1994 161 170 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was partially supported by a Grant-In-Aid from the Ministry of Education, Science and Culture of Japan and by a grant for the Research for the Future Program from the Japan Society for the Promotion of Science (JSPSRFTF96L00403). * Corresponding author; e-mail [email protected]; fax 81–11–706–2739. Copyright © 1999 American Society of Plant Physiologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Carranco, Raúl; Almoguera, Concepción; Jordano, Juan

doi: 10.1104/pp.121.3.723pmid: 10557220

Abstract Chimeric constructs containing the promoter and upstream sequences of Ha hsp17.6 G1, a small heat shock protein gene, reproduced in transgenic tobacco (Nicotiana tabacum) its unique seed-specific expression patterns previously reported in sunflower. These constructs did not respond to heat shock, but were expressed without exogenous stress during late zygotic embryogenesis coincident with seed desiccation. Site-directed mutagenesis of its distal and imperfect heat shock element strongly impaired in vitro heat shock transcription factor binding and transgene expression in seeds. Deletion analyses of upstream sequences indicated the contribution of additional cis-acting elements with either positive or negative effects on transgene expression. These results show differences in the transcriptional activation through the heat shock element of small heat shock protein gene promoters in seeds compared with the heat shock response. In addition, they suggest that heat shock transcription factors and other distinct trans-acting factors cooperate in the regulation of Ha hsp17.6 G1 during seed desiccation. Plant heat shock genes are not only expressed in response to heat stress, but also during zygotic embryogenesis and in other developmental stages in the absence of exogenous stress (for review, see Hightower and Nover, 1991; Schöffl et al., 1998). The regulation of heat shock gene expression during embryogenesis has been investigated for the class I small heat shock protein (sHSP) gene family that encodes cytoplasmic proteins (Waters, 1995). Studies of class I sHSP promoters showed that heat shock elements (HSEs), the cis-acting elements necessary for the heat shock response, were also involved in their regulation during zygotic embryogenesis (Coca et al., 1996; Prändl et al., 1995). Synthetic HSEs could even confer developmental regulation in plant seeds to a minimal cauliflower mosaic virus 35S promoter (Prändl and Schöffl, 1996). Site-directed mutagenesis of the sunflower Ha hsp17.7 G4promoter determined that HSEs are required for its developmental regulation, although only during the desiccation stages characteristic of late embryogenesis. This observation demonstrated the seed regulation of Ha hsp 17.7 G4 by both HSE-dependent and -independent transcriptional activation mechanisms (Almoguera et al., 1998). That work also showed that the heat response of chimeric constructs containing the Ha hsp17.7 G4 promoter and 5′-flanking sequences was abolished by point mutations that only partially affected their expression in embryos. This suggested possible differences in the HSE-mediated activation mechanism of the same sHSP promoter in response to heat stress or during development (for discussion, see Almoguera et al., 1998). The effect of the Arabidopsis abi3 mutants on sHSP gene expression in seeds might indicate additional, although more indirect, evidence for such differences. The sHSPs expressed in seeds during embryogenesis did not accumulate to detectable amounts in the null mutant abi3-6, but the same mutation did not affect expression of these proteins in response to heat shock (Wehmeyer et al., 1996). The ABI3 gene encodes a transcription factor that regulates various seed-specific genes (Giraudat et al., 1992; Parcy et al., 1995). Thus, a possible inference from this observation would be that ABI3, together with heat shock factors (HSFs), are involved in transcriptional activation of at least some sHSP promoters in seeds. Such involvement would imply mechanisms that differ from the heat shock response. We also have isolated and initially characterized in sunflower the mRNA accumulation patterns and seed-specific transcriptional activation of a peculiar plant sHSP gene, Ha hsp17.6 G1. The Ha hsp17.6 G1 promoter is, to our knowledge, the sole example for a heat-stress-non-responsive member of the plant class I sHSP gene family. The presence of an imperfect HSE in the 5′-flanking region ofHa hsp17.6 G1 posed an interesting interpretation dilemma. If that HSE were not functional, the promoter should be activated by mechanisms not involving HSFs. Alternatively, in the case of a functional HSE, the transcriptional activation of the Ha hsp17.6 G1 promoter would require HSFs. In that case, activation should mechanistically differ from a typical heat shock response (for discussion, see Carranco et al., 1997). In the present work we found the answer to this dilemma by analyzing the expression effects of site-directed mutagenesis of the imperfect HSE. The HSE is indeed functional and is required for seed expression of Ha hsp17.6 G1. Additional deletion analyses of the 5′-flanking sequences identified other cis-acting elements with positive or negative effects on the promoter. These observations further define models for sHSP gene regulation during plant zygotic embryogenesis. MATERIALS AND METHODS Site Directed Mutagenesis of the Ha hsp17.6 G1 Promoter We used a megaprimer PCR procedure, essentially as described byAlmoguera et al. (1998), but with the following modifications. The megaprimer was a 206-bp DNA fragment that included the Ha hsp17.6 G1 sequences between −109 and +49 (all positions given from the transcription initiation site [Carranco et al., 1997]), followed by the pBluescript SK sequences between HindIII (in the vector polylinker) and the SK (5′-TCTAGAACTAGTGGATC-3′) primer. The megaprimer was amplified after 30 cycles using an annealing temperature of 48°C and the SK and G1 mutagenic primers. The G1 mutagenic primer was: 5′-GTCCAtATAAGTACAtAATATTTCAtAACACTACTACG- 3′, corresponding to the Ha hsp17.6 G1 sequences (coding strand) between −109 and −72, with lowercase letters indicating the three nucleotide substitutions. The megaprimer and the KS (5′-CGAGGTCGACGGTATCG-3′) primer were used to amplify another 242-bp DNA fragment with the Ha hsp17.6 G1 sequences between theHindIII sites at −126 and +49. The second PCR was for 30 cycles with annealing at 52°C. The 175-bp HindIII DNA fragment, including the mutations, was used to construct −1,486(m)::GUS (see below). Electrophoretic Mobility Shift Assays The conditions for DNA probe labeling and for binding and mobility shift assays in agarose gels using recombinant hHSF1 were essentially as described by Carranco et al. (1997). Binding reactions differed only in the amount of poly [dI.dC] used (4 μg/reaction) and in the presence of variable amounts of bacterial extracts containing hHSF1 (from 2–5 μg protein/reaction). The DNA probes were 175-bpHindIII DNA fragments that contained the wild-type and mutant Ha hsp17.6 G1 sequences between −126 and +49. Ha hsp17.6 G1::GUS Chimeric Constructs and Generation of Transgenic Plants We constructed four Ha hsp17.6 G1::GUS chimeric translational fusions between position +121 of Ha hsp17.6 G1 and the SmaI site in the polylinker of pBI 101.2. The Ha hsp17.6 G1 junction sequence was in all cases an end-filled (with Klenow polymerase) StyI site. The chimeric constructs −1,486::GUS, −533::GUS, and −126::GUS, respectively contained 5′-upstream sequences to the EcoRI (−1,486), XhoI (−533), andHindIII (−126) sites present in Ha hsp17.6 G1(Carranco et al., 1997). Each chimeric Ha hsp17.6 G1::GUS::nos cassette also contained different synthetic sequences placed immediately upstream of the Ha hsp17.6 G1 sequences, and derived from the pBluescript SK polylinker coming from intermediate plasmids (details available upon request). To obtain −1,486(m)::GUS, the wild-type Ha hsp17.6 G1 sequences between the HindIII sites at −126 and +49 were deleted and replaced by the mutant sequences in the 242-bp,HindIII digested PCR fragment (see above). The nucleotide sequence at the Ha hsp17.6 G1::GUS junction for all chimeric constructs, as well as the sequence and orientation of the PCR amplified fragment in −1,486(m)::GUS, was verified by dideoxy sequencing using the GUSIII primer. All DNA manipulations were carried out using previously described standard procedures (Coca et al., 1996; Sambrook et al., 1989). The four Ha hsp17.6 G1::GUS translational fusions constructed in pBI 101 (see Fig. 2) were mobilized into transgenic tobacco (Nicotianum tabacum) with Agrobacterium tumefaciens using the standard leaf disc method of transformation (Horsch et al., 1985). A total of at least 10 independent primary transformants for each chimeric construct was obtained (actual numbers of analyzed plants indicated in the legends of Figs. 3-6). These plants were studied after their selection by Southern and PCR analysis (Coca et al., 1996; Almoguera et al., 1998). Such techniques showed the presence of an average of one to three copies of stable-integrated intact transgenes at different integration sites (data not shown). Heat Stress Treatments Transgenic and non-transgenic tobacco plants were subjected to control and heat shock treatments after clonal duplication of the individual plants, as previously described (Coca et al., 1996). For each original plant, three segregants were used in these experiments (see Fig. 5). Stem samples (a piece of approximately 5-cm length per clone) were collected from 4 cm below the apical meristem. Leaf samples included (per each clone) a complete leaf (without the petiole) removed from 5 cm below the apical meristem. For the assays with whole seedlings, we used the segregating progeny of the original transgenic plants (a pool of approximately 100 kanamycin-resistant seedlings per plant) and similar numbers of non-transgenic seedlings. The thermal stress treatments were also as described by Coca et al. (1996). GUS Assays and Statistical Analysis of Data Transgenic tobacco plants were produced and characterized for developmental and heat-induced GUS expression. GUS activity in seedling, leaf, stem, pollen, seed, and embryo samples from the transgenic tobacco plants was histochemically and/or fluorometrically assayed. The statistical distributions of values for plants transgenic for each chimeric construct were compared by analysis of variance (ANOVA) after logarithmic transformation of data. For a detailed description of these procedures, see Almoguera et al. (1998) and references therein. RESULTS Mutagenesis of the HSE in the Ha hsp17.6 G1 Promoter: Effect on in Vitro HSF Binding We previously demonstrated that the Ha hsp17.6 G1 gene was not transcriptionally active in response to heat shock in sunflower, very likely because of the characteristics of its only distal and imperfect HSE (Carranco et al., 1997). However, this HSE might be still involved in the developmental regulation of Ha hsp17.6 G1 during late embryogenesis. We investigated this possibility by a mutagenesis approach analogous to that described forHa hsp17.7 G4, another sunflower sHSP gene with a more complex HSE structure and expression pattern (Coca et al., 1996;Almoguera et al., 1998). The HSE sequences in the Ha hsp17.6 G1 promoter (Carranco et al., 1997; Fig.1) were altered by introducing three nucleotide substitutions (G–T) at a crucial position within the GAA core repeat (see “Materials and Methods” for details). These mutations were designed to severely impair binding of HSFs and subsequent promoter activation, as previously demonstrated for plant sHSP genes by Barros et al. (1992) and Almoguera et al. (1998). Fig. 1. Open in new tabDownload slide Electrophoretic mobility shift assays withHa hsp17.6 G1 probes. Top, Binding of recombinant hHSF1 to 175-bp end-labeled DNA fragments containing the wild-type (HSEwt, left lanes) or mutant (HSEm, right lanes) HSE. Binding reactions contained either no (−) or increasing amounts of hHSF1 (from left to right, 2, 4, or 5 μg, as indicated by the filled triangles for each probe). The arrow points to the specific hHSF::DNA complexes mentioned in the text. Bottom, Nucleotide sequences of the wild-type and mutant HSE. Dots on the sequence indicate agreement with GAA and TTC consensus core repeats. The two perfect (full consensus matching) core repeats are underlined. Mutations are indicated in lowercase. All sequence positions are given from the transcription initiation site of Ha hsp17.6 G1. Fig. 1. Open in new tabDownload slide Electrophoretic mobility shift assays withHa hsp17.6 G1 probes. Top, Binding of recombinant hHSF1 to 175-bp end-labeled DNA fragments containing the wild-type (HSEwt, left lanes) or mutant (HSEm, right lanes) HSE. Binding reactions contained either no (−) or increasing amounts of hHSF1 (from left to right, 2, 4, or 5 μg, as indicated by the filled triangles for each probe). The arrow points to the specific hHSF::DNA complexes mentioned in the text. Bottom, Nucleotide sequences of the wild-type and mutant HSE. Dots on the sequence indicate agreement with GAA and TTC consensus core repeats. The two perfect (full consensus matching) core repeats are underlined. Mutations are indicated in lowercase. All sequence positions are given from the transcription initiation site of Ha hsp17.6 G1. Before performing functional analyses in transgenic plants, we verified in vitro the effect of these mutations. We analyzed binding of recombinant human HSF1 (hHSF1) to fragments of the Ha hsp17.6 G1 promoter containing the HSE. The results of electrophoretic mobility shift assays are summarized in Figure 1. Using a fragment with unaltered HSE sequences, we detected the previously described specific hHSF1-DNA complex (Fig. 1; Carranco et al., 1997). This complex was observed with various amounts of hHSF1, and it could be abolished only by competition with DNA sequences containing the Ha hsp17.6 G1 or more perfect HSEs (Carranco et al., 1997; data not shown). In contrast, using the fragment containing the mutant HSE, the same complex could not be detected, even with the highest amount of hHSF1 (Fig. 1). The faint, higher mobility bands observed with the mutant fragment upon long autoradiograph exposures likely represent minor nonspecific complexes, as they could be abolished by competition with excess nonspecific DNA fragments (data not shown). Expression Patterns of the Full-Length Ha hsp17.6 G1::GUS Construct in Transgenic Tobacco Histochemical GUS assays in embryos dissected at different developmental stages from the −1,486::GUS transgenic plants (see Fig. 2) determined that in transgenic tobacco, the Ha hsp17.6 G1 promoter directs the expression of this chimeric construct from 24 DPA (data not shown). The highest expression level was reached at 28 DPA (Fig.3). A similar expression pattern was also observed in the seed endosperm (Fig. 3). These expression patterns essentially match the Ha hsp17.6 G1 mRNA accumulation patterns coincident with late seed desiccation that were previously reported in sunflower (Carranco et al., 1997). The tobacco heterologous system thus at least reproduced the temporal expression patterns during zygotic embryogenesis of two different sunflower sHSP genes, Ha hsp 17.6 G1 (Fig. 3) and Ha hsp17.7 G4 (Coca et al., 1996; Almoguera et al., 1998). Fig. 2. Open in new tabDownload slide Maps of the Ha hsp17.6 G1::GUS chimeric constructs. The four translational fusions contain identical 5′-untranslated and coding sequences, as well as different upstream sequences from Ha hsp17.6 G1 (both represented as gray boxes). The 5′-flanking ends denoted by numbers were also used for construct names: −1,486::GUS, −1,486(m)::GUS, −533::GUS, and −126::GUS. Numbers indicate the position from the Ha hsp17.6 G1 transcription initiation site (depicted by arrows). The wild-type (HSEwt) or mutant (HSEm) HSE are indicated by small black boxes in each gene. Reference restriction sites are EcoRI (E), XhoI (X), and HindIII (H). Fig. 2. Open in new tabDownload slide Maps of the Ha hsp17.6 G1::GUS chimeric constructs. The four translational fusions contain identical 5′-untranslated and coding sequences, as well as different upstream sequences from Ha hsp17.6 G1 (both represented as gray boxes). The 5′-flanking ends denoted by numbers were also used for construct names: −1,486::GUS, −1,486(m)::GUS, −533::GUS, and −126::GUS. Numbers indicate the position from the Ha hsp17.6 G1 transcription initiation site (depicted by arrows). The wild-type (HSEwt) or mutant (HSEm) HSE are indicated by small black boxes in each gene. Reference restriction sites are EcoRI (E), XhoI (X), and HindIII (H). Fig. 3. Open in new tabDownload slide Histochemical localization of GUS activity in seeds and pollen of the −1,486::GUS transgenic plants. Left, Developmental stages at top correspond to either dissected embryos (top) or endosperm (bottom). Right, Pollen grains from −1,486::GUS (A) or non-transformed tobacco plants (B). A total of 12 different −1,486::GUS plants were analyzed. For seeds, samples were dissected from at least two different capsules per individual plant. Representative results are shown in each case. Histochemical reactions were for 15 h at 28°C. Scale bars correspond to either 300 μm (seed) or 40 μm (pollen). Fig. 3. Open in new tabDownload slide Histochemical localization of GUS activity in seeds and pollen of the −1,486::GUS transgenic plants. Left, Developmental stages at top correspond to either dissected embryos (top) or endosperm (bottom). Right, Pollen grains from −1,486::GUS (A) or non-transformed tobacco plants (B). A total of 12 different −1,486::GUS plants were analyzed. For seeds, samples were dissected from at least two different capsules per individual plant. Representative results are shown in each case. Histochemical reactions were for 15 h at 28°C. Scale bars correspond to either 300 μm (seed) or 40 μm (pollen). Similar analyses of other tissues and organs from the −1,486::GUS transgenic plants cultivated under controlled conditions did not detect the expression of the transgene, with the exception of mature pollen grains (Figs. 3A and 6). This result was not an artifact of our GUS assay conditions (i.e. detection of similar endogenous enzyme activities in tobacco), as demonstrated by negative results using pollen from untransformed tobacco (Fig. 3B). A significant expression in pollen grains, observed both for G1::GUS (Fig. 3A) and G4::GUS (Coca et al., 1996) chimeric constructs, might reflect a natural activation of the corresponding sunflower promoters in this tissue or an ectopic expression in tobacco (for discussion, see Coca et al., 1996; for review, see Mascarenhas et al., 1996). We further analyzed the pollen expression of the G1::GUS constructs to investigate the specificity of the effect of mutations and deletions in Ha hsp17.6 G1 regulatory sequences. These experiments, which were also used to further define the expression patterns of these genes in seeds, were carried out with more sensitive fluorometric assays using larger numbers of transgenic plants (see below). Requirement of the Imperfect HSE for Efficient Seed Expression: Effects of Upstream Sequence Deletion We investigated the possible involvement of the HSE in the developmental regulation of Ha hsp17.6 G1 by comparing the expression patterns of the −1,486::GUS and −1,486(m)::GUS chimeric constructs in seeds of transgenic tobacco. The −1,486(m)::GUS construct differs only from −1,486::GUS in three point mutations at crucial positions within the HSE (Fig. 1). These mutations were incorporated in the context of a full-length promoter fusion that reproduced the developmental regulation of Ha hsp 17.6 G1 in transgenic tobacco (Figs. 2 and 3). Fluorometric assays of GUS activity showed expression of −1,486::GUS in seeds from 20 to 28 DPA (Fig.4). These assays also detected low expression at 16 DPA, but at levels not significantly distinct from those of non-transformed plants (F = 3.399,P = 0.071). These levels, an average of 31.13 ± 23.9 pmol methylumbelliferone (MU) mg−1 protein min−1, were undetectable by histochemical assays (Fig. 3). Expression from the −1,486(m)::GUS gene was significantly reduced at 24 and 28 DPA (F = 9.97,P = 0.002 and F = 18.79,P = 0.001, respectively), although it was unaffected at 20 DPA (F = 0.48, P = 0.49). Histochemical GUS assays with dissected embryos and endosperm from the −1,486(m)::GUS plants did not detect GUS expression in samples from 16 to 28 DPA, confirming the more sensitive fluorometric assays (data not shown; Fig. 4A). These results revealed that the integrity of the HSE in the Ha hsp17.6 G1 promoter is required for its developmental regulation during late zygotic embryogenesis. Fig. 4. Open in new tabDownload slide Fluorometric quantification of GUS activity during seed maturation in transgenic plants for the different Ha hsp17.6 G1::GUS chimeric constructs. A, Comparison between the expression patterns of the wild-type (▪; −1,486::GUS) and mutant (■; −1,486(m)::GUS) constructs. B, Effect of the −533::GUS () and −126::GUS () deletions. GUS assays were performed with protein extracts prepared from whole seeds at each developmental stage. Activities are given in pmol 4-MU mg− 1 protein min− 1. Individual GUS assays were performed in duplicate. The following numbers of primary transformants were analyzed per chimeric construct: −1,486::GUS, 12 plants; −1,486(m)::GUS, 13 plants; −533::GUS, 10 plants; and −126::GUS, 12 plants. Mean values and ses bars are represented. C, Summary of sequences functionally defined in this work by either mutation (HSE, solid black box) or deletion analyses (hatched and dotted boxes). We indicate the observed positive (+) and negative (−) effects on the Ha hsp17.6 G1 promoter. Other symbols as in Figure 2. Fig. 4. Open in new tabDownload slide Fluorometric quantification of GUS activity during seed maturation in transgenic plants for the different Ha hsp17.6 G1::GUS chimeric constructs. A, Comparison between the expression patterns of the wild-type (▪; −1,486::GUS) and mutant (■; −1,486(m)::GUS) constructs. B, Effect of the −533::GUS () and −126::GUS () deletions. GUS assays were performed with protein extracts prepared from whole seeds at each developmental stage. Activities are given in pmol 4-MU mg− 1 protein min− 1. Individual GUS assays were performed in duplicate. The following numbers of primary transformants were analyzed per chimeric construct: −1,486::GUS, 12 plants; −1,486(m)::GUS, 13 plants; −533::GUS, 10 plants; and −126::GUS, 12 plants. Mean values and ses bars are represented. C, Summary of sequences functionally defined in this work by either mutation (HSE, solid black box) or deletion analyses (hatched and dotted boxes). We indicate the observed positive (+) and negative (−) effects on the Ha hsp17.6 G1 promoter. Other symbols as in Figure 2. The functional involvement of distal Ha hsp17.6 G15′-flanking upstream sequences was investigated by analyzing the effects of deletions with chimeric constructs −533::GUS and −126::GUS. Both promoter constructs contain the intact HSE and various lengths of Ha hsp17.6 G1 upstream sequences (Fig. 2). The expression patterns in seeds of transgenic plants for −533::GUS or −126::GUS were compared with those of −1,486::GUS plants (Fig. 4B). Deletion of upstream sequences to −126 significantly affected GUS expression in seeds at 24 DPA (F = 4.51, P = 0.037) and 28 DPA (F = 9.32, P = 0.003), but not at 20 DPA (F = 0.04, P = 0.84). Thus, the effects of either the HSE mutation or this deletion were similar, both resulting in poor reporter gene expression levels during seed desiccation (see above; Fig. 4A). These results demonstrated that the HSE by itself is not sufficient to activate the Ha hsp17.6 G1 promoter from 24 DPA. In contrast, deletion of upstream sequences to −533 in chimeric construct −533::GUS resulted in substantially higher levels (5.6- to 25.8-fold) of reporter gene expression at 20 (F = 38.99, P = 0.0001), 24 (F = 23.36,P = 0.001), and 28 (F = 13.45,P = 0.004) DPA, compared with −1,486::GUS (Fig. 4B). In summary, we determined that distinct Ha hsp17.6 G1 upstream sequences contain cis-acting elements involved in the temporal and quantitative regulation of seed expression during embryogenesis (Fig. 4C). Lack of Heat Shock Response of G1::GUS Chimeric Constructs in Transgenic Tobacco Previously reported gene-specific RNase A protection and nuclear run-on assays determined that in sunflower the Ha hsp17.6 G1mRNAs do not accumulate in response to heat shock, mainly because the promoter is transcriptionally inactive under heat stress, at least in seedlings (Carranco et al., 1997). We tested the heat shock response of the different G1::GUS chimeric constructs in transgenic tobacco. This heterologous system has been successfully used to reproduce the heat stress response of a different sHSP sunflower promoter, Ha hsp17.7 G4, which also contains imperfect, although more complex, proximal and distal HSEs (Coca et al., 1996). In experiments performed with whole plants or seedlings containing the −1,486::GUS construct, fluorometric assays revealed only insignificant levels of GUS activity under control or heat shock treatments. That activity was similar in magnitude to that in non-transgenic plants. Similar results were obtained in different organs and developmental stages after imbibition (Fig.5). Fig. 5. Open in new tabDownload slide Absence of GUS activity in vegetative tissues of transgenic plants. Data in this figure correspond to the progeny of a subset of the original transgenic plants analyzed in Figures 3 and 4. These plants showed similar seed expression patterns as their parents (data not shown). We analyzed protein extracts from stems and leaves of adult plants (2 months post imbibition, top) or from whole seedlings at earlier developmental stages (20 d post imbibition, bottom). Samples from non-transgenic tobacco (NT) were used as a reference for basal levels of GUS activity. Plants containing a chimeric construct with theHa hsp17.7 G4 sequences between −1,132 and +163 (G4,Coca et al., 1996) were used as a positive control for heat induction in the stem samples. The following numbers of plants were used (for denominations see also Fig. 2): −1,486, −1,486(m), −533, and −126, five plants each; G4, three plants; and NT, six plants. Values for control (white bars), and heat shock induced (black bars) GUS activities are represented as indicated in the legend of Figure 4. Fig. 5. Open in new tabDownload slide Absence of GUS activity in vegetative tissues of transgenic plants. Data in this figure correspond to the progeny of a subset of the original transgenic plants analyzed in Figures 3 and 4. These plants showed similar seed expression patterns as their parents (data not shown). We analyzed protein extracts from stems and leaves of adult plants (2 months post imbibition, top) or from whole seedlings at earlier developmental stages (20 d post imbibition, bottom). Samples from non-transgenic tobacco (NT) were used as a reference for basal levels of GUS activity. Plants containing a chimeric construct with theHa hsp17.7 G4 sequences between −1,132 and +163 (G4,Coca et al., 1996) were used as a positive control for heat induction in the stem samples. The following numbers of plants were used (for denominations see also Fig. 2): −1,486, −1,486(m), −533, and −126, five plants each; G4, three plants; and NT, six plants. Values for control (white bars), and heat shock induced (black bars) GUS activities are represented as indicated in the legend of Figure 4. The same results were observed with transgenic plants containing the −1,486(m), −533, or −126 chimeric constructs. Furthermore, these experiments showed that in vegetative tissues the −533 and −126 5′-deletions did not have any effect on either the basal or the heat-induced GUS activities. As previously observed (Coca et al., 1996), we were able to detect heat-shock-induced GUS activity from another chimeric construct with Ha hsp17.7 G4 sequences, in stems of transgenic tobacco plants (Fig. 5, G4). However, in seedlings, we observed the heat-induced accumulation of the chimeric Ha hsp17.7 G4::GUS mRNA (data not shown). The latter is consistent with the reported accumulation of the Ha hsp17.7 G4 mRNA in sunflower seedlings (Carranco et al., 1997). In contrast, theHa hsp17.6 G1 mRNAs did not accumulate in response to heat shock in either sunflower seedlings or different adult organs under various stress conditions (Carranco et al., 1997). Thus, the results in Figure 5 agree with the lack of heat-shock-induced transcriptional activation of the Ha hsp17.6 G1 promoter in seedlings and with the absence of heat-induced Ha hsp17.6 G1 