Plant Physiology

Leustek, Thomas; Saito, Kazuki

doi: 10.1104/pp.120.3.637pmid: 10398698

Sulfur is one of the six macronutrients required by plants and is found in the amino acids Cys and Met and in a variety of metabolites. When one considers that sulfur in plants is only 3% to 5% as abundant as nitrogen, it is perhaps understandable that sulfur assimilation has been less well studied. As a part of the Cys molecule, the sulfur group, called a thiol, is strongly nucleophilic (electron-donating), making it ideally suited for biological redox processes. When oxidized, two Cys molecules can form a covalent linkage called a disulfide bond, which is readily broken by reduction to form two thiol groups. Disulfide ↔ dithiol interchange is so versatile that nearly all aerobic forms of life, including plants, have evolved to use this reaction as the dominant form of redox control. Redox control regulates enzymes and protects against oxidative damage. SULFUR IS THE FUNCTIONAL COMPONENT OF GSH AND OTHER REDOX FACTORS Free Cys is not used for redox control. It is much too readily oxidized to cystine, the disulfide form, which is visible in the laboratory as a white precipitate that is formed within hours after preparing a solution of Cys. A variety of more stable thiol compounds are involved in redox regulation. The most abundant is glutathione, an enzymatically synthesized tripeptide in which Cys is linked via peptide bonds to the γ-carboxyl group of Glu and the α-amino group of Gly. In plants glutathione is thought to be between 3 and 10 mm,and it is present in the major cellular compartments. The reduced form of glutathione is often referred to as GSH, whereas the disulfide form is GSSG. The balance between forms is overwhelmingly maintained in favor of GSH by the enzyme glutathione reductase, using NADPH as an electron source. The result is that the plant cytoplasm, chloroplast stroma, and mitochondrial matrix are highly buffered in the reducing state. Many intracellular enzymes require reducing conditions for activity, just as they require a specific pH or other properties of their chemical environment. The reason is that Cys residues in proteins can also form disulfide bonds, resulting in a disruption of structure and a loss of activity. There are special cases in which specific disulfide bonds are required for formation of tertiary and quaternary structure in a protein, but this is less common, especially for soluble intracellular proteins. REDOX FACTORS REGULATE PLANT METABOLIC PATHWAYS Other factors that use the chemistry of disulfide ↔ dithiol interchange to mediate redox reactions include the proteins thioredoxin, glutaredoxin, and protein disulfide isomerase. These proteins are nearly ubiquitous and play fundamental roles in many different types of regulation (Fig. 1). One of the first and best examples of the function of thioredoxin comes from Buchanan (1991). The dark reactions of CO2 fixation must be strictly coordinated with the light reactions of photosynthesis. The coordination mechanism relies on the reductive activation of specific enzymes by thioredoxin, which is reduced by photosynthetically reduced Fd. New disulfide ↔ dithiol redox regulated processes are being discovered each year, which attests to the prevalent roles that sulfur chemistry plays in biology. Fig. 1. Open in new tabDownload slide Regulation of metabolism by disulfide ↔ dithiol interchange. The diagram shows how thioredoxin functions as a regulation factor through reduction of a regulatory disulfide on a target enzyme. In the case of C assimilation, the source of electrons for thioredoxin reduction is Fd, reduced via the light reactions of photosynthesis. Thioredoxins also exist in the cytoplasm of plants where NADPH + H+ serves as an electron source. Recent evidence shows that thioredoxin has the potential to act as an oxidant mediating the formation of a disulfide bond on a target enzyme (Stewart et al., 1998). This activity could be important for activation of antioxidant enzymes during oxidative stress. Fig. 1. Open in new tabDownload slide Regulation of metabolism by disulfide ↔ dithiol interchange. The diagram shows how thioredoxin functions as a regulation factor through reduction of a regulatory disulfide on a target enzyme. In the case of C assimilation, the source of electrons for thioredoxin reduction is Fd, reduced via the light reactions of photosynthesis. Thioredoxins also exist in the cytoplasm of plants where NADPH + H+ serves as an electron source. Recent evidence shows that thioredoxin has the potential to act as an oxidant mediating the formation of a disulfide bond on a target enzyme (Stewart et al., 1998). This activity could be important for activation of antioxidant enzymes during oxidative stress. GLUTATHIONE IS IMPORTANT IN STRESS MITIGATION Because of its nucleophilic properties, glutathione serves as the first line of defense against the products of oxygen metabolism, reactive oxygen species, and other electrophilic compounds such as toxins (herbicides), xenobiotics, and heavy metals (May et al., 1998). When plants encounter reactive oxygen species, glutathione is a direct source of electrons for stress mitigation by the enzyme glutathione peroxidase or an indirect means to maintain a reduced pool of ascorbate, another antioxidant. Glutathione reacts directly with toxins in a reaction mediated by glutathione S-transferase. In this way the toxins are inactivated and tagged for transport into the vacuole and for degradation (Kreuz et al., 1996). In some plants heavy-metal detoxification is mediated by glutathione derivatives called phytochelatins, which have the general structure (γ-glutamylcysteine)nGly (n = 2–11), and by Cys-rich proteins called metallothioneins. In both molecules thiol groups serve as the metal ion ligand. PLANT SULFUR ASSIMILATION IMPACTS AGRICULTURE AND THE ENVIRONMENT The preceding discussion illustrates sulfur's essential, general biological role. However, plants also incorporate sulfur into a wide range of secondary compounds that have an impact, in varied and subtle ways, on our use of plants and on the way that plants influence the environment. For example, the pungent odor and taste of onions, garlic, and cabbage are caused by sulfur-containing secondary compounds. The same compounds can impart an objectionable flavor to canola oil, diminishing its commercial value; but they also have been attributed to disease prevention in humans (Fahey et al., 1997). Some sulfur-containing phytoalexins such as camalexin may be important in combating plant pathogens (Zhao et al., 1998). Although sulfur was long thought not to limit plant productivity, the recent restrictions on emissions of sulfurous air pollutants, the ingredients of acid rain, have resulted in sulfur deficiency in some agricultural areas of the world. Another example is that sulfur assimilation by plants has been implicated as a potential factor in moderating climate. Marine algae are prodigious producers of dimethylsulfoniopropionate, a sulfur-containing analog of betaine (Cooper and Hanson, 1998). Dimethylsulfoniopropionate degradation releases dimethylsulfide, which volatilizes from the ocean and seeds the formation of clouds in the atmosphere. The global scale of this process is such that algal growth may actually influence climate. OVERVIEW OF THE SO42− ASSIMILATION PROCESS IN PLANTS Sulfur is available to plants primarily in the form of anionic sulfate (SO42−) present in soil. It is actively transported into roots and then distributed, mostly unmetabolized, throughout the plant. SO42− is a major anionic component of vacuolar sap; therefore, it does not necessarily enter the assimilation stream. Gaseous sulfur dioxide (SO2) is readily absorbed and assimilated by leaves, but it is significant as a nutrient source only in industrial areas with air pollution. Sulfur is assimilated in one of two oxidation states. SO42− can be added to a hydroxyl group of an organic molecule. The reaction is referred to as sulfation and it is catalyzed by sulfotransferases. By contrast, Cys contains reduced sulfur, which is produced from SO42− in a multistep pathway in which eight electrons are added to form sulfide (S2−; Reaction 1). The reduction of SO42− to S2−consumes 732 kJ mol−1. By comparison, reduction of nitrate to NH3 requires 347 kJ mol−1. The pathways of SO42− assimilation in plants are depicted in Figure 2. The figure shows only those enzymes that are known with certainty by characterization of the defined activity of a purified enzyme and through gene cloning. SO42−+ATP+8 e−+8 H+→S2−+4H2O+AMP+PPi (Reaction 1) Cys, the end product of the reductive pathway, is the starting material for production of Met, glutathione, and other metabolites containing reduced sulfur. Fig. 2. Open in new tabDownload slide Plant sulfur assimilation pathways showing only those enzymes that have been conclusively demonstrated. The top line shows SO42− activation and reduction. The sulfation pathway is shown in the second line on the left and assimilation of reduced sulfur into Cys on the second line on the right. All enzymes are shown in bold above the reaction arrow, whereas intermediates are shown below the chemical structure or in isolation when the chemical structure is not shown. The R- group in sulfated metabolite refers to the metabolite that is sulfated. Fd indicates the reduced and oxidized forms of Fd. Fig. 2. Open in new tabDownload slide Plant sulfur assimilation pathways showing only those enzymes that have been conclusively demonstrated. The top line shows SO42− activation and reduction. The sulfation pathway is shown in the second line on the left and assimilation of reduced sulfur into Cys on the second line on the right. All enzymes are shown in bold above the reaction arrow, whereas intermediates are shown below the chemical structure or in isolation when the chemical structure is not shown. The R- group in sulfated metabolite refers to the metabolite that is sulfated. Fd indicates the reduced and oxidized forms of Fd. In higher plants sulfation is a relatively minor fate for sulfur when compared with the reductive pathway. However, in marine algae, which produce large amounts of sulfated extracellular polysaccharides such as agar, sulfation accounts for a much greater proportion of the total assimilated sulfur. SO42− UPTAKE IS MEDIATED BY A FAMILY OF TRANSPORTERS WITH SPECIALIZED FUNCTIONS The transport of SO42−occurs across several membrane systems as it enters and is distributed throughout the plant and within cells. Transport across the plasma membrane occurs with protons at a ratio of 1 SO42−:3 H+ (symport) and is driven by a proton gradient maintained by a proton ATPase. Transport across the tonoplast membrane is mediated by an unknown mechanism that is driven by the electrical gradient between the vacuole sap and cytoplasm. The phosphate/triose phosphate translocator of the inner chloroplast membrane or a proton/SO42− symporter may mediate SO42− transport into chloroplasts. The plasma membrane transporters of plants have been characterized (Smith et al., 1997; Takahashi et al., 1997). The sequences of cDNAs cloned from Stylosanthes hamata, Arabidopsis, soybean, barley, maize, resurrection grass, and Indian mustard showed that the plasma membrane transporters of plants are most closely related to fungal and animal proton/SO42−cotransporters. Hydropathy analysis revealed that the plant transporters may span the membrane 12 times, a structural feature that is typical of many types of solute symporters. In most of the species that have been analyzed, SO42− transporters are encoded by a gene family. The situation in S. hamata is probably typical for most plants. In this species the individual transporters may have specialized functions, since they differ widely in affinity for SO42−, and they show distinct spatial and regulated patterns of expression (Smith et al., 1997). High-affinity forms with Km for SO42− of approximately 9 μm are expressed exclusively in roots, whereas the lower-affinity form with Km for SO42− of approximately 100 μm is expressed principally in leaves but also in roots. The steady-state level of mRNA for the high-affinity form increases rapidly after sulfur starvation, whereas the lower-affinity form is unresponsive or responds more slowly to changes in external SO42− supply. These results imply that the increase in SO42− transport activity observed in roots of sulfur-starved plants is due to an increase in the expression of specific transporters. One of the earliest observations of SO42− transport into roots was that the uptake rate varies in relation to the [SO42−] of the bathing solution. The results with S. hamata suggest that this multiphasic behavior may be due to the activities of separate transporters with different affinities for SO42−. What is the function of multiple SO42− transporters? The expression pattern of the high-affinity type suggests that it mediates uptake of SO42− into the plant and is a way to adjust to variation in the external sulfur supply. By contrast, low-affinity transporters could function in SO42− uptake, both from soil and from the apoplast solution that bathes internal cells. Evidence for specialization of function has been obtained from analysis of an Arabidopsis SO42− transporter that is most closely related in sequence to the low-affinity type fromS. hamata (Takahashi et al., 1997). It is expressed exclusively in the vascular parenchyma of roots and leaves and not in endodermal, cortical, or epidermal cells. The spatial expression pattern of this low-affinity-type transporter indicates that it must be responsible for uptake from the internal apoplastic pool of SO42−, not from the soil. SO42− IS AN INERT COMPOUND THAT MUST BE ACTIVATED BEFORE IT CAN BE METABOLIZED The low reactivity of SO42− is a barrier to assimilation that is overcome by formation of a phosphate-SO42−-anhydride bond in the compound APS. The reaction is catalyzed by ATP sulfurylase (Reaction 2) and is the sole entry point for metabolism of SO42−. SO42−+MgATPMgPPi+APS(Reaction 2) The equilibrium of the adenylation reaction favors the production of SO42− and ATP, the reverse reaction. The Keq is 10−7m in vitro, and the forward reaction can be measured only if enzymes that hydrolyze PPi and modify APS are included. Exactly how ATP sulfurylase operates in the forward direction in vivo has yet to be determined, since the conditions do not appear to be in equilibrium. The PPi concentration is approximately 0.3 mm in plant cells. Despite the theoretical difficulty, Arabidopsis ATP sulfurylase is able to produce APS in vivo under apparently nonequilibrium conditions, since it is fully able to substitute for Escherichia coli ATP sulfurylase even though the PPi concentration in the E. colicytoplasm is approximately 0.5 mm (Murillo and Leustek, 1995). The Arabidopsis enzyme does not have an intrinsic ability to overcome PPi inhibition, which indicates that extrinsic mechanisms in E. coli facilitate the forward operation of plant ATP sulfurylase, mechanisms that may also function in plant cells. There are two ATP sulfurylase isoforms in most plants: a major form localized in plastids and a minor form localized in the cytoplasm. Both enzymes have similar kinetic and structural properties. The isoenzymes are encoded by a gene family, and in Arabidopsis there are multiple genes for the plastid enzyme. Arabidopsis contains a cytosolic form of ATP sulfurylase, but the corresponding gene has not yet been identified (Rotte, 1998). The plastid enzyme exists in both leaves and roots and is responsible for initiating the reductive assimilation of SO42-, since isolated chloroplasts can form Cys from SO42− (Schürmann and Brunold, 1980). The cytoplasmic form probably functions by generating APS for sulfation reactions. Whether plant ATP sulfurylase plays a role in regulating sulfur assimilation has been studied by a number of investigators (Logan et al., 1996; Lappartient et al., 1999). In general, the activity and steady-state mRNA levels increase when plants are starved for sulfur and decrease when plants are fed reduced forms of sulfur (Cys or glutathione). However, the changes in activity and mRNA, although reproducible, are relatively small; they increase or decrease by approximately 2-fold or less, and the regulation occurs mainly in roots. Two publications report the use of transgenic plants to explore whether ATP sulfurylase regulates SO42− assimilation. Hatzfeld et al. (1998) concluded that it is not rate limiting, based on an analysis of a transgenic tobacco cell culture that overexpresses an Arabidopsis ATP sulfurylase but that does not show increased sulfur assimilation. By contrast, transgenic Indian mustard lines that overexpress a different ATP sulfurylase isoenzyme from Arabidopsis accumulate glutathione and show increased resistance to SeO42− (Pilon-Smits et al., 1999). SeO42− is a toxic analog of SO42− that Indian mustard can reduce via the sulfur pathway to a nontoxic, volatile form. The opposite results could be due to differences in experimental systems. THE SULFATION ROUTE FOR ASSIMILATION: SO42− IS DIRECTLY INCORPORATED INTO PLANT METABOLITES Plant sulfotransferases have been characterized that catalyze the sulfation of flavonol, desulfoglucosinolate, choline, and gallic acid glucoside (Varin et al., 1997). Sulfotransfer is the terminal step in the biosynthesis of these compounds. The function of sulfated flavonol and choline is