mRNA accumulation in other organs (Carranco et al., 1997). Distal Sequences of Ha hsp17.6 G1 Promoter Show Different Specificity in Seeds and Pollen The results in Figure 5 also suggested that the effects of the tested HSE mutation and 5′-flanking deletions were seed specific. For example, in the −533 deletion, we did not observe increased levels of GUS activity in different organs of corresponding transgenic plants. The specificity of the deletion and mutation effects was further verified by fluorometric quantification of GUS activities in pollen of the −1,486::GUS, −1,486(m)::GUS, −533::GUS, and −126::GUS plants (Fig.6). The average GUS activity in pollen of the −1,486::GUS plants was 1,361 ± 296 pmol MU mg−1 min−1. Compared with this value, only that for the −126::GUS plants was significantly reduced (62.8 ± 13.1 pmol MU mg−1 min−1,F = 5.77, P = 0.02). This result confirmed the seed specificity of the negative and positive effects, respectively, observed for the HSE mutation and the −533 deletion. In contrast to results with the other chimeric constructs, the clear effect of the −126 deletion in pollen indicated that sequences between −126 and −533 contain positive cis-acting elements that might function not only in seeds but also in pollen (compare Figs. 4 and 6). Fig. 6. Open in new tabDownload slide Quantification of GUS activity in pollen grains. GUS assays were performed with protein extracts prepared from pollen from the following numbers of independent transgenic plants for each chimeric construct: −1,486, 11; −1,486(m), 13; −533, 14; and −126, 11. GUS activities are represented as indicated in the legend of Figure4. For chimeric construct denomination see Figure 2. Fig. 6. Open in new tabDownload slide Quantification of GUS activity in pollen grains. GUS assays were performed with protein extracts prepared from pollen from the following numbers of independent transgenic plants for each chimeric construct: −1,486, 11; −1,486(m), 13; −533, 14; and −126, 11. GUS activities are represented as indicated in the legend of Figure4. For chimeric construct denomination see Figure 2. DISCUSSION The HSE in the Ha hsp17.6 G1 Promoter Is Required for Developmental Regulation in Seeds The Ha hsp17.6 G1 promoter contains a HSE, which, compared with those in other plant sHSP genes (including two sunflower promoters), has unique structural characteristics (Carranco et al., 1997). This element could be considered as a relic resembling similar regulatory sequences found in constitutive HSP genes from other families (i.e. HSP70; for review, see Gurley and Key, 1991). Among the structural characteristics of this HSE are its relative distal position from the TATA box and the presence of only five HSE core motifs, only two of which are perfect. The perfect core motifs are adjacent to imperfect ones (Fig. 1). The second core repeat is the most imperfect and it could either be regarded as a five-nucleotide gap (Carranco et al., 1997) or as a very low homology core repeat with only one conserved position in the TTC sequence (Fig. 1). The Ha hsp 17.6 G1 HSE lacks other more proximal HSE core motifs present, for example, in other promoters as Ha hsp17.7 G4 (Carranco et al., 1997). Despite this structure, previous results indicated that the Ha hsp17.6 G1 HSE could be a functional regulatory element. Thus, we showed that in vitro it was able to bind recombinant hHSF1, although with lower affinity than the more extended, complex, and perfect HSEs of Ha hsp17.7 G4(Carranco et al., 1997; see also Fig. 1). By introducing three very specific nucleotide substitutions in the HSE of Ha hsp17.6 G1 (Fig. 1; Barros et al., 1992; Almoguera et al., 1998), we have been able now to abolish the in vitro binding of hHSF1 (Fig. 1). The same mutations drastically impaired expression from the Ha hsp17.6 G1 promoter in desiccating seeds (Fig. 4A). These results demonstrate the necessity of the HSE for the regulation of Ha hsp17.6 G1 during late embryogenesis. The peculiar architecture of the HSE in Ha hsp17.6 G1, and perhaps of other imperfect HSEs as those in Ha hsp17.7 G4(Coca et al., 1996), might contribute to differences in an HSF-mediated transcription activation mechanism. The HSE in Ha hsp17.6 G1has a distal location compared with more proximal elements located in other plant sHSP genes, including different sunflower promoters (Gurley and Key, 1991; Carranco et al., 1997). The fact that this HSE is functional during embryogenesis (Fig. 4A) and its failure to support heat shock induction (Carranco et al., 1997; Fig. 5) might reflect differences in the effect of distance from the HSE to the initiation site. Heat shock induction in vegetative tissues appears to be more dependent on the presence of more proximal HSEs in sunflower (Carranco et al., 1997; Almoguera et al., 1998) and other plant sHSP promoters (Gurley and Key, 1991; Marrs and Sinibaldi, 1997, and refs. therein). In contrast, distal HSEs are required for (this work, Fig. 4A) or substantially contribute to (Almoguera et al., 1998) developmental regulation. Another interesting possibility is that, as observed in yeast, the imperfect structure of HSEs could influence the conformation of DNA-bound HSF(s) and subsequent promoter activation (Santoro et al., 1998). Our observations support models explaining the activation in seeds of the Ha hsp17.6 G1 promoter with participation of HSF(s). As previously proposed for other sHSP gene promoters, such HSF(s) would have a crucial role in promoter activation (Prändl et al., 1995;Coca et al., 1996; Prändl and Schöffl, 1996; Almoguera et al., 1998). In the case of Ha hsp17.6 G1, the involved HSFs might differ in their sequence specificity from those involved in the heat shock response of other plant sHSP genes (previously discussed by Carranco et al. [1997]). However, based on the results reported here (Figs. 2-6), we propose that the promoter context (the structure of HSE and its functional interaction with other cis-elements) is perhaps the most crucial factor for promoter activation by HSFs during zygotic embryogenesis, at least for Ha hsp17.6 G1. Additional cis-Acting Elements Contribute to the Developmental Regulation of Ha hsp17.6 G1 The effects of 5′-flanking sequence deletions (Figs. 2 and 4B) indicated the existence of other cis-acting elements different from the HSE and located upstream of it. These elements have either positive or negative quantitative effects that modulate the seed-specific expression of the Ha hsp17.6 G1 promoter (summarized in Fig.4C). The HSE, although necessary for temporal and quantitative developmental regulation, is not sufficient for full induction of theHa hsp17.6 G1 promoter (see results for −126::GUS in Fig. 4B). We observed a synergism for promoter activation between the HSE and other positive cis-elements located between −126 and −533 (Fig. 4). This might indicate direct or indirect functional interaction(s) among proteins binding to these cis-elements. We propose that the distal cis-acting elements and unidentified trans-acting factors that interact with them cooperate with HSFs in the developmental regulation of this promoter. A conceivable scenario for such hypothetical interaction is that the HSFs could reach only limiting concentrations in developing embryos. The interaction of such HSFs with the Ha hsp17.6 G1 promoter could be facilitated by accessory, seed-specific factors that would bind to more distal promoter sequences. This accessory factor(s) could also facilitate other crucial interactions, as cooperative interactions between distally bound HSFs and TFIID at the TATAA sequence. In vegetative tissues the HSE would be too imperfect and distal to support heat induction of the promoter in absence of the seed-specific accessory factors. The trans-acting factor(s) with negative effects on Ha hsp17.6 G1 promoter activation would balance the activity of those with positive effects on the same promoter. At the desiccation stages of embryogenesis, the action of positive factor(s) and cooperation with HSFs would be dominant. Earlier in embryogenesis, the negative factor(s) would repress the promoter. In other sHSP promoters efficiently expressed before desiccation (e.g. Ha hsp 17.7 G4; Coca et al., 1996; Almoguera et al., 1998), the negative factor(s) would not bind the promoter, or additional factors would compensate for their effects and allow promoter activation at these stages. Our hypothesis could be extended to other plant sHSP promoters and help to explain the paradox of their differential transcriptional activation during embryogenesis (for discussion, see Carranco et al., 1997) despite the presence of functional HSEs (Carranco et al., 1997; this work, Fig. 4A). Crucial aspects of this hypothesis are that in addition to the HSE, other distinct cis-acting elements are also required, and that both work in concert in promoter activation. Because not all sHSP promoters are active during embryogenesis, the HSEs would not be sufficient for developmental regulation in the context of a natural promoter. However, out of context, even a multimerized HSE (20 copies of synthetic core sequences) was shown to activate transcription from a minimal cauliflower mosaic virus 35S promoter in seeds (Prändl and Schöffl, 1996). However, 5′-deletions of the Gm hsp17.3B promoter in its natural context revealed that the truncation to −237 position was not active in developing seeds, nor was it heat inducible in leaves, despite the presence of nine perfect HSE core repeats (Prändl and Schöffl, 1996). These results agree with our observations of the requirement, but insufficiency, of HSE for the developmental regulation of Ha hsp 17.6 G1. Whereas additional cis-elements necessary for the developmental regulation of Gm hsp17.3B might include other distal HSE core repeats (Prändl and Schöffl, 1996), in Ha hsp17.6 G1 the distal sequences do not include HSEs (Carranco et al., 1997; Figs. 2 and 4). Inferences from the previously discussed observations with plant sHSP gene promoters would be also comparable to the regulation of the yeast HSP82 promoter during early meiotic induction (Szent-Gyorgyi, 1995). HSEs are also required for promoter activation and are even able to confer meiotic induction to a different promoter. However, not all yeast HSP promoters are meiotically induced, and this induction requires functional interaction between proteins binding the HSEs and an upstream repression sequence (URS1; Szent-Gyorgyi, 1995). The activation of the Ha hsp 17.6 G1 promoter during embryogenesis differs, however, from the meiotic induction of HSP82. Regulation of Ha hsp17.6 G1, mediated by sequences between −533 and −1,486, would be seed specific, as these sequences are not involved in negative regulation in pollen or vegetative tissues (Figs.5 and 6). In contrast, URS1 functions in yeast as both a vegetative repressor and a meiotic coactivator (Szent-Gyorgyi, 1995). This work did no attempt to directly identify the trans-acting factors involved in the regulation of Ha hsp17.6 G1 promoter. However, the characterization of the HSE as an imperfect but functional cis-acting element and a preliminary delimitation of other positive and negative cis-acting elements allowed us to further define models of developmental regulation of plant sHSP genes. Our results will also help the eventual isolation and characterization of these unknown factors. ACKNOWLEDGMENTS We thank Drs. Eduardo Santero and Sebastián Chávez (Department of Genetics, University of Sevilla) for their comments and critical reading of this manuscript. LITERATURE CITED 1 Almoguera C Prieto-Dapena P Jordano J Dual regulation of a heat shock promoter during embryogenesis: stage-dependent role of heat shock elements. Plant J 13 1998 437 446 Google Scholar Crossref Search ADS PubMed WorldCat 2 Barros MD Czarnecka E Gurley WB Mutational analysis of a plant heat shock element. 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Biochemistry 30 1991 1 12 Google Scholar Crossref Search ADS PubMed WorldCat 7 Hightower L Nover L Heat Shock and Development. 1991 Springer-Verlag Berlin 8 Horsch RB Fry JE Hoffmann NL Eichholtz D Rogers SG Fraley RT A simple and general method for transferring genes into plants. Science 227 1985 1229 1231 Google Scholar PubMed OpenURL Placeholder Text WorldCat 9 Marrs KA Sinibaldi RM Deletion analysis of the maize hsp82, hsp81, and hsp17.9 promoters in maize and transgenic tobacco: contributions of individual heat shock elements and recognition by distinct protein factors during both heat shock and development. Maydica 42 1997 211 226 Google Scholar OpenURL Placeholder Text WorldCat 10 Mascarenhas JP Crone DF Pollen and the heat shock response. 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Plant Mol Biol 31 1996 157 162 Google Scholar Crossref Search ADS PubMed WorldCat 14 Sambrook J Fritsch EF Maniatis T Molecular Cloning: A Laboratory Manual. 1989 Cold Spring Harbor Laboratory Press Cold Spring Harbor, NY 15 Santoro N Johansson N Thiele DJ Heat shock element architecture is an important determinant in the temperature and transactivation domain requirements for heat shock transcription factor. Mol Cell Biol 18 1998 6340 6352 Google Scholar Crossref Search ADS PubMed WorldCat 16 Schöffl F Prändl R Reindl A Regulation of the heat-shock response. Plant Physiol 117 1998 1135 1141 Google Scholar Crossref Search ADS PubMed WorldCat 17 Szent-Gyorgyi C A bipartite operator interacts with a heat shock element to mediate early meiotic induction of Saccharomyces cerevisiae HSP82. Mol Cell Biol 15 1995 6754 6769 Google Scholar Crossref Search ADS PubMed WorldCat 18 Waters E The molecular evolution of the small heat shock proteins in plants. Genetics 141 1995 785 795 Google Scholar Crossref Search ADS PubMed WorldCat 19 Wehmeyer N Hernandez LD Finkelstein RR Vierling E Synthesis of small heat-shock proteins is part of the developmental program of late seed maturation. Plant Physiol 112 1996 747 757 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was funded by the Spanish Comisión Interministerial de Ciencia y Tecnologı́a (grant no. BIO96–0474) and by Junta de Andalucı́a (grant no. CVI148). * Corresponding author; e-mail [email protected]; fax 34–95–4–62–40–02. Copyright © 1999 American Society of Plant Physiologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Shu, Guoping; Pontieri, Vincenza; Dengler, Nancy G.; Mets, Laurens J.

doi: 10.1104/pp.121.3.731pmid: 10557221

Abstract InFlaveria trinervia (Asteraceae) seedlings, light-induced signals are required for differentiation of cotyledon bundle sheath cells and mesophyll cells and for cell-type-specific expression of Rubisco small subunit genes (bundle sheath cell specific) and the genes that encode pyruvate orthophosphate dikinase and phosphoenolpyruvate carboxylase (mesophyll cell specific). Both cell type differentiation and cell-type-specific gene expression were complete by d 7 in light-grown seedlings, but were arrested beyond d 4 in dark-grown seedlings. Our results contrast with those found for another C4 dicot, Amaranthus hypochondriacus, in which light was not required for either process. The differences between the two C4 dicot species in cotyledon cell differentiation may arise from differences in embryonic and post-embryonic cotyledon development. Our results illustrate that a common C4 photosynthetic mechanism can be established through different developmental pathways in different species, and provide evidence for independent evolutionary origins of C4 photosynthetic mechanisms within dicotyledonous plants. In contrast to C3 photosynthesis, in which the whole process of primary CO2 assimilation takes place autonomously within a single cell, C4CO2 assimilation requires metabolic cooperation between bundle sheath cells (BSC) and mesophyll cells (MC) (Edwards and Walker, 1983; Anderson and Deardall, 1991; Suzuki et al., 1999). CO2 is first fixed by phosphoenolpyruvate carboxylase (PEPCase; EC 4.1.1.31) in MC. The C4 acid formed diffuses to BSC, where it is decarboxylated and the CO2 released is refixed by Rubisco (EC 4.1.1.39). The substrate for PEPCase in the initial CO2 fixation, PEP, is regenerated by pyruvate phosphate dikinase (PPDK, EC 2.7.9.1) in MC (Hatch 1987, 1997; Furbank and Taylor, 1995). The foundation of this metabolic cooperation is the cell-type-specific compartmentation of photosynthetic enzymes and expression of the corresponding enzyme-coding genes. Cell type specificity of the proteins and their mRNAs has been demonstrated in several monocot and dicot C4species (Bauwe, 1984; Martineau and Taylor, 1985; Sheen and Bogorad, 1985, 1986; Langdale et al., 1988a, 1988b; Hudson et al., 1992; Wang et al., 1992; Dengler et al., 1995; Ramsperger et al., 1996; Shu, 1996;Drincovich et al., 1998). The genetic mechanisms that control the cell-type- and tissue-specific expression of genes encoding C4 enzymes have been studied with both molecular and transgenic approaches in C4 species (Berry et al., 1985, 1986; Martineau et al., 1989; Sheen, 1991; Matsuoka et al., 1993; Chitty et al., 1994; Bilang and Bogorad, 1996; Furbank et al., 1997; Marshall et al., 1997; Stockhaus et al., 1997; Taylor et al., 1997; Westhoff et al., 1997; Mann, 1999). Light is one of the most important environmental signals regulating leaf development in plants, including leaf cellular differentiation and photosynthetic gene expression (Tobin and Silverthorne, 1985; Nelson and Langdale, 1992; Fankhauser and Chory, 1997). Light perception and the various light signal transduction pathways that regulate leaf development are currently under intensive study in C3 plants (Fankhauser and Chory, 1997). In foliage leaves of the C4 plant maize, light has been shown to play important roles in C4 cell type differentiation, direct activation of C4photosynthetic genes at the transcriptional level, and establishment of cell type specificity of C4 gene expression (Nelson et al., 1984; Sheen and Bogorad, 1987a, 1987b; Langdale, et al., 1988a; Maroco et al., 1998). However, the role of light in cell type differentiation and cell-type-specific gene expression in C4 dicot foliage leaves is largely unknown. Previous studies have investigated the pattern of cell-type-specific gene expression in developing foliage leaves of several C4 dicots (Wang et al., 1992, 1993b; Dengler et al., 1995; Ramsperger et al., 1996; Berry et al., 1997), but the role of light in these processes was not directly addressed. An important experimental approach in studying the light regulation of plant development is to compare changes in developmental and gene expression patterns between dark- and light-grown foliage organs or whole plants. In the C4 grass maize, several immature, primordial foliage leaves are present in the embryo and expand under dark-grown conditions. Thus, the role of light in regulating C4 gene expression and cell type differentiation can be assessed by comparing developing light- and dark-grown leaves (Nelson et al., 1984; Sheen and Bogorad, 1985, 1986,1987a, 1987b; Langdale et al., 1988a, 1988b). However, in most dicot species, foliage leaves will not initiate or expand in dark-grown seedlings, so the comparative ontogenetic analysis used in C4 grasses cannot be applied (Dengler et al., 1997). For this reason, studies of the role of light in the development of photosynthetic tissues in C3 dicots have often exploited post-germination cotyledons, which can develop in both dark- and light-grown conditions and also have simpler developmental patterns than dicot foliage leaves (Tobin and Silverthorne, 1985; Tsukaya et al., 1994; Kretsch et al., 1995; Fankhauser and Chory, 1997). In cotyledons of the dicot C4 plantAmaranthus hypochondriacs (Berry et al., 1985, 1986; Wang et al., 1993a), Wang et al. (1993a) found that light is not required for either C4 cell type differentiation or cell-type-specific gene expression. This finding contrasts with the results from studies of the foliage leaves of the C4 grass maize, in which light is required for differentiation between BSC and MC (Sheen and Bogorad, 1985; Langdale et al., 1988b). It is not clear whether the discrepancy found between maize foliage leaves and amaranth cotyledons reflects differences between foliage leaves and cotyledons, between monocot leaves and dicot leaves in general, or simply species-specific variation in developmental patterns. More comparative studies of cotyledon development among different C4 dicot species are needed to address these questions. In this study, we assessed the role of light in the regulation of cell type differentiation and cell-type-specific expression of C4 photosynthetic genes in cotyledons of a dicot C4 plant, Flaveria trinervia. We examined cotyledon anatomical development and temporal and spatial mRNA accumulation of three C4 photosynthetic genes that encode the small subunit of Rubisco (RbcS), PPDK, and PEPCase in cotyledons of both dark- and light-grown seedlings of F. trinervia from d 0 to d 12 after sowing. We found that light is essential for full morphological differentiation of cotyledon BSC and MC and for cell-type-specific expression of C4genes. Cell-type-specific gene expression was established between d 4 and d 7 after germination, coincident with maturation of Kranz anatomy in cotyledons. The light dependence of bundle sheath and mesophyll maturation in F. trinervia is more similar to that of maize than to that of amaranth. We suspect that the difference in light responsiveness and in patterns of C4 gene expression between the two C4 dicots F. trinervia and amaranth reflect species-specific differences in embryonic and post-embryonic cotyledon development. MATERIALS AND METHODS Plant Material Seeds were harvested from a self-pollinated Flaveria trinervia plant. Seeds for dark- and light-grown seedlings were soaked in deionized, distilled water for 24 h at 24°C in either regular transparent (light-grown d 0) or light-proof glass beakers (dark-grown d 0). The imbibed seeds were sown in pots with commercial potting soil (PRO-MIX, Premier Horticulture, Red Hill, PA), and covered with 5 mm of vermiculite. The light-grown seedlings were grown in a growth room with 14 h of light/d (approximately 180 μE m−2 s−1) at 24°C. Twelve pots for dark-grown seedlings were prepared and sealed in an air-fluent dark box and kept in a dark room at 24°C. One pot was randomly sampled each day (every 24 h) after sowing; the pot was opened and the seedlings were fixed in complete darkness. Tissue Preparation Whole seedlings were fixed in 4% (w/v) paraformaldehyde for 24 h at 4°C and stored at 4°C in 70% (v/v) ethanol. The cotyledons for in situ hybridization were embedded in paraffin (Paraplast, Oxford Labware, St. Louis) and 6-μm sections were mounted on poly-d-Lys (Sigma, St. Louis)-coated slides. Sections from seedlings of different developmental stages were placed side by side on the same slides for hybridization with the same in situ hybridization probes. Cotyledons used for anatomical observations were dehydrated in an ethanol and acetone series, embedded in Spurr's resin, and sectioned at 2 μm with a diamond knife on an ultramicrotome (MT-7, RMC, Tucson, AZ). The sections were mounted on poly-d-Lys-coated slides and stained in 0.5% (w/v) toluidine blue O in 0.1% (w/v) sodium carbonate (O'Brien and McCully, 1981). Sections were photographed using a microscope (Polyvar, Reichert, Vienna, Austria) and Tmax film (Eastman-Kodak, Rochester, NJ). mRNA in Situ Hybridization The DNA templates used for generating sense and antisense RNA probes were as follows: RbcS was from a 400-bp cDNA fragment that covers exons II and III of the RbcS-R1 gene from Flaveria ramosissima; it was cloned in pBlueScript II SK(+) (Shu, 1996), Ppc was a 1.82-kb cDNA fragment from the 3′ region of the gene (3 kb) that encodes a C4 isoform of PEPCase fromF. trinervia (Hermans and Westhoff, 1992) Pdk was from a 1.3-kb cDNA fragment of the 5′ region of the gene (3.1 kb from F. trinervia (Hermans and Westhoff, 1992; Rosche and Westhoff, 1995). Both Pdk and Ppc cDNA clones were kind gifts from Dr. Peter Westhoff. Digoxigenin-labeled sense and antisense RNA probes were generated by in vitro transcription using T3 and T7 RNA polymerase (Boehringer Mannheim, Basel). Probe hydrolysis followed Langdale et al. (1988a). Slide pretreatment, prehybridizaton, and hybridization were modified from Langdale et al. (1988a) and Wang et al. (1992). Proteinase K treatment was 20 μg/mL for 20 min at 37°C, and RNase A treatment was 5 μg/mL for 20 min at 37°C. The prehybridization and hybridization solution (1,000 μL) contained 125 μL of 10× in situ salts (3.0 m NaCl, 0.1 m Tris, pH 6.8, 0.1m sodium phosphate, pH 6.8, and 50 mm EDTA), 500 μL of deionized formamide, 250 μL of 50% (w/v) dextran sulfate, 25 μL of 20 μg/mL tRNA, 60 μL of 5 μg/mL poly(A+), and 40 μL of distilled, deionized water. Hybridization was at 50°C overnight. The highest washing stringency was 0.25× SSC at 42°C for 30 min. Immunodetection and colorimetric reaction followed protocols from Boehringer Mannheim. The color substrates nitroblue tetrazolium and X-phosphate were used. Slides were photographed using dark-field microscopy (Labophoto, Nikon, Tokyo) on Ektachrome 64T or 160T (Kodak). RESULTS Seedling Morphology, Cotyledon Development, and Cell Type Differentiation In the light (see “Materials and Methods” for growth conditions used), the radicle of a F. trinervia seedling emerged from the seed coat by d 3 (48 h after sowing in soil). Two cotyledons emerged from the soil surface by d 4 (after 72 h). Newly emerged cotyledons were 2 mm long and yellowish opaque. Greening was first observed on the upper or adaxial side of the cotyledons and then on the lower or abaxial side by d 5 (after 96 h). The cotyledons expanded to 4 mm long by d 7 (after 144 h) and remained green through d 25 before undergoing senescence. The first foliage leaf was visible by d 10 and became fully expanded by d 15. When the seeds were germinated in the dark, radicle protrusion was delayed by 12 h. The 4-d-old seedling had an elongated hypocotyl and yellowish-white cotyledons about 1.5 mm long that were partly enclosed in the seed coat. Beyond d 4, the hypocotyl remained elongated and hooked, and cotyledon expansion was arrested. The whole seedling started to shrink by d 9 and died by d 18. Foliage leaves did not expand in dark-grown seedlings. Figure 1, A to C, shows cotyledon anatomical development in light-grown seedlings. Kranz anatomy was present in a rudimentary form in the cotyledons of d 0 seedlings (after 24 h of soaking the seeds in distilled, deionized water) (Fig.1A). Cotyledon vasculature was still forming at this stage, but veins already present had a well-defined bundle sheath layer. The mesophyll was five layers thick and strongly dorsiventral, with a well-defined adaxial palisade layer (layer 1). BSC and MC had numerous storage bodies and lacked differentiated plastids (Fig. 1A); proplastids were present but lacked well-developed thylakoids (V. Pontieri and N.G. Dengler, unpublished results). After 4 d in the light, BSC and MC became somewhat enlarged, storage bodies disappeared, and plastids were visible at the light microscope level. The intercellular space also expanded and some stomata appeared mature (Fig. 1B). Fig. 1. Open in new tabDownload slide Transverse sections of post-germination cotyledons of light- and dark-grown seedlings of F. trinervia. A, d 0, Light-grown; B, d 4, light-grown; C, d 7, light-grown; D, d 0, dark-grown; E, d 5, dark-grown; F, d 10, dark-grown. Bar = 50 μm. am, Abaxial MC; bs, BSC; i, intercellular space; pm, palisade MC; s, stomata; sm, spongy MC; unlabeled arrows, chloroplasts. Fig. 1. Open in new tabDownload slide Transverse sections of post-germination cotyledons of light- and dark-grown seedlings of F. trinervia. A, d 0, Light-grown; B, d 4, light-grown; C, d 7, light-grown; D, d 0, dark-grown; E, d 5, dark-grown; F, d 10, dark-grown. Bar = 50 μm. am, Abaxial MC; bs, BSC; i, intercellular space; pm, palisade MC; s, stomata; sm, spongy MC; unlabeled arrows, chloroplasts. BSC showed a clear differentiation from other MC, with larger, centripetally located chloroplasts. This pattern of plastid localization contrasts with the centrifugal location seen in NADP-malic enzyme-type C4 grass species (Hattersley and Browning, 1981; Dengler et al., 1996). By 7 d in the light, the Kranz anatomy became mature, with BSC and MC undergoing further enlargement and intercellular space becoming extensive (Fig. 1C). MC not directly adjacent to BSC did not develop plastids visible under the light microscope. Chloroplasts of both cell types had well-developed thylakoid membranes. Chloroplasts in MC formed grana, while these were inconspicuous or lacking in BSC (V. Pontieri and N.G. Dengler, unpublished results). Figure 1, D and E, shows that in dark-grown