unknown. Glucosinolates are the compounds responsible for the distinctive taste of mustards. Gallic acid glucoside, also known as turgorin or periodic leaf movement factor, is responsible for triggering nictinastic leaf movement in Mimosa pudica. Sulfation may regulate the process by activating the movement factor. The number of sulfated compounds in plants is not known. In contrast, sulfation plays a key role in the production of growth-regulating peptides in animals. However, recently, a sulfated regulator of cell proliferation, phytosulfokine-α, was identified from plants (Matsubayashi et al., 1997). Several common features of sulfotransferases have emerged from analysis of the enzymes and the encoding cDNAs. The sulfotransferases are strictly dependent upon the phosphorylated APS derivative, PAPS, as a SO42− donor, and they all have remarkably high affinity for PAPS. They all contain two highly conserved, sulfotransferase signature sequences that may be involved in PAPS binding. Last, they are all localized in the cytoplasm or, in the case of the gallic acid glucoside sulfotransferase, to the inner surface of the plasma membrane. Enzymes that synthesize PAPS have been described in plants. But how PAPS is supplied to the sulfotransferases is still to be determined. PAPS is formed through ATP-dependent phosphorylation of the 3′-hydroxyl group of APS, catalyzed by APS kinase (Reaction 3). APS+ATP→PAPS+ADP(Reaction 3) Cytoplasmic ATP sulfurylase exists in most plants, but a cytoplasmic form of APS kinase has not yet been specifically identified. However, such an enzyme could exist, or at least cannot be ruled out, because in Arabidopsis there are three different genes that encode APS kinase, and the localization of only one of them has been studied (Lee and Leustek, 1998; Schiffman and Schwenn, 1998). An alternative to in situ, cytoplasmic synthesis of APS and/or PAPS is a system in which the sulfonucleotides are exported from chloroplasts. SO42− IS REDUCED BEFORE INCORPORATION INTO Cys AND THE REDUCTION PATHWAY BEGINS WITH APS Eight electrons are required to reduce SO42− to S2−. The process occurs through the sequential action of two different enzymes. The exact mechanism in plants has been vigorously debated, resulting in a confusing proliferation of hypotheses (Hell, 1997). Rather than reiterating the debate, we discuss here only those enzymes that are known with certainty. Our intention is to simplify a confusing topic. Several lines of evidence conclusively demonstrate that SO42− reduction begins with APS in plants and eukaryotic algae. Schmidt (1972) identified an enzyme he called APS sulfotransferase, which has now been completely purified from a marine red macroalga (Kanno et al., 1996). cDNAs that encode APS sulfotransferase have been cloned from a marine green alga and from several higher plants, most notably Arabidopsis (Bick and Leustek, 1998). That the enzyme encoded by the cloned cDNAs was named APS reductase rather than APS sulfotransferase has inadvertently confounded the subject (Gutierrez-Marcos et al., 1996; Setya et al., 1996). Although there were reasonable arguments for the new name, none of these were conclusive enough to warrant abridgment of the original name. Here we submit to historical precedent and refer to the enzyme as APS sulfotransferase. The two names derive from two possible catalytic mechanisms, neither of which has yet been confirmed. As proposed by Schmidt (1972), the sulfotransferase transfers SO42− from APS to a thiol compound, generating a thiosulfonate. If GSH were used, the product would be S-sulfoglutathione (Reaction 4). APS+GSH→GS−SO3−+H++AM(Reaction 4) By contrast, a reductase would be expected to transfer electrons from two GSH to APS, generating free sulfite (SO32−) and GSSG (Reaction 5). APS+2GSH→SO32−+2 H++GSSG+AMP (Reaction 5) It is clear from the amino acid sequence of APS sulfotransferase that, if it functions as a sulfotransferase, it belongs to a different class than the enzymes described earlier in relation to sulfation. They do not share any common sequence motifs. Rather, based on its amino acid sequence and its function, APS sulfotransferase appears to belong to a family of thiol-dependent reductases and thioltransferases. GSH PROVIDES ELECTRONS FOR THE FIRST REDUCTION STEP Mounting evidence indicates that GSH is the most likely in vivo reactant. APS sulfotransferase shows a Km[GSH] that is consistent with the in vivo concentration of GSH; it appears to be unable to efficiently use other common biological thiol compounds such as thioredoxin (Prior et al., 1999), and a domain of the enzyme is a GSH-dependent reductase that functions in a manner similar to that of glutaredoxin (Bick et al., 1998). Although it is functionally related to glutaredoxin, the amino acid sequence of the domain shows greater homology to thioredoxin. The distinction between thioredoxin and glutaredoxin is more a matter of the preferred electron source than of the sequence. For example, only glutaredoxin is able to use GSH. In this respect, it may be of some significance that glutaredoxin catalyzes reduction of disulfide substrates through a thioltransferase mechanism, i.e. the thiol is transferred with the formation of a glutathione-mixed disulfide intermediate. The analogous reaction for APS sulfotransferase would be the one depicted in Reaction 4. SO32− can be produced under the reducing conditions in the chloroplast becauseS-sulfoglutathione is readily reduced nonenzymatically in the presence of excess GSH (Reaction 6) (Schürmann and Brunold, 1980). Based on these considerations we think it is very likely that APS sulfotransferase functions as a GSH:APS sulfotransferase. This hypothesis is depicted in Figure 2. GS− SO32−+GSH↔SO32−+GSSG+H+(Reaction 6) SO32+ REDUCTASE COMPLETES THE REDUCTION OF SULFUR WITH ELECTRONS FROM REDUCED Fd In considering the next reduction step it is significant thatS-sulfoglutathione and SO32− could be available in plastids. Plant SO32− reductase catalyzes the reduction of SO32− using electrons donated from reduced Fd (Reaction 7). The enzyme has been convincingly demonstrated by purification and cloning of the corresponding gene and cDNA (Bork et al., 1998). SO32− reductase shows a high affinity for SO32−(Km = approximately 10 μm), which would serve well for efficient metabolism of SO32−. SO32−+6 Fdred→S2−+6 Fdox(Reaction 7) An Fd-dependent enzyme that reduces thiosulonate to thiosulfide has been measured in cell lysates, but it has not been purified or unambiguously demonstrated. The hypothesis that sulfur is reduced as a thiol-bound form, called the “carrier-bound pathway,” has been a tenet of the plant SO42−-assimilation field. As indicated by the preceding discussion, the carrier-bound pathway need not be invoked because there is convincing evidence for an efficient SO32− reductase. However, the existence of thiosulfonate reductase should not be dismissed as improbable, especially since there has not been a concerted effort to demonstrate it conclusively. Cloning of a thiosulfonate reductase cDNA would be definitive. It is our opinion that an excellent opportunity exists for devising a cloning strategy based on functional complementation of an E. coliSO32− reductase mutant if the strain is engineered to express plant Fd and NADPH:Fd oxidoreductase. APS SULFOTRANSFERASE MAY BE A REGULATION POINT IN THE SULFUR-REDUCTION PATHWAY APS sulfotransferase is encoded by a gene family in Arabidopsis (Bick and Leustek, 1998), all of whose members appear to encode plastid-localized enzymes. APS sulfotransferase is localized only in plastids and not in other cellular compartments (Rotte, 1998). No specialization of function has yet been ascribed to the APS sulfotransferase isoenzymes. SO32− reductase is also plastid localized. In Arabidopsis there may be a single gene encoding this enzyme (Bork et al., 1998). There is a great deal of evidence indicating that APS sulfotransferase is a prime regulation point in SO42− assimilation (Brunold and Rennenberg, 1997). The activity of this enzyme changes rapidly in a variety of plant species after sulfur starvation, exposure to reduced sulfur compounds, heavy-metal stress, or other stresses. Heavy metals induce the synthesis of phytochelatins, and high concentrations of metal ions significantly increase the demand for Cys. Recent studies indicate that one potential mechanism for regulating APS sulfotransferase activity may involve changes in the steady-state mRNA level. Sulfur starvation (Gutierrez-Marcos et al., 1996; Takahashi et al., 1997) and heavy-metal treatment (Heiss et al., 1999; Lee and Leustek, 1999) induce the accumulation of APS sulfotransferase mRNA, but the response is limited to roots. By contrast, SO32− reductase does not appear to be appreciably regulated at the mRNA level (Bork et al., 1998). The extent to which the regulation of mRNA abundance is responsible for changes in APS sulfotransferase activity has not yet been adequately explored. INCORPORATION OF REDUCED SULFUR INTO Cys: PROTEIN-PROTEIN INTERACTIONS MAY REGULATE THE PROCESS The incorporation of S2− into Cys is the last step in reductive SO42− assimilation. The reaction is catalyzed by O-acetylserine(thiol)lyase from S2− and OAS (Reaction 8). The synthesis of OAS is catalyzed by Ser acetyltransferase (Reaction 9). OAS+S2−→Cys+acetate(Reaction 8) Ser+acetylCoA→OAS+CoA(Reaction 9) Ser acetyltransferase and OAS(thiol)lyase exist in an enzyme complex known as Cys synthase. The stability of the complex is affected by substrates (OAS disrupts it and S2− stabilizes it), and it appears to form through specific protein-protein interactions (Bogdanova and Hell, 1997). Yet, the free form of each enzyme has catalytic activity, and the complex is not required for channeling of OAS (Droux et al., 1998). Moreover, in chloroplasts the ratio of OAS(thiol)lyase to Ser acetyltransferase is 300:1 (Droux et al., 1998, and refs. therein); therefore, only a fraction of the total OAS(thiol)lyase can be associated in the complex. One clue to the function of the complex is that association with OAS(thiol)lyase changes the kinetic behavior of Ser acetyltransferase from the Michaelis-Menten type to positive cooperativity with respect to its substrates, Ser and acetyl-CoA (Droux et al., 1998). This suggests that OAS (thiol)lyase functions as a regulatory subunit that regulates Ser acetyltransferase in response to OAS and S2−. Positive cooperativity is a form of allosteric regulation in which the velocity of a bisubstrate enzyme is highly sensitive to small changes in substrate concentration. One can think of the enzyme as having a hair-trigger control mechanism. The idea is appealing because Cys synthesis requires coordination of two converging pathways. If there is insufficient S2− resulting from low activity of SO42− reduction, the concentration of OAS will increase, causing dissolution of the Cys synthase complex. By contrast, overactivity of SO42− reduction results in overabundance of S2− and a shortage of OAS, a condition that would stabilize the complex. Ser acetyltransferase activity would be regulated, its velocity becoming less or more sensitive to its own substrates. Another possible form of regulation is an increase in the steady-state mRNA level for the plastid form of Ser acetyltransferase after sulfur starvation (Takahashi et al., 1997; Noji et al., 1998). Unlike the earlier steps in the pathway in which mRNA regulation occurs primarily in roots, plastid Ser acetyltransferase mRNA increases primarily in leaves. A third possible regulation mechanism is the feedback inhibition of Ser acetyltransferase by Cys. However, only the cytosolic isoform appears to be regulated in this way (Noji et al., 1998). Ser acetyltransferase and OAS(thiol)lyase are the only sulfur assimilation enzymes localized in three compartments: the plastids, cytosol, and mitochondria. cDNAs have been cloned from a range of plant species encoding all of the different isoenzymes (Saito et al., 1994). The multifarious localization of Cys synthase enzymes presents a problem in that only plastids contain the full complement of enzymes needed for SO42− reduction. It may be that S2− is exported from plastids to supply the substrate required by cytosolic and mitochondrial Cys synthases. Evidence for a specialization of function was obtained by analysis of the cytosolic form of OAS(thiol)lyase from Arabidopsis. It is expressed predominantly in roots, in the vascular parenchyma, and in cortical cells (Gotor et al., 1997). Expression in leaves is concentrated in trichomes. Gotor et al. (1997) noted that the toxic heavy-metal Cd is known to accumulate in trichomes of exposed plants, so it could be that the cytosolic OAS(thiol)lyase has a specialized role, supplying Cys for a detoxification mechanism. NEGATIVE AND POSITIVE SIGNALS REGULATE SO42− ASSIMILATION From the preceding discussion it is evident that SO42− reduction and assimilation into Cys is regulated in plants by a range of mechanisms that include substrate availability, modulation of enzyme activity, and gene expression. Since the exclusive function of SO42− reduction is to produce Cys, future studies must concentrate on the question of how the range of regulation mechanisms for individual enzymes is coordinated. Current thinking focuses on both negative and positive signals. It has been known for some time that reduced sulfur compounds such as Cys and glutathione, when applied to plants, repress the activity of the sulfur-assimilation enzymes. Conversely, SO42− starvation induces the activity of certain enzymes. With the availability of DNA probes it was discovered that the steady-state levels of mRNA for SO42− transporter, ATP sulfurylase, and APS sulfotransferase decline after the application of glutathione or Cys to plant roots (Smith et al., 1997; Lappartient et al., 1999; Lee and Leustek, 1999). In contrast, SO42− starvation increases the steady-state level of mRNA for these proteins. Whether these responses are due to transcriptional or posttranscriptional mechanisms is not yet known. One form of reduced sulfur, glutathione, could act as an endogenous signal because it is known to be transported through the phloem of plants and its level in phloem sap is markedly reduced after short-term sulfur starvation. Further support for this hypothesis comes from the “split-root” experiments of Lappartient et al. (1999) in which a portion of the root system was sulfur starved. In another portion of the root system fed normal levels of sulfate, the steady-state mRNA level and activity for SO42−transporter and ATP sulfurylase increased at precisely the time that the level of transported glutathione declined. The implication is that glutathione acts as a negative signal or repressor. Positive regulation could be a derepression phenomenon caused by a decrease in repressor. Recent experiments indicate, however, that a positive signal may also exist. When OAS was fed to roots of barley, the steady-state mRNA level for the high-affinity transporter increased coordinately with SO42−-transport activity (Smith et al., 1997). The response was more rapid than when plants were starved for sulfur, but the magnitude of the increase was smaller. OAS feeding also caused the level of reduced sulfur compounds to increase, possibly explaining why the OAS-induced increase in transporter expression was attenuated compared with SO42− starvation. 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Z Pflanzenphysiol 100 1980 257 268 Google Scholar Crossref Search ADS WorldCat 32 Setya A Murillo M Leustek T Sulfate reduction in higher plants: molecular evidence for a novel 5′-adenylylphosphosulfate (APS) reductase. Proc Natl Acad Sci USA 93 1996 13383 13388 Google Scholar Crossref Search ADS PubMed WorldCat 33 Smith FW Hawkesford MJ Ealing PM Clarkson DT Vanden Berg PJ Belcher AR Warrilow AG Regulation of expression of a cDNA from barley roots encoding a high affinity sulphate transporter. Plant J 12 1997 875 884 Google Scholar Crossref Search ADS PubMed WorldCat 34 Stewart EJ Åslund F Beckwith J Disulfide bond formation in the Escherichia coli cytoplasm: an in vivo role reversal for the thioredoxins. EMBO J 17 1998 5543 5550 Google Scholar Crossref Search ADS PubMed WorldCat 35 Takahashi H Yamazaki M Sasakura N Watanabe A Leustek T de Almeida-Engler J Engler G Van Montagu M Saito K Regulation of cysteine biosynthesis in higher plants: a sulfate transporter induced in sulfate-starved roots plays a central role in Arabidopsis thaliana. Proc Natl Acad Sci USA 94 1997 11102 11107 Google Scholar Crossref Search ADS PubMed WorldCat 36 Varin L Marsolais F Richard M Rouleau M Biochemistry and molecular biology of plant sulfotransferases. FASEB J 11 1997 517 525 Google Scholar Crossref Search ADS PubMed WorldCat 37 Zhao JM Williams CC Last RL Induction of Arabidopsis tryptophan pathway enzymes and camalexin by amino acid starvation, oxidative stress, and an abiotic elicitor. Plant Cell 10 1998 359 370 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the U.S. National Science Foundation (grant nos. IBN 9601146 and MCB 9728661 to T.L.), by Grants-in-Aid for Scientific Research from the Ministry of Education, Science, Sports and Culture, Japan, and by the Research for the Future Program (grant no. 96I00302) from the Japan Society for the Promotion of Science (to K.S.). * Corresponding author; e-mail [email protected]; fax 1–732–932–0312. 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)

Wang, Xuemin

doi: 10.1104/pp.120.3.645pmid: 10398699

Phospholipases hydrolyze phospholipids, which are the backbones of biological membranes. The activities of these enzymes not only have a profound impact on the structure and stability of cellular membranes but also play a pivotal role in regulating many critical cellular functions. The activation of phospholipases is involved in many cell-signaling cascades. These enzymes often execute their regulatory functions through the generation of second messengers that transduce biotic and abiotic cues into physiological responses. Three classes of phospholipases, PLD, PLC, and PLA2 (Fig.1), have been studied extensively for their roles in generating lipid and lipid-derived messengers. PLC and PLA2 are two well-documented signaling enzymes in animal systems. PLC produces the second messengers DAG and inositol phosphate, and PLA2 catalyzes the rate-limiting step in eicosanoid synthesis and regulation. In the last several years, PLD has been identified as an important signaling enzyme that produces both PA and a free-head group such as choline (Fig. 1). This activity has been proposed to play a role in mediating a wide range of cellular processes, including hormone action, membrane trafficking, cell proliferation, cytoskeletal organization, defense responses, differentiation, and reproduction (Rose et al., 1995; Cockcroft, 1997;Colley et al., 1997; Exton, 1997; Fan et al., 1997; Ritchie and Gilroy, 1998). Fig. 1. Open in new tabDownload slide Sites of cleavage of phospholipids by PLD, PLC, and PLA2, and the products of PLD, PA, and head group (H). Fig. 1. Open in new tabDownload slide Sites of cleavage of phospholipids by PLD, PLC, and PLA2, and the products of PLD, PA, and head group (H). Significant progress was made recently toward understanding the structure, regulation, and function of PLD. Plant PLD is an “old” enzyme receiving renewed attention, mainly because of its potential role in transmembrane signaling. It was discovered in plants about one-half century ago, and some distinct and perplexing properties of its activity were soon noted (for review, see Heller, 1978; Wang, 1997). This conventional PLD is widespread in plant tissues and has been purified from several species; however, its regulation and physiological function remained an enigma. The molecular cloning of the first eukaryotic PLD from plants helped to propel the investigation to the molecular realm (Wang et al., 1994; Hammond et al., 1995; Rose et al., 1995). The identification, cloning, and expression of novel types of plant PLDs established that they are a family of heterogeneous enzymes that differ in catalytic and regulatory properties (Pappan et al., 1997a,1997b, 1998; Qin et al., 1997). In addition, the regulated activation of PLD was recently documented in several plant systems, including wounding, hormone action, and plant-pathogen interactions (Ryu and Wang, 1996; Fan et al., 1997; Lee et al., 1997; Ritchie and Gilroy, 1998). The genetic manipulation of PLD in the cell was achieved in plants, mammals, and yeast, and this has provided new insights into the involvement of PLD in cellular functions (Rose et al., 1995; Colley et al., 1997; Fan et al., 1997). With these developments, the role of PLD in signaling cascades has become a topic that attracts increasing attention in various systems (for review, see Wang, 1997; Chapman, 1998; Munnik et al., 1998). THE PLD MULTIPLE GENE FAMILY Eukaryotic intracellular PLD, which was first cloned from castor bean (Wang et al., 1994), is a highly conserved gene family. The conservation of several regions of the PLD amino acid sequences led to the identification and cloning of PLDs from yeast and animals (Hammond et al., 1995; Rose et al., 1995). All cloned PLDs contain two HxKxxxxD motifs, which are separated by approximately 320 amino acids in plant PLDs. The conserved His, Lys, and Asp residues form a catalytic triad responsible for catalysis. The HxKxxxxD motif was also observed in two phospholipid-synthesizing enzymes, bacterial PS synthase and cardiolipin synthase, in endonucleases, and in other proteins of unknown function in viruses and bacteria. The characteristics of the HxKxxxxD motif are used to define the PLD superfamily (Sung et al., 1997). Plant PLD is encoded by a multiple heterologous gene family. Four PLD cDNAs, designated PLDα, PLDβ, PLDγ, and PLDγ2, were isolated from Arabidopsis (Qin et al., 1997). The Arabidopsis genome project also yielded two PLD genes, for which cDNAs are not yet isolated. Multiple PLDs were cloned in rice and cabbage (Morioka et al., 1997;Pannenberg et al., 1998). Database searches in October 1998 found 15 complete PLD cDNA/gene sequences isolated from eight plant species (castor bean, Arabidopsis, cowpea, cabbage, tobacco, Pimpinella brachycarpa, rice, and maize). Alignments of these PLD sequences revealed several distinct clusters. Cluster I included Arabidopsis PLDα and all of the cDNA cloned to date from other plant species whose sequence identity was 75% to 90%. Therefore, PLDs of cluster I are grouped as PLDα, and this classification takes into account sequence similarity, catalytic properties (described in a later section), and gene structure. Cluster II consists of Arabidopsis PLDβ and the two PLD isologs on chromosomes II and IV (tentatively named PLDβ2 and PLDβ3), which share approximately 75% amino acid sequence identity. Cluster III has two members, Arabidopsis PLDγ1 and PLDγ2, which share more than 85% sequence identity. The overall sequence identity shows that PLDβ and PLDγ are more similar to each other than either is to PLDα. Multiple PLD genes also occur in other systems. In mammalian cells, two distinct PLDs were cloned, PLD1 and PLD2. PLD1 has two alternative splicing variants, PLD1a and PLD1b (Hammond et al., 1995; Colley et al., 1997). Two PLDs were reported in yeast, but only the sequence of PLD1 was identified (Rose et al., 1995; Waksman et al., 1997). The overall domain structures of plant PLDs are similar, but important differences occur in some of the motifs (Fig.2). A C2 domain is present in all cloned plant PLDs, but not in animal or yeast PLDs. C2 is a Ca2+/phospholipid-binding fold, and Ca2+ binding is coordinated by four to five amino acid residues provided by bipartite loops (Ponting and Parker, 1996). PLDγ and PLDβ conserve all of the Ca2+-coordinating acidic amino acids (Qin et al., 1997), whereas two of the acidic residues in the C2 domain of PLDα are substituted by either positively charged or neutral amino acid residues, indicating a possible change of affinity for Ca2+ in PLDα. A PPI-binding motif (RxxxxKxRR) and an inverted sequence (RKxRxxxxR) are present in PLDβ near the catalytic domain of the C terminus (Qin et al., 1997). Three of these four basic consensus residues are conserved in PLDγ, whereas PLDα shows the least conservation of residues (some are replaced by acidic residues). PLDγ possesses a myristoylation consensus sequence that is not present in PLDα or PLDβ (Qin et al., 1997). Fig. 2. Open in new tabDownload slide Domain structures of PLDα, PLDβ, and PLDγ in Arabidopsis. XX in the PLDα C2 marks the loss of two acidic residues potentially involved in Ca2+ binding; XX in the PPI-binding motifs marks the loss of the number of basic residues potentially required for PPI binding. Fig. 2. Open in new tabDownload slide Domain structures of PLDα, PLDβ, and PLDγ in Arabidopsis. XX in the PLDα C2 marks the loss of two acidic residues potentially involved in Ca2+ binding; XX in the PPI-binding motifs marks the loss of the number of basic residues potentially required for PPI binding. DISTINCT CATALYTIC PROPERTIES OF DIFFERENT PLDs Molecular analyses have documented not only the occurrence of multiple PLDs but also the structural variations that may underlie distinct biochemical properties. PLD activities from plants can be divided into three groups based on their differing requirements for Ca2+ in vitro. The first group is the conventional plant PLD that displays a striking Ca2+ requirement; it is most active at millimolar concentrations of Ca2+, with the optimal concentration ranging from 20 to 100 mm (Heller, 1978). PLDα expressed from the castor bean PLDα cDNA exhibits the characteristic activity of conventional PLD purified from plants (Dyer et al., 1994; Wang et al., 1994; Pappan et al., 1998). Antisense suppression of PLDα in Arabidopsis led to the loss of this conventional PLD activity (Pappan et al., 1997a), so the PLDα gene product must have been responsible for it. Additionally, three isoforms and two cDNAs of the conventional PLD were also identified in some plant species (Dyer et al., 1994; Young et al., 1996; Pannenberg et al., 1998). The second group of PLDs includes those that are the most active at micromolar levels of Ca2+. The presence of such PLD activity was documented in transgenic Arabidopsis, in which the expression of PLDα was suppressed by an antisense gene (Pappan et al., 1997a). The cloning and analysis of PLDβ from Arabidopsis provided unequivocal, molecular evidence for the new type of PLD (Pappan et al., 1997b). The PLDγ that was cloned later also exhibited a Ca2+ dependence similar to that of PLDβ (Qin et al., 1997). These PLDs are PPI dependent and are stimulated by PIP2 and to a lesser extent by PIP, but not by other acidic phospholipids such as PI, PS, PG, and PA. Although the above PLDs require Ca2+ for activity, a third type that is independent of cations was reported inCatharanthus roseus suspension cells (Wissing et al., 1996). Another unique property of this PLD is its lack of transphosphatidylation activity: Two membrane-associated and two soluble variants of this activity have been noted. However, to our knowledge, it has not been purified to homogeneity, and no PLD cloned thus far exhibits such activity. This third type of PLD also differs in substrate specificity and preferences. Conventional PLD uses more than one phospholipid as a substrate. In general, PC, PE, and PG are good substrates, whereas PI, PS, cardiolipin, and plasmalogens are much less efficiently used, if at all (Heller, 1978; Dyer et al., 1994; Abousalham et al., 1997). PLDα, PLDβ, and PLDγ all use PC, PE, and PG as substrates, but the reaction conditions required for PLDβ and PLDγ are strikingly different from those for PLDα (Pappan et al., 1998). PLDβ and PLDγ, but not PLDα, can use PS and NAPE as substrates. Although PLDβ and PLDγ hydrolyze the same substrates, PLDγ prefers ethanolamine-containing PE and NAPE to other lipids, but PLDβ does not. The Ca2+-independent PLD from C. roseus exhibits a unique substrate specificity (Wissing et al., 1996). It is PI specific, which is in contrast to cloned PLDα, PLDβ, and PLDγ, which do not hydrolyze PI. These varied substrate specificities and preferences suggest that the activation of different PLDs may result in selective hydrolysis of membrane phospholipids. REGULATION AND ACTIVATION OF PLD Their distinct structural and biochemical properties suggest that PLD isoenzymes are subject to unique controls and activation mechanisms. The different Ca2+ requirements could mean that changes in the levels of cytoplasmic Ca2+ activate PLD isoenzymes differentially. However, the fact that the conventional PLD (PLDα) requires millimolar levels of Ca2+ in vitro casts doubt upon the significance of Ca2+ in controlling its activity in vivo. It is important to note that the optimal Ca2+ concentration was determined by using a single class of lipid substrate often in the presence of organic solvents or detergents such as SDS, which are artificial conditions. A recent study showed that PLDα was active at nearly physiological Ca2+ concentrations when it was assayed at an acidic pH (4.5–5.0) and in the presence of mixed lipid vesicles containing PIP or PIP2 (K. Pappan and X. Wang, unpublished data). This suggests that even though the effect of Ca2+ on PLDα is complex, its activity can be increased by elevating cellular Ca2+ levels. On the other hand, PLDβ and PLDγ were inactive at that pH and were most active at a neutral pH. These distinct pH optima may mean that changes in cellular pH have a different effect on PLD isoforms. At near-physiological concentrations of Ca2+, PLDβ and PLDγ are neutral phospholipases, whereas PLDα is an acidic phospholipase that may be activated by cellular acidification. The presence of a C2 domain in plant PLDs points to a specific mode of activation by Ca2+. C2 domains were identified in a number of signal transduction and membrane trafficking proteins, such as PKC, PLC, and PLA2 (Ponting and Parker, 1996). This domain is important in the Ca2+-regulated translocation of proteins to membranes. Indeed, in the wound activation of PLD in castor bean, the Ca2+-mediated translocation of PLD from the cytosol to the membranes had already been proposed before the presence of a C2 domain on PLDs was recognized (Ryu and Wang, 1996). There is also data suggesting Ca2+-mediated activation of PLD in vivo (Munnik et al., 1998). Another potential regulator of plant PLD is PPI. Not only do PLDβ and PLDγ require PPIs for activity, PLDα activity is also stimulated by PPIs when low levels of Ca2+ are present (Qin et al., 1997). Binding assays have shown that PLDβ, PLDγ, and PLDα are able to bind PIP2. Two PLD regions may be involved in PIP2 binding: one is the near N-terminal C2 domain and the other is the near C-terminal PPI-binding motifs that are missing in PLDα. PPIs are minor lipids and their levels are regulated dynamically. The activation of PLD is likely to be interconnected with the metabolism and signaling of PPIs. It was also suggested that plant PLD is regulated by trimeric G-proteins based on its stimulation by mastoparan, cholera toxin, and alcohol (Munnik et al., 1995; Chapman, 1998). By comparison, the activation by small G-proteins such as ARF (ADP-ribosylation factor) and Rho is the best-characterized mechanism of regulation for mammalian PLD. ARF, Rho, and PKC synergistically activate PLD1 but not PLD2. These proteins may promote PLD1 activity via direct protein-to-protein interactions. Animal PLD is also stimulated by other factors, including Ca2+ flux, PKC, receptor-linked Tyr kinases, PIP2, and gelsolin, and it is down-regulated by fodrin, clathrin assembly protein 3, synaptojanin, ceramide, and some lysophospholipids (Fig.3; Cockcroft, 1997; Exton, 1997). LysoPE was also suggested to be a negative regulator of plant PLD (Ryu et al., 1997). The formation of DAG-PPi from PA is thought to attenuate plant PLD activation (Munnik et al., 1998). In addition, PLD gene expression and the differential appearance of PLD isoforms are involved in the long-term regulation of PLD (Dyer et al., 1994; Young et al., 1996; Fan et al., 1997). Fig. 3. Open in new tabDownload slide Schematic diagram of up- and down-regulation of PLD in plants and animals, showing that PLD signaling can be regulated by modulating PLD activity or by removing PA. The proteinaceous stimulators and inhibitors identified are mainly from animal systems. Fig. 3. Open in new tabDownload slide Schematic diagram of up- and down-regulation of PLD in plants and animals, showing that PLD signaling can be regulated by modulating PLD activity or by removing PA. The proteinaceous stimulators and inhibitors identified are mainly from animal systems. The activation of PLD in animal systems was first identified just over 10 years ago, and it is now documented in more than 30 cell types stimulated by receptor-directed agonists and by other stimuli such as Ca2+ ionophores and phorbol esters (Cockcroft, 1997; Exton, 1997). Although, historically, the activation of PLD was observed first in plants, studies of PLD activation in plants now lag behind those in animals. It has long been known that wounds and other stresses stimulate a rapid increase in PA and other lipid metabolites. These increases were regarded initially as autolysis resulting from the release of PLD and other lipolytic enzymes during cell damage. Recent studies have shown that wounding a tissue triggers a rapid activation of PLD-mediated phospholipid hydrolysis not only at the wound site but also at undamaged areas (Ryu and Wang, 1996). Stimulation of plant PLD has also been shown in response to treatments with ABA, light, fungal elicitors, and bacterial pathogens (Young et al., 1996; Fan et al., 1997; Chapman, 1998; Munnik et al., 1998; Ritchie and Gilroy, 1998). DOWNSTREAM TARGETS OF PLD-DERIVED MESSENGERS Identification of the downstream events of PLD activation is important to our understanding of PLD function. PA stimulation of signaling proteins is the most-studied mechanism of action in animals. One group of such proteins is protein kinases, including Ca2+-dependent and independent kinases such as PKC, mitogen-activated protein kinases, and Raf-kinases (Cockcroft, 1997; Exton, 1997). PA can bind to Raf-kinase, but it is unclear how this binding may activate the enzyme. Recent reports indicate the presence of a PA-specific protein kinase that mediates the activation of NADPH oxidase (Waite et al., 1997). Other enzymes activated by PA are PIP-5 kinase, PLC, and PLA2, which are involved in lipid-signaling cascades. In addition to performing as a direct messenger, PA can be metabolized further to other lipid mediators (DAG, lysoPA, and free fatty acids; Fig.4). The head group released by PLD can also have regulatory functions. The formation ofN-acylethalonamine by PLD was implicated in the responses of plants to fungal elicitation (Chapman, 1998). Fig. 4. Open in new tabDownload slide Direct and derived products of PLD activation. LysoPA and free fatty acid (FA) can be formed from PA by nonspecific acyl hydrolase or by PLA. PA is dephosphorylated to DAG by PA phosphatase. CDP-DAG is the precursor for the synthesis of PS, PI, and PG. XOH, Primary alcohol used for transphosphatidylation; Ptd, phosphatidyl; NAE, N-acylethanolamine. Fig. 4. Open in new tabDownload slide Direct and derived products of PLD activation. LysoPA and free fatty acid (FA) can be formed from PA by nonspecific acyl hydrolase or by PLA. PA is dephosphorylated to DAG by PA phosphatase. CDP-DAG is the precursor for the synthesis of PS, PI, and PG. XOH, Primary alcohol used for transphosphatidylation; Ptd, phosphatidyl; NAE, N-acylethanolamine. Some of the cellular roles of PA may result from its effect on membrane properties and configuration rather than from its direct effect on proteins. PA is a nonbilayer lipid favoring hexagonal phase formation, particularly in the presence of Ca2+ (Cornell and Arnold, 1996). Thus, a rapid increase in the local concentration of PA may destabilize membranes. The activities of a number of signaling proteins, including G-proteins, PKC, PLC, PLA, PA phosphatase, DAG kinase, and PLDs, are sensitive to changes in membrane conformation (Cornell and Arnold, 1996; Pappan et al., 1998). An increase in PA also increases the net negative charge of membranes, which may alter protein-to-membrane interactions and the flux of ions such as Ca2+. In addition, PA-mediated changes in membrane properties may be produced by altering membrane lipid composition, because PA is a central precursor in glycerolipid biosynthesis (Fig. 4). INVOLVEMENT OF PLD IN SIGNALING PATHWAYS It has been suggested that PLD plays a role in a broad range of cellular responses, but the requirement of PLD for a particular cellular function was not documented conclusively until recently. The molecular cloning of plant PLD helped to identify the first definitive requirement of PLD in a physiological process. It was noted that the sequence of the yeast sporulation-defective mutant SPO14 contains several regions of sequence similarity to the then newly cloned castor bean PLD, and this gene was later found to encode PLD1 (Rose et al., 1995). Both PLD1 activity and its presence in the nucleus are necessary for signaling the completion of meiosis (Sung et al., 1997). Whether plant PLDs are involved in a similar process is not known. Antisense suppression of plant PLD resulted in a loss of more than 90% of the PLDα in Arabidopsis flowers. But the fertility of PLDα-suppressed plants was not affected, indicating that a high level of PLDα is not essential for reproduction (Fan et al., 1997). Recent studies provide strong evidence of a role for PLDα in ABA action. The expression of PLDα is up-regulated by ABA, as indicated by the increased levels of PLDα promoter activity, mRNA, protein, and membrane-associated activity in response to ABA treatments (Fan et al., 1997; Wang, 1997; Xu et al., 1997). Senescence of the leaves detached from the PLDα-deficient transgenic plants was retarded when they were incubated with ABA (Fan et al., 1997). These data indicate that PLDα is a mediator in ABA actions; the loss of PLDα activity in transgenic plants renders Arabidopsis less sensitive to ABA. A role for PLD/PA in ABA signaling was also indicated in an independent study that used a different system (Ritchie and Gilroy, 1998). ABA increased PLD activity after it was applied to barley aleurone protoplasts. Direct application of PA to aleurone protoplasts suppressed the production of α-amylase and increased the synthesis of an amylase inhibitor in a manner that mimicked the ABA antagonism of GA-induced events in barley aleurone. The fact that an ABA-mediated physiological process is changed by the genetic and pharmacologic alteration of PLD activity suggests that PLD constitutes an early step in mediating ABA action. PLD has also been implicated in the action and production of ethylene. Antisense suppression of PLDα decreases the rate of ethylene-promoted senescence in detached Arabidopsis leaves (Fan et al., 1997). In cultured carrot cells, PLD activation is thought to constitute a signaling step in the perception of an ethylene burst that occurs at the early stage of Glc starvation (Lee et al., 1998). LysoPE is proposed to retard senescence by blocking PLDα activity, which may be involved in promoting the burst of ethylene (Ryu et al., 1997). The involvement of PLD in injury-induced lipid hydrolysis is perhaps the earliest result connecting PLD to a cellular process. PLD can be activated rapidly by stress injuries such as mechanical wounding, frost, and γ-irradiation (Voisine et al., 1993; Ryu and Wang, 1996). Apparently, wound activation of PLD results from its translocation to membranes, which is mediated by an increase in cytoplasmic Ca2+upon wounding (Ryu and Wang, 1998). PLD activation is proposed to be an early event in the response of the plant to stress injuries, and the PLD-generated PA may serve as an effector or as a substrate for the production of other mediators such as DAG, polyunsaturated fatty acids, and oxylipins in defense signaling (Ryu and Wang, 1996, 1998). The role of PLD in defense signaling extends to plant-pathogen interactions. In rice leaves challenged with the bacterial pathogenXanthomonas oryzae pv oryzae, PLDα clustered at the region of the plasma membranes that came into contact with bacteria during hypersensitive interactions but not in the susceptible interactions (Young et al., 1996). In tobacco cells treated with the fungal elicitor xylanase, a rapid release of N-acyl ethanolamine was noted (Chapman, 1998), which probably resulted from hydrolysis by PLDγ or PLDβ but not by PLDα, because the former two hydrolyze NAPE, and PLDγ prefers NAPE or PE over other phospholipids (Pappan et al., 1998). One potential mechanism by which PLD participates in plant-defense responses is the regulation of NADPH oxidase, which is involved in reactive oxygen production. In neutrophils, the activation of PLD is known to mediate an oxidative burst, and PA is a potent activator of NADPH oxidase (Waite et al., 1997). NADPH oxidase is a complex composed of membrane-bound and cytosolic proteins. It becomes active when its cytosolic subunits translocate to the membrane, and the translocation of p47-phox is prompted by phosphorylation. Recent studies show that p47-phox is a substrate for the newly identified PA-activated protein kinase in animals (Waite et al., 1997). Plant NADPH oxidase and neutrophil NADPH oxidase seem to have the same subunit components. In addition, phosphorylation and translocation of plant p47-phox and p67-phox also occur in tomato cells treated with race-specific fungal elicitors (Xing et al., 1997). However, whether PLD and PA play a role in regulating plant NADPH oxidase activity is unclear. One study using soybean suspension-cultured cells failed to obtain evidence for the involvement of PLD in the pathogen-elicited production of hydrogen peroxide (Taylor and Low, 1997). FUNCTIONAL HETEROGENEITY AND CROSSTALK OF LIPID SIGNALING PATHWAYS The occurrence of multiple PLDs with distinct regulatory and catalytic properties in the same organism suggests that each may have unique functions. Some evidence for distinct functions was obtained from the genetic manipulation of PLD in plant, animal, and yeast systems. The transfer of a PLDα antisense cDNA into Arabidopsis resulted in the loss of more than 95% of PLDα activity, but PLDβ and PLDγ activities in the PLDα-deficient leaves were not reduced significantly (Pappan et al., 1997a). The PLDα antisense leaves displayed a marked retardation in ABA- or ethylene-promoted senescence, indicating that the loss of PLDα was not compensated for by PLDβ and PLDγ (Fan et al., 1997). The yeast SPO14 mutant was found to contain another PLD activity, designated PLD2, and thus disruption of the PLD1 function was not compensated for by the PLD2 gene (Waksman et al., 1997). Overexpression of mammalian PLD2 resulted in cytoskeletal reorganization, whereas an increase in PLD1 expression did not alter cell morphology (Colley et al., 1997). It is now evident that more than one phospholipase is often involved in mediating a specific cellular response. PLD is thought to function as an integral part of a network involving other lipid-signaling enzymes such as PLA2 and PLC (Fig.5). Mitogenic signaling in animal cells, for example, involves both PIP2-PLC and PC-PLD, and the activation of PLC results in the initial rise of DAG, whereas PLD coupled with PA phosphatase provides the sustained supply of DAG required for cell proliferation (Exton, 1997). On the other hand, PA is a stimulator of PLC, PLA2, and PKC. Fig. 5. Open in new tabDownload slide A working model depicting the networking of PLD activation with other lipid mediators and signaling enzymes. Plus signs indicate stimulation, and minus signs denote inhibition. IP3, Inositol 1,4,5-trisphosphate; ptase, phosphatase; PUFAs, polyunsaturated fatty acids. Fig. 5. Open in new tabDownload slide A working model depicting the networking of PLD activation with other lipid mediators and signaling enzymes. Plus signs indicate stimulation, and minus signs denote inhibition. IP3, Inositol 1,4,5-trisphosphate; ptase, phosphatase; PUFAs, polyunsaturated fatty acids. The network of PLD, PLC, and PLA2 generates several potent lipid mediators, such as PA, lysophospholipids, DAG, and free polyunsaturated fatty acids. Stimulus-induced increases in these lipid metabolites have also been found in some plant systems. Moreover, the formation of PA was shown to precede that of DAG, lysophospholipids, and free fatty acids, suggesting a possible PLD-led activation of acyl hydrolases, PLC, and/or PA phosphatase (Voisine et al., 1993; Ryu and Wang, 1996, 1998; Lee et al., 1997). In addition, PA is a stimulator of PIP-5 kinase, which is responsible for the synthesis of PIP2 (Fig. 5). On the other hand, PIP2 is an activator of plant PLDβ, PLDγ, and some PLDs from animals and yeast (Cockcroft, 1997; Qin et al., 1997). It has been proposed that activation of PLD and PIP-5 kinase in mammalian cells forms a positive feedback loop that leads to rapid generation of PA and PIP2, which are involved in vesicular trafficking. In addition, crosstalk can occur within the PLD family, and the activation of one PLD may stimulate or attenuate the function of another. OUTSTANDING QUESTIONS AND PROSPECTS Recent advances in the investigation of PLDs in plants, animals, and yeast point to an important role for PLD in the mediation of cellular processes; however, many questions remain and a comprehensive understanding of PLD function is yet to be achieved. One major question addresses the molecular and cellular mechanisms by which PLD mediates the cellular functions. An answer requires identification of the cellular targets of PLD activation and the molecules that interact with PLD. Very little, if anything, is known about the downstream reactions or processes (e.g. kinases, phosphatases, ion channels, adapter proteins, and other targets) of PA, PA-derived mediators, and head groups in plant signaling. The paucity of information of the cellular effects of lipid messengers is the major impediment in lipid-signaling research in plants. The finding of multiple PLD proteins indicates that the cellular regulation and the functioning of PLDs are complex. The limited biochemical and genetic data have suggested that the different PLDs may have unique functions. Defining the biochemical properties of each PLD is important to the understanding of its catalysis and regulation in the cell. Arabidopsis contains (potentially) six active PLDs; only three of them, PLDα, PLDβ, and PLDγ, have been analyzed. Important insights into the cellular function of different PLDs can be obtained by determining the spatial and temporal expression and intracellular and cell/tissue localization. However, such information is presently available only for PLDα. Although this article is concerned primarily with the role of PLD in signaling cascades, it is important to note that PLD can participate in other cell functions, such as membrane degradation and remodeling. The early studies of plant PLD functions dealt only with phospholipid breakdown during senescence, aging, and stress injuries; it was suggested that increases of PLD initiated a phospholipid degradation pathway (Voisine et al., 1993; Fan et al., 1997, and refs. therein). This catabolic role could be carried out by different PLD proteins, or the same PLD could exert both degradation and signaling functions, depending on the severity of the stress. With the availability of molecular information for the various PLDs, PLD isoenzyme-specific antibodies and DNA/RNA probes should be forthcoming and will be instrumental in addressing some of the above questions. In addition, the function of different PLDs can be studied effectively by generating and characterizing PLD antisense and knockout transgenic plants. Because of possible genetic redundancy, particularly for PLDβ and PLDγ, producing double or triple mutants may be necessary for an unambiguous determination of the role of PLD in cellular metabolism. With our present knowledge of the molecular biology and biochemistry of this class of enzyme, we are poised for major advancements in the long-sought understanding of the physiological functions of PLDs and the membrane-lipid involvement in plant-signaling cascades. ACKNOWLEDGMENTS I thank Dr. L. Zheng for her comments on and assistance with the figures and apologize to the colleagues whose work was not directly cited because of space limitations. 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This is contribution no. 99-226-J of the Kansas Agricultural Experiment Station. * E-mail [email protected]; fax 1–785–532–7278. 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)