seedlings, cotyledon anatomical development was similar to that of the light-grown seedlings between d 0 and d 4. In the dark, development is arrested at this stage, and from d 7 to 10, dark-grown BSC and MC remained small, with small and undifferentiated pro-plastids (Fig. 1F). The cellular expansion characteristic of the maturation phase of Kranz anatomy development in the light did not occur in the dark. There was also very little overall cotyledon expansion in dark-grown seedlings. While MC expansion was evident in both light- and dark-grown cotyledons, it occured earlier and to a greater extent in the light (Fig. 1). The length to width ratio of palisade-like MC decreased from d 4 to d 7 and was more pronounced in light-grown cotyledons (Fig. 1). The isodiametric form of non-palisade MC was maintained during cell expansion through d 7 (Fig. 1). The dorsiventral mesophyll was moderately developed in the foliage leaves and cotyledons of F. trinervia, and was also evident in both cotyledons and foliage leaves of a C3 species in the same genus,Flaveria pringlei (Fig. 2F; V. Pontieri and N.G. Dengler, unpublished results). This dorsiventral mesophyll pattern was less conspicuous in the cotyledons and foliage leaves of the C4 dicot Amaranthus hypochondriacus (Wang et al., 1992, 1993a). Fig. 2. Open in new tabDownload slide RbcS mRNA accumulation patterns detected by mRNA in situ hybridization to transverse sections of the post-germination cotyledons of light- and dark-grown seedlings of F. trinervia (C4) (A–E) and F. pringlei (C3) (F). A, d 4, Light-grown, RbcS RNA antisense; B, d 7, light-grown, RbcS RNA antisense; C, d 9, light-grown, RbcS RNA antisense; D, d 4, dark-grown, RbcS RNA antisense, the seed coat is visible; E, d 7, dark-grown, RbcS RNA antisense; F, d 9, light-grown, RbcS RNA antisense, a section fromF. pringlei (C3). Cotyledon pairs were appressed while embedding in paraffin so that their adaxial sides were adjacent to each other. Transverse sections (6 mm) were taken from a region midway between the apex and base of cotyledons. Bar = 320 μm. Fig. 2. Open in new tabDownload slide RbcS mRNA accumulation patterns detected by mRNA in situ hybridization to transverse sections of the post-germination cotyledons of light- and dark-grown seedlings of F. trinervia (C4) (A–E) and F. pringlei (C3) (F). A, d 4, Light-grown, RbcS RNA antisense; B, d 7, light-grown, RbcS RNA antisense; C, d 9, light-grown, RbcS RNA antisense; D, d 4, dark-grown, RbcS RNA antisense, the seed coat is visible; E, d 7, dark-grown, RbcS RNA antisense; F, d 9, light-grown, RbcS RNA antisense, a section fromF. pringlei (C3). Cotyledon pairs were appressed while embedding in paraffin so that their adaxial sides were adjacent to each other. Transverse sections (6 mm) were taken from a region midway between the apex and base of cotyledons. Bar = 320 μm. Light Induction of BSC-Specific RbcS mRNA Accumulation We examined the temporal and spatial accumulation patterns of RbcS mRNA in the cotyledons of both dark- and light-grown seedlings d 0 to d 12 after seed sowing, and the results are shown in Figure 2. The cotyledons of light- and dark-grown seedlings from d 0 to d 3 lacked detectable RbcS mRNA by our in situ hybridization techniques (data not shown). In light-grown seedlings, accumulation of RbcS mRNAs in BSC was observed at d 4 and the level continued to increase up to d 7 (Fig. 2, A and B). Abundant accumulation of RbcS mRNA was also detected in MC at d 4 (Fig. 2A), but the level continuously decreased and became undetectable in the cotyledons of d 7 and older seedlings (Fig. 2, B and C). At d 4, dark-grown seedlings showed a low level of RbcS mRNA in both BSC and MC (Fig. 2D); the level of RbcS mRNA accumulation remained unchanged by d 6 and was found to decline slightly in both BSC and MC by d 7 (Fig. 2E). RbcS mRNA was not detectable in the cotyledons of d 9 and older seedlings under dark-grown conditions (data not shown). The RbcS mRNA accumulation patterns observed in our in situ hybridization studies fell into a similar temporal scheme to that reported for light- and dark-grown foliage leaves of maize seedlings (Nelson et al., 1984). For comparison, we also examined the pattern of RbcS mRNA accumulation in the cotyledons of a C3 Flaveria species,F. pringlei. We detected abundant RbcS mRNA accumulation in all cotyledon photosynthetic cell types of light-grown seedlings, with the highest level in adaxial palisade MC (Fig. 2F). This resembles the pattern observed in the dark-grown cotyledons of F. trinervia (Fig. 2, D and E), although the accumulation level was much higher. The above results suggest that in the cotyledons of F. trinervia, the expression of RbcS genes is developmentally controlled, so that RbcS mRNAs accumulate in both cell types independent of light conditions. However, in the presence of light, RbcS gene expression was either down-regulated or repressed in MC and up-regulated in BSC in d 5 and older cotyledons. The RbcS mRNA accumulation pattern in dark-grown cotyledons was C3 like in terms of cell specificity and remained unchanged after d 5. Light Induction of MC-Specific Pdk and Ppc mRNA Accumulation We also examined the accumulation of Pdk and Ppc mRNA by mRNA in situ hybridization and daily sampling from d 0 to d 12 (Fig.3). In MC of the light-grown cotyledons, we detected a dorsiventral gradient of mRNA accumulation for both Pdk and Ppc at d 4: high levels of mRNA accumulated in palisade-like MC (layer 1) and very low levels in other MC (layers 2–4) (Figs. 1 and 3, A and D). By d 5, high levels were detected in the MC of layers 2 to 4 (Fig. 3B). The level of mRNA accumulation decreased after d 5 and reached the steady state at d 7. From d 7 to d 12, the mRNA transcripts were detected in all MC except in the abaxial mesophyll (layer 5), which is adjacent to the lower epidermal cells and lacks direct contact with the BSC (Figs. 1C and 3, C and E). The absence of Pdk and Ppc expression in this mesophyll layer has also been observed in the foliage leaves of this species (Shu, 1996). The absence of Ppc mRNA accumulation in MC distal from BSC has also been reported for husk leaves of maize and foliage leaves of Atriplex rosea(Langdale et al., 1988b; Dengler et al., 1995). Fig. 3. Open in new tabDownload slide Pdk and Ppc mRNA accumulation patterns inF. trinervia cotyledons. A to C and F to H, Pdk mRNA accumulation patterns. D, E, I, and J, Ppc mRNA accumulation patterns. A, d 4, Light-grown, Pdk RNA antisense; B, d 5, light-grown, Pdk RNA antisense; C, d 7, light-grown, Pdk RNA antisense; D, d 4, light-grown, Ppc RNA antisense; E, d 7, light-grown, Ppc RNA antisense; F, d 4, dark-grown, Pdk RNA antisense; G, d 4, dark-grown, Pdk RNA sense probe; H, d 7, dark-grown, Pdk RNA antisense; I, d 7, dark-grown, Ppc RNA antisense; J, d 7, light-grown, Ppc RNA sense. Bar = 320 μm. For details, see Figure 2 legend. Under dark-field microscopy, the abundance of mRNA accumulation corresponds to the color saturation: yellowish red indicates low abundance and dark red indicates high abundance. Fig. 3. Open in new tabDownload slide Pdk and Ppc mRNA accumulation patterns inF. trinervia cotyledons. A to C and F to H, Pdk mRNA accumulation patterns. D, E, I, and J, Ppc mRNA accumulation patterns. A, d 4, Light-grown, Pdk RNA antisense; B, d 5, light-grown, Pdk RNA antisense; C, d 7, light-grown, Pdk RNA antisense; D, d 4, light-grown, Ppc RNA antisense; E, d 7, light-grown, Ppc RNA antisense; F, d 4, dark-grown, Pdk RNA antisense; G, d 4, dark-grown, Pdk RNA sense probe; H, d 7, dark-grown, Pdk RNA antisense; I, d 7, dark-grown, Ppc RNA antisense; J, d 7, light-grown, Ppc RNA sense. Bar = 320 μm. For details, see Figure 2 legend. Under dark-field microscopy, the abundance of mRNA accumulation corresponds to the color saturation: yellowish red indicates low abundance and dark red indicates high abundance. In BSC of light-grown cotyledons, low levels of accumulation were observed from d 5 to d 6 for Pdk mRNAs (Fig. 3B), but were undetectable by d 7 (Fig. 3C). A low level of RNA accumulation remained detectable in BSC of the d 7 and older seedlings using both the sense and antisense endogenous Ppc probes (Fig. 3E) at the same high hybridization stringency (Fig. 3, E and J). We found that the temporal and spatial patterns of Ppc mRNA accumulation were very similar to those of Pdk mRNA, with two exceptions: (a) the steady-state level of Ppc mRNA in the MC of the expanded cotyledons (from d 7 to d 12) was much higher than that of Pdk mRNA (Fig. 3, C and E), which is consistent with the estimates from RNA blotting analysis of mRNA from the maize leaf blade (Sheen and Bogorad, 1987b; Langdale and Kidner, 1994); (b) a low level of accumulation of unknown RNA transcripts was detected in the BSC of light-grown cotyledons with both sense and antisense endogenous Ppc probes, whereas the mRNA accumulation detected with endogenous Pdk gene probes was clearly MC specific in the 7 d and older cotyledons of light-grown seedlings. In the cotyledons of dark-grown seedlings, no mRNA accumulation for either gene was detected between d 0 and d 3. The level of Pdk and Ppc mRNA accumulation in the palisade mesophyll was higher than in the other mesophyll layers or in BSC between d 4 and d 7 (Fig. 3, F, H, and I). The accumulation patterns of RbcS, Pdk, and Ppc mRNAs in the expanded light-grown cotyledons of 7 d and older seedlings were essentially the same as those detected in the expanded foliage leaves of this species (Shu, 1996). Studies of both anatomy and photosynthetic physiology have shown that the foliage leaves of F. trinervia are typical C4 organs (Ku et al., 1991). Although no data on photosynthetic physiology are available for the cotyledons, our results concerning anatomy and gene expression patterns indicate that the expanded cotyledons of light-grown F. trinervia could function as typical C4photosynthetic organs and undergo C4CO2 assimilation. DISCUSSION Light Is Required for Full Differentiation of Photosynthetic Cell Types in C4 F. trinerviaCotyledons Post-germination cotyledon development in F. trinerviaappears to include two phases: a light-independent phase, or post-germination I, and a light-dependent phase, or post-germination II. From d 0 to d 4, both light- and dark-grown seedlings shared similar developmental patterns, such as nutrient reserve breakdown, repositioning and differential growth of plastids so that BSC have enlarged, and centripetally placed plastids, while MC had small, peripherally positioned plastids and C3-like photosynthetic gene expression. Light is therefore apparently not required for these developmental processes. We refer to this phase as post-germination I (Fig.4A). Fig. 4. Open in new tabDownload slide Differences in cotyledon and seedling development between two C4 dicotyledonous species, F. trinervia (A) and A. hypochondriacus(B). Embryogenesis, Formation of the embryo proper and endosperm; the two species represent two types of embryogenesis patterns in angiosperms. Post-embryogeny, Absorption of endosperm and seed formation in both species; cotyledon expansion in A, perisperm formation in B; cotyledon cell differentiation and light-induced chloroplast formation take place. Maturation, Nutrient reserve deposition; lipid and protein deposition in cotyledons for both species, starch deposition in cotyledons for A and in perisperm for B. Desiccation, Dehydration and dormancy of seedlings. Germination, From seed imbibition to rupture of testa by radicle. Post-germination I, C3-like development such as mobilization of cotyledon nutrient reserves, resumption of cell type differentiation, and non-cell-type-specific C3 and C4 gene expression. Post-germination II, Full C4 development such as maturation of Kranz anatomy and establishment and maintenance of cell-type-specific C4 gene expression. The two species employ two different subtypes of decarboxylation pathways and show different developmental patterns in light- and dark-grown conditions (shaded). C4 competence, Hypothetical period or developmental window when essential light-induced signals for C4 development and cell-type-specific C4 gene expression are generated. Fig. 4. Open in new tabDownload slide Differences in cotyledon and seedling development between two C4 dicotyledonous species, F. trinervia (A) and A. hypochondriacus(B). Embryogenesis, Formation of the embryo proper and endosperm; the two species represent two types of embryogenesis patterns in angiosperms. Post-embryogeny, Absorption of endosperm and seed formation in both species; cotyledon expansion in A, perisperm formation in B; cotyledon cell differentiation and light-induced chloroplast formation take place. Maturation, Nutrient reserve deposition; lipid and protein deposition in cotyledons for both species, starch deposition in cotyledons for A and in perisperm for B. Desiccation, Dehydration and dormancy of seedlings. Germination, From seed imbibition to rupture of testa by radicle. Post-germination I, C3-like development such as mobilization of cotyledon nutrient reserves, resumption of cell type differentiation, and non-cell-type-specific C3 and C4 gene expression. Post-germination II, Full C4 development such as maturation of Kranz anatomy and establishment and maintenance of cell-type-specific C4 gene expression. The two species employ two different subtypes of decarboxylation pathways and show different developmental patterns in light- and dark-grown conditions (shaded). C4 competence, Hypothetical period or developmental window when essential light-induced signals for C4 development and cell-type-specific C4 gene expression are generated. From d 5 to d 7, light- and dark-grown seedlings showed divergent development patterns. In light-grown seedlings, we observed arrest of hypocotyl elongation, continued BSC and MC expansion and redifferentiation, and high-level, cell-type-specific mRNA accumulation of the three C4 photosynthetic genes in the cotyledons. In contrast, we observed continuous hypocotyl elongation, arrest of cell expansion and redifferentiation, and low and nonspecific mRNA accumulation for all three genes in dark-grown seedlings. Therefore, light is essential for the full differentiation of cotyledon BSC and MC. We refer to this phase as post-germination II (Fig. 4A). Our division of post-germination cotyledon development into two phases is based solely on the observation that some development can proceed in both light- and dark-grown seedlings. It is known that regulatory proteins and mRNAs synthesized in the post-embryogeny phase can be stored in various forms in mature and desiccated seeds and be mobilized in the post-germination phases to regulate gene activities (Goldberg et al., 1989; Kretsch et al., 1995). Thus, it is possible that light-independent developmental processes in dark-grown seedlings of post-germination I are actually light dependent and regulated by light-induced signals synthesized during the post-embryogeny phase of seed development. We found that some veins with morphologically distinctive BSC and associated mesophyll layers were already present in d 0 imbibed embryos, but that fully mature Kranz anatomy, including substantial cell expansion and the formation of extensive intercellular space in the abaxial mesophyll did not develop until d 7. Thus, it is likely that C4 cell type development is initiated in either late embryogenesis or in the post-embryogeny phase of the parental plant, but that differentiation processes are arrested by nutrient storage and seed desiccation. Further development of Kranz anatomy is resumed during the post-germination I phase and is completed during the post-germination II phase only in light-grown cotyledons. Based on our observations, the completion of the transition from post-germination I to post-germination II in the cotyledons of F. trinervia requires light-induced signals that contribute to the competence for further C4 development (Fig. 4). C4 competence is likely attained around d 5 under light-grown conditions. Previous studies indicate that C4 competence is also required for foliage leaf development in both monocot and dicot C4 species, since young leaves that have not obtained C4competence always show C3-like gene expression patterns even in light-grown conditions (Sheen and Bogorad, 1985;Langdale et al., 1988b; Wang et al., 1992, 1993b; Dengler et al., 1995). Establishment of Cell-Type-Specific C4 Gene Expression Is Light Dependent In the present study, we found that light is essential for inducing both high-level and cell-type-specific accumulation of RbcS, Pdk, and Ppc mRNAs in post-germination cotyledons of light-grown seedlings. The two events, up-regulation and the cell-type-specificity of C4 gene expression, are likely regulated by different light-induced signals. While up-regulation of the three C4 genes was observed by d 4 (right after seedling emergence), initial mRNA accumulation patterns were not cell type specific. Complete cell-type-specific patterns for RbcS and Pdk genes were only finalized at d 7. The difference of 3 d between the two events indicates that the cell type specificity of C4 gene expression is likely regulated by light-induced developmental signals or conditions, rather than by direct light action on C4 gene transcription. A study of maize foliage leaves using biolistic gene transfer methods showed that red light, though important in inducing rapid up-regulation of C4 genes and chloroplast development, is not sufficient to suppress RbcS gene transcription in improper cell types such as maize epidermal cells (Bilang and Bogorad, 1996). Recently, transgenic studies using GUS fusion constructs from different sequences at the 5′ and 3′ ends of Pdk genes, Ppc, and NADP-malic enzyme genes have shown that control of cell type specificity and the level of gene expression use different cis-acting elements (Taylor et al., 1997; Westhoff et al., 1997). The cis-acting elements involved in controlling cell-type-specific expression of the C4-enzyme-coding genes are not found in the genes that encode enzymes of C3 isoforms (Sheen, 1991;Chitty et al., 1994; Stockhaus et al., 1994, 1997; Furbank et al., 1997; Marshall et al., 1997). All of the above results suggest that the regulation of C4 gene expression is 2-fold: regulation of the steady-state level of gene activity, a common theme shared with C3 plants, and regulation of cell type specificity of gene expression, a unique feature of C4 genes. We detected a low level of hybridization signal in cotyledon BSC with the antisense Ppc mRNA probe (Fig. 3E). We also detected a similar level of hybridization signal in both BSC and MC at the same high hybridization stringency with the sense riboprobe of Ppc (Fig. 3J). Therefore, it is likely that the signal detected in BSC with both Ppc sense and antisense probes are background noise, and that the accumulation of Ppc and Pdk mRNAs are MSC specific. We found that an increase in Pdk and Ppc mRNA levels in MC of light-grown cotyledons coincided with cotyledon greening. High levels of mRNA accumulation for both genes begins in the palisade MC (layer 1) by d 4 and extends to the abaxial side of the cotyledons by d 5. Dorsiventral chlorophyll gradients that cause the dorsiventral cotyledon greening patterns have also been reported in the post-germination cotyledons of a C3 plant,Cucurbita pepo (Knapp et al., 1988), and have been suggested to be due to differential exposure to blue light along the dorsiventral axis of the cotyledons (Knapp et al., 1988). It remains to be investigated whether the Pdk and Ppc mRNA gradients seen in F. trinervia might have been caused by a blue light gradient. Cell-Type-Specific C4 Gene Expression Occurs during Maturation of Morphological Cell Type Differentiation An important question in understanding C4development is whether light induction of C4 gene expression is coupled with leaf development and leaf cell type differentiation (Nelson and Dengler, 1992; Liu and Dengler, 1994; Berry et al., 1997). Our results show that in F. trinervia, accumulation of mRNA for genes involved in C4carbon metabolism occurs as morphological cell type differentiation becomes mature. BSC and MC differentiation proceeded simultaneously in post-germination cotyledons and full differentiation of BSC and MC were observed by d 7, which is also the time when cell-type-specific mRNA accumulation for all three genes takes place. In contrast to our findings for F. trinervia cotyledons, in young foliage leaves of this species the establishment of a MC-specific pattern of Ppc and Pdk expression was conspicuously delayed in relation to the establishment of a BSC-specific pattern of RbcS gene expression. The same delay of Ppc and Pdk expression relative to that of RbcS is seen in the young foliage leaves of maize, amaranth, and Atriplex rosea (Langdale et al., 1988a; Schäffner and Sheen, 1992;Wang et al., 1992; Dengler et al., 1995; Shu, 1996). This distinction between cotyledons and foliage leaves may reflect a difference in the timing of cell proliferation relative to differentiation. In cotyledon development, cell proliferation only occurs during embryogenesis, and post-germination cotyledon cells do not divide (Tsukaya et al., 1994;Kretsch et al., 1995). In contrast, developing foliage leaves undergo considerable cell proliferation, and BSC surrounding the veins may be delimited before cell division within the MC ceases (Dengler et al., 1996). Our data support the view that signals involved in cell-type-specific regulation of C4 gene expression are positional relative to bundle sheath or vascular tissues (Langdale and Nelson, 1991; Nelson and Langdale, 1992). It remains unclear whether the up-regulation of Pdk and Ppc and the suppression of RbcS genes in the same MC are regulated by the same positional signals. In maize husk leaves, the absence of Ppc mRNAs and the presence of rbcL and RbcS mRNAs are found to coincide in the MC that do not have direct contact with the BSC (Langdale, 1988b; Langdale and Nelson, 1991). We did not observe the same coincidence in the cotyledons of F. trinervia. No RbcS mRNA accumulation was detected in the fifth layer of MC, where the Pdk and Ppc mRNAs are absent. This observation suggests that up-regulation of Pdk and Ppc and suppression of RbcS genes in MC, though normally coincident, may be controlled by different signals. Results from promoter analyses, mutant studies, and transgenic studies using different C4 photosynthetic genes in Flaveria sp. also suggest that there is no universal regulation mechanism among different C4photosynthetic genes (Langdale and Kidner, 1994; Furbank et al., 1997;Taylor et al., 1997; Westhoff et al., 1997). F. trinervia and A. hypochondriacus Differ in C4 Gene Expression Pattern and Cotyledon Development Our results demonstrate that in F. trinervia, cell-type-specific expression of genes involved in C4 carbon metabolism in the post-germination cotyledons only takes place in light-grown seedlings, not in dark-grown seedlings. This is true in spite of the existence of clear morphological differentiation between BSC and MC in the cotyledons. This finding emphasizes the fact that light is essential for the establishment and maturation of differential expression of these genes. The light requirement for differential expression of rbcS, PPDK, and PEPCase mRNA in F. trinervia is in marked contrast to the observed light independence of their differential expression in the cotyledons of another C4 dicot, A. hypochondriacus (Amaranthaceae) (Wang et al., 1993a). Wang et al. (1993a) found that the accumulation of PPDK mRNA and protein and PEPCase protein are MC specific in cotyledons of both dark- and light-grown 2-d-old seedlings. RbcS mRNAs and polypeptides were not cell type specific at d 2, but became BSC specific at d 5 in both dark- and light-grown cotyledons. Thus, light is not required for the establishment of cell-type-specific C4 gene expression in this species (Wang et al., 1993a). The underlying mechanisms that lead to this difference between the two dicot species is unclear. It is possible that the apparent difference in light requirement for cell-type-specific C4 gene expression between the two species is associated with their difference in seedling development and cotyledon cell type differentiation. Previous studies have shown that F. trinervia and A. hypochondriacus have strikingly different patterns of embryogenesis, post-embryogenic development, and post-germination development, and these are summarized in Figure 4. Embryogenesis in angiosperms is classified into six types (Johri et al., 1992): F. trinervia is an Asterad type, whereas A. hypochondriacus is a Chenopodiad type (Misra, 1964; Coimbra and Salema, 1994). Dicot plants are also classified into at least three types based on seed nutrient storage location: cotyledon storage, such as A. hypochondriacus endosperm storage, or perisperm storage. The three types show distinctive temporal patterns of cotyledon development (Johri et al., 1992; Bewley and Black, 1994;Kaplan and Cooke, 1997). Both Flaveria and Amaranthus spp. have nuclear endosperm that starts to form at the heart stage of embryogenesis. At the post-embryogeny phase, endospermis is gradually absorbed by the growing embryo. In amaranth, a portion of the nucellus converts into a perisperm storage tissue (Fig. 4B). Carbohydrates are stored as starch grains in perisperm plastids, but starch grains are absent from cotyledon plastids (Coimbra and Salema, 1994). In Flaveria, cotyledons and hypocotyl convert to nutrient storage organs during post-embryogenic development, and cotyledon cells are filled with protein bodies, lipids, and plastids with starch grains (Misra, 1964). Previous studies have shown that the presence of carbohydrates suppresses C4 gene transcription in isolated maize leaf MC (Sheen, 1990). In developing foliage leaves of A. hypochondriacus, a tight coordination between cell-type-specific C4 gene expression and the state of carbon metabolism or sink-source transition have been reported (Wang et al., 1993b; Berry et al., 1997). It is plausible that amaranth cotyledons, which lack starch deposition, could have faster photosynthetic development than the Flaveria cotyledons in the post-embryogeny phase, to become more leaf-like and obtain light-induced C4 competence before seed desiccation (Fig. 4). Further detailed comparative study is needed to confirm this speculation. The temporal differences in cotyledon development and C4 gene expression between A. hypochondriacus and F. trinervia are likely an evolutionary adaptation of each species to its germination environment. The great diversity among different plant species in seed structure, germination strategy, and corresponding temporal control of light-regulated genes is well documented (Stebbins, 1974; Johri et al., 1992; Bewley and Black, 1994). The differences between these two C4 dicot species in C4 development of cotyledons provide evidence that independent evolutionary origins of C4 photosynthetic mechanisms need not arise by the evolution of common developmental control pathways. A third distinctive developmental pattern has been recognized in species of Haloxylon (Chenopodiaceae) that show C4photosynthesis in the single large subepidermal BSC/MC ring surrounding assimilatory shoots (Pyankov et al., 1999). The reduced leaves of these plants are nonphotosynthetic, but the cotyledons are C3 and show no morphological differentiation among the photosynthetic cells at any stage of development (Pyankov et al., 1999). Evidence of independent evolution of C4photosynthetic pathways has been reported and reviewed for grass species (Hattersley and Browning, 1981; Hattersley, 1984; Dengler et al., 1985, 1986; Sinha and Kellogg, 1996). Extensive comparative studies of C4 gene expression and post-germination cotyledon development across different taxa of C4 dicots are promising to bring new insight into the evolution of developmental mechanisms in general and the evolution of this complex adaptation in particular. ACKNOWLEDGMENTS We are grateful to Peter Westhoff for providing the Pdk and Ppc cDNA clones and Jane Langdale and Jing-liang Wang for the in situ hybridization protocols. Our thanks also go to our colleagues Hewson Swift, Gayle Lamppa, Martin Kreitman, and Manfred Ruddat for helpful discussion. Aida Pascual, Mary Crane, and Maya Moody provided valuable technical assistance. We thank Sue Yamins and her greenhouse staff for taking care of seedstock plants. We thank J. Sheen and E.A. 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Copyright © 1999 American Society of Plant Physiologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Noguchi, Takahiro; Fujioka, Shozo; Choe, Sunghwa; Takatsuto, Suguru; Yoshida, Shigeo; Yuan, Heng; Feldmann, Kenneth A.; Tax, Frans E.