Facchini, Peter J.; Yu, Min; Penzes-Yost, Catherine

doi: 10.1104/pp.120.3.653pmid: 10398700

Abstract Tyrosine decarboxylase (TYDC) is a common plant enzyme involved in the biosynthesis of numerous secondary metabolites, including hydroxycinnamic acid amides. Although a definite function has not yet been determined, amides have been proposed to form a physical barrier against pathogens because they are usually found as integral cell wall components. Canola (Brassica napus) was independently transformed with chimeric genes (35S::TYDC1 and35S::TYDC2) under the transcriptional control of the cauliflower mosaic virus 35S promoter, and encoding two TYDC isoforms from opium poppy (Papaver somniferum). All T0 plants displayed a suppressed level of wild-type TYDC activity, and transgene mRNAs were not detected. Silencing of 35S::TYDC1 was overcome in the T1 progeny of self-pollinated T0 plants, since high levels of TYDC1 mRNAs were detected, and TYDC activity increased up to 4-fold compared with wild-type levels. However, TYDC1 mRNA levels decreased in T2 plants and were not detected in the T3 progeny. TYDC activity also gradually declined in T2 and T3 plants to nearly wild-type levels. In contrast, silencing of 35S::TYDC2 was maintained through four consecutive generations. T1 plants with a 3- to 4-fold increase in wild-type TYDC activity showed a 30% decrease in cellular tyrosine pools and a 2-fold increase in cell wall-bound tyramine compared with wild-type plants. An increase in cell wall-bound aromatic compounds was also detected in these T1plants by ultraviolet autofluorescence microscopy. The relative digestibility of cell walls measured by protoplast release efficiency was inversely related to the level of TYDC activity. Plant responses to pathogens include the induction of numerous metabolic pathways that comprise an arsenal of biochemical and physical defenses. Induction of hydrolytic enzymes such as chitinases and glucanases and the production of low-Mrantimicrobial compounds known as phytoalexins are common biochemical defense responses that increase disease resistance in plants by directly inhibiting the growth of pathogens (Hahlbrock and Scheel, 1989). The deposition of lignin and the cross-linking of Hyp-rich glycoproteins within the polysaccharide matrix of the cell wall are examples of physical defense mechanisms that reduce plant cell susceptibility to penetration by invading pathogens (Showalter et al., 1985; Matern and Kneusel, 1988). TYDC (EC 4.1.1.28) catalyzes the decarboxylation of Tyr to tyramine (Fig. 1) and is widespread in higher plants (Hosoi et al., 1970; Tocher and Tocher, 1972; Marques and Brodelius, 1988; Kawalleck et al., 1993; Trezzini et al., 1993;Facchini and De Luca, 1994). The rapid and transient induction of TYDC mRNAs in response to elicitors and/or pathogens in parsley (Schmelzer et al., 1989; Kawalleck et al., 1993), Arabidopsis (Trezzini et al., 1993), and opium poppy (Papaver somniferum; Facchini et al., 1996) suggests that tyramine serves as a precursor to an important class of plant defense-response metabolites. In opium poppy, TYDC is encoded by a family of 10 to 15 genes that can be categorized into two subgroups based on sequence identity (Facchini and De Luca, 1994; Facchini et al., 1998). Each subgroup consists of approximately six members that share approximately 90% identity at the nucleotide and amino acid levels. In contrast, comparison of subgroup members (represented by TYDC1 and TYDC2) reveals sequence identities of <75%. Although the catalytic properties of the different TYDC isoforms are similar (Facchini and De Luca, 1994), theTYDC gene family exhibits differential and organ- and temporal-specific expression (Facchini et al., 1998). Fig. 1. Open in new tabDownload slide Reactions in the biosynthesis of hydroxycinnamic acid amides that are catalyzed by TYDC and THT. Fig. 1. Open in new tabDownload slide Reactions in the biosynthesis of hydroxycinnamic acid amides that are catalyzed by TYDC and THT. Recent studies have shown that the biosynthesis of hydroxycinnamic acid amides of tyramine and their subsequent polymerization in the cell wall by oxidative enzymes are integral and ubiquitous components of the plant defense response to pathogen challenge (Clarke, 1982;Negrel and Martin, 1984; Negrel and Jeandet, 1987; Negrel and Lherminier, 1987; Negrel et al., 1993a; Schmidt et al., 1998). These amides, together with other cell wall-bound phenolics, are believed to create a barrier against pathogens by reducing the digestibility of the cell wall and/or by directly inhibiting the growth of fungal hyphae. Hydroxycinnamic acid amides, which have been found in a variety of plants (Martin-Tanguy et al., 1978), are formed by the condensation of hydroxycinnamoyl-CoA esters with various amines such as polyamines (e.g. putrescine and spermidine) or tyramine. THT (EC 2.3.1.110) catalyzes the condensation of tyramine and select derivatives of hydroxycinnamoyl-CoA (Fig. 1) and is induced in response to pathogens (Fleurence and Negrel, 1987), elicitor treatment (Villegas and Brodelius, 1990; Schmidt et al., 1998; Yu and Facchini, 1999), and wounding (Negrel et al., 1993a). The enzyme was first isolated from tobacco leaves (Negrel and Martin, 1984) and has been purified to homogeneity from potato (Hohlfeld et al., 1995, 1996), tobacco (Negrel and Javelle, 1997), and opium poppy (Yu and Facchini, 1999). The use of transgenic plants with altered levels of a specific enzyme is a powerful technique with which to study metabolic regulation and to refine our understanding of the physiological roles for secondary metabolic pathways. For example, the co-suppression of PAL activity in transgenic tobacco demonstrated that this enzyme is a rate-determining step in the biosynthesis of phenylpropanoid derivatives, including lignin, and showed that phenolic metabolites are crucial for the resistance of plants to pathogens (Bate et al., 1994; Maher et al., 1994). Introduction of a foreign TDC(Trp decarboxylase) (EC 4.1.1.25) gene into canola (Brassica napus) resulted in the redirection of Trp into tryptamine rather than into indole glucosinolates (Chavadej et al., 1994). In contrast, expression of the same TDC gene in transgenic potato resulted in altered aromatic amino acid biosynthesis and increased susceptibility of the plants to pathogen infestation (Yao et al., 1995). In the present study, we tested the hypothesis that an increase in TYDC activity in canola transformed with chimeric TYDC transgenes would increase the incorporation of tyramine and/or hydroxycinnamic acid amides into cell walls and result in a corresponding decrease in cell wall digestibility. MATERIALS AND METHODS Growth and Transformation of Canola Two TYDC cDNAs from opium poppy (Papaver somniferum cv Marianne) were placed under the transcriptional control of the CaMV35S promoter. pBI35S::TYDC1 was constructed by replacement of GUS between the KpnI andSacI sites of pBI122 with a KpnI/SacI fragment from pBluescript (Stratagene) containing the TYDC1 cDNA (Facchini and De Luca, 1994). pBI35S::TYDC2 was constructed by replacement of GUS between the XbaI andSacI sites of pBI122 with a XbaI/SacI fragment from pBluescript containing the TYDC2 cDNA (Facchini and De Luca, 1994). pBI122 is a modified version of pBI121 (CLONTECH, Palo Alto, CA) containing the restriction sites ApaI,XhoI, and KpnI in an adapter fragment that was inserted into the SmaI site between the 35Spromoter and GUS. pBI122 also harbors the NPT IIgene for kanamycin resistance under the control of the constitutiveNOS (nopaline synthase) promoter. Plasmids were sequenced through the 35Spromoter-TYDC junction to verify construct assembly. pBI35S::TYDC1 and pBI35S::TYDC2 were mobilized in the disarmed Agrobacterium tumefaciens strain LB4404 by direct DNA transfer (An, 1987) and used to transform canola (Brassica napus cv Westar) by the cotyledonary petiole method (Moloney et al., 1989). Plants were maintained in a growth chamber at a PPFD of 400 μE m−2s−1 and a light/dark regime of 16 h (21°C)/8 h (15°C). Regenerated plants were tested for integration of chimeric TYDC and NPT II genes into the canola genome, TYDC and NPT II enzyme activities, and the presence of TYDC mRNAs. Nucleic Acid Isolation and Analysis Genomic DNA was extracted by grinding 100 mg of leaf tissue in 400 μL of 200 mm Tris-HCl, pH 7.8, 250 mm NaCl, 0.5% SDS, and 25 mm EDTA. Debris were removed by centrifugation, and DNA was precipitated with an equal volume of isopropanol and recovered by centrifugation. The pellet was rinsed in 70% ethanol, dried, and redissolved in water. Fragments ofTYDC1 and TYDC2 were amplified by 30 PCR cycles from 100 ng of genomic DNA using a primer-annealing temperature of 55°C and specific oligonucleotides designed from published sequences (TYDC1 sense primer, AGGGACTACTTGTGAAGCCA; TYDC1 antisense primer, ACTGATTCAAGCAATTTCGC; TYDC2 sense primer, ACTTCTTAGCTGATTATTAT; TYDC2 antisense primer, ACGGCATGAGTCATGTAAAC; Facchini and De Luca, 1994). PCR products were analyzed on 1% agarose gels containing ethidium bromide. Total RNA was isolated according to the method of Logemann et al. (1987), and 15 μg was fractionated onto 1.0% formaldehyde agarose gels before transfer to nitrocellulose membranes (Sambrook et al., 1989). RNA blots were hybridized to random-primer32P-labeled (Feinberg and Vogelstein, 1984) full-length TYDC1 or TYDC2 cDNAs at 65°C in 0.25 m sodium phosphate, pH 8.0, 7% SDS, 1% BSA, and 1 mm EDTA. Blots were washed at 55°C, twice with 2× SSC containing 0.1% SDS and twice with 0.2× SSC containing 0.1% SDS (Sambrook et al., 1989) (1× SSC: 0.15m NaCl and 0.015 m sodium citrate, pH 7.0). RNA blots were autoradiographed with an intensifying screen on Kodak X-OMAT film at −80°C. TYDC, THT, and NPT II Assays For TYDC and THT assays, plant tissues were frozen under liquid nitrogen and ground to a fine powder with a mortar and pestle. Powdered tissues were extracted with 200 mm Tris-HCl, pH 7.8, debris was removed by centrifugation, and the supernatant was desalted using a PD-10 column (Pharmacia). TYDC activity was assayed by measuring the release of 14CO2 froml-[carboxyl-14C]Tyr (Facchini and De Luca, 1994). The TYDC assay contained 50 mm Tris-HCl, pH 7.8, 1 mmEDTA, 25 μm pyridoxal-1-phosphate, 0.1 μCi (specific activity = 55 mCi mmol−1) [14C]Tyr, and 500 μL of protein extract (total volume = 1 mL) in an airtight vial. Reactions were incubated for 60 min at 35°C with constant agitation. Enzymatically liberated 14CO2 was trapped on quaternary ammonium-saturated GF/D filters suspended above the reaction solution. Reactions were stopped by the addition of 0.2n HCl and agitated for an additional 1 h before scintillation counts from GF/D filters were determined. 4-Coumaroyl-CoA for the THT assay was enzymatically synthesized using total protein extract from Escherichia coli harboring pQE19, which expresses a recombinant tobacco 4-coumarate:coenzyme A ligase (4CL) (Lee and Douglas, 1996). The synthesis reaction consisted of 0.1 mm CoA, 0.2 mm 4-coumaric acid (Sigma), 2.5 mm ATP, 1 mm DTT, and 300 mg of total bacterial protein extract (Meng and Campbell, 1997). After 1 h of incubation, the synthesized 4-coumaroyl-CoA was purified using a Sep-Pak C18 column (Waters). The 4-coumaroyl-CoA was concentrated, and its identity and purity were confirmed by TLC and comparison of the UV spectrum with that of an authentic standard. THT activity was measured as described previously (Yu and Facchini, 1999). Ninety microliters of desalted enzyme extract in 50 mm Tris-HCl, pH 7.8, was incubated for 1 h with 0.5 μCi of [8-14C]tyramine and 100 nmol of 4-coumaroyl-CoA. Reactions were stopped by the addition of 1.0m HCl, and 20 μL was applied to a silica gel 60 F254 TLC plate that was subsequently developed in chloroform:methanol (5:4). The developed TLC plate was autoradiographed for 12 h. Radiolabeled spots corresponding to 4-coumaroyltyramine (RF = 0.82) were scraped off the plate and radioactivity was quantified by liquid scintillation counting. A dot-blot assay was used to determine the level of NPT II activity (Radke et al., 1988). Leaf tissue (100 mg) was extracted in 100 μL of 50 mm sodium phosphate, pH 7.0, 14 mmβ-mercaptoethanol, 10 mm EDTA, 0.1% Sarcosyl, and 0.1% Triton X-100. The soluble protein extract was incubated in 15 mm Tris-maleate, pH 7.0, 10 mmMgCl2, 100 mmNH4Cl, and 0.5 mm DTT with 10 μCi of [γ-32P]ATP (specific activity, 3000 Ci mmol−1) in the presence or absence of 0.1 mg mL−1 kanamycin at 37°C for 1 h. Radiolabeled products were immobilized on P-81 paper (Whatman, Maidstone, UK), which was then washed at 65°C for 1 h in 10 mm sodium phosphate, pH 7.0, containing 1.0% SDS. The P-81 paper was dried and autoradiographed for 24 h. The total protein concentration of plant extracts was determined by the method ofBradford (1976). Extraction and Analysis of Amino Acids Leaves (1 g) were freeze-dried and ground in 100% methanol (10:1 [v/w]). The homogenate was incubated at 60°C for 30 min and then centrifuged for 15 min at 12,000g. The supernatant was collected, and the pellet was extracted once more with 50% methanol. The combined extracts were reduced to dryness and redissolved in 75 μL of dilution buffer containing 100 mmNaHCO3 and 100 mmH3BO3, pH 8.5. Twenty microliters of the resuspended solution was mixed with 20 μL of 9-fluorenylmethyl chloroformate (20.7 mg mL−1) (Varian, Sugarland, TX) and incubated at room temperature for 10 min to generate fluorescent amino acid derivatives. After extraction of the free fluorescent dye in 70 μL of pentane ethyl acetate (80:20), 20 μL of the aqueous phase containing the amino acid derivatives was subjected to HPLC (Amino Tag column and Fluorichrome detector, Varian). Each amino acid was quantified as a percentage of total amino acids. Alkaline Hydrolysis and Analysis of Cell Walls Leaves (1 g) were ground in 100% methanol (1:1 [w/v]). The homogenate was incubated at 60°C for 30 min and then centrifuged for 15 min at 12,000g. The pellet was extracted two more times with 50% methanol, ensuring that no soluble aromatic compounds were present. The pellet was then hydrolyzed in 1.0 mNaOH for 4 h at 37°C. Insoluble debris were removed by centrifugation. The supernatant was acidified (pH 2.0) with 6.0m HCl and extracted three times with equal volumes of ethyl acetate. The pooled ethyl acetate fractions were reduced to dryness and the residue was recovered in methanol. Extracted samples were applied to a silica gel 60 F254 TLC plate, which was developed in chloroform:methanol (5:4). Authentic standards displayed the following RF values: tyramine, 0.23; and 4-coumaroyltyramine, 0.82. Tyramine in hydrolyzed cell walls extracts was quantified by HPLC on a liquid chromatography system (BioSys 500, Beckman) and a photodiode array detector (System Gold 168, Beckman) using a C18 reverse phase column (4.6 × 250 mm; Ultrasphere, Beckman) at 1200 psi with an isocratic gradient of methanol:water (8:2) containing 0.1% triethylamine and a flow rate of 0.5 mL min−1. The tyramine peak was identified from its UV spectrum and by comparison of its retention time with that of an authentic standard. Cell Wall Digestibility Measurement Cell wall digestibility was determined by measuring the number of protoplasts released after digestion of leaf tissue with hydrolytic enzymes (Brisson et al., 1994; Yao et al., 1995). Leaf sections (5 mm) were placed in a Petri dish containing 10 mL of plasmolysis solution (50 mm Hepes, pH 5.5, 50 mmCaCl2, and 500 mm mannitol). After 2 h of incubation, the plasmolysis solution was replaced with enzymatic solution (50 mm Hepes, pH 5.5, 50 mmCaCl2, 500 mm mannitol, 25 mg mL−1 cellulase, and 3 mg mL−1 pectinase). After 3 h of incubation, the enzyme solution was removed, and 10 mL of high-density solution (50 mm Hepes, pH 5.5, 50 mmCaCl2, and 500 mm Suc) was added to allow the protoplasts to float. The high-density solution was centrifuged at 3000g for 5 min, and the top 9 mL was removed. Protoplasts in the remaining 1 mL were counted in a hemocytometer. Histochemical Analysis of Cell Walls Cross-sections of leaves (approximately 500 μm thick) were prepared by hand-sectioning, and aromatic compounds were monitored by UV autofluorescence using a microscope (Aristoplan, Wild-Leitz, Wetzlar, Germany). RESULTS TYDC and THT Activities in Wild-Type Canola The basal levels of TYDC and THT activity in various organs of wild-type canola were determined. The highest levels of TYDC activity were detected in roots (Fig. 2A). TYDC activity in young leaves was approximately 17% of that in roots but was considerably lower in mature leaves, stems, and flower buds. THT activity was abundant in roots but was highest in mature leaves (Fig.2B). Substantial THT activity was detected in flower buds but was found at lower levels in stems and young leaves. Fig. 2. Open in new tabDownload slide TYDC and THT activities in wild-type canola. Bars represent the means ± se of three experiments. Fig. 2. Open in new tabDownload slide TYDC and THT activities in wild-type canola. Bars represent the means ± se of three experiments. Transformation of Canola with Chimeric TYDC Genes Approximately 2000 canola cotyledonary petioles were treated withA. tumefaciens harboring either pBI35S::TYDC1 or pBI35S::TYDC2. Two putative 35S::TYDC1(BN10 and BN11) and 13 putative 35S::TYDC2 (BN23 through BN219) transgenic plants were regenerated under kanamycin selection (Fig. 3). Regeneration efficiencies were 0.2% for 35S::TYDC1 and 1.3% for 35S::TYDC2. As a control, canola cotyledonary petioles were treated with A. tumefaciens harboring pBI122 and a regeneration efficiency of 18% was obtained. In comparison, the regeneration efficiency for canola cotyledonary petioles treated withA. tumefaciens harboring pCGN 783 was reported at 83%, and the transformation efficiency was estimated at 55% (Moloney et al., 1989). Furthermore, 85 putative transgenic canola plants were regenerated after treatment of stem segments with A. tumefaciens harboring a chimeric TDC gene in pBI 121, of which 11 were confirmed as transgenic (Chavadej et al., 1994). Fig. 3. Open in new tabDownload slide PCR assay demonstrating the integration of chimeric opium poppy TYDC1 and TYDC2transgenes into the canola genome. Genomic DNA from two canola lines transformed with 35S::TYDC1 (BN10 and BN11) and 13 canola lines transformed with35S::TYDC2 (BN23 through BN219) were used as templates for PCR with either TYDC1- orTYDC2-specific primers. Cloned opium poppy TYDC1 (pTYDC1) and TYDC2 (pTYDC2) cDNAs were used as positive control templates, whereas extracted wild-type canola genomic DNA (WT) was used as a negative control template. Fig. 3. Open in new tabDownload slide PCR assay demonstrating the integration of chimeric opium poppy TYDC1 and TYDC2transgenes into the canola genome. Genomic DNA from two canola lines transformed with 35S::TYDC1 (BN10 and BN11) and 13 canola lines transformed with35S::TYDC2 (BN23 through BN219) were used as templates for PCR with either TYDC1- orTYDC2-specific primers. Cloned opium poppy TYDC1 (pTYDC1) and TYDC2 (pTYDC2) cDNAs were used as positive control templates, whereas extracted wild-type canola genomic DNA (WT) was used as a negative control template. Regenerated kanamycin-resistant plants were allowed to flower and set seed. Each T0 plant was tested for the presence of the chimeric TYDC1 or TYDC2 transgenes. PCR results suggested that all regenerated plants were transgenic (Fig. 3). No PCR products were amplified with either TYDC1- orTYDC2-specific primers using wild-type genomic DNA as a template. Further evidence for the transformation of regenerated plants was obtained by direct assay for NPT II activity. Plants transformed with NOS::NPT II are resistant to kanamycin because NPT II phosphorylates and, consequently, detoxifies the antibiotic. The dot-blot assay shown in Figure4 illustrates the relative levels of NPT II activity in regenerated T0 plants. No radioactivity was immobilized on the P-81 paper in the absence of kanamycin. NPT II activity was not detected in wild-type plants but was detected in 14 putative transformants (Fig. 4). NPT II activity in T0 plants confirms that transgenes were inserted into transcriptionally active genomic regions. Fig. 4. Open in new tabDownload slide NPT II dot-blot assay of leaf extracts from putative T0 transgenic canola plants. Labeling reactions were performed with [γ-32P]ATP and plant extracts in the presence (+Kan) or absence (−Kan) of kanamycin. The negative control was wild-type (WT) canola. BN10 and BN11 were transformed with35S::TYDC1, whereas all other plants were transformed with 35S::TYDC2. Fig. 4. Open in new tabDownload slide NPT II dot-blot assay of leaf extracts from putative T0 transgenic canola plants. Labeling reactions were performed with [γ-32P]ATP and plant extracts in the presence (+Kan) or absence (−Kan) of kanamycin. The negative control was wild-type (WT) canola. BN10 and BN11 were transformed with35S::TYDC1, whereas all other plants were transformed with 35S::TYDC2. TYDC Activity in Consecutive Generations of Transgenic Canola TYDC activity levels in young leaves of T0plants are shown in Figure 5. Despite detectable NPT II activity in all but one primary transformant, most T0 plants showed suppressed levels of wild-type TYDC activity. In BN10 and BN213, TYDC activity was similar to that in wild-type plants, but in BN11, BN212, and BN214, it was less than 20% of the wild-type level. The mean TYDC activity of all primary transformants was approximately 50% of that in wild-type plants. TYDC1 and TYDC2 mRNAs were not detected in T0 plants (data not shown), suggesting that a trans-silencing mechanism might be responsible for the suppressed35S::TYDC expression and endogenous TYDC activity (Matzke and Matzke, 1995). Although a canola TYDC cDNA was not available to measure endogenous TYDC mRNA levels, the apparent homology among known TYDC genes across species is sufficiently low (Facchini and De Luca, 1994) to ensure that opium poppy TYDC probes did not hybridize with canola TYDC mRNAs under high-stringency conditions. Although no definite conclusions about the silencing of endogenous canola TYDC genes can be drawn, our data clearly show the specific silencing of35S::TYDC transgenes. Fig. 5. Open in new tabDownload slide Relative TYDC activity in young leaves of T0 canola transformed with35S::TYDC1 (BN10 and BN11) and35S::TYDC2 (BN23 through BN219). The relative TYDC activity in wild-type (WT) leaves is shown for comparison. Bars represent the means ± se of three experiments. TYDC specific activity in wild-type leaves was approximately 20 pkat mg−1 protein. Fig. 5. Open in new tabDownload slide Relative TYDC activity in young leaves of T0 canola transformed with35S::TYDC1 (BN10 and BN11) and35S::TYDC2 (BN23 through BN219). The relative TYDC activity in wild-type (WT) leaves is shown for comparison. Bars represent the means ± se of three experiments. TYDC specific activity in wild-type leaves was approximately 20 pkat mg−1 protein. Two transgenic canola lines were selected to analyze the inheritance of TYDC suppression. BN11 (35S::TYDC1) and BN214 (35S::TYDC2) showed the most severely suppressed TYDC activity among T0 plants (Fig. 5) but exhibited high levels of NPT II activity (Fig. 4). All BN11 and BN214 T1 plants tested positive for the presence ofTYDC transgenes by PCR analysis. This departure from the expected segregation ratio for a hemizygous single-copy gene suggests that TYDC transgenes were present in multiple copies in T0 plants. TYDC activity in the T1 progeny of BN214 was suppressed relative to wild-type plants (Fig. 6) and was within a range similar to that displayed by T0 plants transformed with 35S::TYDC2 (Fig. 5). Among 11 tested BN214 T1 plants, the mean TYDC activity was approximately 55% of the wild-type level. In