doi: 10.1104/pp.121.3.743pmid: 10557222

Abstract Seven dwarf mutants resembling brassinosteroid (BR)-biosynthetic dwarfs were isolated that did not respond significantly to the application of exogenous BRs. Genetic and molecular analyses revealed that these were novel alleles ofBRI1 (Brassinosteroid-Insensitive 1), which encodes a receptor kinase that may act as a receptor for BRs or be involved in downstream signaling. The results of morphological and molecular analyses indicated that these represent a range of alleles from weak to null. The endogenous BRs were examined from 5-week-old plants of a null allele (bri1-4) and two weak alleles (bri1-5 and bri1-6). Previous analysis of endogenous BRs in several BR-biosynthetic dwarf mutants revealed that active BRs are deficient in these mutants. However,bri1-4 plants accumulated very high levels of brassinolide, castasterone, and typhasterol (57-, 128-, and 33-fold higher, respectively, than those of wild-type plants). Weaker alleles (bri1-5 and bri1-6) also accumulated considerable levels of brassinolide, castasterone, and typhasterol, but less than the null allele (bri1-4). The levels of 6-deoxoBRs in bri1 mutants were comparable to that of wild type. The accumulation of biologically active BRs may result from the inability to utilize these active BRs, the inability to regulate BR biosynthesis in bri1 mutants, or both. Therefore,BRI1 is required for the homeostasis of endogenous BR levels. Based on their wide distribution in the plant kingdom, their diverse physiological effects at nanomolar levels, and the discovery of mutants deficient in their biosynthesis, it is now widely accepted that brassinosteroids (BRs) are important hormones that regulate growth and development (Fujioka and Sakurai, 1997a, 1997b; Sakurai and Fujioka, 1997; Yokota, 1997; Altmann, 1998; Clouse and Sasse, 1998). During a relatively short time, a number of BR mutants were isolated from Arabidopsis, pea, and tomato (for review, see Clouse and Feldmann, 1999). Many of the BR mutants have been well characterized by genetic, molecular, and biochemical studies. All of the BR mutants are dwarfs in that they exhibit a short, robust stature and dark-green, round leaves. In addition, most of the mutants have reduced fertility, a prolonged lifespan, and display abnormal skotomorphogenesis when grown in the dark. These BR dwarf mutants are divided into two classes based on their phenotypic response to exogenously supplied BRs (Clouse and Feldmann, 1999). One class of BR dwarf mutants is impaired in BR biosynthesis. The phenotype of this class of BR mutants can be rescued by exogenous application of BRs, but not by treatment with any other plant hormone. At present, six BR-biosynthetic mutants in Arabidopsis have been characterized (det2: Chory et al., 1991; Li et al., 1996,1997; Fujioka et al., 1997; Noguchi et al., 1999; cpd:Szekeres et al., 1996; Mathur et al., 1998; dwf4: Azpiroz et al., 1998; Choe et al., 1998; dwf1: Feldmann et al., 1989;Takahashi et al., 1995; Klahre et al., 1998; Choe et al., 1999a;dwf7/ste1: Choe et al., 1999b; and sax1:Ephritikhine et al., 1999). In addition, two BR-deficient mutants have been isolated and characterized from pea (lkb: Nomura et al., 1997, 1999; lk: Yokota et al., 1997) and two from tomato (dwarf: Bishop et al., 1996, 1999; dpy:Koka et al., 1999). A second class of BR dwarf mutants resembles the biosynthetic mutants in morphology, but cannot be rescued by BR feeding. This class of mutants is predicted to be blocked in the perception of BRs or in essential components of BR signaling downstream of perception. BR-insensitive mutants have been identified for Arabidopsis (bri1: Clouse et al., 1996; Kauschmann et al., 1996; Li and Chory, 1997), pea (lka: Nomura et al., 1997, 1999), and tomato (cu-3: Koka et al., 1999). Clouse et al. (1996) first identified a BR-insensitive mutant in Arabidopsis. This mutant,bri1-1 (brassinosteroid-insensitive 1-1), did not respond to BRs in root-inhibition assays, but did retain sensitivity to other plant hormones, including auxin and GA. Genetic analysis showed that bri1-1 was caused by a recessive mutation in a single gene. Kauschmann et al. (1996) also identified a BR-insensitive mutant they called cbb2 (cabbage 2), which was allelic to bri1-1. Li and Chory (1997) identified 18 additional alleles ofbri1, and isolated the BRI1 gene by mapping, isolation of DNA near BRI1, and the identification of a mutation in one allele that resulted in a detectable RFLP. TheBRI1 gene was predicted to encode a membrane-bound Leu-rich repeat (LRR) receptor kinase (RK), which appeared to be constitutively expressed throughout the plant, both in the light and in the dark. The predicted protein showed striking similarities to other plant LRR-RK gene products, such as those of CLAVATA1 (Clark et al., 1997) and Xa21 (Song et al., 1995), which are involved in developmental signaling pathways and in pathogen response, respectively. Although physiological and molecular data raise the possibility that BRI1 is a receptor for BRs, direct biochemical evidence has not yet been described. The natural occurrence of BRs in Arabidopsis was first demonstrated byFujioka et al. (1996). Castasterone, 6-deoxocastasterone, typhasterol, and 6-deoxotyphasterol were identified from extracts made from Arabidopsis shoots. A subsequent study expanded the BR profiles in Arabidopsis. From fully expanded siliques of Arabidopsis, six BRs, including brassinolide, castasterone, typhasterol, 6-deoxocastasterone, 6-deoxotyphasterol, and 6-deoxoteasterone, were identified (Fujioka et al., 1998). All BRs identified in Arabidopsis are important components of either the early or late C6-oxidation pathways. These pathways were previously established with studies using cultured cells of Catharanthus roseus (Fujioka and Sakurai, 1997b; Sakurai and Fujioka, 1997). The studies in Arabidopsis suggested that both the early and late C6-oxidation pathways were functional in this species as well. The analysis of BR levels in the BR-biosynthetic mutants have thus far substantiated the predicted BR pathways in Arabidopsis. The accumulation of precursor molecules in the BR biosynthetic pathway has been observed for several of these mutants. In addition, the BR-biosynthetic mutants of Arabidopsis have been shown to be deficient in endogenous BRs or sterols downstream of the blocked step (det2: Fujioka et al., 1997; dwf1: Klahre et al., 1998; Choe et al., 1999a; dwf7/ste1: Choe et al., 1999b). However, no information about endogenous BRs in BR-insensitive mutants of Arabidopsis is available. In GA-insensitive mutants such asDwarf-8 in maize (Fujioka et al., 1988), gai in Arabidopsis (Talon et al., 1990), and Rht3 in wheat (Appleford and Lenton, 1991), the accumulation of bioactive C19-GAs such as GA1 and GA20 has been reported. Furthermore, transcription of one GA-biosynthetic gene is increased in thegai mutant (Peng et al., 1997). The accumulation of GAs and the increased level of GA-biosynthetic gene transcription have led to the conclusion that GAI, which may encode a transcription factor, is also required for proper feedback regulation of the GA-biosynthetic pathway (Peng et al., 1997; Harberd et al., 1998). There is little known about how BR biosynthesis and signaling are coordinated. BRs are synthesized using sterols as precursors, but much less is understood about how sterols are funneled into BR biosynthesis. Within the BR-specific pathway, the 22α-hydroxylase supplied byDWF4 appears to be rate limiting (Choe et al., 1998). However, there is currently nothing known about how DWF4transcription is regulated or spatially controlled. Analysis ofCPD, a Cyt P450 (CYP) 90A one step downstream ofDWF4 in the current BR-biosynthetic pathway, indicated thatCPD mRNA levels are decreased by BR treatment (Mathur et al., 1998). This suggests that BR biosynthesis is regulated by a negative-feedback loop. Although the RK encoded by BRI1 is likely a key component of BR signal transduction, little is known about the regulation of BRI1 or about the nature of additional proteins that function downstream of BRI1. In animals, for both nuclear steroid hormone receptors and for some membrane-spanning ligands and receptors, feedback regulation occurs as a result of receptor activation (Wilkinson et al., 1994; Blumberg et al., 1998). We studied the BR profile of bri1 mutants to understand the regulation of BRs in Arabidopsis. In this study, we describe the isolation of seven bri1alleles and present morphometric analysis of plants homozygous for one severe allele and two weak alleles. We have also identified mutations within the BRI1 gene for all seven alleles, and provide data suggesting that the severe alleles represent the null phenotype ofBRI1. We also determined the levels of endogenous BRs in three bri1 alleles with gas chromatography-selective ion monitoring (GC-SIM) analysis using 2H-labeled internal standards. We show here that mutations in BRI1cause very large accumulations of 6-oxoBRs such as brassinolide, castasterone, and typhasterol, and that the level of BRs is positively correlated with allele and phenotypic severity. MATERIALS AND METHODS Isolation and Mapping of bri1 Alleles bri1-3 (dwf2-32) and bri1-4(dwf2-2074) were found in a screen for dwarf mutants from a population of plants of the Wassilewskija-2 (Ws-2) ecotype transformed with the T-DNA plasmid of Agrobacterium tumefaciens using the seed transformation method (Feldmann and Marks, 1987; Feldmann and Azpiroz, 1994). However, the dwarf phenotype did not cosegregate with kanamycin resistance, and therefore bri1-3 andbri1-4 represent untagged alleles (data not shown).bri1-5(dwf2-W41), bri1-7, bri1-8, andbri1-9 were isolated after ethyl methanesulfonate (EMS) mutagenesis of Ws-2 seeds, and bri1-6(dwf2-399) was identified among the dwarf mutants obtained from the Arabidopsis Biological Resource Center (Ohio State University, Columbus) (CS399, http://aims.cps.msu.edu/aims/). CS399 had been isolated previously using an unknown mutagen. All seven alleles were classified as insensitive to BRs based on their lack of a significant response in feeding studies performed on inflorescences and seedlings (data not shown, see Choe et al., 1998, 1999a). Three of these mutations were initially designated as Ws-2 EMS (WM) mutants (see TableI) but have been renamed asbri1 alleles. Table I. bri1 mutant alleles Allele . Former Designation . Mutagen . DNA Sequence Change . Predicted Change in BRI1 Protein1-a . bri1-3 dwf2-32 T-DNA bp 2,745, 4-bp deletion 44 Amino acids, then stop codon after amino acid 914 bri1-4 dwf2-2074 T-DNA bp 459, 10-bp deletion 13 Amino acids, then stop codon after amino acid 153 bri1-5 dwf2-w41 EMS TGC-TAC, bp 206 C69-Y69 bri1-6 dwf2-399 EMS GGT-GAT, bp 1,931 G644-D644 bri1-7 dwf2-WM3-2 EMS GGT-AGT, bp 1,838 G613-S613 bri1-8 dwf2-WM6-2 EMS CGG-CAG, bp 2,948 R983-N983 bri1-9 dwf2-WMB19 EMS CTT-TTT, bp 1,985 S662-F662 Allele . Former Designation . Mutagen . DNA Sequence Change . Predicted Change in BRI1 Protein1-a . bri1-3 dwf2-32 T-DNA bp 2,745, 4-bp deletion 44 Amino acids, then stop codon after amino acid 914 bri1-4 dwf2-2074 T-DNA bp 459, 10-bp deletion 13 Amino acids, then stop codon after amino acid 153 bri1-5 dwf2-w41 EMS TGC-TAC, bp 206 C69-Y69 bri1-6 dwf2-399 EMS GGT-GAT, bp 1,931 G644-D644 bri1-7 dwf2-WM3-2 EMS GGT-AGT, bp 1,838 G613-S613 bri1-8 dwf2-WM6-2 EMS CGG-CAG, bp 2,948 R983-N983 bri1-9 dwf2-WMB19 EMS CTT-TTT, bp 1,985 S662-F662 F1-a The total length of BRI1 is 1,196 amino acids (Li and Chory 1997). Open in new tab Table I. bri1 mutant alleles Allele . Former Designation . Mutagen . DNA Sequence Change . Predicted Change in BRI1 Protein1-a . bri1-3 dwf2-32 T-DNA bp 2,745, 4-bp deletion 44 Amino acids, then stop codon after amino acid 914 bri1-4 dwf2-2074 T-DNA bp 459, 10-bp deletion 13 Amino acids, then stop codon after amino acid 153 bri1-5 dwf2-w41 EMS TGC-TAC, bp 206 C69-Y69 bri1-6 dwf2-399 EMS GGT-GAT, bp 1,931 G644-D644 bri1-7 dwf2-WM3-2 EMS GGT-AGT, bp 1,838 G613-S613 bri1-8 dwf2-WM6-2 EMS CGG-CAG, bp 2,948 R983-N983 bri1-9 dwf2-WMB19 EMS CTT-TTT, bp 1,985 S662-F662 Allele . Former Designation . Mutagen . DNA Sequence Change . Predicted Change in BRI1 Protein1-a . bri1-3 dwf2-32 T-DNA bp 2,745, 4-bp deletion 44 Amino acids, then stop codon after amino acid 914 bri1-4 dwf2-2074 T-DNA bp 459, 10-bp deletion 13 Amino acids, then stop codon after amino acid 153 bri1-5 dwf2-w41 EMS TGC-TAC, bp 206 C69-Y69 bri1-6 dwf2-399 EMS GGT-GAT, bp 1,931 G644-D644 bri1-7 dwf2-WM3-2 EMS GGT-AGT, bp 1,838 G613-S613 bri1-8 dwf2-WM6-2 EMS CGG-CAG, bp 2,948 R983-N983 bri1-9 dwf2-WMB19 EMS CTT-TTT, bp 1,985 S662-F662 F1-a The total length of BRI1 is 1,196 amino acids (Li and Chory 1997). Open in new tab The original bri1 allele, bri1-1, showed tight linkage to a marker on the bottom of chromosome IV (1/126 chromosomes recombinant for the CAPS marker DHS1, Clouse et al., 1996).DHS1 is tightly linked to SSLP marker nga1107, andBRI1 is physically located between these two markers (Li and Chory, 1997). We established mapping populations by crossingbri1 alleles to wild-type plants of the Columbia ecotype and selecting dwarf plants among the F2 progeny. DNA was isolated from single leaves or flowers as described previously (Dellaporta et al., 1983; Krysan et al., 1996). PCR reactions were prepared as described by Bell and Ecker (1994). Since five of these seven BR-insensitive dwarf mutants mapped to the same general location as bri1-1, we performed crosses between all seven of these mutants to determine if they were alleles of the same gene. All combinations tested generated dwarf plants in the F1, and thus these represent alleles of the same gene (data not shown). Morphometric Analysis Approximately 20 seeds were planted in round pots (10 cm in diameter) with soil (Metromix 350, Grace Sierra, Miltipas, CA) presoaked in water. Flats containing the pots were covered in plastic wrap and cold treated for 3 to 4 d before transfer to a growth chamber (16 h of light [240 μmol m−2s−1] at 22°C and 8 h of dark at 21°C, with 75%–90% humidity). The plastic wrap was removed after the seedlings were established (5–7 d), and the seedlings were thinned so that there were four or five well-spaced seedlings per pot. The pots were subirrigated with water or Hoagland's nutrient solution as necessary. When the plants were 5 weeks of age, the morphological traits listed in Table II were measured. Plant height was measured to the nearest millimeter, and the length of siliques and the length and width of leaves were measured to the nearest half-millimeter using a ruler. The length of siliques and the distance between siliques on the main inflorescence were measured to the nearest 10th of a millimeter using a ruler in the ocular of a dissecting microscope. Table II. Morphometric analysis of Ws-2 and bri1 mutants at 5 weeks of age Parameter . Ws-2 . bri1-4 . bri1-5 . bri1-7 . Height (cm) 29.7 ± 2.2 1.26 ± 0.43 4.02 ± 1.12 3.18 ± 1.15 Distance between siliques on main inflorescence2-a(cm) 0.86 ± 0.37 0.05 ± 0.03 0.14 ± 0.08 0.18 ± 0.10 Length of leaf blade2-b(cm) 1.29 ± 0.26 0.22 ± 0.06 1.02 ± 0.14 0.58 ± 0.12 Width of leaf blade2-b(cm) 0.91 ± 0.12 0.30 ± 0.08 1.09 ± 0.11 0.69 ± 0.11 Length of siliques2-c(cm) 1.38 ± 0.06 0.20 ± 0.02 0.62 ± 0.14 0.32 ± 0.14 No. of seeds2-c 50.9 ± 4.6 0.13 ± 0.35 22.9 ± 14.3 3.73 ± 6.6 Parameter . Ws-2 . bri1-4 . bri1-5 . bri1-7 . Height (cm) 29.7 ± 2.2 1.26 ± 0.43 4.02 ± 1.12 3.18 ± 1.15 Distance between siliques on main inflorescence2-a(cm) 0.86 ± 0.37 0.05 ± 0.03 0.14 ± 0.08 0.18 ± 0.10 Length of leaf blade2-b(cm) 1.29 ± 0.26 0.22 ± 0.06 1.02 ± 0.14 0.58 ± 0.12 Width of leaf blade2-b(cm) 0.91 ± 0.12 0.30 ± 0.08 1.09 ± 0.11 0.69 ± 0.11 Length of siliques2-c(cm) 1.38 ± 0.06 0.20 ± 0.02 0.62 ± 0.14 0.32 ± 0.14 No. of seeds2-c 50.9 ± 4.6 0.13 ± 0.35 22.9 ± 14.3 3.73 ± 6.6 Measurements of plant height were made on at least 15 plants. Each value represents the mean ± sd. F2-a The distances between the first 15 pedicels on the main bolt were measured from a single plant. F2-b The second pair of leaves was measured and averaged for 15 plants. F2-c The first 15 siliques on a single plant were measured, and the seeds were removed and counted. Open in new tab Table II. Morphometric analysis of Ws-2 and bri1 mutants at 5 weeks of age Parameter . Ws-2 . bri1-4 . bri1-5 . bri1-7 . Height (cm) 29.7 ± 2.2 1.26 ± 0.43 4.02 ± 1.12 3.18 ± 1.15 Distance between siliques on main inflorescence2-a(cm) 0.86 ± 0.37 0.05 ± 0.03 0.14 ± 0.08 0.18 ± 0.10 Length of leaf blade2-b(cm) 1.29 ± 0.26 0.22 ± 0.06 1.02 ± 0.14 0.58 ± 0.12 Width of leaf blade2-b(cm) 0.91 ± 0.12 0.30 ± 0.08 1.09 ± 0.11 0.69 ± 0.11 Length of siliques2-c(cm) 1.38 ± 0.06 0.20 ± 0.02 0.62 ± 0.14 0.32 ± 0.14 No. of seeds2-c 50.9 ± 4.6 0.13 ± 0.35 22.9 ± 14.3 3.73 ± 6.6 Parameter . Ws-2 . bri1-4 . bri1-5 . bri1-7 . Height (cm) 29.7 ± 2.2 1.26 ± 0.43 4.02 ± 1.12 3.18 ± 1.15 Distance between siliques on main inflorescence2-a(cm) 0.86 ± 0.37 0.05 ± 0.03 0.14 ± 0.08 0.18 ± 0.10 Length of leaf blade2-b(cm) 1.29 ± 0.26 0.22 ± 0.06 1.02 ± 0.14 0.58 ± 0.12 Width of leaf blade2-b(cm) 0.91 ± 0.12 0.30 ± 0.08 1.09 ± 0.11 0.69 ± 0.11 Length of siliques2-c(cm) 1.38 ± 0.06 0.20 ± 0.02 0.62 ± 0.14 0.32 ± 0.14 No. of seeds2-c 50.9 ± 4.6 0.13 ± 0.35 22.9 ± 14.3 3.73 ± 6.6 Measurements of plant height were made on at least 15 plants. Each value represents the mean ± sd. F2-a The distances between the first 15 pedicels on the main bolt were measured from a single plant. F2-b The second pair of leaves was measured and averaged for 15 plants. F2-c The first 15 siliques on a single plant were measured, and the seeds were removed and counted. Open in new tab Determination of the DNA Sequence of bri1 Alleles The specific mutations in these seven bri1 alleles were identified by first designing primers based on the genomic sequence (Li and Chory, 1997; GenBank accession no. AF017056). Using oligonucleotides synthesized by Genosys Biotechnologies (The Woodlands, TX), a fragment of DNA corresponding to the coding region ofBRI1 was amplified using XTaq (Panvera, Madison, WI), the F1 and R1 primers (F1 [79] agagataggtggttgggggtaaaatgtat, R1 [4366] aaaaatagacccaaggaaaatcggactga), and DNA isolated from a single leaf or flower from each mutant (Krysan et al., 1996). The bracketed numbers refer to the nucleotide position (Li and Chory, GenBank accession no. AF017056). DNA fragments were purified (Prep-A-Gene DNA purification system, Bio-Rad, Hercules, CA) and sequenced at the Arizona Research Laboratory (University of Arizona, Tucson). For each mutant, the entire fragment was sequenced in one strand using the following additional oligonucleotides: F2(431) taatcagaagaagaggtaac, F3 (896) tgcagagacgacaaagttac, F4 (1368) tttctcgatgtttcctccaa, F5 (1836) ccggaatctctgacgaatct, F6 (2298) agaatctcgctatcctcaag, F7 (2741) caatttgggtcataacgata, F8 (3236) gaagctgactggtgtgaaag, F9 (3702) catatcatccacagagacat, and F10 (4208) aatagaggatggagggttca. Putative mutations were confirmed by sequencing the second strand using the following primers: R2 (1437) ttcccggagatgtcaagatg, R3 (3503) caagaagaggcacaagattt, and R4 (2862) tccgtaagcatagtaagagc, and repeating the sequencing on fragments isolated from independent PCR reactions. Quantitative Analysis of Endogenous BRs Plants were grown for 5 weeks on soil. The aerial portions (rosette leaves, inflorescences, flowers, and siliques) were harvested and frozen. The tissue was lyophilized (Lyphlick 12, LabConco, Kansas City, MO) and ground to a fine powder using a mortar and pestle. Lyophilized plant material (40 g fresh weight equivalent) was extracted with 400 mL of MeOH-CHCl3 (4:1) twice, and [2H6]brassinolide, [2H6]castasterone, [2H6]typhasterol, [2H6]teasterone, [2H6]6-deoxocastasterone, [2H6]6-deoxotyphasterol, and [2H6]6-deoxoteasterone (100 ng each) were added to the extract as internal standards. After evaporation of the solvent in vacuo, the extract was partitioned between CHCl3 and water three times. The CHCl3-soluble fraction was subjected to silica gel chromatography (Sep-Pak Vac Silica, 35 mL, Waters, Milford, MA). The column was subsequently eluted with 100 mL of CHCl3, 2% (v/v) MeOH in CHCl3, and 7% (v/v) MeOH in CHCl3. The 2% (v/v) MeOH and 7% (v/v) MeOH fractions were purified by Sephadex LH-20 column chromatography (column volume of 200 mL). The column was eluted with MeOH-CHCl3 (4:1). The effluents of elution volume to total column volume 0.6 to 0.8 were collected as the BR-containing fraction. After purification with an ODS cartridge (Sep-Pak Plus C18, Waters) with 20 mL of MeOH, eluates were subjected to ODS-HPLC (Senshu Pak Pegasil ODS, 10 × 30 mm + Senshu Pak Pegasil ODS, 20 × 250 mm; Senshu Scientific, Tokyo) at a flow rate of 8 mL min−1. Ninety percent acetonitrile was used as a solvent for the eluate derived from the 2% (v/v) MeOH fraction, and 70% (v/v) acetonitrile was used for the eluate derived from the 7% MeOH fraction. HPLC purification from the 7% MeOH fraction yielded a brassinolide fraction (retention time [Rt] from 8–10 min), a castasterone fraction (Rt from 12–14 min), a teasterone fraction (Rt from 17–20 min), a typhasterol fraction (Rt from 26–32 min), and a 6-deoxocastasterone fraction (Rt from 38–44 min). HPLC purification from the 2% (v/v) MeOH fraction yielded a 6-deoxoteasterone fraction (Rt from 32–36 min) and a 6-deoxotyphasterol fraction (Rt from 48–52 min). Each fraction was analyzed by GC-SIM after derivatization. Quantitative Analysis of Endogenous Sterols For sterol analysis, lyophilized plant material (2 g fresh weight equivalent) from wild-type and bri1 mutant alleles was used. Plant material was extracted with 50 mL of MeOH-CHCl3 (4:1) twice, and [2H7]24-methylenecholesterol (3 μg/g fresh weight), [2H6]campesterol (30 μg/g fresh weight), and [2H6]campestanol (1 μg/g fresh weight) were added to the extract as internal standards. After evaporation of the solvent in vacuo, the extract was partitioned between CHCl3 and water three times. The CHCl3-soluble fraction was purified with a silica gel cartridge column (Sep-Pak Vac Silica, 12 mL, Waters), which was eluted with 20 mL of CHCl3. The eluate was purified with an ODS cartridge (Sep-Pak Plus C18, Waters), which was eluted with 20 mL of MeOH. The eluent was subjected to ODS-HPLC (Senshu Pak ODS 4150-N; 10 × 150 mm, Senshu Scientific) at a flow rate of 2 mL min−1with MeOH. Fractions were collected every 0.5 min (Rt between 10 and 20 min). The main fractions of each sterol were as follows: 24-methylenecholesterol (Rt of 13 to 13.5 min), campesterol (Rt of 15.5 to 16 min), and campestanol (Rt of 16.5 to 17 min). Each fraction was analyzed by full-scan gas chromatography-mass spectrometry (GC-MS) after derivatization. 2H Standards [2H6]Campesterol was kindly supplied by Tama Biochemical (Tokyo). [2H6]Brassinolide, [2H6]castasterone, [2H6]typhasterol, [2H6]teasterone (Takatsuto and Ikekawa, 1986), [2H6]6-deoxocastasterone (Choi et al., 1996), [2H6]6-deoxotyphasterol, [2H6]6-deoxoteasterone (Choi et al., 1997), [2H7]24-methylenecholesterol (Takatsuto et al., 1998), and [2H6]campestanol (Noguchi et al., 1999) were chemically synthesized. GC-MS Analysis GC-MS analysis was carried out under the following conditions: a mass spectrometer (Automass JMS-AM150, JEOL, Tokyo) was connected to a gas chromatograph (model 5890A-II, Hewlett-Packard, Wilmington, DE), electron ionization (70 eV) with a source temperature of 210°C, a DB-5 column (J&W Scientific, Folsom, CA; 15-m × 0.25-mm, 0.25-μm film thickness), and an injection temperature of 250°C. The column temperature program was: 80°C for 1 min, raised to 320°C at a rate of 30°C min−1, and held at this temperature for 5 min. The interface temperature was 250°C and the carrier gas was He at a flow rate of 1 mL min−1 with splitless injection. BR fractions were analyzed by GC-SIM after derivatization as below. Fractions containing brassinolide, castasterone, and 6-deoxocastasterone were derivatized to bis-methaneboronate, and fractions of teasterone, typhasterol, 6-deoxoteasterone, and 6-deoxotyphasterol were derivatized to methaneboronate-trimethylsilyl ether. Monitored ions in the analysis of each BR were as follows: brassinolide, m/z 534, 528, 338, 332, 161, and 155; castasterone, m/z 518, 512, 287, 161, and 155; typhasterol and teasterone, m/z 550, 544, 535, and 529; 6-deoxocastasterone, m/z 504, 498, 489, and 483; and 6-deoxotyphasterol and 6-deoxoteasterone, m/z 536, 530, 521, 515, and 215. The endogenous levels of BRs, except for brassinolide, were determined as the ratio of the peak area of molecular ions for the internal standard to that of the endogenous steroid. The endogenous levels of brassinolide were determined as the ratio of the peak areas of fragment ions of m/z 338 and m/z 332. Sterols were analyzed by full-scan GC-MS after derivatization to the trimethylsilyl ether, and the endogenous levels were determined as the ratio of the peak areas of molecular ions for the internal standard to that of the endogenous sterol. Molecular ions of the internal standard and the endogenous sterol were as follows: 24-methylenecholesterol,m/z 477 and 470; campesterol, m/z 478 and 472; and campestanol, m/z 480 and 474. RESULTS Identification and Morphological Characterization of Sevenbri1 Mutants We have screened plants mutagenized by T-DNA insertional mutagenesis and EMS for dwarf mutants displaying the typical characteristics of BR biosynthetic mutants. These characteristics include short stature, small, dark green leaves, and reduced fertility. These mutants were divided into two categories based on the responses of the inflorescences to exogenous application of brassinolide. Nine mutants with no response or a reduced response in these feeding experiments were mapped to see if they were linked tobri1-1, a previously identified brassinolide-insensitive mutant. For bri1-3, zero of 44 recombinants were detected with nga1107; for bri1-5, two of 88 recombinant chromosomes were detected with nga1107 and zero of 88 with DHS1; and forbri1-6, two of 108 recombinants were detected with nga1107. For bri1-7 and bri1-9, no recombinants for nga1107 were detected out of 32 chromosomes for each allele.bri1-4 and bri1-5 were also crossed tobri1-1, and the resulting F1generation were dwarf, indicating that these mutants were alleles ofbri1. The results of the mapping and complementation tests were supported by the finding of a unique DNA sequence alteration in each of our seven bri1 alleles within the BRI1coding region (Table I). The two other mutants did not map to the same location as bri1 and will be described elsewhere (S. Choe, F.E. Tax, and K.A. Feldmann, unpublished data). These seven bri1 mutants can be divided into severe, intermediate, and weak alleles based on their morphological characteristics. bri1-3 and bri1-4 represent severe alleles. bri1-4 plants, as shown in Figure1 and Table II, are extremely small, with all major above-ground organs reduced in size and dark-green in color, and rarely produce seeds. Plants of the intermediate allelebri1-8 are slightly larger than bri1-3 andbri1-4 plants, and are more fertile (data not shown). Four alleles comprise the weak class of bri1 alleles;bri1-5 and bri1-6 shown in Figure 1 are typical of this class. Plants from the four weak alleles were between 3 and 6 cm in height at 5 weeks of age, resemble the wild type in color, and are reasonably fertile, although not as fertile as the wild type (TableII). There are some interesting differences among these weak alleles. For example, bri1-5 plants have very short internodes along the inflorescence, develop leaves that are wider than those of the wild type (data for the third and fourth leaves are shown in Table II), and are the most fertile of these bri1 mutants. In contrast,bri1-9 plants have the longest internode length of these four weak alleles, yet have the smallest rosette leaves (data not shown). Fig. 1. Open in new tabDownload slide Wild type and four representative alleles ofbri1 at 5 weeks of age. A, Wild type; B,bri1-3; C, bri1-4; D,bri1-5; and E, bri1-6. Fig. 1. Open in new tabDownload slide Wild type and four representative alleles ofbri1 at 5 weeks of age. A, Wild type; B,bri1-3; C, bri1-4; D,bri1-5; and E, bri1-6. DNA Sequence Analysis of bri1 Mutants To determine the specific DNA sequence alterations responsible for the phenotypes of these bri1 alleles, we amplified the coding region of BRI1 from these mutants using PCR and performed DNA sequence analysis on each allele. For each allele, a single alteration was identified within the sequenced region.bri1-3 and bri1-4, the most severe alleles, both contain small deletions that are predicted to alter the BRI1ORF, resulting in a premature stop codon. Deletions of this size are common in untagged mutations resulting from T-DNA mutagenesis (S. Choe and K.A. Feldmann, unpublished results). The location of the deletions in these two severe alleles may indicate that these represent null alleles of bri1 (see Fig. 2). Fig. 2. Open in new tabDownload slide Schematic of the BRI1 locus including the positions of the bri1 mutations. aa, Amino acid. Fig. 2. Open in new tabDownload slide Schematic of the BRI1 locus including the positions of the bri1 mutations. aa, Amino acid. The intermediate and weak alleles each contained a single base pair change resulting in an amino acid substitution. These mutations were distributed throughout the BRI1 gene: four mutations were located in the extracellular domains of BRI1, and one (bri1-8) altered a conserved residue in the intracellular kinase domain. Two mutations (bri1-6 and bri1-7)were changes of Gly in a region located between two LRR domains. This island has been postulated to be a ligand-binding domain (Li and Chory, 1997), and these two mutations result in weak alleles. The remaining two mutations include a change of a Cys near the amino terminus and a mutation in a LRR just carboxy-terminal of the island. Interestingly, all of the missense mutations we have identified in the extracellular domain are weak alleles. Quantitative Analysis of Endogenous Sterols and BRs Quantitative analysis was performed to determine the BR and sterol levels in several bri1 alleles.2H-labeled BRs and sterols were used as internal standards to determine the endogenous levels of BRs and sterols. Plant materials were from 5-week-old plants of the null allelebri1-4, the two weaker alleles bri1-5 andbri1-6, and corresponding wild-type plants. Thebri1-4 and bri1-5 mutations are in the Ws-2 background and the bri1-6 mutation is in the Enkheim-2 (En-2) background. First, bri1-4 and bri1-5 (Ws-2 background) were examined, and the results are shown in Figure3. The endogenous levels of sterols such as 24-methylenecholesterol, campesterol, and campestanol in both alleles were comparable to those of wild type (Ws-2). The levels of 6-deoxo-BRs such as 6-deoxotyphasterol and 6-deoxocastasterone inbri1-4 were only slightly higher than those of the wild type. In bri1-5, the levels of 6-deoxoBRs were comparable to those of the wild type. However, striking differences were observed in the levels of 6-oxoBRs. The levels of brassinolide, castasterone, and typhasterol in bri1-4 were 57-, 128-, and 33-fold higher, respectively, than those of wild-type (Ws-2) plants (Fig. 3). The level of teasterone in bri1-4 was significantly higher than that of wild type. In addition, the weaker allele, bri1-5, accumulated considerable levels of brassinolide, castasterone, and typhasterol (22-, 51-, and 17-fold, respectively). Fig. 3. Open in new tabDownload slide The proposed brassinolide biosynthetic pathway and the quantification of endogenous sterols and BRs frombri1-4 (a null allele), bri1-5 (a weaker allele), and wild type (Ws-2). Values in top, middle, and bottom represent endogenous levels (per gram fresh weight) inbri1-4, bri1-5, and the wild type, respectively. Most of the data for the wild type have been already published (Choe et al., 1999b). Data quantifying teasterone were not available in our previous study because of low recovery. In this study, we repeated the analysis using the same plant materials. Endogenous teasterone was not detected, while recovery of the internal standard ([2H6]teasterone) was very good. Fig. 3. Open in new tabDownload slide The proposed brassinolide biosynthetic pathway and the quantification of endogenous sterols and BRs frombri1-4 (a null allele), bri1-5 (a weaker allele), and wild type (Ws-2). Values in top, middle, and bottom represent endogenous levels (per gram fresh weight) inbri1-4, bri1-5, and the wild type, respectively. Most of the data for the wild type have been already published (Choe et al., 1999b). Data quantifying teasterone were not available in our previous study because of low recovery. In this study, we repeated the analysis using the same plant materials. Endogenous teasterone was not detected, while recovery of the internal standard ([2H6]teasterone) was very good. To further confirm our findings, endogenous sterols and BRs were examined from 5-week-old plants of another weak allele,bri1-6, which is in a different background frombri1-5. Both bri1-5 and bri1-6 show similar phenotypes, although they are in different backgrounds (see Fig. 1). There was no significant difference in sterol and BR levels in Ws-2 and En-2 (see Fig. 3; Table III).bri1-6 contained the same pattern of sterols and BRs asbri1-5. bri1-6 also accumulated brassinolide, castasterone, and typhasterol (26-, 70-, and 10-fold higher than those of wild-type [En-2] plants, respectively). The accumulation of brassinolide and castasterone appears to be related to BRI1 gene dosage. However, the levels of 6-deoxoBRs such as 6-deoxocastasterone, 6-deoxotyphasterol, and 6-deoxoteasterone inbri1-6 were comparable to wild type. Our findings inbri1-4, bri1-5, and Ws-2 were confirmed in an allele isolated from a different ecotypic background. Table III. Endogenous levels of sterols and BRs in bri1-6 and wild-type plants (En-2) Sterol/BR . En-2 . bri1-6 . μg/g fresh wt Sterol  24-Methylenecholesterol  3.5  2.3  Campesterol 40 31  Campestanol  0.63  0.52 ng/g fresh wt BR  6-Deoxoteasterone  0.08  0.05  6-Deoxotyphasterol  3.0  1.7  6-Deoxocastasterone  2.1  2.7  Teasterone  0.07  0.08  Typhasterol  0.26  2.5  Castasterone  0.23 16.1  Brassinolide  0.10  2.6 Sterol/BR . En-2 . bri1-6 . μg/g fresh wt Sterol  24-Methylenecholesterol  3.5  2.3  Campesterol 40 31  Campestanol  0.63  0.52 ng/g fresh wt BR  6-Deoxoteasterone  0.08  0.05  6-Deoxotyphasterol  3.0  1.7  6-Deoxocastasterone  2.1  2.7  Teasterone  0.07  0.08  Typhasterol  0.26  2.5  Castasterone  0.23 16.1  Brassinolide  0.10  2.6 Open in new tab Table III. Endogenous levels of sterols and BRs in bri1-6 and wild-type plants (En-2) Sterol/BR . En-2 . bri1-6 . μg/g fresh wt Sterol  24-Methylenecholesterol  3.5  2.3  Campesterol 40 31  Campestanol  0.63  0.52 ng/g fresh wt BR  6-Deoxoteasterone  0.08  0.05  6-Deoxotyphasterol  3.0  1.7  6-Deoxocastasterone  2.1  2.7  Teasterone  0.07  0.08  Typhasterol  0.26  2.5  Castasterone  0.23 16.1  Brassinolide  0.10  2.6 Sterol/BR . En-2 . bri1-6 . μg/g fresh wt Sterol  24-Methylenecholesterol  3.5  2.3  Campesterol 40 31  Campestanol  0.63  0.52 ng/g fresh wt BR  6-Deoxoteasterone  0.08  0.05  6-Deoxotyphasterol  3.0  1.7  6-Deoxocastasterone  2.1  2.7  Teasterone  0.07  0.08  Typhasterol  0.26  2.5  Castasterone  0.23 16.1  Brassinolide  0.10  2.6 Open in new tab DISCUSSION Theoretically, hormone-insensitive mutants can be predicted to show the same phenotype as hormone-deficient mutants. In fact,bri1 mutants are dwarfed and the phenotypes are similar to BR-deficient mutants such as cpd, det2,dwf1, and dwf7/ste1. The one major difference is that the dwarfism and other growth characteristics of bri1mutants are not rescued by BRs. The bri1-1 phenotype is the result of a recessive mutation in a gene located on chromosome IV (Clouse et al., 1996). The gene affected in BR-insensitive mutants was isolated and shown to encode a putative membrane-bound LRR-RK (Li and Chory, 1997). From these molecular and genetic studies, BRI1has been suggested to encode a BR receptor or an essential component involved in BR signaling. We have described the isolation of seven additional bri1 alleles ranging from nulls to weak alleles, and report that BRI1 plays a role in the homeostasis of BRs. Isolation and Characterization of a Broad Spectrum ofbri1 Alleles The bri1 alleles described in this study possess phenotypes that range from severe dwarf mutants resembling the originalbri1-1 dwarf mutant (Clouse et al., 1996) to semi-dwarfs. Molecular analysis indicated that the two severe alleles,bri1-3 and bri1-4, contain small deletions predicted to cause a frame shift, and introduce a premature stop codon into BRI1. The positions of these deletions, in the fourth LRR and in the second of 11 conserved kinase subdomains, predict that these two severe alleles should produce truncated forms ofBRI1. The two severe bri1 alleles are similar tocpd mutants in their overall size and morphology.cpd and bri1 mutants are the smallest dwarfs of the eight BR dwarf loci isolated thus far (Clouse and Feldmann, 1999). Based on the description of the 18 mutants isolated by Li and Chory (1997) and the observation that the infertility of bri1mutants is positively correlated with their severely reduced stature (see Table II), 17 of these are likely also severe mutants, but little morphological description of these mutants has been presented. The intermediate and weak bri1 alleles we have isolated have at least a 7-fold reduction in plant height compared with their respective wild type, and thus are still classified as dwarf mutants. These resemble loss-of-function alleles of dwf1 ordwf7/ste1 in their overall morphology (Feldmann et al., 1989; Choe et al., 1999). Most morphological parameters of the plant are altered proportionally in these mutants (see Fig. 1; Table II), but there are some exceptions. Leaves from bri1-5 mutants are wider than the wild type, and the internode distance inbri1-5 mutants is short compared with other bri1weak alleles such as bri1-7 (see Fig. 1; Table II), even though bri1-5 plants are taller than bri1-7plants. bri1-9 mutants, which have small, rosette leaves, have longer inflorescences and longer internodes than would be predicted from the size of the leaves (data not shown). The five intermediate and weak mutants generated by EMS mutagenesis each had a single base change leading to an amino acid substitution within the BRI1 coding region. These mutations are dispersed throughout the BRI1 protein, both within the kinase domain and in different domains of the extracellular domain. The intermediate allele, bri1-8, is caused by a change of a conserved Arg in subdomain VIa that is present in many RKs identified to date (Walker 1993; Li and Chory, 1997). However, the phenotype of bri1-8is not as strong as would be expected for a change in such a conserved residue. The extracellular region of BRI1 is composed of several different domains, including a putative leucine zipper, two sets of paired Cys residues, 25 LRRs, and a domain nestled within the LRRs called the island domain. Two weak alleles (bri1-6 andbri1-7) contain mutations that change different Gly residues within this island domain. A mutant with a change of a different Gly within this island domain was reported by Li and Chory (1997); however, their mutant was not fertile and therefore was probably a severe allele. Two other mutations were identified in the extracellular domain: a change in a Cys to a Tyr in the paired Cys domains located in the amino terminus of the extracellular domain (bri1-5; see Fig. 2), and a change in a Ser to a Phe in the first LRR after the island domain (bri1-9). These two weak alleles are the first mutations reported in the extracellular regions of BRI1 not in the island domain. Quantitative Analysis of BRs in bri1 Mutants At the biochemical level, several dwarf mutants of Arabidopsis have been shown to be blocked in specific steps of the BR-biosynthetic pathway. In addition, the endogenous BRs present in each mutant have been shown to be dependent on the positions of the genetic blocks in the pathway (det2: Fujioka et al., 1997; Noguchi et al., 1999; dwf1: Klahre et al., 1998; Choe et al., 1999a;dwf7/ste1: Choe et al., 1999b). bri1 mutants have phenotypes that mimic those of BR biosynthetic mutants, butbri1 mutants do not respond significantly to BRs. This raises the possibility that bri1 is either a BR-receptor mutant or a mutant acting in a downstream step. If so, bri1mutants should contain the same endogenous BRs found in wild type, but they may accumulate BRs, especially castasterone and brassinolide. Our data show that both of the above predictions are correct. All BRs found in the wild type are present in bri1. Brassinolide, the presumptive terminal biologically active BR for the growth of Arabidopsis, accumulates in bri1 mutants. The ratio of brassinolide for a null allele (bri1-4), a weaker allele (bri1-5), and wild type (Ws-2) was 57:22:1. Thus, the accumulation of brassinolide is related to the amount of functional BRI1. A similar trend was observed for typhasterol and castasterone, the biosynthetic precursors of brassinolide. The ratios were 33:17:1 and 128:51:1, respectively. The pattern and accumulation of BRs suggests that the BRI1 gene controls either a step associated with the binding of brassinolide to a receptor or a subsequent downstream step. The presence of abnormally high levels of brassinolide and castasterone in BR-insensitive dwarf mutants indicates a link between BR biosynthesis and BR action. In mutants with impaired response to BR, BR biosynthesis might be normal or activated. Our preliminary study suggested that the abundance of transcripts of biosynthesis enzymes might be increased in bri1 mutants (S. Choe and K.A. Feldmann, unpublished data). Detailed northern analyses and metabolic studies will answer the question of whether BR biosynthesis inbri1 mutants is activated or not. Models for the Regulation of BR Synthesis and Signaling End-product feedback regulation of biosynthetic genes is common among plant hormones. For example, treatment of plants by exogenous ethylene inhibits further ethylene production (Yang and Hoffman, 1984). For GA and BRs, there is also evidence for negative regulation after application of hormone, and a mechanism involving regulation at the transcriptional level has also been demonstrated (Phillips et al., 1995; Mathur et al., 1998). For example, treatment with specific BR intermediates or end products such as brassinolide and castasterone caused a reduction in the mRNA levels of the CPD gene, a gene required for necessary structural modification of brassinolide. Interestingly, this repression is inhibited by cycloheximide, indicating that at least one factor needs to be newly synthesized for this repression to occur (Mathur et al., 1998). A second theme in these hormone biosynthetic and signaling pathways is that mutants deficient in receptors or other downstream components fail to regulate hormone levels. For example, the etr1-1 mutant in Arabidopsis lacks the ability to suppress ethylene synthesis after treatment with exogenous ethylene, although normal amounts of ethylene are made in untreated plants (Bleecker et al., 1988). This suggests that feedback regulation operates through a functional signaling pathway. Accumulation of bioactive GAs has also been observed in several mutants that are insensitive to GA treatment (Fujioka et al., 1988; Talon et al., 1990; Appleford and Lenton, 1991). Of these three GA insensitive genes in various plant species, only the molecular nature of the GAI gene is known, and it encodes a product related to transcription factors (Peng et al., 1997). In thegai mutant, there is also an increase in the transcription of GA5, a key GA-biosynthetic gene, indicating, as with BRs, that there is feedback regulation at the transcriptional level. The results presented here indicate that the BRI1 gene is required for feedback regulation of BR biosynthesis. There is substantial accumulation of brassinolide and other intermediates in both a null allele and two weak alleles of bri1. One possibility is that BRI1, which encodes a predicted transmembrane RK, phosphorylates cellular proteins that directly or indirectly regulate the activity or transcription of BR-biosynthetic proteins such as CPD. Alternatively, BRI1 phosphorylation could result in the transcription and/or translation of a repressor of BR biosynthesis, as has been proposed for the regulation of CPD (Mathur et al., 1998). Interestingly, the lka mutant from pea, which is BR insensitive, also accumulates bioactive BRs (Nomura et al., 1997,1999). However, the degree of the accumulation is not so high in thelka mutant. Presumably, the role of LKA in BR signaling pathway could be different from that of BRI1, or lka may be a weak allele of a pea BRI1 homolog. Molecular characterization of theLKA gene will provide important information about its role in the BR signaling pathway. ACKNOWLEDGMENTS We thank Steve Clouse for sending seeds of bri1-1 and the Arabidopsis Biological Resource Center at Ohio State University for supplying CS399 (bri1-6) seeds. We thank Alice Traut, Amanda Ross, and Brian Gregory for technical assistance with the mutant isolation and mapping. 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Trends Plant Sci 2 1997 137 143 Google Scholar Crossref Search ADS WorldCat 55 Yokota T Nomura T Kitasaka Y Takatsuto S Reid JB Biosynthetic lesions in brassinosteroid-deficient pea mutants. The 24th Proceedings of Plant Growth Regulation Society of America, Atlanta, Georgia, August, 1997. Latimer JG 1997 The Plant Growth Regulation Society of America LaGrange , GA, p 94 Author notes 1 This work was supported by a Grant-in-Aid for Scientific Research (B) from the Ministry of Education, Science, Sports and Culture of Japan (grant no. 10460050 to S.F.), by the National Science Foundation (grant no. 9604439 to K.A.F.), and by the U.S. Department of Agriculture (grant no. 97–353044708 to F.E.T.). * Corresponding author; e-mail [email protected]; fax 81–48–462–4674. Copyright © 1999 American Society of Plant Physiologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Leng, Qiang; Mercier, Richard W.; Yao, Weizhe; Berkowitz, Gerald A.