contrast, TYDC activity increased in BN11 T1 progeny relative to the wild-type level (Fig. 6). The mean TYDC activity among BN11 T1 plants was 3-fold higher than the wild-type level and was 15-fold higher than the BN11 T0parent. Fig. 6. Open in new tabDownload slide Relative TYDC activity in young leaves of the T1 progeny of BN11 (35S::TYDC1) and BN214 (35S::TYDC1) canola lines. The relative TYDC activity in wild-type (WT) leaves is shown for comparison. Bars represent the means ± se of three experiments. TYDC specific activity in wild-type leaves was approximately 20 pkat mg−1 protein. Fig. 6. Open in new tabDownload slide Relative TYDC activity in young leaves of the T1 progeny of BN11 (35S::TYDC1) and BN214 (35S::TYDC1) canola lines. The relative TYDC activity in wild-type (WT) leaves is shown for comparison. Bars represent the means ± se of three experiments. TYDC specific activity in wild-type leaves was approximately 20 pkat mg−1 protein. TYDC1 mRNAs were detected in young leaves of all T1 progeny of BN11 (Fig.7). In contrast, TYDC2 mRNAs were not detected in any BN214 T1 progeny, and homologous TYDC mRNAs were not detected in wild-type leaves (Fig. 7). These data show that silencing of 35S::TYDC1 was overcome in BN11 T1 plants, but35S::TYDC2 silencing was maintained in BN214 T1 plants. The T1progeny of two other T0 plants, BN10 (35S::TYDC1) and BN212 (35S::TYDC2), were tested to verify the reproducibility of transgene inheritance and expression. TYDC1 mRNAs were detected in all BN10 T1 progeny, which also exhibited higher levels of TYDC activity compared with wild-type plants (data not shown). In contrast, all BN212 T1progeny showed suppressed levels of TYDC activity compared with wild-type plants, and TYDC1 mRNAs were not detected (data not shown). Thus, the reversion of transgene suppression from the T0 to T1 generations in plants transformed with 35S::TYDC1 (BN10 and BN11), and the continued transgene silencing in T1 plants transformed with35S::TYDC2 (BN212 and BN214), occurred in independent transgenic lines. Fig. 7. Open in new tabDownload slide Gel-blot analysis of RNA from young leaves of the T1 progeny of BN11 (35S::TYDC1) and BN214 (35S::TYDC1) canola lines. RNA extracted from young leaves of wild-type (WT) canola plants was used as a control. Total RNA was extracted and 15 μg was fractionated on 1.0% formaldehyde agarose gels, transferred to nylon membranes, and hybridized at high stringency with 32P-labeledTYDC1- or TYDC2-specific probes. Gels were stained with ethidium bromide before blotting to ensure equal loading. Fig. 7. Open in new tabDownload slide Gel-blot analysis of RNA from young leaves of the T1 progeny of BN11 (35S::TYDC1) and BN214 (35S::TYDC1) canola lines. RNA extracted from young leaves of wild-type (WT) canola plants was used as a control. Total RNA was extracted and 15 μg was fractionated on 1.0% formaldehyde agarose gels, transferred to nylon membranes, and hybridized at high stringency with 32P-labeledTYDC1- or TYDC2-specific probes. Gels were stained with ethidium bromide before blotting to ensure equal loading. TYDC activity did not increase above wild-type levels through four successive generations of the BN214 line (Fig.8). Mean TYDC activity among BN214 progeny increased from approximately 50% of wild-type levels in T1 plants to near-wild-type levels in T3 plants. TYDC mRNAs were not detected in the T2 or T3 progeny of BN214 (data not shown). Mean TYDC activity among BN11 progeny decreased from maximum levels found in T1 plants to nearly wild-type levels in T3 plants (Fig. 8). TYDC mRNA levels also decreased in T2 plants to less than 20% of those found in T1 plants and, ultimately, to undetectable levels in T3 plants (data not shown). All T2 and T3plants derived from BN11 and BN214 lines tested positive for the presence of TYDC transgenes by PCR analysis. The failure to recover the parental genotype through three generations is consistent with the suggested multiple insertion of TYDC transgenes in T0 plants. Fig. 8. Open in new tabDownload slide Relative mean TYDC activity in young leaves of successive self-pollinated generations (T1, T2, and T3 plants) derived from BN11 (35S::TYDC1) and BN214 (35S::TYDC1) canola lines. The relative TYDC activity in wild-type (WT) canola leaves is shown for comparison. Bars represent the means ± se. TYDC specific activity in wild-type leaves was equal to approximately 20 pkat mg−1protein. Fig. 8. Open in new tabDownload slide Relative mean TYDC activity in young leaves of successive self-pollinated generations (T1, T2, and T3 plants) derived from BN11 (35S::TYDC1) and BN214 (35S::TYDC1) canola lines. The relative TYDC activity in wild-type (WT) canola leaves is shown for comparison. Bars represent the means ± se. TYDC specific activity in wild-type leaves was equal to approximately 20 pkat mg−1protein. Aromatic Amino Acid and Cell Wall-Bound Amine Analysis In two T1 plants, BN11-6 and BN11-9 (35S::TYDC1), TYDC activity in young leaves was 3- to 4-fold higher than wild-type levels (Fig. 6). In two other T1 plants, BN214-2 and BN214-6 (35S::TYDC2), TYDC activity in young leaves was only 30% of wild-type levels (Fig. 6). Internal cellular pools of amino acids were measured in young leaves of these T1 plants and compared with wild-type plants. Although the concentration of most amino acids did not vary (data not shown), Tyr pools were somewhat lower and higher in the tested T1 progeny of BN11 and BN214, respectively, relative to wild-type controls (Table I). In contrast, Phe pools were not significantly different in the tested T1 progeny of BN11 and BN214 compared with wild-type plants (Table I). Differences in TYDC activity and amino acid levels between wild-type and transgenic plants were much less apparent in mature leaves. Table I. Analysis of aromatic amino acids in young leaves and cell wall-bound tyramine levels in mature leaves from wild-type and transgenic canola plants Plant . Amino Acid . Tyramine . Tyr . Phe . % total μg g−1 fresh wt Wild type 18.3  ± 5.6 2.0  ± 0.6 7.2  ± 4.5 BN11 (35S∷TYDC1)-a 12.0  ± 3.4 2.7  ± 0.8 14.7  ± 3.6 BN214 (35S∷TYDC2)-b 24.9  ± 5.8 1.8  ± 0.4 5.9  ± 4.4 Plant . Amino Acid . Tyramine . Tyr . Phe . % total μg g−1 fresh wt Wild type 18.3  ± 5.6 2.0  ± 0.6 7.2  ± 4.5 BN11 (35S∷TYDC1)-a 12.0  ± 3.4 2.7  ± 0.8 14.7  ± 3.6 BN214 (35S∷TYDC2)-b 24.9  ± 5.8 1.8  ± 0.4 5.9  ± 4.4 Values represent the means ± se of four independent measurements. F0-a Plants tested were T1 progeny that showed the highest TYDC activity (BN11-6 and BN11-9). F0-b Plants tested were T1 progeny that showed the lowest TYDC activity (BN214-2 and BN214-6). Open in new tab Table I. Analysis of aromatic amino acids in young leaves and cell wall-bound tyramine levels in mature leaves from wild-type and transgenic canola plants Plant . Amino Acid . Tyramine . Tyr . Phe . % total μg g−1 fresh wt Wild type 18.3  ± 5.6 2.0  ± 0.6 7.2  ± 4.5 BN11 (35S∷TYDC1)-a 12.0  ± 3.4 2.7  ± 0.8 14.7  ± 3.6 BN214 (35S∷TYDC2)-b 24.9  ± 5.8 1.8  ± 0.4 5.9  ± 4.4 Plant . Amino Acid . Tyramine . Tyr . Phe . % total μg g−1 fresh wt Wild type 18.3  ± 5.6 2.0  ± 0.6 7.2  ± 4.5 BN11 (35S∷TYDC1)-a 12.0  ± 3.4 2.7  ± 0.8 14.7  ± 3.6 BN214 (35S∷TYDC2)-b 24.9  ± 5.8 1.8  ± 0.4 5.9  ± 4.4 Values represent the means ± se of four independent measurements. F0-a Plants tested were T1 progeny that showed the highest TYDC activity (BN11-6 and BN11-9). F0-b Plants tested were T1 progeny that showed the lowest TYDC activity (BN214-2 and BN214-6). Open in new tab Neither tyramine nor hydroxycinnamic acid amides of tyramine were detected in methanol extracts from young or mature leaves, stems, or roots of wild-type or transgenic plants. However, cell wall-bound tyramine levels increased 2-fold in mature BN11-6 and BN11-9 leaves compared with wild-type plants (Table I). In contrast, cell wall-bound tyramine extracted from mature BN214-2 and BN214-6 leaves was similar to wild-type levels (Table I). No difference in cell wall-bound tyramine was detected in young leaves from wild-type and transgenic plants. The difference in cell wall-bound tyramine levels between some transgenic and wild-type plants prompted a microscopic examination of UV autofluorescence in corresponding leaf sections. This method has been used for the detection of cell wall-bound phenolic amides (Clarke, 1982), ferulic acid (Nicholson, 1992; Kato et al., 1994), and lignin and suberin (Monties, 1989; Schmutz et al., 1993). As shown in Figure9, the autofluorescence intensity of mesophyll cells from mature BN11-9 leaves (Fig. 9B) was stronger than that from mature, wild-type leaves (Fig. 9A). In contrast, the autofluorescence intensity of mesophyll cells from mature BN214-6 leaves (Fig. 9C) was weaker than that from mature wild-type leaves (Fig. 9A). Relative autofluorescence intensity was consistent in plants that showed similar levels of TYDC activity. No difference in autofluorescence intensity was detected in young leaves from wild-type or transgenic plants. Fig. 9. Open in new tabDownload slide UV autofluorescence microscopy of leaf mesophyll cross-sections from wild-type (A) and transgenic (B and C) canola plants. B, BN11-9 leaf; C, BN214-6 leaf. Tissues were photographed immediately after hand sectioning. Magnification, ×150. Fig. 9. Open in new tabDownload slide UV autofluorescence microscopy of leaf mesophyll cross-sections from wild-type (A) and transgenic (B and C) canola plants. B, BN11-9 leaf; C, BN214-6 leaf. Tissues were photographed immediately after hand sectioning. Magnification, ×150. Fig. 10. Open in new tabDownload slide Comparison of relative TYDC activity with protoplast release efficiency in mature leaves of individual T2 and T3 progeny of BN11 (35S::TYDC1) and BN214 (35S::TYDC2) transgenic canola lines. TYDC activity and protoplast release efficiency were measured in the same leaf. Wild-type TYDC activity (100%) was equal to approximately 10 pkat mg−1 protein. Fig. 10. Open in new tabDownload slide Comparison of relative TYDC activity with protoplast release efficiency in mature leaves of individual T2 and T3 progeny of BN11 (35S::TYDC1) and BN214 (35S::TYDC2) transgenic canola lines. TYDC activity and protoplast release efficiency were measured in the same leaf. Wild-type TYDC activity (100%) was equal to approximately 10 pkat mg−1 protein. Cell Wall Digestibility Oxidative crosslinking of hydroxycinnamic acid amides of tyramine in the cell wall has been suggested to increase cell wall strength and provide a barrier against microbial penetration (Clarke, 1982; Negrel et al., 1993a). The cell wall digestibility of leaves from wild-type and transgenic plants was tested by measuring the number of protoplasts released after incubation with cellulase and pectinase (Brisson et al., 1994; Yao et al., 1995). This approach assumes that the efficiency of protoplast release is directly proportional to cell wall digestibility. No significant difference in protoplast release efficiency was found in young leaves from wild-type or transgenic plants (data not shown). However, BN11-6 and BN11-9 leaf sections released an average of only 42% of the protoplasts released by wild-type leaf sections (TableII). In contrast, BN214-2 and BN214-6 leaf sections released an average of 87% more protoplasts than wild-type canola leaf sections (Table II). Table II. Release of protoplasts from wild-type and transgenic mature canola leaves Plant . Protoplasts Released . % Wild Type . ×105 g−1 fresh wt Wild type 10.3  ± 1.4 — BN11 (35S∷TYDC1)1-a 4.3  ± 0.5  42 BN214 (35S∷TYDC2)1-b 19.3  ± 2.3 187 Plant . Protoplasts Released . % Wild Type . ×105 g−1 fresh wt Wild type 10.3  ± 1.4 — BN11 (35S∷TYDC1)1-a 4.3  ± 0.5  42 BN214 (35S∷TYDC2)1-b 19.3  ± 2.3 187 Values represent the means ± se from three independent experiments. F1-a Plants tested were T1 progeny that showed the highest TYDC activity (BN11-6 and BN11-9). F1-b Plants tested were T1 progeny that showed the lowest TYDC activity (BN214-2 and BN214-6). Open in new tab Table II. Release of protoplasts from wild-type and transgenic mature canola leaves Plant . Protoplasts Released . % Wild Type . ×105 g−1 fresh wt Wild type 10.3  ± 1.4 — BN11 (35S∷TYDC1)1-a 4.3  ± 0.5  42 BN214 (35S∷TYDC2)1-b 19.3  ± 2.3 187 Plant . Protoplasts Released . % Wild Type . ×105 g−1 fresh wt Wild type 10.3  ± 1.4 — BN11 (35S∷TYDC1)1-a 4.3  ± 0.5  42 BN214 (35S∷TYDC2)1-b 19.3  ± 2.3 187 Values represent the means ± se from three independent experiments. F1-a Plants tested were T1 progeny that showed the highest TYDC activity (BN11-6 and BN11-9). F1-b Plants tested were T1 progeny that showed the lowest TYDC activity (BN214-2 and BN214-6). Open in new tab Data presented in Table II suggest a correlation between TYDC activity and cell wall digestibility in select T1 (i.e. those with the highest and lowest levels of TYDC activity) and wild-type plants. To further test this relationship, protoplast release efficiency was directly compared with relative TYDC activity in mature leaves from randomly selected T2 and T3 progeny of the BN11 and BN214 transgenic lines (Fig. 10). The mean TYDC activity approached wild-type levels in successive progeny of both lines (Fig. 8), but individual T2 and T3 plants displayed a wide range of relative TYDC activity. As shown in Figure 10, high levels of TYDC activity correlated with reduced protoplast release efficiency, whereas low levels of TYDC activity relative to wild-type plants resulted in an increased recovery of protoplasts. DISCUSSION Canola was transformed with chimeric transgenes under the transcriptional control of the CaMV 35S promoter and encoding two TYDC isoforms from opium poppy. The transgenic plants were used to test the hypothesis that higher levels of TYDC activity will promote the conversion of the cellular Tyr pool to tyramine, which in turn will increase cell wall-bound hydroxycinnamic acid amide formation. Elevated amide levels would be expected to decrease cell wall susceptibility to digestion by hydrolytic enzymes. Increased TYDC activity in transgenic plants is potentially harmful because of the phytotoxicity of hydroxyphenylethylamines such as tyramine (Negrel et al., 1993b). Detoxification of tyramine was reported in cultured tobacco cells only under conditions that favored THT induction. The exclusive recovery of T0plants with suppressed TYDC activity relative to wild-type plants (Fig.5) suggests that expression of 35S::TYDCtransgenes inhibited the plant-regeneration process. Moreover, the low transformation efficiency is consistent with the suggestion that only T0 plants that exhibited specifictrans-silencing of 35S::TYDC1 and35S::TYDC2, but not NOS::NPT II, were recovered. In contrast, tobacco transformed with35S::TDC (Songstad et al., 1990) was reported to accumulate both soluble tryptamine and tyramine because of a proposed modification in the substrate specificity of Trp decarboxylase in tobacco (Songstad et al., 1991). Similarly, canola (Chavadej et al., 1994) and potato (Yao et al., 1995) transformed with35S::TDC accumulated high levels of tryptamine without any apparent adverse effects; therefore, we cannot rule out the possibility that the transformation efficiency was low for unknown reasons. In the BN10 and BN11 lines, 35S::TYDC1 was active in T1 plants, whereas35S::TYDC2 remained silent in all transgenic progeny of the BN212 and BN214 lines. The consistent results obtained for independent lines transformed with each construct suggests that transgene expression was related to the specific nucleotide sequences of TYDC1 and TYDC2. Expression of35S::TYDC transgenes might have been affected by homologous TYDC genes in canola (Matzke and Matzke, 1995).TYDC genes from opium poppy (Facchini and De Luca, 1994) and parsley (Kawalleck et al., 1993) share >60% nucleotide identity. Similar homology between TYDC genes from opium poppy and canola is probable. Reactivation of a silenced transgene after passage through successive generations has been reported previously. For example, transformation of tobacco with the bean PAL2 gene resulted in transgenic plants with severely reduced PAL activity (Elkind et al., 1990). The sense suppression of PAL in T0 plants was progressively overcome in pedigrees of homozygous progeny (Bate et al., 1994). A β-1,3-glucanase transgene (gn1) in tobacco driven by the CaMV 35Spromoter was silent during seed germination and vegetative growth but was activated in meiotically derived seed tissues (Castresana and Balandin, 1997). Expression of gn1 was not reactivated in plantlets regenerated mitotically from leaf explants ofgn1-suppressed plants. Reactivation of 35S::TYDC1 in canola might also require passage through meiosis. The expression ofNOS::NPT II in T0 plants shows that T-DNA insertions occurred in transcriptionally active genomic regions (Fig. 4). NPT II activity was detected in consecutive generations of all transgenic lines except BN219 (data not shown); thus, the silencing of 35S::TYDC1 in T0 plants, its reactivation in T1 plants, and its subsequent and gradual re-suppression in T2 and T3plants are related to the TYDC transgene. Canola might be prone to silencing aromatic amino acid decarboxylase transgenes. Expression of a 35S::TDC transgene in canola, which was not initially silenced in T0 plants (Chavadej et al., 1994), also became completely suppressed over four successive generations (V. De Luca, personal communication). Such meiotically heritable alterations in transgene activity have been demonstrated for trans-silencing mechanisms mediated by reading frame (Matzke and Matzke, 1995) and promoter homology (Park et al., 1996). TYDC1 mRNA levels in BN11 T1 progeny did not always translate into a proportional increase in TYDC activity (Figs. 6 and 7), suggesting that posttranslational regulation of heterologous TYDC activity might have also occurred in plants that expressed 35S::TYDC1. The increase in TYDC activity in young leaves of BN11-6 and BN11-9 T1 plants resulted in decreased cellular Tyr pools and increased cell wall-bound tyramine in corresponding mature leaves compared with wild-type plants (Table I). In contrast, the increased Tyr pools in young leaves of BN214-2 and BN214-6 plants are consistent with the reduced TYDC activity relative to wild-type levels. TYDC activity levels were not much different in mature leaves of transgenic plants, so little difference was found between the cellular Tyr pools of mature leaves. In contrast, increased cell wall-bound tyramine levels were only detected in mature leaves that also showed high THT activity. A temporal discrepancy between the optimum expression of TYDC transgenes and the maximum insolubilization of tyramine was apparent; therefore, TYDC activity was altered most abundantly in young leaves, but cell wall modifications (i.e. tyramine levels and digestibility) were not detectable until leaves matured. This discrepancy might be related to the transient nature of heterologous TYDC activity in canola and the low endogenous THT activity in young leaves. Our data provide in vivo evidence that the formation of tyramine represents a sink for Tyr in canola and that TYDC activity appears to determine, at least in part, the level of cell wall-bound tyramine in mature leaves. This conclusion is supported by the kinetic properties of THT from various species. THT follows Michalis-Menton kinetics in the presence of low concentrations of hydroxycinnamoyl-CoA derivatives but exhibits negative cooperativity at concentrations above 2 to 3 μm, resulting in a decrease in the affinity for tyramine (Hohlfeld et al., 1995; Negrel and Javelle, 1997; Yu and Facchini, 1999). This negative cooperativity has been proposed as a physiological mechanism involved in the regulation of amide biosynthesis (Negrel and Javelle, 1997). The cellular concentrations of hydroxycinnamoyl-CoA derivatives are probably much lower than the level of tyramine (Hahlbrock and Scheel, 1989). Negative cooperativity implies that an increase in the cellular tyramine pool will lead to an increase in amide formation, even at constant levels of hydroxycinnamoyl-CoA derivatives (Negrel and Javelle, 1998). An increase in the cellular pool of hydroxycinnamoyl-CoA esters would result in increased amide formation only if tyramine levels also increase; thus, the level of TYDC activity should play a role in the regulation of cell wall-bound amide biosynthesis. Tyramine released by alkaline hydrolysis was probably insolubilized in cell walls as hydroxycinnamic acid amides. The harsh treatment required to extract tyramine would also be expected to hydrolyze amide bonds. In addition, most amides are highly cross-linked in the cell wall, preventing their extraction by alkaline hydrolysis. Numerous bond types between cell wall components and tyramine derivatives have been demonstrated (Borg-Olivier and Monties, 1993). UV autofluorescence examination of leaf sections from transgenic canola confirmed that cell wall modification had occurred (Fig. 9). The stronger autofluorescence of mature mesophyll cell walls from BN11-6 and BN11-9 T1 plants relative to wild-type leaves suggests an increased content of aromatic residues. In contrast, the weaker autofluorescence of mature mesophyll cell walls from BN214-2 and BN214-6 T1 plants suggests a reduced incorporation of aromatic compounds. These data are consistent with the relative levels of tyramine solubilized from cell walls of wild-type and transgenic plants (Table I). However, the alkaline-hydrolyzed tyramine might not reflect the entire hydroxycinnamic acid amide content of the cell walls; thus, UV autofluorescence intensity might not always have been proportional to extracted tyramine levels. A change in the availability of tyramine could be expected to result in an alteration of cell wall strength, because tyramine-derived amides are purported to become oxidatively cross-linked, together with other phenolics, in the cell wall (Clarke, 1982; Negrel and Lherminier, 1987; Matern and Kneusel, 1988; Iiyama et al., 1994). An inverse relationship between the level of TYDC activity and the susceptibility of cell walls to enzymatic hydrolysis was revealed by measuring protoplast release efficiency (Fig. 10). Protoplast recovery from mature leaf sections was reduced by 60% in BN11-6 and BN11-9 T1 plants, with a mean TYDC activity that was 3- to 4-fold higher than that in wild-type plants (Table II). In contrast, the 50% reduction in mean TYDC activity in BN214-2 and BN214-6 T1 plants corresponded to an increase in the cell wall digestibility of mature leaves. The similar protoplast release efficiency of young leaf tissue from wild-type and transgenic plants was consistent with our inability to detect differences in cell wall-bound tyramine levels in young leaves with altered TYDC activity. We have shown that 35S::TYDC transgenes in canola are subject to transcriptional silencing mechanisms. In transgenic plants with elevated TYDC activity, internal Tyr pools decreased and cell wall-bound tyramine levels increased compared with wild-type plants. Increased TYDC activity also correlated with decreased enzymatic digestibility of cell walls. Overall, our data suggest thatTYDC and tyramine affect cell wall properties that might have implications in plant-pathogen interactions. 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Plant Cell 7 1995 1787 1799 Google Scholar Crossref Search ADS PubMed WorldCat 52 Yu M, Facchini PJ (1999) Purification, characterization, and immunolocalization of hydroxycinnamoyl-CoA: tyramineN-(hydroxycinnamoyl)transferase from opium poppy. Planta (in press) Author notes 1 This work was supported by grants from the Natural Sciences and Engineering Research Council of Canada and the Alberta Agricultural Research Institute to P.J.F. * Corresponding author; e-mail [email protected]; fax 1–403–289–9311. 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)

Matsuyama, Takashi; Satoh, Hidetaka; Yamada, Yasuyuki; Hashimoto, Takashi

doi: 10.1104/pp.120.3.665pmid: 10398701

Abstract The root cap functions in the perception of gravity, the protection of the root apical meristem, and facilitation of the passage of roots through the soil, but the genes involved in these functions are poorly understood. Here we report the isolation of a root-specific gene from the cap of maize (Zea mays L.) primary root by cDNA subtraction and differential screening. The gene zmGRP4(Z.maysglycinerich protein 4) encodes a member of the glycine-rich proteins with a putative signal peptide at the amino terminus. The deduced molecular mass of mature zmGRP4 is 14.4 kD. In situ-hybridization analysis has shown zmGRP4 to be strongly expressed in the lateral root cap and weakly expressed in the root epidermis. A polyclonal antibody raised against recombinant zmGRP4 detected a protein of 36 kD in the insoluble protein fraction extracted from the root tip and the root proper, indicating posttranslational modification(s) of zmGRP4. Immunohistochemical analysis showed the accumulation of zmGRP4 in the mucilage that covers the root tip. These results indicate that lateral root-cap cells secrete modified zmGRP4 into the mucilage to which the protein may contribute to its characteristic physical properties. The root cap covers the root meristem at the root apices of vascular plants but is absent in nonvascular plants such as liverworts and mosses. It is proposed that the root cap perceives gravity and protects the root apical meristem (Sievers and Braun, 1996). The root cap has a high regenerative capacity. When the root cap, which is sharply delineated from the root proper, is surgically removed from a maize (Zea mays L.) root, it regenerates completely within a few days (Barlow, 1975). Anatomical studies suggest that the root cap consists of several distinct regions (Moore and McClelen, 1983; Dolan et al., 1993). The maize root cap, for example, can be divided into three regions: the calyptrogen, the columella root cap, and the lateral root cap. The calyptrogen faces the distal end of the quiescent center of the root apical meristem, is composed of approximately four cell layers, and serves as a root-cap meristem. The columella cells are generated by periclinal cell division from the central region of the calyptrogen. Sedimented large amyloplasts containing well-developed starch granules are characteristic of the columella cells. These amyloplasts function as statoliths in root gravitropism (Sievers and Braun, 1996). The lateral root cap surrounds the columella root cap. In maize roots with a closed-type construction, the lateral cap cells originate from the calyptrogen (Barlow, 1996). However, in Arabidopsis roots, which have an open-type construction, there is no discrete boundary between the root proper and the cap, and the lateral cap cells are derived from the same initials as the root epidermal cells (Dolan et al., 1993). The lateral root-cap cells are rich in the hypertrophied dictyosome cisternae that form large secretory vesicles (Mollenhauer et al., 1961). These cisternae may reflect the massive secretion of mucilage from the lateral cap cells, because the vesicle content was observed to be deposited between the plasma membrane and the outer tangential walls of the lateral cap cells (Morré et al., 1967). In addition to these three tissues, sloughed-off cap cells and root mucilage may also be included as components, which together make up the cap region. Root-cap cells are continuously pushed toward the root-cap periphery and finally slough off into the external root environment. These detached cells are found at the root periphery, even at some distance from the root cap (Vermeer and McCully, 1982); they are metabolically active and have unique patterns of gene expression (Brigham et al., 1995). Several functions of the sloughed-off cap cells as a root-soil interface have been proposed (Hawes et al., 1998). The root mucilage typically covers the root apex, is an amorphous and uneven gel, and ranges in thickness from 50 μm to 1 mm. The mucilage is secreted largely from the root cap, but the root epidermis is also covered by a thin film of mucilage, which is histochemically distinct from the cap-derived mucilage (Greaves and Darbyshire, 1972; Clarke et al., 1979; Foster, 1982; Vermeer and McCully, 1982). The matrix of maize mucilage consists of 95% polysaccharides and 5% protein (Harris and Northcote, 1970; Bacic et al., 1986). To understand the functions of the root cap at the molecular level, we have identified genes that are specifically or predominantly expressed in maize root cap. One such gene is expressed strongly in the lateral root cap, and its gene product is secreted into and accumulates in the mucilage. MATERIALS AND METHODS Plant Material Maize (Zea mays L. cv Merit) was supplied by the Asgrow Seed Company (Kalamazoo, MI). Seeds were soaked in tap water for 72 h in the dark at 30°C. After imbibition seeds were germinated on paper towels saturated with tap water for 1 to 2 d in the dark at 30°C. When the primary roots were 2 to 3 cm long, the cap and selected portions of the root were removed by a scalpel under a magnifying glass and immediately frozen in liquid N2. The tip region used in this study was the apical 5 mm of the root and included the root apical meristem and the whole root cap. The region of the root proper was between 1 and 3 cm from the distal end of the root. Maize plantlets were grown under 18-h light/6-h dark or 24-h dark conditions at 30°C on layered wet paper towels in plastic pots. RNA Isolation, cDNA Synthesis, and Subtractive Hybridization Poly(A+) RNA was extracted directly from the root cap and the root proper of maize primary roots using Dynabeads oligo(dT25) (Dynal, Oslo, Norway). Several hundred nanograms of poly(A+) RNA were used to construct double-stranded cDNA using a cDNA synthesis kit (Pharmacia). Subtractive hybridization was done essentially as described by Wang and Brown (1991) and Hashimoto et al. (1993). The double-stranded cDNAs were fragmented by AluI and RsaI and ligated to a PCR linker. cDNA fragments of 0.2 to 2.0 kb were amplified by PCR. The cDNA fragments from the root proper were then biotinylated with Photoprobe biotin (Vector Laboratories, Burlingame, CA) and used as the driver DNA. One subtraction cycle consisted of five steps: hybridization of the excess driver DNA to the tracer DNA from the root cap for 20 h at 68°C; removal of nonhybridizing driver DNA by binding to streptavidin and extraction with organic solvent; another hybridization of the excess driver DNA to the remaining tracer DNA once again for 2 h at 68°C; removal of driver DNA as above; and PCR amplification of the tracer DNA. This subtraction cycle was repeated twice to produce subtracted root-cap cDNA fragments. Subtracted root-proper cDNA fragments were also generated in the same way, except that cDNA fragments from the root cap and the root proper were used, respectively, as the driver and the tracer DNAs. Screening of Differentially Expressed cDNAs The subtracted root-cap cDNA fragments were digested withEcoRI, which cleaved the PCR linker, and inserted into theEcoRI site of pBluescript II SK(−) (Stratagene). These plasmids were introduced into the bacterial strain DH5α to construct a root-cap cDNA library, and 386 independent colonies were grown overnight in Luria-Bertani medium containing 50 μg mL−1 ampicillin at 37°C. From each culture a 50-μL aliquot was blotted in duplicate onto a membrane (Hybond N+, Amersham) using a filtration manifold system (GIBCO-BRL). After denaturation and neutralization, the duplicate filters were hybridized at 42°C for 16 h with either a32P-labeled, subtracted root-cap cDNA probe or a32P-labeled, nonsubtracted root-proper cDNA probe, in a hybridization buffer containing 50% formamide, 10% dextran sulfate, 1% SDS, 5× SSPE (1× SSPE: 180 mm NaCl, 1 mm EDTA, and 10 mmNa2HPO4, pH 7.5), 5× Denhardt's solution (1× Denhardt's solution: 0.02% [w/v] BSA, 0.02% [w/v] Ficoll, and 0.02% [w/v] PVP), and 100 μg mL−1 salmon testis DNA, and washed at 65°C in 0.1× SSPE and 0.1% SDS. Seventy-two positive cDNA clones, which hybridized only to the root-cap cDNA probe, were obtained. To group the positive clones, the 72 recombinant bacterial cultures containing positive cDNA clones were blotted onto a Hybond N+ membrane and processed as described above. One cDNA clone, which had hybridized specifically and strongly to the root-cap cDNA probe, was chosen, labeled with32P, and hybridized to the membrane. Positive clones were regarded as members of the same group. Next, another strongly and specifically hybridizing cDNA clone other than the members of this group was chosen and processed as above. Four hybridizations were done, resulting in four independent groups and 29 remaining cDNA clones. Representative clones from the four groups and the extra 29 cDNA clones were partially sequenced by a DNA sequencer (model 373A, Perkin-Elmer), using M13 reverse and universal primers. The sequence analysis classified the root-cap-positive cDNA clones into 23 groups. Subtracted cDNA fragments from the root cap, nonsubtracted cDNA fragments from the root cap, and nonsubtracted cDNA fragments from the root proper were blotted in amounts of 0.05, 0.5, and 5 μg per slot onto a Hybond N+ membrane, as described above. Representative cDNA fragments from the 23 groups were used as the probes for hybridization. Ten cDNA fragments hybridized to the subtracted and nonsubtracted root-cap cDNA pools or to the subtracted root-cap cDNA pool, but not to the root-proper cDNA pool, and will be referred to as “root-cap abundant.” The other 13 clones either hybridized to the root-proper cDNA pool or did not hybridize to any cDNA pools. A cDNA library of the maize primary root-tip region from within 1 mm of the distal-tip end was made in λZAPII (Stratagene) (Matsuyama et al., 1999). A total of 4 × 106 recombinants were independently screened with the 10 root-cap-abundant cDNA fragments as probes. Hybridization and other procedures were done as described above. Positive plaques were identified with 3 of the 10 root-cap-abundant probes. In this report, one cDNA representing six phage clones was analyzed. These positive recombinant phages were converted to pBluescript SK(−) plasmids by in vivo excision using the manufacturer's protocol (Stratagene). Both DNA strands of the longest insert of the six clones were sequenced. DNA and predicted amino acid sequences were analyzed with GeneWorks software (IntelliGenetics, Campbell, CA). Genomic-DNA-Hybridization Analysis Total genomic DNA was isolated from 3-d-old etiolated maize seedlings by cetyl-trimethyl-ammonium bromide extraction (Murray and Thompson, 1980). Genomic DNA (30 μg) was digested with restriction enzymes, electrophoresed on a 1% agarose gel, and blotted onto a Hybond N+ membrane. The membrane was hybridized to the full-length zmGRP4(Z.maysGRP4) cDNA probe and washed under the conditions described above. Northern-Hybridization and RT-PCR Analysis Total RNA was isolated from several tissues, including the root tip, root proper, young leaves from 2-week-old plants, and shoots from 3-d-old light-grown and etiolated plants using phenol:chloroform extraction and LiCl precipitation (Mohnen et al., 1985). Poly(A+) RNA was purified from total RNA using Oligotex-dT30 Super (Takara Shuzo, Tokyo, Japan). Poly(A+) RNA (1.5 μg per lane) was electrophoresed on a 1.2% formaldehyde agarose gel, blotted onto a Hybond N+ membrane, and hybridized to the full-length zmGRP4 cDNA probe under the conditions described above. After stripping the probe from the membrane by incubating at 67°C in a buffer containing 50% formamide, 10 mm Tris-HCl, and 10 mmEDTA, pH 8.0, the 32P-labeledPstI-SacI fragment of a ubiquitin cDNA (Christensen and Quail, 1989) was hybridized to the same membrane. For RT-PCR analysis, total RNA was isolated from approximately 100 mg of root tip, root proper, shoot, and etiolated shoot using the RNeasy Plant Mini kit (Qiagen, Chatsworth, CA) and used to construct first-strand cDNA using the SUPERSCRIPT preamplification system (GIBCO-BRL). PCR primers for zmGRP4 were 5′-TTGTATCTCACAATGGCAGGC and 5′-GCGTTGGAATTCCAAGAACC (Fig. 1) and PCR primers for maize α-tubulin1 (Montoliu et al., 1989) were 5′-CTTGATCGCATCAGGAAGC and 5′-TCAGCAGAGATGACTGGAGC. PCR amplification was carried out for 18 cycles of denaturation at 94°C for 1 min, annealing at 60°C for 30 s, and elongation at 72°C for 2 min. Amplified zmGRP4 fragments were electrophoresed on a 1.2% agarose gel, blotted onto a Hybond N+ membrane, and hybridized to the full-length zmGRP4 cDNA probe as described above. Representative amplified DNA fragments were partially sequenced to confirm their identity. Fig. 1. Open in new tabDownload slide Nucleotide and deduced amino acid sequences ofzmGRP4. A putative signal peptide is double underlined. The sequence between the vertical arrowheads was used as both antisense and sense probes for in situ-hybridization analysis. The two arrows indicate the positions of PCR primers used for RT-PCR analysis. The stop codon is shown by an asterisk. Fig. 1. Open in new tabDownload slide Nucleotide and deduced amino acid sequences ofzmGRP4. A putative signal peptide is double underlined. The sequence between the vertical arrowheads was used as both antisense and sense probes for in situ-hybridization analysis. The two arrows indicate the positions of PCR primers used for RT-PCR analysis. The stop codon is shown by an asterisk. In Situ-Hybridization Analysis Maize primary root tips were fixed in 3% paraformaldehyde and 2% glutaraldehyde for 12 h at 4°C. After samples were dehydrated in a graded ethanol series and cleared in a graded xylene series, they were embedded in wax (Histoprep 580, Wako, Osaka, Japan) and sectioned at 10 μm by using a rotary microtome. Digoxigenin-labeled antisense and sense RNA probes were prepared from a 3′-untranslated region ofzmGRP4 (see Fig. 1) using an RNA labeling kit (DIG, Boehringer Mannheim). Samples were incubated with the RNA probes at 50°C for 16 h, treated with RNase A (2.5 μg mL−1 in 0.5 m NaCl, 10 mm Tris-HCl, and 1 mm EDTA, pH 7.5) at 37°C for 30 min, and washed with several changes of 2× SSC (1× SSC: 150 mm NaCl and 15 mmNa3C6H5O7) and once with 0.1× SSC at 50°C. Signals were detected by a nucleic acid detection kit (DIG, Boehringer Mannheim). The color reaction was stopped with 10 mm Tris-HCl and 1 mm EDTA, pH 8.0. Sections were passed through an ethanol series and mounted for microscopic observation. zmGRP4 Antiserum A portion of the zmGRP4 cDNA encoding the carboxyl-terminal amino acid residues from 132 to 192 was subcloned into the EcoRI site of pET-32b(+) (Novagen), which would then express a fusion protein consisting of the N-terminal region of thioredoxin and the C-terminal region of zmGRP4. This plasmid was introduced into the BL21 (DE3) bacterial strain (Novagen), and the expression of the fusion protein was induced by 1 mm isopropyl β-d-thiogalactopyranoside at 37°C for 3 h in Luria-Bertani medium. Bacterial cells were harvested by centrifugation, suspended in 50 mm potassium phosphate buffer, pH 8.0, containing 1% Triton X-100 and 1 μg mL−1 lysozyme, incubated at 30°C for 15 min, and then ruptured by three cycles of freeze/thaw treatment and sonication for 1 min. After centrifugation of the homogenate, the fusion protein in the supernatant was separated by preparative SDS-PAGE using a PrepCell (model 491, Bio-Rad). The eluate fractions containing the fusion protein were concentrated with a YM-10 membrane filter (Amicon). The buffer of the concentrated protein solution was exchanged by using a PD-10 column (Pharmacia) equilibrated with buffer A (20 mm Tris-HCl and 10% glycerol, pH 7.0). The above solution was loaded onto a Mono-Q fast-protein liquid-chromatography column (Pharmacia) previously equilibrated with buffer A and eluted with a linear gradient of KCl from 0 to 0.5m in buffer A. The fractions containing the fusion protein were concentrated with a YM-10 filter and desalted using a PD-10 column. The fusion protein had an approximate purity of 99.9%, as determined by staining with Coomassie Brilliant Blue R-250 after SDS-PAGE, and was used to raise antiserum in mice. The reactivity of the antiserum against zmGRP4 was confirmed as follows. A BamHI-SmaI fragment of thezmGRP4 cDNA encoding the carboxyl-terminal amino acid residues from 137 to 192 was subcloned into pGEX-2T (Pharmacia). The resultant pGEX-zmGRP4 plasmid or pGEX-2T was introduced into the BL21 (DE3) bacterial strain, and the expression of either a chimeric protein consisting of zmGRP4 and GST, or GST alone, respectively, was induced at 37°C for 3 h with 1 mm isopropyl β-d-thiogalactopyranoside. Approximately 100 ng of total protein extracts from these bacterial cells was separated on a 15% SDS-polyacrylamide gel and transferred to an Immobilon PVDF membrane (Millipore). The membrane was blocked at room temperature in buffer B (100 mm Tris-HCl and 150 mm NaCl, pH 7.5) containing 5% skim milk powder for 1 h. The antiserum was diluted 1:5000 in buffer B and incubated with the membranes at room temperature for 1 h. After washing the antiserum several times with buffer B containing 0.2% Tween 20, the following procedures, including secondary antibody treatment and immunodetection, were performed by using an ECL Plus kit (Amersham) according to the manufacturer's instructions. The antiserum reacted strongly with the fusion protein consisting of zmGRP4 and GST, but not with GST alone. Immunohistochemical Analysis Fixed sections were prepared as described for in situ hybridization and blocked at room temperature in buffer B containing 0.2% Tween 20 and 3% BSA for 1 h. Anti-zmGRP4 serum or preimmune mouse serum was diluted 1:300 in blocking buffer and incubated with the sections at room temperature for 1 h. After washing the primary antibody several times in buffer B containing 0.2% Tween 20, anti-mouse IgG conjugated with alkaline phosphatase (Kirkegaard and Perry Laboratories, Gaithersburg, MD) was diluted 1:1000 in buffer B and incubated with the sections at room temperature for 1 h. After briefly washing the slides with buffer B, immunodetection was performed as described for in situ-hybridization analysis. Immunoblot Analysis Approximately 100 maize root tips were frozen with liquid N2 and ground to a fine powder with a pestle. For some sample preparations used in Figure 6B, after root mucilage including sloughed-off cap cells was gently wiped from the root tip with a paper towel, the root tips were immediately collected in liquid nitrogen. The pulverized root cells were extracted with buffer (50 mm Tris-HCl, 3 mm EDTA, 3 mm DTT, and 3 mm PMSF), and the suspension was centrifuged at 15,000g for 5 min. The supernatant was referred to as the soluble fraction. The pellet was resuspended in sample buffer (125 mm Tris-HCl, pH 6.8, containing 1% SDS) and used as the insoluble fraction. A total protein fraction was prepared by directly extracting the pulverized cells with sample buffer. Protein concentration was determined using the BCA protein assay reagent (Pierce). Fig. 6. Open in new tabDownload slide Immunoblot analysis of zmGRP4. Each lane was loaded with 10 μg of protein extracted from the root tip or root proper. A, Total protein fraction (lanes T), Tris-buffer-soluble fraction (lanes S), and Tris-buffer-insoluble but SDS-buffer-soluble fraction (lanes I) were separated on a 15% SDS-polyacrylamide gel. B, The Tris-buffer-soluble fraction (S) and Tris-buffer-insoluble but SDS-buffer-soluble fraction (I) were extracted from intact root tips (+) or root tips from which mucilage had been removed (−). Fig. 6. Open in new tabDownload slide Immunoblot analysis of zmGRP4. Each lane was loaded with 10 μg of protein extracted from the root tip or root proper. A, Total protein fraction (lanes T), Tris-buffer-soluble fraction (lanes S), and Tris-buffer-insoluble but SDS-buffer-soluble fraction (lanes I) were separated on a 15% SDS-polyacrylamide gel. B, The Tris-buffer-soluble fraction (S) and Tris-buffer-insoluble but SDS-buffer-soluble fraction (I) were extracted from intact root tips (+) or root tips from which mucilage had been removed (−). Protein preparations (10 μg per lane) were separated on a 15% SDS-polyacrylamide gel. The remaining steps were performed as described above, except that 3% BSA was substituted for 5% skim milk powder in the blocking buffer. RESULTS Isolation of a Maize GRP cDNA That Is Highly Expressed in Root Cap The root cap and the root proper are sharply delineated in the maize primary root, which has a closed-type construction. This anatomical feature is used to facilitate excision of maize root-cap tissues from the root proper using a scalpel (Barlow, 1975). We collected approximately 500 root caps