doi: 10.1104/pp.121.3.753pmid: 10557223

Abstract Cyclic nucleotide-gated (cng) non-selective cation channels have been cloned from a number of animal systems. These channels are characterized by direct gating upon cAMP or cGMP binding to the intracellular portion of the channel protein, which leads to an increase in channel conductance. Animal cng channels are involved in signal transduction systems; they translate stimulus-induced changes in cytosolic cyclic nucleotide into altered cell membrane potential and/or cation flux as part of a signal cascade pathway. Putative plant homologs of animal cng channels have been identified. However, functional characterization (i.e. demonstration of cyclic-nucleotide-dependent ion currents) of a plant cng channel has not yet been accomplished. We report the cloning and first functional characterization of a plant member of this family of ion channels. The Arabidopsis cDNA AtCNGC2 encodes a polypeptide with deduced homology to the α-subunit of animal channels, and facilitates cyclic nucleotide-dependent cation currents upon expression in a number of heterologous systems. AtCNGC2 expression in a yeast mutant lacking a low-affinity K+ uptake system complements growth inhibition only when lipophilic cyclic nucleotides are present in the culture medium. Voltage clamp analysis indicates that Xenopus laevis oocytes injected with AtCNGC2 cRNA demonstrate cyclic-nucleotide-dependent, inward-rectifying K+ currents. Human embryonic kidney cells (HEK293) transfected with AtCNGC2 cDNA demonstrate increased permeability to Ca2+ only in the presence of lipophilic cyclic nucleotides. The evidence presented here supports the functional classification of AtCNGC2 as a cyclic-nucleotide-gated cation channel, and presents the first direct evidence (to our knowledge) identifying a plant member of this ion channel family. Cyclic nucleotides (cAMP and cGMP) are important (secondary) signaling molecules in both eukaryote and prokaryote cells (Reggiani, 1997). They are typically involved in sensing extracellular stimuli and the transduction of the signal into altered metabolic responses (Zagotta and Siegelbaum, 1996; Reggiani, 1997). Cyclic nucleotide involvement in sensory perception, at least in animal systems, often occurs through the action of cell-membrane-localized cyclic nucleotide-gated (cng), non-selective cation channel proteins (Zagotta and Siegelbaum, 1996). Cng channels involved in light (i.e. in rod and cone cells), taste (gustatory receptors), and smell (olfactory receptors) perception and in chemotaxis (in sperm) have been recently cloned from a variety of animal systems (Goulding et al., 1992; Bonigk et al., 1993; Weyand et al., 1994; Misaka et al., 1997). These cDNAs encode pore-forming (i.e. α) subunits of channel proteins that facilitate the conductance of cations (typically K+, Ca2+, and Na+) across cell membranes upon the direct binding of cAMP or cGMP to the intracellular portion of the polypeptide (Zagotta and Siegelbaum, 1996). Cng cation channel α-subunits share some sequence homology and secondary structure similarity with α-subunits of animal voltage-gated outward-rectifying K+-selective ion channel (Shaker) proteins. Like Shaker α-subunits, cng-gated cation channels cloned to date from animal systems have six membrane-spanning regions and a P (pore) region, with intracellular hydrophilic N and C termini (Zagotta and Siegelbaum, 1996). The pore region of (both animal and plant) K+-selective voltage-gated channels retains a highly conserved signature pore sequence that determines K+ selectivity (Heginbotham et al., 1994; Ketchum and Slayman, 1996). The pore region of cng-gated channels (which, as noted above, do not display K+ selectivity) retains some but not all of the K+ channel signature pore sequence. Cng channels are defined functionally as ligand-gated channels that are activated by ligand (cyclic nucleotide) binding to the channel protein. Conductance facilitated by some plant (Hoshi, 1995) and animal (Bruggemann et al., 1993) voltage-gated K+channels is also affected by cyclic nucleotide, but in a different manner. In this case, the rectified conductance of channels is activated by voltage, but direct binding of cyclic nucleotide to the protein modulates the voltage:current relationship. Plant homologs (KAT1, AKT1, and KST1) of animal Shaker K+channels that have been cloned and functionally characterized have cyclic-nucleotide-binding sites (Anderson et al., 1992; Sentenac et al., 1992; Muller-Rober et al., 1995). However, these channels are structurally and functionally distinct from animal cng channels. Binding of cyclic nucleotide to this class of channels results in a reduction of current at a given voltage, but voltage is the primary determinant of conductance (Hoshi, 1995). Cytosolic cyclic nucleotides are also known to modulate the conductance of other classes of K+-selective channels, but in an indirect fashion (Zagotta and Siegelbaum, 1996). In this third category of cyclic nucleotide regulation of channel function, the effect is mediated by cyclic-nucleotide-dependent protein kinase phosphorylation of the ion channel. Cyclic nucleotide binding to protein kinase allows for kinase-dependent phosphorylation of some K+ channels; cyclic-nucleotide-dependent phosphorylation modulates conductance of the channel (Rudy et al., 1991; Wang and Giebisch, 1991). Assmann and co-workers (Li et al., 1994) have demonstrated that K+ currents across some native plant cell membranes are modulated by cAMP-dependent protein kinase phosphorylation of the channel. Preliminary reports (Kamasani et al., 1997) indicated that induced currents upon expression of the plant K+ channel KAT1 in X. laevis oocytes are inhibited by kinase-dependent phosphorylation. Understanding the molecular basis for regulation of cation transport across animal cell membranes has been facilitated by the cloning and functional characterization of cDNAs encoding ion channels representing many different protein families. This work has led to a rather complex picture of diverse channel families with different functions (i.e. involvement in action potentials, maintenance of membrane potential, ion transport, and sensory signal transduction, etc.) in animal systems. Our understanding of the involvement of ion channels in plant cell function is just beginning, and will be advanced by the ongoing discovery of new classes of plant ion channels and the subsequent characterization of the mechanisms regulating ion conductance facilitated by these transport proteins. cDNAs have been cloned from plants which encode inward-rectifying (Schachtman et al., 1992;Sentenac et al., 1992; Muller-Rober et al., 1995) and outward-rectifying (Czempinski et al., 1997; Gaymard et al., 1998) voltage-gated K+ channels. However, no cDNA has yet been cloned from plants that encodes a protein functionally demonstrated to be a member of the cng ion channel family of proteins. The objective of the work described in this report was to undertake this effort. MATERIALS AND METHODS Isolation of AtCNGC2 The Arabidopsis expressed sequence tag (EST) database (dbEST) was screened using an animal cng channel sequence (the chick cone photoreceptor; GenBank accession no. X89598). An EST (stock no. 38D12T7; GenBank accession nos. T04542 and T13368) encoding a partial-length putative cng channel was obtained from the DNA Stock Center at the Arabidopsis Biological Resource Center (The Ohio State University, Columbus). A cDNA library CD 4-7 obtained from the Arabidopsis Biological Resource Center (D'Alessio et al., 1992) was screened using the EST 38D12T7 sequence as a probe. Several positive clones were identified. One clone was sequenced to completion utilizing a DNA sequencing system (Silver Sequence, Promega, Madison, WI) and was shown to encode a full-length putative plant cng cation channel called AtCNGC2. The plasmid construct is labeled pZL-cngc. All standard molecular biology procedures for library screening, subcloning, and DNA sequencing were performed essentially as described in Ausubel et al. (1987). All enzymes were obtained from Gibco-BRL (Cleveland) unless otherwise noted. Computational Analysis The Expressed Sequence Tag (EST) database (dbEST) and the non-redundant (NR) database at GenBank were screened using BLAST software through the Internet (Madden et al., 1996). Other sequence databases utilized include: The Institute for Genomic Research (TIGR) and the University of Minnesota-EST Arabidopsis database. DNA sequence analyses were undertaken using the Genetics Computer Group (GCG) Version 9.1 software package (Madison, WI) run on an open VMS workstation at The University of Connecticut Biotechnology Center (Telnet address: clone3.mcb.uconn.edu). The computer-assisted hydropathy plots were done using the program RAOARGOSfrom the PC/Gene Computational Software Package (IntelliGenetics, University of Geneva) under a DOS environment. The sequence alignment analysis was undertaken using Clustal W 1.7 software (Higgins and Sharp, 1988) running under the University of Connecticut UNIX workstation. Expression of cDNA and Synthesis of cRNA Encoding AtCNGC2 The cDNA encoding AtCNGC2 was subcloned into the yeast (Saccharomyces cerevisiae) expression vector pYES2 (Invitrogen, Carlsbad, CA). The resultant plasmid, labeled pYES-cngc, was used for the yeast transformation and subsequent complementation experiments. The plasmid construct pZL-cngc, containing AtCNGC2, was used to generate full-length sense RNA encoding methylated, capped runoff transcripts following the protocol and procedures outlined in the Epicentre AmpliScribe T7 High Yield Transcription Kit manual (Epicentre Technologies, Madison, WI). The resultant purified sample was used directly for injection into oocytes (50 nL/oocyte containing 50 ng cRNA). The cDNA encoding AtCNGC2 was subcloned into the mammalian expression vector pcDNA3 (Invitrogen). The resultant plasmid, labeled pcDNA3-cngc, was used for mammalian cell transformation and subsequent expression studies. All standard molecular biology procedures for DNA manipulation, cDNA expression, and cRNA synthesis were performed essentially as described in Ausubel et al. (1987) unless otherwise noted. All enzymes were obtained from Gibco-BRL unless otherwise noted. Yeast Complementation Studies pYES-cngc and pYES2 (empty cassette) were transformed into the K+-uptake deficient Saccharomyces cerevisiae yeast mutant strain CY162 (MATα ura3–52, trk1Δ, his3Δ200, his4–15, and trk2Δ::pck64 provided by Dr. L. Kochian, Cornell University, Ithaca, NY) following the lithium acetate transformation protocol precisely as described in Ausubel et al. (1987). Positive transformants were selected by uracil prototrophy on synthetic minimal media (SMM) (i.e. YNB agar without amino acids [BIO 101, Vista, CA] with the addition of CSM-URA [BIO 101], 100 mm KCl, and 2% [w/v] dextrose). The resultant plates were incubated at 30°C for 4 d and positive colonies isolated for subsequent experiments. To test for complementation of the trk1Δ and trk 2Δ mutation, yeast colonies (transformed with pYES-cngc or pYES2) were transferred to fresh SMM plates supplemented with 2 mm KCl (substituting 2% [w/v] Gal and 0.5% [w/v] Suc for the dextrose). The transformed yeast cultures were also plated on SMM agar (2 mm KCl, Gal, and Suc) with the addition of 10 μm dibutyryl-cAMP or dibutyryl-cGMP (Sigma, St. Louis). Complementation of K+ uptake in the yeast mutant was also assayed in liquid medium. Single colonies were isolated from appropriate plates and transferred to a liquid culture consisting of a modification of the synthetic minimal media described above (YNB formulation substituting NaI for KI and NaH2PO4 for KH2PO4, with the addition of 0.1 mm KCl). Growth rates were monitored by determiningA 595 of the developing cultures after 24 h. Absorbance measurements of the developing cultures were converted to protein concentrations from a standard curve generated using a Bradford protein assay kit (Sigma). Functional Expression in Oocytes Standard methods (Very et al., 1995) were used to express AtCNGC2 cRNA in X. laevis oocytes. For each experiment, 50 nL of water containing 50 ng of cRNA encoding AtCNGC2 (50 nL of water for controls) was injected into stage 5 X. laevis oocytes prepared and cultured by standard methods. Two-electrode voltage clamp recordings in the whole-cell configuration were performed 5 d after injection utilizing an amplifier (GeneClamp 500, Axon Instruments, Foster, CA). Voltage stimuli were generated and currents were recorded using pClamp 6.04 software (Axon Instruments) in a bath solution (similar to that used for recording KAT1 currents from oocytes; Schachtman et at., 1992) containing 96 mm KCl, 1.8 mmCaCl2, 1.8 mmMgCl2, and 10 mm HEPES-KOH, pH 7.5. Current recordings were filtered at 2 kHz. Leak currents were measured without cNMP and were subtracted from each trace (Baumann et al., 1994; Eismann et al., 1994; Crary et al., 1998). After leak currents were measured, oocytes were perfused with 10 μm dibutyryl-cNMP for 30 min prior to obtaining a second series of voltage clamp recordings (E hold = −60 mV, step voltages between +80 and −160 mV in 20-mV increments). The recording chamber was perfused at a rate of 2 mL/min with bath solutions with or without ligand. Pipettes were pulled from KIMAX-51 capillaries (KIMBLE Products, Vineland, NJ). Functional Expression in HEK293 Cells pcDNA3-cngc was co-transfected with CD8 into the human embryonic kidney cell line HEK293 (American Type Tissue Culture Collection, Rockville, MD) following an electroporation and selectivity method described in Jurman et al. (1994). The cNMP-induced rise in [Ca2+]i was measured in individual cells by fluorescence spectroscopy using the Ca2+ sensitive dye FURA-2. The components associated with the photometry system included an inverted microscope (Nikon, Tokyo), a computerized MAC2000 modular system, and a 150-W Xenon lamp (Ludl Electronic Products, New York). This system allows for single cell measurements by directing 100% of the emitted light to a photometer. The dye was excited at 340 and 380 λ with emission intensity (510 λ) measured in volts. A Ca2+imaging calibration kit with FURA-2 pentasodium salt (Molecular Probes, Eugene, OR) was used to calibrate the system, and the ratio of the two signals (340/380) was plotted as a function of [Ca2+]i according to the formula described by Grynkiewicz et al. (1985). Fluorescence imaging of cNMP-induced Ca2+ entry into AtCNGC2 transfected and control cells (non-transfected) was performed according to the method described by Baumann et al. (1994)with the following modifications. Cells were loaded with the Ca2+-sensitive fluorescent dye FURA-2 by incubation in loading buffer (120 mm NaCl, 3 mmKCl, 1 mm CaCl2, 3 mmMgCl2, 50 mm Glc, 5 mmNaOH, and 10 mm HEPES, pH 7.4) with 2 μmFURA-2/AM (Molecular Probes) for 30 min at 37°C (under 5% CO2). The experimental protocol used by Baumann et al. (1994) to demonstrate functional expression of theDrosophila retinal cng was followed to evaluate AtCNGC2 currents in this system. Dye-loaded AtCNGC2-transfected and non-transfected control cells were incubated for 30 min with 50 μm dibutyryl-cNMP in loading buffer (except that the [Mg2+] was 10 mm); this high level of divalent cation blocks cNMP-activated Ca2+ entry (Baumann et al., 1994). [Ca2+]i of HEK293 cells was monitored (cells were exposed to cNMP and Ca2+ throughout the time course of the experiment) in the presence of high [Mg2+] (blocking cNMP-activated Ca2+ currents), after perfusion in Mg2+-free loading buffer (removing the Mg2+ block of cng channel currents), and then after the block was restored by perfusion again with loading buffer containing high [Mg2+]. RESULTS AND DISCUSSION The amino acid sequence of an animal cng ion channel (the chick cone photoreceptor; GenBank accession no. X89598) was used to search the Arabidopsis EST database. Several homologous EST clones were identified. One clone (GenBank accession no. T04542) encoding a 1.2-kb insert was used to screen an Arabidopsis cDNA library (CD4–7, Ohio State University Arabidopsis Resource Center). A full-length cDNA clone corresponding to the EST probe was identified. The isolated cDNA, AtCNGC2 (GenBank accession no. AF067798), is 2,374 bp in length and harbors an open reading frame of 2,178 bp encoding a polypeptide of 726 amino acids with a predicted molecular mass of 83.3 kD. The presence of the gene encoding AtCNGC2 in the Arabidopsis genome was confirmed by Southern-blot analysis (data not shown). Another group submitted an identical sequence to the database (accession no. Y16328, Köhler and Neuhaus, 1998). The deduced amino acid sequence encoded by AtCNGC2 (Fig.1) was aligned with cng channels cloned from several plant (AtCNGC1 a second putative Arabidopsis cng clone, and HvCBT1, a barley homolog) and animal (CRET and BOLF) species. A computer-assisted hydropathy plot (results not shown) of the deduced AtCNGC2 polypeptide indicated the presences of six putative transmembrane domains. This is consistent with animal cng channels and plant inward-rectifying K+ channels (for review, see Zagotta and Siegelbaum, 1996; Maathuis et al., 1997). In addition, sequence analysis indicates that AtCNGC2 contains a potential pore-forming region with lower hydrophobicity (relative to the membrane spanning domains) between S5 and S6, as well as putative cyclic nucleotide and calmodulin-binding domains. Fig. 1. Open in new tabDownload slide Multiple amino acid sequence alignment of AtCNGC2 with several other animal and (putative) plant cng channel proteins. The amino acid sequences were aligned using the Clustal W 1.7 multiple-alignment program (Higgins and Sharp, 1988). Identical, strongly, and weakly conserved amino acid residues are denoted with asterisks (*), colons (:), and periods (.), respectively, based on the Gonnet Pam250 scoring matrix as described in the documentation provided with Clustal W 1.7. Proposed domains corresponding to the plant sequences are overlined and denoted as follows: S1 to S6 indicate the putative transmembrane domain, P indicates the pore region, CNBD indicates the cyclic-nucleotide binding domain, and CaMBS (double-overlined) indicates the calmodulin binding site as determined utilizing programs from the PC/Gene computational software package (IntelliGenetics, University of Geneva) as well as data presented in several manuscripts cited in the results section. The corresponding regions displayed in the two animal sequences are underlined and similarly annotated. GenBank accession numbers corresponding to the various peptides are as follows: AtCNGC1 from Arabidopsis (Y16327); HvCBT1 from barley (AJ002610); CRET from a chick retinal channel (X89598); and BOLF from a bovine olfactory channel (X55010). Fig. 1. Open in new tabDownload slide Multiple amino acid sequence alignment of AtCNGC2 with several other animal and (putative) plant cng channel proteins. The amino acid sequences were aligned using the Clustal W 1.7 multiple-alignment program (Higgins and Sharp, 1988). Identical, strongly, and weakly conserved amino acid residues are denoted with asterisks (*), colons (:), and periods (.), respectively, based on the Gonnet Pam250 scoring matrix as described in the documentation provided with Clustal W 1.7. Proposed domains corresponding to the plant sequences are overlined and denoted as follows: S1 to S6 indicate the putative transmembrane domain, P indicates the pore region, CNBD indicates the cyclic-nucleotide binding domain, and CaMBS (double-overlined) indicates the calmodulin binding site as determined utilizing programs from the PC/Gene computational software package (IntelliGenetics, University of Geneva) as well as data presented in several manuscripts cited in the results section. The corresponding regions displayed in the two animal sequences are underlined and similarly annotated. GenBank accession numbers corresponding to the various peptides are as follows: AtCNGC1 from Arabidopsis (Y16327); HvCBT1 from barley (AJ002610); CRET from a chick retinal channel (X89598); and BOLF from a bovine olfactory channel (X55010). The six putative transmembrane segments (S1–S6), the pore (P) region, the cyclic nucleotide-binding domain (CN), and the calmodulin-binding domain (CaM) of AtCNGC2 are shown in Figure 1. AtCNGC2 shows a relatively low overall sequence identity (22% or less) with the animal cng channels shown in Figure 1; however, the displayed sequences share significant homology in several positions restricted predominately to the C terminus. The most conserved regions are the S6 transmembrane domain, the pore region, and the cyclic nucleotide binding domain. As expected, AtCNGC2 shows greater overall identity with the putative plant cng channels (46% for AtCNGC1; 32% for HvCBT1). Moreover, the structural domains between the plant cng channels share a relatively high degree of homology. The alignment between plant and animal membrane spanning domains upstream from the pore region is significantly hampered by the fact that plant cng channels include about 50 more amino acids between S5 and S6. The putative cyclic-nucleotide-binding domain located in AtCNGC2 has structural features consistent with corresponding domains in animals cng channels (compare with Ludwig et al., 1990; Bonigk et al., 1993). A key amino acid residue associated with nucleotide binding is present as a conservative substitution in AtCNGC2. Specifically, the Asn at position D600 replaces a Glu residue in the other plant and animal sequences. Other invariant residues are distinguished with asterisks in Figure 1. Although the overall sequence identity between plant and animal CN-binding domains is low, their homology to the animal sequences is sufficient to distinguish this region as a cyclic-nucleotide-binding site. The denoted S4 domain for AtCNGC2, commonly described as the voltage-sensing region in voltage-gated K+channels, contains a number of evenly spaced basic residues indicative of that motif (Jan and Jan, 1992). However, the plant and animal cng channels have fewer positively charged residues in this domain. The putative pore region for AtCNGC2 shows significant homology to both the animal and other putative plant cng channel sequences. GYGD, the consensus sequence for K+ channels, is not present in the animal cng channels. Animal cng channels contain an acidic Glu residue shown to play a critical role in binding both monovalent and divalent cations (Eismann et al., 1994) in this region. AtCNGC1 and HvCBT1 share a GQNL in this position. The neutral Gln is characteristic of (animal) nonspecific cation channels (Kerr and Sansom, 1995). AtCNGC2 displays an ANDL (amino acid residue positions 415–418) at this aligned region; the acidic Asp distinguishes it from the other plant channels. A putative CaM binding domain for AtCNGC2 located within the proposed CN binding site is identified in Figure 1. The proposed site contains two major hydrophobic (Y647 and Y660) anchors separated by 12 amino acid residues. It also includes two minor evenly spaced hydrophobic (A650 and L654) residues and a positively charged conserved Arg (R659). This region is able to form a basic amphiphilic α-helix indicative of known CaM binding sites (O'Neil and DeGrado, 1990; Ikura et al., 1992). These features are loosely conserved in the barley sequence and the other Arabidopsis sequence shown in Figure 1. One manner in which the functional characterization of AtCNGC2 was undertaken involved heterologous expression in the K+-uptake-deficient yeast (S. cerevisiae) mutant CY162 (Gaber et al., 1988; Ko and Gaber, 1991). The Trk1 and Trk2 K+ transporter deletions in this mutant are lethal at low external [K+]; complementation of this mutation (i.e. growth at low K+) has been used to demonstrate function of a number of cloned plant K+ transporters (Anderson et al., 1992; Sentenac et al., 1992; Schachtman and Schroeder, 1994;Quintero and Blatt 1997; Schachtman et al., 1997; Fu and Luan, 1998). In the series of experiments shown in Figure2 and TableI, the ability of the AtCNGC2 translation product to facilitate K+ transport (i.e. K+ uptake into the yeast mutant) was evaluated by monitoring growth of the yeast at low external [K+]. The CY162 yeast did not grow on solid medium when transfected with the empty plasmid (pYES2) either in the presence or absence of the lipophilic cyclic nucleotide analog dibutyryl-cAMP (Fig. 2). Transfection with AtCNGC2 alone also did not complement the K+ uptake mutation. However, when lipophilic cAMP was supplied to the growth medium, transfection with AtCNGC2 did allow for growth of the mutant yeast at low external [K+]. Fig. 2. Open in new tabDownload slide Complementation (evaluated as growth on solid media in the presence of lipophilic cAMP) of K+-uptake mutation in CY162 yeast by transfection with AtCNGC2. Yeast was transformed with either plasmid containing AtCNGC2 (A and B) or empty pYES2 plasmid (C and D), and grown on solid medium containing 2 mm KCl in the presence (A and C) or absence (B and D) of 10 μm dibutyryl-cAMP. At high (100 mm) KCl, yeast transformed with either the empty plasmid or AtCNGC2 grew well in the absence or presence of cyclic nucleotide (data not shown). Pictures were taken after growth for 7 d at 30°C. This experiment was repeated twice. Fig. 2. Open in new tabDownload slide Complementation (evaluated as growth on solid media in the presence of lipophilic cAMP) of K+-uptake mutation in CY162 yeast by transfection with AtCNGC2. Yeast was transformed with either plasmid containing AtCNGC2 (A and B) or empty pYES2 plasmid (C and D), and grown on solid medium containing 2 mm KCl in the presence (A and C) or absence (B and D) of 10 μm dibutyryl-cAMP. At high (100 mm) KCl, yeast transformed with either the empty plasmid or AtCNGC2 grew well in the absence or presence of cyclic nucleotide (data not shown). Pictures were taken after growth for 7 d at 30°C. This experiment was repeated twice. Table I. Transfection with AtCNGC2 enhances growth in liquid culture of the K+-uptake-deficient yeast mutant in the presence of (lipophilic) cyclic nucleotide. CY162 yeast transfected with either pYES2 (control) or pYES-cngc were grown in liquid cultures containing 0.1 mm KCl as the sole K+ source in the absence or presence of 10 μm lipophilic (dibutyryl) analogs of cyclic nucleotides. Treatment means (shown ±se) represent four independent cultures grown under each experimental condition. Results of four independent experiments are shown. Experiment . Growth . Treatment . Control . +AtCNGC2 . % Increase . μg protein/mL  1 +cAMP 3.23  ± 0.46 22.4  ± 0.80 593  1 −cAMP 1.67  ± 0.38 2.06  ± 0.44 23  2 +cAMP 3.16  ± 0.75 18.6  ± 0.91 488  3 +cAMP 3.49  ± 0.71 21.2  ± 1.01 507  4 +cAMP 2.10  ± 0.42 21.5  ± 1.25 923  4 +cGMP 4.66  ± 0.62 19.2  ± 0.55 312 Experiment . Growth . Treatment . Control . +AtCNGC2 . % Increase . μg protein/mL  1 +cAMP 3.23  ± 0.46 22.4  ± 0.80 593  1 −cAMP 1.67  ± 0.38 2.06  ± 0.44 23  2 +cAMP 3.16  ± 0.75 18.6  ± 0.91 488  3 +cAMP 3.49  ± 0.71 21.2  ± 1.01 507  4 +cAMP 2.10  ± 0.42 21.5  ± 1.25 923  4 +cGMP 4.66  ± 0.62 19.2  ± 0.55 312 Open in new tab Table I. Transfection with AtCNGC2 enhances growth in liquid culture of the K+-uptake-deficient yeast mutant in the presence of (lipophilic) cyclic nucleotide. CY162 yeast transfected with either pYES2 (control) or pYES-cngc were grown in liquid cultures containing 0.1 mm KCl as the sole K+ source in the absence or presence of 10 μm lipophilic (dibutyryl) analogs of cyclic nucleotides. Treatment means (shown ±se) represent four independent cultures grown under each experimental condition. Results of four independent experiments are shown. Experiment . Growth . Treatment . Control . +AtCNGC2 . % Increase . μg protein/mL  1 +cAMP 3.23  ± 0.46 22.4  ± 0.80 593  1 −cAMP 1.67  ± 0.38 2.06  ± 0.44 23  2 +cAMP 3.16  ± 0.75 18.6  ± 0.91 488  3 +cAMP 3.49  ± 0.71 21.2  ± 1.01 507  4 +cAMP 2.10  ± 0.42 21.5  ± 1.25 923  4 +cGMP 4.66  ± 0.62 19.2  ± 0.55 312 Experiment . Growth . Treatment . Control . +AtCNGC2 . % Increase . μg protein/mL  1 +cAMP 3.23  ± 0.46 22.4  ± 0.80 593  1 −cAMP 1.67  ± 0.38 2.06  ± 0.44 23  2 +cAMP 3.16  ± 0.75 18.6  ± 0.91 488  3 +cAMP 3.49  ± 0.71 21.2  ± 1.01 507  4 +cAMP 2.10  ± 0.42 21.5  ± 1.25 923  4 +cGMP 4.66  ± 0.62 19.2  ± 0.55 312 Open in new tab The addition of (non-lipophilic) cAMP to the solid growth medium did not facilitate growth of the mutant yeast transfected with either AtCNGC2 or the empty plasmid (data not shown), suggesting that the cAMP binding domain of AtCNGC2 is cytosolic; animal cng channels have been shown to have this protein architecture (Zagotta and Siegelbaum, 1996). We observed that on solid medium, growth of the mutant yeast transfected with AtCNGC2 was less robust than growth that occurred when other (e.g. KAT1, data not shown) plant K+transporters were used to complement the K+uptake mutations of this yeast strain. Additional CY162 yeast complementation experiments were undertaken using liquid culture conditions (Table I). In liquid culture with low (i.e. the same as in the solid growth medium; 2 mm) [K+], we found that the CY162 yeast strain grew well without transfection, as did yeast transformed with AtCNGC2 (data not shown). Further experiments were undertaken with liquid culture made up with synthetic medium formulated such that the sole K+source provided 0.1 mm K+. Even under these conditions, the yeast mutant transfected with the empty plasmid (i.e. control) displayed a basal level of growth in liquid culture (Table I). In the absence of dibutyryl-CNMP, the mutant yeast transfected with either the empty plasmid or AtCNGC2 displayed the same level of basal growth (experiment 1 in Table I). However, as shown in four independent experiments with this liquid medium, transfection of the yeast with AtCNGC2 resulted in a significant increase in the growth rate in the presence of dibutyryl-cAMP (Table I). Growth of the yeast transfected with AtCNGC2 was found to be increased by either (lipophilic) cAMP or cGMP, indicating that AtCNGC2 is responsive to either of these cyclic nucleotides (experiment 4 in Table I). Further functional studies of AtCNGC2 were undertaken by expression of this putative plant cng channel in X. laevis oocytes. Voltage clamp studies were performed on control (water-injected) oocytes and oocytes expressing AtCNGC2 (Fig.3). No K+ currents were observed in control oocytes in the absence (data not shown) or presence of cyclic nucleotides (cNMP). In contrast to control oocytes, the addition of dibutyryl-cNMP (10 μm cAMP or cGMP) to the recording bath solution resulted in cNMP-dependent K+ currents. Moreover, depolarizing voltages resulted in no current with these oocytes, confirming that AtCNGC2 is an inwardly rectified cNMP gated ion channel. Fig. 3. Open in new tabDownload slide Voltage clamp analysis of cyclic nucleotide-activated currents in oocytes expressing AtCNGC2. Recordings were made at −60 mV holding potential and command potentials between +80 and −160 mV (in 20-mV increments) on water-injected oocytes in the presence of 10 μm dibutyryl-cAMP (○;n = 6), and on oocytes injected with AtCNGC2 cRNA in the presence of 10 μm dibutyryl-cAMP (●;n = 10) and 10 μm dibutyryl-cGMP (▾; n = 9). In the main body of the figure, current:voltage relationships (current values are shown ±se) are portrayed. In the inset, representative time-dependent currents are shown for an oocyte expressing AtCNGC2 in the presence of dibutyryl-cAMP with command potentials precisely as described above. Fig. 3. Open in new tabDownload slide Voltage clamp analysis of cyclic nucleotide-activated currents in oocytes expressing AtCNGC2. Recordings were made at −60 mV holding potential and command potentials between +80 and −160 mV (in 20-mV increments) on water-injected oocytes in the presence of 10 μm dibutyryl-cAMP (○;n = 6), and on oocytes injected with AtCNGC2 cRNA in the presence of 10 μm dibutyryl-cAMP (●;n = 10) and 10 μm dibutyryl-cGMP (▾; n = 9). In the main body of the figure, current:voltage relationships (current values are shown ±se) are