and extracted poly(A+) RNA directly from the root cap and also from the root proper. The cDNAs specifically present in the root cap were enriched by subtracting the root-proper cDNA fragment pool from the root-cap cDNA fragment pool. Subsequently, the subtracted root-cap cDNA fragment library was duplicated and hybridized independently with the above root-proper cDNA fragment pool or the root-cap cDNA fragment pool as the probes. This differential screening recovered 72 cDNA fragments that hybridized specifically to the root-cap cDNA fragment pool, and these clones were classified into 23 groups by cross-hybridization and partial DNA sequencing. Further slot-blot hybridization with the above two probes confirmed that 10 cDNA groups were much more abundant in the root cap than in the root proper. Representative cDNA fragments from these 10 clones were used as the probes to screen a maize root-tip cDNA library, and three distinct cDNA clones were obtained. One of the three cDNA clones is reported here. The other two clones encoded a novel protein (Matsuyama et al., 1999) and a maize expansin. Figure 1 shows the nucleotide and deduced amino acid sequences of a cDNA clone. The cDNA was 821 bp long and contained an open reading frame encoding a 16.9-kD polypeptide of 192 amino acids. The predicted protein had a hydrophobic putative signal peptide with a potential cleavage site between 22 and 23 amino acid residues (von Heijne, 1985) and was a member of the cell wall GRPs (Showalter, 1993). We will refer to this protein as zmGRP4. zmGRP4, excluding the putative signal peptide, was rich in Gly (40%), Ser (19%), Asn (7%), Ala (7%), and Tyr (6%). The high content of Gly, Ser, and Ala of zmGRP4 is consistent with the general characteristics of Gly-rich cell wall proteins. Genomic DNA blot-hybridization analysis was done with a full-lengthzmGRP4 cDNA probe at high-stringency conditions (Fig.2). BamHI and EcoRI digested the zmGRP4 cDNA once, whereas BglII andXhoI did not digest the cDNA. Two to three strong bands and two to four weak bands were detected when the maize genome was digested with BamHI, BglII, EcoRI, orXhoI. Therefore, a small number of genes homologous tozmGRP4 are likely to exist in maize. In support of this, the amino acid sequence of another maize GRP cDNA (accession no.AF031083) and zmGRP4 share an 82% identical region of approximately 90 amino acid residues (data not shown). At the nucleotide sequence level, this region is 83% identical between these twoGRPs. Fig. 2. Open in new tabDownload slide Genomic DNA-hybridization analysis ofzmGRP4. Full-length zmGRP4 cDNA was used as a probe. Maize genomic DNA (20 μg) was digested withBamHI, BglII, EcoRI, orXhoI. The positions of the molecular markers are shown on the left. Fig. 2. Open in new tabDownload slide Genomic DNA-hybridization analysis ofzmGRP4. Full-length zmGRP4 cDNA was used as a probe. Maize genomic DNA (20 μg) was digested withBamHI, BglII, EcoRI, orXhoI. The positions of the molecular markers are shown on the left. zmGRP4 Is Expressed Strongly in the Lateral Root Cap and Weakly in the Epidermis of the Root Proper RNA blot-hybridization analysis with the full-length cDNA ofzmGRP4 as the probe detected zmGRP4 expression in the root tip but not in the root proper, etiolated shoot, shoot, or mature leaf (Fig. 3A). Since RNA-blot hybridization may not detect low levels of zmGRP4 expression and may detect expression of zmGRP4-related gene(s) as well, RT-PCR was done to specifically amplify zmGRP4 RNA (Fig.3B). Expression of zmGRP4 was detected in the root tip and root proper but not in the shoot and etiolated shoot. Although quantitative analysis is often difficult with RT-PCR, repeated RT-PCR analyses in which different amplification cycles were used (data not shown) confirmed that zmGRP4 is expressed more strongly in the root tip than in the root proper. Amplification of maize α-tubulin RNA indicated that approximately equal amounts of cDNA were used for each tissue sample. Fig. 3. Open in new tabDownload slide Expression of zmGRP4 in maize tissues. A, Northern-hybridization analysis. Poly(A+) RNA (1.5 μg) was isolated from 2- to 3-cm-long roots, 3-d-old etiolated shoots, and 3-d-old shoots, and leaves of 2-week-old plants. A maize ubiquitin probe served as a control to estimate the relative loading of RNA in each lane. B, RT-PCR analysis. S, Shoot; ES, etiolated shoot; root proper, 0- to 0.5-, 0.5- to 1.0-, and 1.0- to 1.5-cm regions distal from the excision site; RT, root tip; NT, negative control without reverse transcription. A maize α-tublin gene (TUA) served as a positive control. Fig. 3. Open in new tabDownload slide Expression of zmGRP4 in maize tissues. A, Northern-hybridization analysis. Poly(A+) RNA (1.5 μg) was isolated from 2- to 3-cm-long roots, 3-d-old etiolated shoots, and 3-d-old shoots, and leaves of 2-week-old plants. A maize ubiquitin probe served as a control to estimate the relative loading of RNA in each lane. B, RT-PCR analysis. S, Shoot; ES, etiolated shoot; root proper, 0- to 0.5-, 0.5- to 1.0-, and 1.0- to 1.5-cm regions distal from the excision site; RT, root tip; NT, negative control without reverse transcription. A maize α-tublin gene (TUA) served as a positive control. The 3′-untranslated region of the zmGRP4 cDNA was used as a probe for in situ-hybridization analysis to study the detailed expression pattern of zmGRP4 in maize primary root. The antisense probe detected strong zmGRP4 expression in the lateral root-cap cells and rather weak expression in epidermal cells of the root proper (Fig. 4A). The sense probe did not detect any hybridization signals (Fig. 4B). Peripheral cells that had been or were being detached from the lateral root cap showed little zmGRP4 expression (Fig. 4C, arrowheads), whereas weak zmGRP4 expression extended to several peripheral cells toward the central region of the root cap (Fig. 4D).zmGRP4 expression in epidermal cells of the root proper terminated in the region where zmGRP4 expression in the lateral root cap ended (Fig. 4C, arrow). Fig. 4. Open in new tabDownload slide In situ-hybridization analysis ofzmGRP4. Longitudinal sections of maize primary root tip were hybridized with antisense (A, C, and D) or sense (B) digoxigenin-labeled zmGRP4-specific probes. The images in C and D were enlarged from the rectangles in A. An arrow shows the end of zmGRP4 expression in the epidermis, whereas arrowheads indicate sloughed-off cap cells. Co, Columella; LRC, lateral root cap; Ep, epidermis; RAM, root apical meristem. Scale bars = 100 μm. Fig. 4. Open in new tabDownload slide In situ-hybridization analysis ofzmGRP4. Longitudinal sections of maize primary root tip were hybridized with antisense (A, C, and D) or sense (B) digoxigenin-labeled zmGRP4-specific probes. The images in C and D were enlarged from the rectangles in A. An arrow shows the end of zmGRP4 expression in the epidermis, whereas arrowheads indicate sloughed-off cap cells. Co, Columella; LRC, lateral root cap; Ep, epidermis; RAM, root apical meristem. Scale bars = 100 μm. zmGRP4 Accumulates in Root Mucilage A polyclonal antibody was raised against a truncated zmGRP4 protein that contained amino acid residues 132 to 192. This carboxy-terminal region of zmGRP4 includes amino acid stretches of low Gly abundance and is expected to be specific for zmGRP4. The closest homolog of zmGRP, encoded by a maize expressed sequence tag (AF031083), is 58% identical in this region (data not shown). Immunohistochemical analysis using this antiserum showed that zmGRP4 is present specifically in the mucilage that covers the root tip (Fig.5A). A preimmune mouse antiserum detected no signals (Fig. 5B). Longer exposure detected a relatively small amount of zmGRP4 in the lateral root-cap cells (Fig. 5C). Sloughed-off cap cells appeared to contain little zmGRP4 (Fig. 5C, red arrowheads). A weak signal was also observed in epidermal cells of the root proper in the distal 1 cm of the root tip (Fig. 5D). The presence of mucilage at the periphery of the root epidermis was not apparent because the layer of root epidermal mucilage is expected to be very thin (Foster, 1982). Fig. 5. Open in new tabDownload slide Immunolocalization of zmGRP4. Longitudinal sections of maize primary root tip were incubated with anti-zmGRP4 serum (A, C, and D) or preimmune serum (B). The lateral root-cap region adjacent to the root proper is shown with a higher magnification in C, whereas the root proper region 1 cm distal to the cap is shown in D. Alkaline-phosphatase reactions were done for 1 h in A and B, and for 3 h in C and D. Red arrowheads indicate sloughed-off cap cells. RC, Root cap; RAM, root apical meristem; M, mucilage; RP, root proper; LRC, lateral root cap; Cx, cortex; Ep, epidermis. Scale bars = 100 μm. Fig. 5. Open in new tabDownload slide Immunolocalization of zmGRP4. Longitudinal sections of maize primary root tip were incubated with anti-zmGRP4 serum (A, C, and D) or preimmune serum (B). The lateral root-cap region adjacent to the root proper is shown with a higher magnification in C, whereas the root proper region 1 cm distal to the cap is shown in D. Alkaline-phosphatase reactions were done for 1 h in A and B, and for 3 h in C and D. Red arrowheads indicate sloughed-off cap cells. RC, Root cap; RAM, root apical meristem; M, mucilage; RP, root proper; LRC, lateral root cap; Cx, cortex; Ep, epidermis. Scale bars = 100 μm. zmGRP4 May Be Posttranslationally Modified Immunoblot analysis using the anti-zmGRP4 serum revealed that zmGRP4 exists in the maize root as a major band with an apparent molecular mass of 36 kD and a minor band with an apparent molecular mass of 34 kD in the insoluble fraction that was extracted with the SDS-containing buffer (Fig. 6A). Extraction with Tris buffer without SDS did not recover any zmGRP4 protein. The 36-kD form was much more abundant in the root tip than in the root proper. A few faint bands of 27 and 25 kD were also detected in the insoluble fraction from the root tip. Since the deduced molecular mass of the mature zmGRP4 is 14.4 kD, posttranslational modifications of zmGRP4 in maize root are suspected. Manual removal of root mucilage and detached cap cells from the root tip markedly reduced the abundance of the 36-kD zmGRP4 in the preparation (Fig. 6B). This strongly suggests that zmGRP4 accumulates in root mucilage mainly as a modified 36-kD protein. DISCUSSION zmGRP4 Is a New Member of Maize GRPs Many structural cell wall proteins that have putative signal peptides and no catalytic domains have been reported in various plants (Showalter, 1993).These cell wall proteins are characterized by a high abundance of a single amino acid, repetitive sequence motifs, and a tendency to become insolubilized within the cell wall. The three major plant cell wall protein classes include HRGPs, PRPs, and GRPs. GRP cDNAs have been isolated from several plants, including three from maize. zmGRP is expressed in the epidermal cells of embryo, scutellar tissue, and young leaf, and induced by ABA, water stress, and wounding in leaves (Gómez et al., 1988). Since zmGRP does not have an amino-terminal signal peptide, it may be a cytosolic protein. Besides being rich in Gly, zmGRP has a putative RNA-binding motif (Gómez et al., 1988). Therefore, zmGRP and zmGRP4 belong to different subclasses of the GRP family. zmGRP3 (Goddemeier et al., 1998) has an N-terminal signal peptide but shows no significant homology to zmGRP4, except for abundant Gly residues. The expression ofzmGRP3 was root specific, with the highest expression level in the meristematic and elongation regions (Goddemeier et al., 1998). RNA-blot analysis of zmGRP3 indicates thatzmGRP3 and zmGRP4 are expressed in different regions of the maize root. zmGRP4 Expression in the Root The expression of cell wall proteins depends on cell type, developmental stage, and stress responses (Showalter, 1993).zmGRP4 is expressed strongly in the lateral root cap and weakly in root epidermis but scarcely in sloughed-off cap cells (Fig.4). The immunohistochemical localization of zmGRP4 (Fig. 5) strongly indicates that zmGRP4 is synthesized in lateral root-cap and root-epidermal cells and then secreted into the mucilage. Lateral root-cap cells develop considerable hypertrophied Golgi cisternae and are the main site of mucilage secretion. Maize root epidermal cells are also reported to contain hypertrophied dictyosome cisternae and release mucilage (Clarke et al., 1979; Foster, 1982). Detached cap cells, however, have dictyosomes that are no longer hypertrophied (Clowes and Juniper, 1968). This close correlation between zmGRP4expression and differentiation of secretion machinery suggests that zmGRP4 is secreted via hypertrophied Golgi cisternae into the mucilage. Likewise, bean GRP 1.8 was localized to dictyosomes of xylem parenchyma cells and was suggested to be exported into the walls of neighboring protoxylem vessels (Ryser and Keller, 1992). Sloughed-off cells did not express zmGRP4, whereas the outermost cap periphery cells did express zmGRP4 (Fig. 4, C and D). A notable switch in gene expression was also reported to occur upon cap-border cell differentiation in pea (Brigham et al., 1995). zmGRP4 Is Posttranslationally Modified Many cell wall proteins are modified posttranslationally. For example, Pro residues of HRGP are enzymatically converted into Hyp residues, which are then glycosylated to various degrees (Cassab, 1998). zmGRP4 mainly existed as a 36-kD protein, whereas the deduced molecular mass of mature zmGRP4 is 14.4 kD. The high Gly content in GRPs may cause aberrant electrophoretic migration on SDS gels. When zmGRP4 was expressed in Escherichia coli as a GST-fusion protein, the recombinant fusion protein detected was approximately 2-kD larger than expected by SDS-PAGE analysis (T. Matsuyama and T. Hashimoto, unpublished results). However, this aberrant migration alone does not explain the more than 20-kD difference between the expected and observed size of zmGRP4 extracted from maize root tips. Insolubilization of cell wall proteins has been observed in various developmental or stress-responsive processes (Cassab, 1998). Insolubilization of bean GRP 1.8 occurs during hypocotyl development (Keller et al., 1989). H2O2generated by fungal elicitor or glutathione treatment of bean or soybean cells causes oxidative cross-linking and therefore the insolubilization of PRP (Bradley et al., 1992). Recovery of isodityrosine after hydrolysis of cross-linked HRGP indicates that the Tyr hydroxy groups in HRGP undergo intermolecular condensation via H2O2 (Fry, 1986). zmGRP4 contains a relatively high percentage of Tyr residues. Oxidative cross-linking between zmGRP4s themselves or between zmGRP4 and other proteins via Tyr residues might result in insolubilization and increased molecular mass of zmGRP4. It should also be noted that PRPs insolubilized by H2O2 were not extracted even in SDS-containing buffer (Brisson et al., 1994), and potential cross-linking of xylem GRPs with the aromatic residues of lignin has also been proposed (Showalter, 1993; Cassab, 1998). The absence of lignin and polyphenolics in root mucilage suggests that the cross-linking partners of zmGRP4 may be at least partly different from those of previously reported cell wall proteins. Glycosylation is a common posttranslational modification found in secreted proteins. However, there are few reports of the potential glycosylation of GRPs (Showalter, 1993). Exceptions include a 30-kD GRP purified from soybean aleurone layers, which was reported to contain approximately 9% (w/w) sugars, including Man, Ara, Glc, Xyl, and Gal (Matsui et al., 1995). Purified soybean GRP showed a broad band after SDS-PAGE separation, indicating a microheterogeneity in the sugar component (Matsui et al., 1995). On the other hand, zmGRP4 extracted from maize root tips migrated as discrete bands on SDS-PAGE (Fig. 6). Since the deduced zmGRP4 amino acid sequence has no canonicalN-glycosylation sites, the modification could beO-glycosylation with homogeneous sugar side chains, if zmGRP4 were to be glycosylated. Possible Functions of zmGRP4 in Root Mucilage Soil and sand sheaths usually cling tightly to the roots of field-grown grasses such as maize root. The sheath is thought to be formed by the binding of soil particles in mucilage originating from the root (Vermeer and McCully, 1982, and the refs. therein). Root hairs are probably not primarily responsible for the adhesion of soil aggregates. Mucilage, soil particles, sloughed-off root-cap cells, and some soil bacteria form the rhizosphere, and the chemical and physical properties of the mucilage should be very important in determining the nature of the rhizosphere. Root mucilage is composed of 99.9% water (Guinel and McGully, 1986). The dry mass of mucilage consists mainly of polysaccharides and polyuronic acids (Jones and Morré, 1967; Floyd and Ohlrogge, 1970; Paull et al., 1975). Although proteins have been detected in maize mucilage (Chaboud, 1983), their properties and possible roles have attracted little attention. Previous chemical analysis of maize mucilage, from which detached cap cells have been mostly removed, showed that the amino acid composition is rich in Gly (13.8% of total amino acids) (Bacic et al., 1986). We have shown here that zmGRP4 is a mucilage protein and possibly the major component of the protein fraction. Other well-characterized GRPs are localized in the vascular system, and in xylem in particular (Ryser and Keller, 1992, and the refs. therein). Ultrastructural localization, however, has demonstrated that bean GRP 1.8 is localized to unlignified primary walls of protoxylem cells, and a correlation between GRP 1.8 deposition and lignification was evidently lacking in bean hypocotyls (Ryser and Keller, 1992; Ryser et al., 1997). An apparent positive correlation of GRP deposition with expansive growth and an inverse correlation with lignification have been reported for petunia ptGRP1, which is deposited at the cell wall/membrane interface, rather than within the cell wall (Condit, 1993). Thus, these GRPs may provide elasticity to the stretching wall or some protective environment to cells under frictional stress. Some GRP sequences are predicted to adopt β-pleated sheets composed of varying numbers of antiparallel strands; such a structure could provide elasticity and tensile strength during vascular development (Showalter, 1993). The soil sheath adhering along the entire length of field-grown maize roots is mostly permeated by mucilage, which is histochemically similar to that produced by the root cap (Vermeer and McCully, 1982). An experiment designed to measure the penetration resistance showed that maize roots receive much less frictional resistance than metal probes when growing into the soil (Bengough and McKenzie, 1997). One function of root mucilage, working together with sloughing root-cap cells, may be to decrease the frictional resistance during growth in the soil and to protect growing roots from abrasion by soil particles. If zmGRP4 has physical properties similar to other GRPs, it may provide elasticity to the root mucilage and may complement other mucilage components (e.g. polysaccharides and pectin) for a lubricant function. Large amounts of fixed C are secreted into the rhizosphere from the surface of grass roots (Russell, 1977). The secreted C is mostly in the form of sugar, but a wide range of amino acids, organic acids, vitamins, and auxins are either released from the roots or synthesized by microorganisms in the root environment (Bar-Yosef, 1996). These organic compounds may support survival and growth of detached cap cells and soil bacteria. Some of the compounds even may be involved in interactions between particular plant genotypes and soil microorganisms. Secreted proteins in the rhizosphere may play similar roles. In this regard, distribution of GRPs in root mucilages of other maize genotypes and other plant species, and the stability of zmGRP4 in the rhizosphere should be interesting to examine in the future. The accession number for the sequence described in this article isAB014475. ACKNOWLEDGMENTS We thank Dr. Katsumi Ueda for valuable advice on in situ-hybridization techniques and Dr. Robert Winz for critical reading of the manuscript. 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Proc Natl Acad Sci USA 88 1991 11505 11509 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported in part by a Grant-in-Aid for Scientific Research on Priority Areas (“The Molecular Basis of Flexible Organ Plans in Plants,” no. 06278103) from the Ministry of Education, Science, Sports and Culture of Japan to T.H. T.M. was supported by a Japan Society for the Promotion of Science Research Fellowship for Young Scientists (no. 5487). * Corresponding author; [email protected]; fax 81–743–72–5489. 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)