portrayed. In the inset, representative time-dependent currents are shown for an oocyte expressing AtCNGC2 in the presence of dibutyryl-cAMP with command potentials precisely as described above. As pointed out by Assmann (1995), the level of cyclic nucleotide used to invoke a physiological response by plant proteins is an important consideration, due to the extremely low levels (up to approximately 1.5 μm; but the localized concentration may be greater in specific cell types or cell compartments) of cyclic nucleotides thought to be present in plant cells. It should be noted that the level of (lipophilic) cyclic nucleotide used in these experiments, and those shown in Figure 2 and Table I (i.e. 10 μm) is substantially lower than that used to characterize animal cng channels upon expression in oocytes (e.g. 0.2 mm, Kaupp et al., 1989; 2 mm, Yao et al., 1995). The concentration of lipophilic cyclic nucleotide used in our experiments was also at or below that used in other studies with plant cells to elicit ion currents across native membranes (Kurosaki et al., 1994; Kurosaki 1997;Volotovski et al., 1998) as well as numerous other physiological responses (Ichikawa et al., 1997; Reggiani, 1997, and refs. therein). We cannot know the exact cNMP level in the oocyte cytosol that is maintained in the presence of 10 μm lipophilic cAMP/cGMP in the recording bath solution. The activation by cNMP of AtCNGC2 currents can be best evaluated using a detached cell, patch/voltage clamp configuration for current recordings from oocyte membranes. We are currently setting up such a system to continue this line of investigation. Monitoring cyclic nucleotide activation of AtCNGC2 currents in this configuration would also allow for the evaluation of calmodulin/Ca2+ interaction with cyclic nucleotide gating of AtCNGC2 currents. In addition, patch clamp analyses would provide confirmation that gating of AtCNGC2 currents by cyclic nucleotide is mediated by direct binding of the ligand to the channel. Functional expression of AtCNGC2 was also performed in the human embryonic kidney cell line HEK293 using fluorescence spectroscopy. Animal cng channels are known to be both permeable to and blocked by divalent cations to different extents depending on their role in various signal transduction pathways (Zagotta and Siegelbaum, 1996).Baumann et al. (1994) used an experimental protocol making use of this effect in their functional characterization of a Drosophilacng channel. They found that high external [Mg2+] blocked inward Ca2+ currents through this cng channel. We followed a similar strategy in our functional expression of AtCNGC2 in HEK293 cells. Ca2+ permeability (i.e. increased [Ca2+]i) of AtCNGC2 transfected cells was observed only in the presence of dibutyryl-cAMP or dibutyryl-cGMP (Fig. 4). Fig. 4. Open in new tabDownload slide Cytosolic Ca2+ rise in HEK293 cells expressing AtCNGC2. A, Dibutyryl-cAMP- or dibutyryl-cGMP-activated Ca2+ influx are depicted in top two panels, respectively. The bottom panel shows the change in cytosolic [Ca2+] of non-transfected (CK) cells in the presence of cyclic nucleotide. The closed arrows designate the transition from high [Mg2+] (10 mm) to Mg2+-free perfusion buffer (the absence of Mg2+ intiates Ca2+ influx); open arrows designate the transition from Mg2+-free buffer to high-[Mg2+] buffer. B, Histogram representing peak cytosolic Ca2+ values after the addition of dibutyryl cNMP (D-cAMP n = 6; D-GMP n = 7) or membrane-impermeable cNMP (cAMP n = 5; cGMPn = 4) in AtCNGC2-transfected cells and non-transfected cells (CK) (treated with dibutyryl-cAMP;n = 7). Fig. 4. Open in new tabDownload slide Cytosolic Ca2+ rise in HEK293 cells expressing AtCNGC2. A, Dibutyryl-cAMP- or dibutyryl-cGMP-activated Ca2+ influx are depicted in top two panels, respectively. The bottom panel shows the change in cytosolic [Ca2+] of non-transfected (CK) cells in the presence of cyclic nucleotide. The closed arrows designate the transition from high [Mg2+] (10 mm) to Mg2+-free perfusion buffer (the absence of Mg2+ intiates Ca2+ influx); open arrows designate the transition from Mg2+-free buffer to high-[Mg2+] buffer. B, Histogram representing peak cytosolic Ca2+ values after the addition of dibutyryl cNMP (D-cAMP n = 6; D-GMP n = 7) or membrane-impermeable cNMP (cAMP n = 5; cGMPn = 4) in AtCNGC2-transfected cells and non-transfected cells (CK) (treated with dibutyryl-cAMP;n = 7). Upon removal of external Mg2+, [Ca2+]i of HEK293 cells expressing AtCNGC2 rose in the presence of either dibutyryl-cAMP (Fig.4A, top panel) or dibutyryl-cGMP (Fig. 4A, middle panel); no Ca2+ rise was observed in AtCNGC2 transfected cells in the absence of cNMP in the perfusion bath (Fig. 4A, bottom panel). As shown in Figure 4A, replacement of the Mg2+ block prevented further Ca2+ entry, and the action of an endogenous HEK293 cell Ca-ATPase efflux system (Baumann et al., 1994) reduced [Ca2+]i back down close to basal levels (Fig. 4A, top and middle panels). The results of a series of such experiments are summarized in Figure 4B. Similar to results (not shown) obtained when AtCNGC2 was functionally expressed in yeast and oocytes, external cNMP (i.e. cAMP and cGMP, in contrast to the lipophilic analogs) did not activate AtCNGC2 currents in HEK293 cells (Fig. 4B). Inward Ca2+ rise was not observed when HEK293 cells (non-transfected) were exposed to lipophilic cNMP (Fig. 4B). Preliminary experiments examining ion selectivity of AtCNGC2 expressed in oocytes using voltage clamp analysis (not shown) also indicated that AtCNGC2 is permeable to other monovalent cations (except, interestingly, Na+); such selectivity is not typically observed in studies of cloned animal cng channels (Zagotta and Siegelbaum, 1996). In summary, sequence analysis indicates that AtCNGC2 shows homology to animal cng channels. Our functional analyses of AtCNGC2 in a yeast mutant, X. laevis oocytes, and cultured HEK293 cells indicate that AtCNGC2 is an inwardly rectified ion channel that conducts a number of cations and is activated by internal but not external cAMP and cGMP, and that high external divalent cations block channel conductance. Sequence analysis also identified a calmodulin binding site on AtCNGC2. These are all properties shared with cloned animal cng channels. Some evidence is present in the published literature that is consistent with the presence of cng channels in plants. cNMP-dependent inward Ca2+ and K+ flux has been observed across the plant cell plasmalemma (Kurosaki et al., 1994; Kurosaki, 1997; Volotovski et al., 1998). However, these studies of native membranes did not identify a specific transport protein as mediating ion conductance, and, further, could not rule out ion channels other than members of the cng channel family. In addition, a number of studies have recently reported the cloning of plant cDNAs encoding polypeptides with sequence homology to animal cng channels. Köhler and Neuhaus (1998) reported the sequence of AtCNGC2 and another putative Arabidopsis cng channel (AtCNGC1 in Fig.1) in the Plant Gene Register (accession nos. Y16328 and Y16327, respectively). Schuurink et al. (1998) cloned a barley cDNA (HvCBT1 in Fig. 1) encoding a putative cng channel. However, ion transport functions of the translation products of these plant cDNAs were not reported. HvCBT1 was found not to complement growth of the K+ uptake yeast mutant used in our work, although the barley protein was shown to bind calmodulin (Schuurink et al., 1998). Thus, the results presented here, including demonstration of cyclic-nucleotide-dependent cation transport by AtCNGC2 in three different heterologous expression systems, to our knowledge represent the first functional characterization of a cloned plant member of the cng family of ion channels. The cytosolic secondary messengers cAMP, cGMP, calmodulin, and Ca2+, in addition to inward K+ and Ca2+ currents across the plasmalemma, are well known to be involved in numerous signal transduction pathways in plants. Our studies suggest that AtCNGC2 function in planta may be related to some of these signal transduction cascades. Further functional analysis of AtCNGC2 in heterologous expression systems focusing on the interactive effects these secondary messengers have on AtCNGC2 channel currents may provide an excellent context to extend our understanding of the molecular basis for at least some signaling pathways in plants. ACKNOWLEDGMENTS The pursuit of knowledge can be akin to drinking from a “magic” bottle of wine; the more you pour, the more that is left in the bottle. This work is fondly dedicated to the memories of two colleagues who, as plant physiologists, drank with gusto from that bottle during their too-short careers, and left us all richer and wiser from their efforts: Dr. Bruce Wasserman (Rutgers University, New Brunswick, NJ) and Dr. Richard Crain (University of Connecticut). 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Annu Rev Neurosci 19 1996 235 263 Google Scholar Crossref Search ADS PubMed WorldCat NOTE ADDED IN PROOF After submission of this manuscript, a paper appeared in print that reported the characterization of AtCNGC2, AtCNGC1, as well as sequences encoding several other (putative) Arabidopsis cng channels (Köhler et al., 1999). Functional characterization (i.e. cyclic nucleotide-dependent cation flux) of these cDNAs was not reported, although the translation products of AtCNGC1 and AtCNGC2 were demonstrated to bind calmodulin. Author notes 1 This material is based on work supported by the National Science Foundation (grant nos. MCB–9513921 and BIR–9512977) and by the Department of Energy (grant no. DE–FG02–95ER20202). This is Storrs Agricultural Experiment Station publication no. 1,886. 2 Present address: M.D. Anderson Cancer Center U–79, Section of Molecular and Cellular Biology, University of Texas, Houston, TX 77030. * Corresponding author; e-mail [email protected]; fax 860–486–0682. Copyright © 1999 American Society of Plant Physiologists This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)

Kuriyama, Hideo

doi: 10.1104/pp.121.3.763pmid: 10557224

Abstract A tracheary element (TE) is a typical example of a cell type that undergoes programmed cell death in the developmental processes of vascular plants. The loss of the selective permeability of the tonoplast, which corresponds to tonoplast disintegration, occurred after the cells commenced secondary wall thickening and played a pivotal role in the programmed cell death of TEs in a zinnia (Zinnia elegans L.) cell culture. A search for events specifically associated with the TE vacuole provided an important clue to the understanding of the cell death mechanism. The transport of fluorescein, a fluorescent organic anion, across the tonoplast declined drastically in differentiating TEs. The capacity of the vacuole to accumulate the probe was also impaired. Treatment with probenecid, an inhibitor of organic anion transport, caused rapid cell death of TEs and led to the ultimate disruption of the vacuole even in other types of cultured cells. These changes in vacuolar properties during TE development were suppressed by cycloheximide. Specific mRNA accumulation in cells cultured in a TE differentiation-inductive condition was abolished by probenecid. These results suggest that a change in vacuolar membrane permeability promotes programmed cell death in TEs. In recent years, much attention has been directed to the characterization of programmed cell death (PCD) mechanisms in plant cells (Greenberg, 1996; Jones and Dangl, 1996). As in animals, PCD is indispensable for the integral development or maintenance of various tissues and organs in multicellular plant species (Pennel and Lamb, 1997). Plant PCD occurs in leaf cells undergoing the hypersensitive response to preclude pathogen spread (e.g. Greenberg, 1997), in senescent leaf cells for the translocation of their components to other younger, growing parts (e.g. Smart, 1994), in reproductive organ cells to support fertilization or to supply nutrients for gametophyte and zygote development (e.g. Greenberg, 1996), and in root cortex cells to form aerenchyma for the efficient internal diffusion of oxygen (e.g.Justin and Armstrong, 1987). It remains unclear whether common regulatory mechanisms mediate these varied examples of PCD in plants (Fukuda, 1997b; Pennel and Lamb, 1997). In the vascular systems, PCD plays a role in the construction of conduits such as vessels and tracheids that supply water. These conductive tissues consist of dead tracheary element (TE) cells that are highly differentiated to form a rigid, waterproof structure for long-distance water transport. Differentiation into such cells requires various genetically controlled mechanisms, including those involved in PCD (e.g. Jones and Dangl, 1996; Fukuda, 1997a). Transdifferentiation of isolated zinnia (Zinnia elegans) mesophyll cells has been used to analyze TE differentiation both at the physiological and the molecular level (Fukuda and Komamine, 1980). In suspension culture, nearly one-half of all cells can be induced to become TEs semisynchronously (Fukuda and Komamine, 1980). This system very precisely reflects most of the genetic events that occur during the progression of TE differentiation in plants. For example, genes expressed prior to the initiation of secondary wall thickening (SWT) (TED2-4, CCoAMT, CAOMT, and ZePel) in cultured cells are also detected around the site of TEs of zinnia hypocotyls or roots in a temporally and spatially regulated manner (Demura and Fukuda, 1993, 1994; Ye et al., 1994; Ye and Varner, 1995; Domingo et al., 1998). Furthermore, Sato et al. (1993, 1995) characterized several peroxidases of zinnia, one of which was specifically activated in the differentiation-inductive culture. They proposed that this protein was selectively involved in lignin deposition in the secondary wall of TEs. The p48h-17 gene that encodes a Cys protease (Ye and Varner, 1996) and the ZRNaseI gene (Ye and Droste, 1996) were isolated as genes specifically expressed in the inductive condition and were demonstrated to localize around the vessels in situ. Such hydrolytic enzymes are assumed to function in the autolytic processes that produce hollow TEs (e.g. Fukuda, 1997a). The cascade of gene expression underlying the cytological phenotype of cultured TEs can represent TE differentiation in situ (Fukuda, 1996). One of the TE-associated events, vacuole disruption, is known to occur prior to heavy lignin deposition and drastic autolysis (Burgess and Linstead, 1984a). The mixing of the vacuolar contents with the cytoplasm disorganizes the whole intracellular structure. Burgess and Linstead (1984b) confirmed its consistency by comparing TEs in culture with those in situ, and it has since been widely recognized as the critical event that leads to cell death and subsequent autolysis (Fukuda, 1992; Groover et al., 1997). Various hydrolytic enzymes are synthesized for the progression of autolysis (Thelen and Northcote, 1989; Minami and Fukuda, 1995; Ye and Droste, 1996; Ye and Varner, 1996; Beers and Freeman, 1997; Fukuda, 1997a; Aoyagi et al., 1998). The enzymes are thought to accumulate in the TE vacuole until disruption of the vacuole, so that they do not begin to destroy the working molecular apparatus in the cytoplasm. The maintenance of vacuolar compartmentation should be necessary for differentiating TE cells to produce a complete set of these enzymes (Fukuda et al., 1998). If this is the case, some unknown mechanisms that bring about the disintegration of the vacuolar membrane and the release of enzymes must have a key role in the cell death program. A major question, therefore, is whether the vacuolar membrane of TEs exhibits specific visible characteristics leading to the disintegration. To address this question, a simple and efficient method to detect and quantify vacuolar malfunction and disruption in TEs is needed. In this report, the transport of a fluorescent organic anion probe across the tonoplast was assayed in cultured cells to monitor vacuolar function. A drastic change in the distribution and thus the transport kinetics of the organic anion could be observed in cells before and after the initiation of SWT. This effect was controlled by some genetic program expressed in TEs, and interestingly was very similar to that caused by probenecid, an inhibitor of organic anion transport (Cole et al., 1990; Oparka et al., 1991; Wright and Oparka, 1994). Treatment with this compound was followed by acceleration of TE cell death and by the disruption of the vacuole even in other cultured zinnia cells. Probenecid affected the accumulation of mRNA of a marker gene for SWT in culture. This intrinsic inhibitory effect on the organic anion transport was coupled with the cell death program of TE differentiation. MATERIALS AND METHODS Plant Material and Culture The first leaves of 14-d-old seedlings of zinnia (Zinnia elegans L. cv Canary Bird [Takii Shubyo, Kyoto]) were used for the isolation of mesophyll cells in suspension culture according to the method of Fukuda and Komamine (1980). All experiments were performed with cells cultured in inductive D medium that contained 0.1 mg/L α-naphthylacetic acid and 0.2 mg/L benzyladenine as hormones. The percentages of total (T), living (L), or dead (D) TEs were defined as follows: T = TEs/(TEs + other living cells) × 100, L = living TEs/(TEs + other living cells) × 100, D = dead TEs/(TEs + other living cells) × 100, where living cells mean those stained with fluorescein diacetate (FDA) so that their cytoplasm and vacuole were distinguishable. More than 500 cells were examined as one sample for the determination of these values. All assays were performed in triplicate within each experiment. Microscopy Observations were carried out with an epifluorescence microscope (model BH2-RFL-T2 or BX-50-FLA, Olympus, Tokyo) and an inverted microscope (IX 70, Olympus). To visualize the fluorescence of fluorescein, an excitation filter of 490 nm and a dichroic mirror of 500 nm were used. The filter and mirror were changed to 545 and 570 nm, respectively, for the observation of FM 4-64 fluorescence. The autofluorescence of chloroplasts was eliminated with a 460-nm barrier filter if necessary. Photographs were taken using a model PM-CBSP or PM-30 camera (Olympus) with black and white film (PREST 400, Fuji Photo Film, Tokyo), color film (Super G 400, Fuji Photo Film), or reversal film (Ektachrome 400, Kodak, Rochester, NY), and were selected to best illustrate the phenomenon described. Treatment with Reagents and Inhibitors FDA (Aldrich, Milwaukee, WI) and FM 4-64 (Molecular Probes, Eugene, OR) were added to the medium of each sample to a final concentration of 0.1 and 1 μg/mL, respectively. Probenecid (Wako Pure Chemical Industry, Osaka) was used at 100 μm unless otherwise specified. Cycloheximide (CHX) (Wako Pure Chemical Industry) was used at a concentration of 50 μm based on previous results (Fukuda and Komamine, 1983). The effects of the chemicals were assessed by microscopic analyses. For the determination of the number of cells excluding fluorescein from their vacuole, cells were subjected to a further 1-h incubation with FDA at the same temperature of the culture condition before observation. RNA Gel-Blot Analysis Zinnia cells were lysed by snap-freezing and subsequent incubation with phenol in buffer (200 mm Tris-Cl, 100 mm NaCl, and 10 mmEDTA, pH 8.0) in immediate vigorous vortexing. Total RNA was isolated by mixing the lysate with an equal volume of extraction buffer (50 mm Tris-Cl, 300 mm NaCl, and 5 mm EDTA, pH 8.0) followed by the microcentrifugation and precipitation of the contents in aqueous phase with ethanol. Remnant DNA was removed by RNA precipitation with one-third volume of 10 m LiCl at 4°C. RNA gel-blot analyses were carried out according to the method of Sambrook et al. (1989). The TED3 and ZCP4 probes were generated by PCR and labeled with digoxigenin-11-dUTP (Boehringer Mannheim, Basel) using cloned cDNAs as templates (Demura and Fukuda, 1993; Yamamoto et al., 1997). For signal detection, blocking reagent, anti-digoxigenin Fab fragment, and CDP-Star were used according to the manufacturer's instructions (Boehringer Mannheim). Signals were recorded by the exposure of membranes to radiographic films (Hyperfilm ECL, Amersham International, Buckinghamshire, Little Chalfont, UK). RESULTS Selective Permeability of the Tonoplast in Differentiating TEs TEs are highly specialized, mortal cells that form the rigid, lignified secondary wall. Electron microscopic observations by Burgess and Linstead (1984a, 1984b) showed that there are many cells with both SWT and apparently intact subcellular components, indicating that TEs can survive for a certain period after the initiation of SWT. Since cells within the secondary wall always die, PCD has already been or is being expressed at this stage (Fukuda, 1997a). To analyze the kinetics of living TEs and to determine their longevity in zinnia cell culture, FDA, a vital dye, was applied. FDA is de-esterified in living cells to become fluorescein that can fluoresce. When FDA was added to culture at 55 h, three characteristic staining patterns of differentiating TE cells could be observed (Fig. 1A). In contrast with other intact cells, stained TEs could be categorized into those with green fluorescence in the cytoplasm (Fig. 1A, b, t1), those with yellow fluorescence in the whole cell (Fig. 1A, d, t2), and those with no fluorescence (Fig. 1A, b, t3), which correspond to living, dying, and dead TEs, respectively. These characteristics were designated as type 1 (Fig. 1A, b, t1), type 2 (Fig. 1A, d, t2), and type 3 (Fig. 1A, b, t3) for further analyses. FDA can diffuse across the plasma membrane, causing cytoplasmic fluorescence to appear immediately (Fig. 1A), whereas, in general, vacuole staining occurs very gradually (e.g. Yoshida, 1994) due to the physicochemical properties of the dye and membrane lipids (see “Discussion”). TEs immediately filled with yellow fluorescence (Fig. 1A, d, t2) indicate the lack of a functional vacuole in these cells. Fig. 1. Open in new tabDownload slide Light (a, c, and e) and fluorescence (b, d, and f) images of zinnia cells stained with FDA, some of which are differentiating into TEs. A, The staining patterns of TEs were categorized into living type 1 (t1, a and b), dying type 2 (t2, c and d), and dead type 3 (t3, a and b). After a 1-h incubation with FDA, fluorescein was transported to the vacuole of non-TE cells but remained in the cytoplasm of TEs (e and f). co, Cell that either has not yet differentiated or will never differentiate into a TE; cy, cytoplasm; va, vacuole. B, After a further 2-h incubation, non-TE cells (nte) sequestered fluorescein completely in their vacuole (a and b) but many TE cells (te) still excluded the dye (a). The red autofluorescence of the chloroplasts in the cytoplasm is visible in the non-TE cell. The bars indicate 20 μm. Fig. 1. Open in new tabDownload slide Light (a, c, and e) and fluorescence (b, d, and f) images of zinnia cells stained with FDA, some of which are differentiating into TEs. A, The staining patterns of TEs were categorized into living type 1 (t1, a and b), dying type 2 (t2, c and d), and dead type 3 (t3, a and b). After a 1-h incubation with FDA, fluorescein was transported to the vacuole of non-TE cells but remained in the cytoplasm of TEs (e and f). co, Cell that either has not yet differentiated or will never differentiate into a TE; cy, cytoplasm; va, vacuole. B, After a further 2-h incubation, non-TE cells (nte) sequestered fluorescein completely in their vacuole (a and b) but many TE cells (te) still excluded the dye (a). The red autofluorescence of the chloroplasts in the cytoplasm is visible in the non-TE cell. The bars indicate 20 μm. After incubation with 0.1 μg/mL FDA for about 1 h, fluorescein in the cytoplasm usually began to enter the vacuole. Figure 1A, e and f, show the staining pattern of a TE and non-TE cell 1 h after FDA treatment. While non-TE cells allowed the dye to enter their vacuole, many TEs still had fluorescein only in their cytoplasm. More than one-half of all TEs exhibited this staining pattern. Because the cytoplasm and the vacuole were differently stained by this method, the vacuolar compartment was discernible even after the entrance of the dye into the vacuole (Fig. 1A, f). Further incubation resulted in the disappearance of cytoplasmic green fluorescence in all non-TE cells, because fluorescein was sequestered into vacuoles (Fig. 1B). However, many TEs still had fluorescein in their cytoplasm only (Fig. 1B). TEs therefore appeared to have a defect on fluorescein transport into the vacuole. The time course of the appearance of each cell type was investigated through the differentiation process. Figure2A shows how the percentages of TE cells with the characteristic staining changed. Until 4 h after the first appearance of TE cells, most of the TEs corresponded to type 1 cells. The percentage of type 1 cells then decreased, while the percentage of type 3 cells increased. At a maximum (at 56 h), about 20% of living TEs could be seen at a time. The number of type 2 cells was always small and constant throughout the entire culture process. All type 1 cells at each point were transformed to type 3 cells between 4 and 8 h, indicating that the longevity of living TEs in culture was about 6 h after the onset of SWT. Consequently, the percentage of living TEs in culture decreased gradually from 44 to 72 h (Fig. 2B). Fig. 2. Open in new tabDownload slide A, Time course of the appearance of TE types 1 to 3. Zinnia cells cultured for the indicated times were stained with FDA and examined immediately. The percentages of living (type 1), dying (type 2), and dead (type 3) TEs were calculated. B, The percentages of living TEs in the culture were recorded. Error bars representsd. Fig. 2. Open in new tabDownload slide A, Time course of the appearance of TE types 1 to 3. Zinnia cells cultured for the indicated times were stained with FDA and examined immediately. The percentages of living (type 1), dying (type 2), and dead (type 3) TEs were calculated. B, The percentages of living TEs in the culture were recorded. Error bars representsd. Inhibition of Fluorescein Transport into the Vacuole of TEs Since the above result (Fig. 1A, e and f) suggested that the tonoplast of TEs became less permeable to fluorescein, the number of TE and non-TE cells that excluded the dye from the vacuole was counted following FDA administration. Cells at 0, 50, or 58 h of culture were loaded with FDA and the percentages of TEs and non-TE cells excluding fluorescein from their vacuole were calculated separately (Fig. 3, A–C). As a control, cells incubated with FDA from the start of the culture were also examined (Fig. 3D). Figure 3A shows that almost all zinnia cells began to transport fluorescein into their vacuoles within 30 min after the addition of FDA to the medium. At 50 and 58 h (Fig. 3, B and C), when a substantial percentage of TEs had become apparent (Fig. 3E), fluorescein uptake also occurred but at a much slower rate. About 40% (B) or 60% (C) of living TEs excluded the dye from the vacuole even at 3 h. Thus, the accumulation of the dye into the TE vacuole was significantly inhibited. If FDA was loaded at the start of culture, the dye was completely sequestered in the vacuole of most mesophyll cells, but the majority of TEs exhibited both green and yellow fluorescence in the cytoplasm and vacuole, respectively. It was striking that 10% of TEs in the culture containing FDA from the beginning also excluded the dye at 50 and 58 h, albeit once they had sealed the dye in the vacuole before SWT (at 18 h, Fig. 3D). Because these TEs still contained fluorescein in their cytoplasm, the exclusion of fluorescein from the vacuole occurred concurrently with the inhibition of uptake. The difference in fluorescein exclusion of TEs at almost the same developmental stage (50 or 58 h) in Figure 3, B to D, strongly suggested that fluorescein can fluoresce even in the TE vacuole and that the lack of fluorescein in TEs (Fig. 3, B and C) was due to an inhibition of dye transport and not to degradation or the influence of vacuolar pH. Experiments designed to examine the direct uptake of fluorescein molecules by these cells through acid loading gave the same results (data not shown). The results shown in Figure 3E suggest that the amount of FDA used here had no effect on TE differentiation. Fig. 3. Open in new tabDownload slide Exclusion of fluorescein from the TE vacuole. Cells cultured for 0 (A), 50 (B), or 58 h (C) were stained with FDA for the indicated times at 26°C. The percentages of TEs and non-TE cells that do not contain fluorescein in their vacuole at all were calculated. Cells from the same culture but from another batch containing FDA from the start were sampled at indicated times and examined for the presence of fluorescein in vacuoles (D). The percentages of total TEs and the ratios of dead to living TEs in cultures in the presence or absence of FDA are shown in E. Asterisks (*) indicate that the parameter could not be defined at this time because TE formation had not yet occurred. Error bars represent sd. Fig. 3. Open in new tabDownload slide Exclusion of fluorescein from the TE vacuole. Cells cultured for 0 (A), 50 (B), or 58 h (C) were stained with FDA for the indicated times at 26°C. The percentages of TEs and non-TE cells that do not contain fluorescein in their vacuole at all were calculated. Cells from the same culture but from another batch containing FDA from the start were sampled at indicated times and examined for the presence of fluorescein in vacuoles (D). The percentages of total TEs and the ratios of dead to living TEs in cultures in the presence or absence of FDA are shown in E. Asterisks (*) indicate that the parameter could not be defined at this time because TE formation had not yet occurred. Error bars represent sd. Effects of Probenecid on the Viability of TE The effect of probenecid on TE development was tested because it is known to inhibit the transport of organic anions such as fluorescein derivatives across the tonoplast (Wright and Oparka, 1994). Probenecid altered the structure of cultured cells, and some laterally located cytoplasm disappeared (Fig. 4A). To compare non-TEs with TEs, cells were stained with FDA in the presence or absence of 100 μm probenecid at 55 h. Figure 4B, a and b, shows the fluorescent cytoplasm of non-TE cells. Like most plant mesophyll cells, those of zinnia leaves have a large central vacuole without any intervening cytoplasmic strands and basically maintain such an intracellular structure until about 72 h. The cytoplasm was distributed in a relaxed mode around the central vacuole of non-TE cells (Fig. 4B, a and b). But the cytoplasm of living TEs was compressed to form a very thin layer (Fig. 4B, c and d) as differentiation progressed. This layer ran along the inward protrusion of the thickened secondary walls. After probenecid treatment, the cytoplasm of non-TE cells became thinner, tense, and compressed around the vacuole due to vacuolar swelling, similar to those of TEs (Fig. 4B, e and f). Because all probenecid-treated non-TE cells continued to exclude fluorescein from their vacuole for more than 3 h, a concentration of 100 μm probenecid was also effective in blocking fluorescein transport across the tonoplast in zinnia cells. This concentration was significantly lower than those used in other cases (Oparka et al., 1991). Curiously, under these conditions, type 1 TEs were very rare even at 55 h, when they are usually abundant (Fig. 2). In fact, the percentage of living to total TEs was drastically decreased by probenecid treatment (Fig.5A). When 100 μm probenecid was added 51 h after induction, almost all living TEs were dead within 1 h (Fig. 5A), in contrast to controls (Fig. 2B). Non-TE cells gradually died beginning at 3 h, and were completely dead by 25 h. Probenecid can therefore efficiently kill differentiating TE cells. The percentage of TEs to total cells at each time point was almost equal to that before addition (Fig. 5B), indicating that further TE formation was consequently aborted by probenecid-induced TE cell death. Fig. 4. Open in new tabDownload slide A, Alteration of cellular structure by probenecid. Zinnia cultured cells before (a) and after (b–d) treatment with probenecid were observed using an inverted microscope and a Petri dish. Several 1-μL drops of 0.1 mg/mL probenecid were carefully added to the cell suspension cultured for 50 h until visible changes occurred. Micrographs are arranged sequentially (a, before probenecid addition; b, 15 min after probenecid addition; c, 1 h after probenecid addition; d, after a prolonged incubation). Upon higher magnification of other cell images (e and f), the structural change is obvious and the cytoplasm before probenecid addition (e) became localized to the pole of cells by the treatment (f). B, Comparison of the morphological features of TEs with those of non-TE cells treated with probenecid. Light (a, c, and e) and fluorescence (b, d, and f) micrographs of zinnia cells cultured for 55 h and then stained with FDA for less than 10 min. Non-TE cells (a and b), TEs (c and d), and non-TE cells treated with probenecid (e and f) accumulated fluorescein in their cytoplasm, but the morphology of the cytoplasm and the vacuole of control non-TE cells was different from those of TEs and non-TE cells treated with probenecid. Swelling of the vacuole was evident in TEs and non-TE cells treated with probenecid. The bars indicate 20 μm. Fig. 4. Open in new tabDownload slide A, Alteration of cellular structure by probenecid. Zinnia cultured cells before (a) and after (b–d) treatment with probenecid were observed using an inverted microscope and a Petri dish. Several 1-μL drops of 0.1 mg/mL probenecid were carefully added to the cell suspension cultured for 50 h until visible changes occurred. Micrographs are arranged sequentially (a, before probenecid addition; b, 15 min after probenecid addition; c, 1 h after probenecid addition; d, after a prolonged incubation). Upon higher magnification of other cell images (e and f), the structural change is obvious and the cytoplasm before probenecid addition (e) became localized to the pole of cells by the treatment (f). B, Comparison of the morphological features of TEs with those of non-TE cells treated with probenecid. Light (a, c, and e) and fluorescence (b, d, and f) micrographs of zinnia cells cultured for 55 h and then stained with FDA for less than 10 min. Non-TE cells (a and b), TEs (c and d), and non-TE cells treated with probenecid (e and f) accumulated fluorescein in their cytoplasm, but the morphology of the cytoplasm and the vacuole of control non-TE cells was different from those of TEs and non-TE cells treated with probenecid. Swelling of the vacuole was evident in TEs and non-TE cells treated with probenecid. The bars indicate 20 μm. Fig. 5. Open in new tabDownload slide Cell death caused by probenecid. Cells were cultured for 51 h and then treated with 100 μmprobenecid. A, The percentages of living TEs and non-TE cells were recorded separately. At 51 h, T and L values (%) were as follows: T(51) = 30.68 ± 6.08,L(51) = 24.18 ± 1.30. B, The percentage of TEs to total cells (including dead cells) in the culture was shown. In this case, the conventional T value does not make sense because non-TE cells were also killed by this treatment. c25, Sample of cells in a control batch in which cells have been cultured for 76 h in the absence of probenecid. Error bars represent sd. Fig. 5. Open in new tabDownload slide Cell death caused by probenecid. Cells were cultured for 51 h and then treated with 100 μmprobenecid. A, The percentages of living TEs and non-TE cells were recorded separately. At 51 h, T and L values (%) were as follows: T(51) = 30.68 ± 6.08,L(51) = 24.18 ± 1.30. B, The percentage of TEs to total cells (including dead cells) in the culture was shown. In this case, the conventional T value does not make sense because non-TE cells were also killed by this treatment. c25, Sample of cells in a control batch in which cells have been cultured for 76 h in the absence of probenecid. Error bars represent sd. Probenecid is a weak acid and its permeation into cells is influenced by medium pH, so the effect of pH on TE viability was investigated. Cells cultured for 53 h were incubated in the presence of 100 μm probenecid for 1 to 3 h at either pH 4.5 or 5.5 (Fig. 6A). The medium pH at 53 h of culture declined to 4.8 (Roberts and Haigler, 1994) and was adjusted with 1 n KOH just after the addition of probenecid. More TEs were killed by 100 μm probenecid at pH 4.5 than at pH 5.5, due to the abundance of the undissociated form of probenecid at pH 4.5 (Fig. 6A). Therefore, probenecid may only be effective once it has entered TEs. Under the same pH conditions, an increase in probenecid concentration had a more severe effect on TEs within 1 h, while the viability of non-TE cells was almost unaffected up to 3 h (Fig. 6B). The culture medium was replaced with 20 mm2-(N-morpholino)-ethanesulfonic acid (MES) medium (pH 4.7) just before the experiment to prevent the differential effect of probenecid by medium pH. The probenecid effect on living TEs is therefore dose dependent. Fig. 6. Open in new tabDownload slide Effect of pH and probenecid concentration on cell viability. A, Dependence of the probenecid effect on medium pH. Probenecid at 100 μm was added at 53 h to culture media, the pH of which was adjusted to 4.5 or 5.5 with the appropriate amounts of 1 n KOH just after probenecid addition. The percentages of TE and non-TE cells were examined separately at 1 and 3 h after the addition of probenecid. T(53) = 23.62 ± 4.69, L(53) = 18.34 ± 3.64. B, Dependence of the probenecid effect on the probenecid concentration under the same pH (4.7) medium. The medium of zinnia cell culture was buffered with 20 mm MES (pH 4.7) at 53 h after induction and supplied with 100 to 1 μm probenecid. The percentages of TE and non-TE cells were calculated separately 1 and 3 h after the treatment with probenecid. T(53) = 23.62 ± 4.69, L(53) = 18.34 ± 3.64. Error bars represent sd. Fig. 6. Open in new tabDownload slide Effect of pH and probenecid concentration on cell viability. A, Dependence of the probenecid effect on medium pH. Probenecid at 100 μm was added at 53 h to culture media, the pH of which was adjusted to 4.5 or 5.5 with the appropriate amounts of 1 n KOH just after probenecid addition. The percentages of TE and non-TE cells were examined separately at 1 and 3 h after the addition of probenecid. T(53) = 23.62 ± 4.69, L(53) = 18.34 ± 3.64. B, Dependence of the probenecid effect on the probenecid concentration under the same pH (4.7) medium. The medium of zinnia cell culture was buffered with 20 mm MES (pH 4.7) at 53 h after induction and supplied with 100 to 1 μm probenecid. The percentages of TE and non-TE cells were calculated separately 1 and 3 h after the treatment with probenecid. T(53) = 23.62 ± 4.69, L(53) = 18.34 ± 3.64. Error bars represent sd. To determine whether probenecid also killed non-TE cells by disrupting the vacuole, vacuole morphology was examined using FM 4-64, which specifically stains the yeast vacuolar membrane (Vida and Emr, 1995). FM 4-64 selectively stained the tonoplast of zinnia cells (Fig.7) when added to the culture from the start, although it significantly retarded and reduced TE differentiation (data not shown). The tonoplast of TEs formed a wave along the pattern of secondary walls (Fig. 7, a and b). When the vacuole disrupted, it shrank and fragmented (Fig. 7, c and d). The swollen vacuole of non-TE cells (Fig. 7, e and f) also ruptured and shrank after 6 h of incubation in 100 μm probenecid, similar to TEs (Fig. 7, g and h). Fig. 7. Open in new tabDownload slide Vacuole disruption visualized by FM 4-64. FM 4-64 was added to the culture from the beginning for the selective staining of the vacuole. The light (a, c, e, and g) and fluorescence (b, d, f, and h) micrographs of zinnia cells are shown. TE (a and b) and non-TE cells treated with 100 μm probenecid for 1 h (e and f) have a large central vacuole. The vacuole of TEs ruptured to form small, fragmented vacuoles (arrowhead, c and d). Similarly, treatment with probenecid for 6 h caused the rupture, fragmentation, and shrinkage of the vacuole of non-TE cells (arrowhead, g and h). The bar indicates 20 μm. Fig. 7. Open in new tabDownload slide Vacuole disruption visualized by FM 4-64. FM 4-64 was added to the culture from the beginning for the selective staining of the vacuole. The light (a, c, e, and g) and fluorescence (b, d, f, and h) micrographs of zinnia cells are shown. TE (a and b) and non-TE cells treated with 100 μm probenecid for 1 h (e and f) have a large central vacuole. The vacuole of TEs ruptured to form small, fragmented vacuoles (arrowhead, c and d). Similarly, treatment with probenecid for 6 h caused the rupture, fragmentation, and shrinkage of the vacuole of non-TE cells (arrowhead, g and h). The bar indicates 20 μm. The relationship between tonoplast selective permeability and vacuole shape was also investigated using probenecid and FDA. Although the dye was confined to the cytoplasm in all living cells treated with 100 μm probenecid at this time of culture (55 h), incubation for another 6 h killed non-TEs significantly (Fig. 5A). Many non-TE cells were first filled evenly with fluorescein, after which the fluorescence soon faded away. Among such cells, one cell filled with fluorescein, one cell precluded dye entry into the vacuole, and a third cell was dead as shown at the right, center, and left, respectively in Figure 8, a and b. The tonoplast of the cell at the right clearly lost selective permeability, like type 2 TE cells (Fig. 1A, c and d), whereas that of the middle cell retained selective permeability to fluorescein. In a bright-field image, the large compartment of the central vacuole was not seen in the cell (at the right), and a spherical, balloon-like structure was observed instead. This was composed of the fragmented tonoplast and was probably the same as those found in dying TE cells (Fig. 8, e and f; Groover et al., 1997). These morphological data indicate that probenecid can also induce cell death in non-TEs via the loss of tonoplast selective permeability, and that the resultant fragmented vacuole loses membrane integrity. Furthermore, another cell shown in Figure 8, c and d, had lost its tonoplast selective permeability during probenecid treatment, although the shape of its central vacuole apparently re-mained intact. Figure 8, e and f, shows dying TEs that still had the boundary between the cytoplasm and the central vacuole. The existence of these cells strongly suggested that the loss of tonoplast selective permeability occurred before the physical fragmentation of the vacuolar membrane both in TEs and in probenecid-treated non-TE cells. Fig. 8. Open in new tabDownload slide Tonoplast disintegration of cells treated with probenecid. Light (a, c, e, and g) and fluorescence (b, d, f, and h) micrographs of zinnia cells. Non-TE cells that were treated with 100 μm probenecid for 6 h after 55 h of culture (a–d) were stained with FDA for 1 h. Large arrows point to the tonoplast (a, c, and f). The arrowhead indicates the fragmented, spherical vacuole formed after the disruption of the central vacuole (a). TEs incubated with FDA for 1 h (e and f) lost their tonoplast selective permeability without any drastic changes in the boundary between the cytoplasm and the vacuole (large arrow). Dead non-TE cells and differentiating TE cells killed by 100 μm probenecid (g and h) did not autolyze their contents. Small arrows indicate the slightly thickened secondary wall of a very young TE cell killed by probenecid. The bar indicates 20 μm. Fig. 8. Open in new tabDownload slide Tonoplast disintegration of cells treated with probenecid. Light (a, c, e, and g) and fluorescence (b, d, f, and h) micrographs of zinnia cells. Non-TE cells that were treated with 100 μm probenecid for 6 h after 55 h of culture (a–d) were stained with FDA for 1 h. Large arrows point to the tonoplast (a, c, and f). The arrowhead indicates the fragmented, spherical vacuole formed after the disruption of the central vacuole (a). TEs incubated with FDA for 1 h (e and f) lost their tonoplast selective permeability without any drastic changes in the boundary between the cytoplasm and the vacuole (large arrow). Dead non-TE cells and differentiating TE cells killed by 100 μm probenecid (g and h) did not autolyze their contents. Small arrows indicate the slightly thickened secondary wall of a very young TE cell killed by probenecid. The bar indicates 20 μm. In contrast to the formation of TE (Fukuda 1997a, 1997b; Groover et al., 1997), non-TE cells killed by probenecid plasmolyzed and did not autolyze (Fig. 8, g and h). This probenecid treatment produced many dead cells with only slight SWT, the contents of which also remained undigested (Fig. 8, g and h). Therefore, probenecid could disrupt the vacuole but could not induce autolysis. Cell Death Suppressed by Cycloheximide (CHX) To examine whether the inhibition of organic anion transport into the TE vacuole was regulated by the cell death program, CHX, an inhibitor of cellular protein synthesis, was applied to the culture. The influence of CHX on TE formation differed depending on the timing of inhibitor added to the culture. When zinnia mesophyll cells were treated with CHX from the start of the culture (Fukuda and Komamine, 1983), SWT was completed prevented. At 50 h, when about 20% of cells had commenced wall synthesis, they responded to this antibiotic in a different manner. As shown in Figure9A, CHX treatment up to 9 h prevented the differentiation of cells into TEs, as shown by the low percentage of TE cells (22%) relative to the untreated control (40%). Moreover, the increase in dead TEs was also blocked from 3 to 9 h. The proportion of TEs excluding fluorescein from their vacuole decreased dramatically among the total living TEs. Therefore, differentiation, cell death, and fluorescein exclusion from the vacuole of TEs were all suppressed by CHX treatment. The parallel changes in cell death and vacuolar exclusion of fluorescein suggest that biochemical changes had occurred, resulting in a loss of tonoplast integrity. Fluorescein exclusion by the vacuole of living TEs also clearly requires new or continuous expression of a certain protein(s) associated with TE formation. Fig. 9. Open in new tabDownload slide Effect of CHX on TE differentiation, cell death, and the inhibition of fluorescein transport across the tonoplast. A, CHX at 50 μm was added to culture at 50 h. The percentages of TEs, living TEs, and living TEs whose vacuole was not fluorescent were determined. The parameters presented were dead TEs, living TEs containing fluorescein in their vacuole, living TEs excluding fluorescein from the vacuole. B, The percentages of living non-TE cells were not affected by the 50 μm CHX treatment of this experiment. c9 and c21, Patterns of differentiating TEs in control batches in which cells have been cultured for 59 and 71 h in the absence of CHX, respectively. Error bars representsd. Fig. 9. Open in new tabDownload slide Effect of CHX on TE differentiation, cell death, and the inhibition of fluorescein transport across the tonoplast. A, CHX at 50 μm was added to culture at 50 h. The percentages of TEs, living TEs, and living TEs whose vacuole was not fluorescent were determined. The parameters presented were dead TEs, living TEs containing fluorescein in their vacuole, living TEs excluding fluorescein from the vacuole. B, The percentages of living non-TE cells were not affected by the 50 μm CHX treatment of this experiment. c9 and c21, Patterns of differentiating TEs in control batches in which cells have been cultured for 59 and 71 h in the absence of CHX, respectively. Error bars representsd. The effect of CHX was attenuated by 21 h possibly due to partial CHX detoxification as seen in yeast growth (Meyers et al., 1992), such that differentiation and cell death could progress to nearly the control level (Fig. 9A). The percentage of living non-TE cells in culture was almost constant irrespective of CHX treatment (Fig. 9B). CHX at 50 μm did not have any significant nonspecific toxic effect on zinnia cells. The ratio of non-TE cells excluding fluorescein from their vacuole to total living non-TE cells was much smaller than that of TEs throughout this experiment (data not shown). The treatment did not affect the fluorescein uptake into the vacuole of non-TE cells. Accumulation of Marker Genes for TE Differentiation Could Be Altered by Probenecid To investigate the effect of probenecid on mRNA accumulation in cultured cells, RNA gel-blot analyses were performed using probes for two marker genes, TED3 and ZCP4. Although in situ studies showed RNA expression of the TED3 and the Cys protease genes (p48-17 and ZCP4) associated with TE cells (Demura and Fukuda, 1994; Ye and Varner, 1996; Igarashi et al., 1998; A. Minami and H. Fukuda, unpublished results), the regulation of expression appears to differ markedly. TED3 mRNA is steadily expressed after 36 h, but ZCP4 mRNA only appears from 48 to 72 h in this culture system (Fukuda, 1997a). All known TE-associated genes exhibit one of these expression patterns in cultured cells (Fukuda, 1997a). The expression of these genes represents two distinct regulatory points in the TE differentiation program, although visible changes such as SWT or the production of other hydrolytic enzymes start temporally together with the latterZCP4 expression. Total RNA samples from cells incubated for 1 h with or without 100 μm probenecid from 57 h after induction, when transcripts of these genes had accumulated as described previously (Fukuda, 1997a), were probed with TED3 and ZCP4. The results shown in Figure10indicate that ZCP4 mRNA accumulation was abolished by a 1-h treatment with probenecid, whereas TED3 mRNA was maintained at a constant level in 2 μg of total RNA. These data strongly suggest that ZCP4 mRNA accumulated mainly in cells that could be selectively killed by probenecid treatment, and strongly support the observation of selective TE cell death that resulted from this treatment (Fig. 5A). Meanwhile, TED3 was equally expressed in these cells irrespective of the difference in susceptibility to probenecid. Probenecid therefore affects the cells that have expressed the latter TE-differentiation program, suggesting the involvement of the latter regulatory point in the changes in the vacuolar properties of TEs. These results are summarized in Figure 11. Fig. 10. Open in new tabDownload slide RNA gel-blot analyses using digoxigenin-labeled cDNA probes for TED3 and ZCP4 genes. The total RNA of cells cultured for 57 h and subsequently treated with (+) or without (–) 100 μm probenecid for 1 h was extracted. In each lane 2 μg of total RNA was loaded. rRNA was displayed by staining with methylene blue (Sambrook et al., 1989). Fig. 10. Open in new tabDownload slide RNA gel-blot analyses using digoxigenin-labeled cDNA probes for TED3 and ZCP4 genes. The total RNA of cells cultured for 57 h and subsequently treated with (+) or without (–) 100 μm probenecid for 1 h was extracted. In each lane 2 μg of total RNA was loaded. rRNA was displayed by staining with methylene blue (Sambrook et al., 1989). Fig. 11. Open in new tabDownload slide A simple model of events occurring during TE PCD. A, Related events occurring in cultured cells (Fukuda, 1997a) after hormonal induction (0 h) are displayed with time periods selected for two experiments studying the effect of probenecid and cycloheximide (Figs. 5 and 9). B, Events occurring in a single, differentiating TE are shown with an emphasis on the changes in the vacuole. A general model of TE PCD was made by Groover et al. (1997). Shaded areas indicate the distribution of fluorescein when FDA is added to the culture at stage c. CHX suppresses progression to stage b, c, and d (Fig. 9). Related events and vacuolar properties are also shown. a, Before TE differentiation; b, secondary wall formation; c, vacuole swelling, inhibition of fluorescein transport into the vacuole, and fluorescein exclusion from the vacuole; d, loss of the selective permeability of the tonoplast; e, tonoplast shrinkage and fragmentation; f, a dead TE. Fig. 11. Open in new tabDownload slide A simple model of events occurring during TE PCD. A, Related events occurring in cultured cells (Fukuda, 1997a) after hormonal induction (0 h) are displayed with time periods selected for two experiments studying the effect of probenecid and cycloheximide (Figs. 5 and 9). B, Events occurring in a single, differentiating TE are shown with an emphasis on the changes in the vacuole. A general model of TE PCD was made by Groover et al. (1997). Shaded areas indicate the distribution of fluorescein when FDA is added to the culture at stage c. CHX suppresses progression to stage b, c, and d (Fig. 9). Related events and vacuolar properties are also shown. a, Before TE differentiation; b, secondary wall formation; c, vacuole swelling, inhibition of fluorescein transport into the vacuole, and fluorescein exclusion from the vacuole; d, loss of the selective permeability of the tonoplast; e, tonoplast shrinkage and fragmentation; f, a dead TE. DISCUSSION Change in Tonoplast Permeability during TE Formation FDA has been used as a probe to monitor vacuolar function and/or properties during TE differentiation. This dye is highly lipophilic and readily passes through the plasma membrane of a cell; its acetyl ester groups are then cleaved by intracellular esterase activity to generate fluorescein. Most of the fluorescein molecules in the cytoplasm are in the polar, dissociated form and are therefore retained within the cell (Oparka, 1991). The selective permeability of the membrane restricts free passage of electrolytes across the membrane and maintains cell integrity. This dye was used to identify living and dead TEs in the zinnia cell culture (Fig. 1A, a–d). Three characteristic staining patterns were recognized: cells with green fluorescence in their cytoplasm, cells with yellow fluorescence in the whole cell, and cells with no fluorescence. The difference in these fluorescence spectra reflects the variant molecular forms of fluorescein affected by the pH of the milieu where the dye is present (Martin and Lindqvist, 1975). The cytoplasmic and vacuolar pH are generally about 7.0 and 5.5, respectively. At physiological range, the fluorescence of the yellow wavelength area becomes conspicuous as fluorescein is exposed to the solutions of lower pH. The first and the last cell types obviously correspond to the living and dead states of TEs, respectively. The second type of TE has just lost the tonoplast selective permeability to both fluorescein and H+, and only accounted for a small number of dying cells undergoing tonoplast disintegration (Fig.2A). Closer examination indicated that after the ester groups of FDA were cleaved, the dye was transported to the vacuole of non-TE cells, but remained in the cytoplasm of differentiating live TEs (Fig. 1A, f, and Fig. 1B). The time course of the appearance of cells with or without fluorescein in their vacuoles revealed that the inhibition of fluorescein transport across the tonoplast was correlated with the formation of TEs (Fig. 3, B and C). Moreover, since the number of TEs excluding fluorescein from their vacuoles was variable depending on the timing of FDA addition to culture (Fig. 3, B–D), one can conclude that both the inhibition of uptake across the tonoplast and the exclusion of fluorescein from the vacuole accompany TE formation. Oparka et al. (1991) and Wright and Oparka (1994) determined that a probenecid-sensitive, ATP-dependent organic anion transporter is responsible for the transport of fluorescein derivatives into the vacuoles of plant cells. Organic anion transporters that can be affected by probenecid are also detected in vitro in the tonoplast of barley or rye leaf cells (Fig. 1; Blake-Kalff and Coleman, 1996; Klein et al., 1997). Probenecid addition appeared to promote and reinforce the inhibition of transport in differentiating, living TEs, whereas it caused the initiation of transport inhibition in non-TE cells. Thus, probenecid killed living TEs rapidly and selectively and then killed non-TE cells after a significant time lag (Fig. 5A). The possible target of probenecid in TE cells is not on the outer surface but in the cytoplasm (Fig. 6A), and the deleterious effect of probenecid is dose dependent (Fig. 6B). Thus, the probenecid effect could at least in part mimic developmental changes in the TE vacuole. In addition, many features of differentiating, living TEs are similar to those of probenecid-treated cells. In contrast to fluorescein, Lucifer Yellow, a hydrophilic dye with a much lowerpK a value, did not leak from the vacuole following probenecid treatment (Oparka et al., 1991). When the dye was added to culture at the beginning, substantial accumulation could be observed in the vacuole of all cells but not in the cytoplasm. TEs could be formed even in such a situation but, unlike fluorescein, Lucifer Yellow did not leak from the TE vacuole until the very moment of its disruption (data not shown). Also, the vacuoles of TEs became swollen as differentiation progressed (Fig. 4B, c and d). This vacuole swelling also occurred in non-TE cells treated with probenecid (Fig. 4B, e and f; Oparka et al., 1991). Enhanced vesiculation in the cytoplasm is reported to occur both in TEs (Groover et al., 1997) and in onion epidermal cells fed with probenecid (Oparka et al., 1991). Furthermore, Groover et al. (1997) showed that the vacuolar membrane shrinks inwardly when the TE vacuole collapses (Fig. 7, c and d). Non-TE cells treated with probenecid also died from the disintegration of their tonoplasts (Fig. 7, e–h; Fig. 8, a–d) in the same manner as TEs (Fig. 8, e and f). The small, spherical structure of the fragmented tonoplast that results from shrinkage was formed in dying non-TE cells (Fig. 7, g and h; Fig. 8, a and b) in the same manner as TEs (Fig. 7, c and d). These findings show that all of the known effects of probenecid can occur in differentiating, living cells, and suggest that these effects play an important role in vacuole disruption. It is possible that the effects of probenecid on TE development are inevitable consequences of anion transport inhibition. For example, the unbalanced distribution of electrolytes across the membrane could perturb the membrane potential and the total transport system. Disruption of cellular osmoregulation could thus have deleterious effects on cellular homeostasis. Probenecid might alter the water potential gradient across the tonoplast by blocking organic anion transporters and possibly other transport systems, leading to tonoplast disintegration. Probenecid has been shown to perturb the osmoregulatory mechanisms of guard cells, forcing them to remain swollen (Schwartz et al., 1995). Furthermore, a certain class of ATP-dependent drug transporters prevent the swelling-induced membrane disintegration of animal cells (Roman et al., 1997). This class of transporters on plant cell vacuoles (Rea et al., 1998) may perform a similar function in zinnia cultured cells, so that probenecid inhibition of those transporters can lead to a loss of membrane integrity. Developing TEs have intrinsic inhibition of organic anion transport (Figs. 1 and 3). Probenecid may reinforce this effect and cause rapid cell death of TEs through tonoplast disintegration. These functions are distinct from those of proton pumps or aquaporins. Organic anion uptake by the vacuoles is independent from vacuolar pH and H+-ATPase inhibition (Blake-Kalff and Coleman, 1996; Klein et al., 1997), and the alkalization of the vacuole does not seem to cause vacuole disruption in cultured cells (Matsuoka et al., 1997). Aquaporins (Maurel, 1997) cannot continue to transport water against a water potential gradient until a membrane compartment disrupts. One criterion to discriminate PCD processes from necrotic death is the dependence of cell death on specific gene expression. Figure 9A shows the response of living TEs to CHX, which suppressed SWT, death, and the exclusion of fluorescein molecules from the vacuole of TEs. These findings suggest that not only vacuole disruption but also the change in permeability of the tonoplast resulted from the function of a certain gene product(s) that appears at TE formation. The possibility that some necrotic effects such as spontaneous lipid deterioration are responsible for the tonoplast disintegration was completely ruled out. Even after the initiation of SWT, the vacuole disruption of TEs is still under the control of a genetic program. Transport Inhibition and TE Cell Death Program Possible regulatory mechanisms of plant PCD include Ca2+-mediated processes (Greenberg, 1997;O'Brien et al., 1998) and oligosaccharide-dependent processes (McCann, 1997). These molecules also function in TE differentiation (Roberts and Haigler, 1989, 1990; Roberts et al., 1997; Domingo et al., 1998). In particular, Ca2+ influx and extracellular proteolysis mediate the death of zinnia cells (Groover and Jones, 1999). However, events directly related to the vacuolar membrane of TEs had not been found. As indicated above, the organic anion transport system across the tonoplast was impeded in cultured zinnia cells that began to show SWT. Since cells that develop SWT eventually die from vacuole disruption, events that occur specifically in the TE vacuole must be important in the cell death of TEs. The inhibitory effect on organic anion transport could lead to vacuole disruption (Figs. 5A, 7, and 8) even in non-TE cells. Moreover, the effects of transport inhibition are distinct from those of the reagents tested by Groover and Jones (1999), which killed all zinnia cultured cells immediately. Probenecid may act on another, novel event in the process of vacuole disruption. RNA gel-blot analyses revealed that probenecid abolishedZCP4 mRNA accumulation within 1 h (Fig. 10). Cells expressing ZCP4 were killed rapidly and selectively, suggesting that the intrinsic inhibitory effect of fluorescein transport, which could be promoted by probenecid treatment, occurred in parallel with ZCP4 expression. It is unlikely, however, that probenecid, a xenobiotic compound, has a key role in the actual differentiation process of TEs in vivo. Some genetic mechanism(s) may exist that can bring about the same effect. The result of the experiment with CHX (Fig. 9) strongly supports this idea. In mammalian hepatocytes, probenecid treatment invokes the expression of a particular gene (Gant et al., 1995). However, probenecid drastically decreased ZCP4 mRNA in culture and did not induce ZCP4 expression in non-TE cells (Fig. 10). Probenecid probably does not induce specific genes to exert its disruptive effect, much less activate the entire genetic program of TE formation. Not all of the TE formation processes (Groover et al., 1997) were mimicked by probenecid-treated non-TE cells. Non-TE cells and immature TEs killed by probenecid did not autolyze their contents (Fig. 8, g and h) but, rather, morphologically resembled those reported on PCD in a diluted carrot suspension culture (McCabe et al., 1997; Pennell and Lamb 1997). The coordinated progress of the program of TE formation may require other regulatory mechanisms that would establish a background such as sufficient production of hydrolytic enzymes to function in autolysis. The inhibition of organic anion transport into the vacuole alone does not regulate total TE formation. However, given the critical role of vacuole collapse as the trigger of TE cell death, further analysis of intrinsic transport inhibition at the molecular level may reveal the fatal signal generated by the TE cell death program. ACKNOWLEDGMENTS Many thanks to Prof. H. Fukuda (University of Tokyo) for critical reading of the manuscript and helpful discussions and to his colleagues for their kind gifts of cDNA clones. 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