Scott, Derek B.; Jin, Wei; Ledford, Heidi K.; Jung, Hou-Sung; Honma, Mary A.

doi: 10.1104/pp.120.3.675pmid: 10398702

Abstract We have identified a new locus that regulates vegetative phase change and flowering time in Arabidopsis. An early-flowering mutant, eaf1(earlyflowering1) was isolated and characterized.eaf1 plants flowered earlier than the wild type under either short-day or long-day conditions, and showed a reduction in the juvenile and adult vegetative phases. When grown under short-day conditions, eaf1 plants were slightly pale green and had elongated petioles, phenotypes that are observed in mutants altered in either phytochrome or the gibberellin (GA) response.eaf1 seed showed increased resistance to the GA biosynthesis inhibitor paclobutrazol, suggesting that GA metabolism and/or response had been altered. Comparison of eaf1 to other early-flowering mutants revealed that eaf1 shifts to the adult phase early and flowers early, similarly to thephyB(phytochromeB) and spy(spindly) mutants. eaf1 maps to chromosome 2, but defines a locus distinct from phyB, clf(curlyleaf), and elf3(early-flowering3). These results demonstrate thateaf1 defines a new locus involved in an autonomous pathway and may affect GA regulation of flowering. The transition from vegetative to reproductive development is of vital importance for the survival of virtually all flowering plants. The vegetative phase of Arabidopsis can be subdivided into juvenile and adult phases that can be distinguished by physiological and morphological markers (Martinez-Zapater et al., 1995; Chien and Sussex, 1996; Telfer et al., 1997). Juvenile leaves are small and round in shape, with adaxial trichomes and no abaxial trichomes (Telfer et al., 1997). As development continues, emerging adult leaves are more elongated and lanceolate in shape, and trichomes begin to appear on the abaxial surfaces (Chien and Sussex, 1996; Telfer et al., 1997). Application of the hormone GA3 will induce abaxial trichomes on leaves where they are not normally present (Chien and Sussex, 1996; Telfer et al., 1997), although the earliest arising leaves do not respond to this induction. At the appropriate stage in development and in response to specific cues, the shoot apical meristem becomes reprogrammed and generates a reproductive shoot that carries either an inflorescence or a single flower. Daylength is a key regulator of flowering in many plant species, and in Arabidopsis flowering is hastened under LD conditions (Koornneef et al., 1998). Vernalization accelerates flowering in some Arabidopsis ecotypes, and the shoot meristem itself is thought to be the site of perception of the vernalization signal (Dennis et al., 1996; Wilson and Dean, 1996). Genetic studies with Arabidopsis and pea have identified a large number of genes that regulate flowering (Reid et al., 1996; Koornneef et al., 1998). In Arabidopsis at least two flowering pathways are thought to exist, a photoperiod-sensitive pathway and an autonomous pathway. GA may function as part of the autonomous pathway or could define a third pathway. Each of the flowering pathways includes both activator and repressor genes, and epistasis tests and physiological experiments have led to the idea that these pathways function in parallel. In Arabidopsis several genes that serve as promoters of flowering have been recently cloned, and the sequences of the proteins encoded as well as their expression patterns have suggested possible functions for these genes. For example, the genes CO (Putterill et al., 1995) and LD (Lee et al., 1994) both appear to encode transcription factors, whereasFCA possibly affects RNA metabolism (Macknight et al., 1997), suggesting that a cascade of gene activation events likely controls flowering. Genes that function to repress the transition to adult and/or reproductive phases have been identified in Arabidopsis (Koornneef et al., 1998). elf3(early-flowering3) (Hicks et al., 1996; Zagotta et al., 1996) and lhy (longhypocotyl) (Schaffer et al., 1998) are photoperiod-insensitive mutants that show alterations in circadian clock function and appear to be involved in the repression of flowering in the LD pathway. The HST(HASTY) gene is thought to promote a juvenile pattern of development, and loss-of-function mutations in this gene result in early transition to the adult phase and early flowering (Telfer and Poethig, 1998). Arabidopsis mutants defective in phytochrome synthesis (Reed et al., 1994; Koornneef et al., 1995) or phytochrome function (Ahmad and Cashmore, 1996) flower early, indicating that this light receptor is involved in repression of floral initiation. At least some of the effects of phytochrome may be mediated by GA, because recent studies (Reed et al., 1996) show that the hypocotyl tissue of phyB(phytochromeB) mutant seedlings is more responsive to exogenous GA than wild-type seedlings. The tfl(terminalflower) mutant flowers early and produces a determinate inflorescence, often generating only one or a few terminal flowers. Application of GA accelerates the onset of the adult phase, and induces early flowering and the production of larger, slightly pale-green leaves in wild-type Arabidopsis (Jacobsen and Olszewski, 1993; M. Honma, unpublished data). The GA-deficient mutant ga1 of Arabidopsis flowers later when grown in LD and does not flower in SD conditions, indicating that GA is required for the photoperiod-insensitive (autonomous) flowering pathway (Wilson et al., 1992). GA levels and response to the hormone are sensitive to changes in photoperiod (Pharis et al., 1987; Zeevaart and Gage, 1993), suggesting a role for GA in photoperiodic induction of flowering. Suppressors of the ga1 mutant have been identified and shown to suppress the late-flowering phenotype of ga1. These suppressors known as spy(spindly) (Jacobsen and Olszewski, 1993) and rga(repressor ofga1-3) (Silverstone et al., 1997) are thought to have partially activated the GA response.spy flowers early, and spy mutant seed show increased resistance to the GA biosynthesis inhibitor paclobutrazol. The double mutant rga ga1 flowers earlier thanga1 alone. The phenotypes of spy andrga suggest that activating the GA response can cause earlier flowering. We have identified a new gene, eaf1(earlyflowering1), that appears to function in the autonomous pathway to repress the transition from juvenile to adult development. eaf1 mutant plants exhibit truncated juvenile and adult phases, resulting in early flowering. eaf1 mutant plants exhibit phenotypes similar to the GA response mutants spy and rga, andeaf1 seed show increased resistance to paclobutrazol. Our results are consistent with the notion that the eaf1 mutant is altered in either GA biosynthesis or response to the hormone, and that this change is responsible for the alteration in flowering time. MATERIALS AND METHODS Arabidopsis Seed Stocks Seed stocks were obtained from the Arabidopsis Biological Resource Center at Ohio State University, Columbus, or from individual researchers (Drs. D.R. Meeks-Wagner, J. Reed, G. Coupland, T.-p. Sun, and R. Amasino). Acst and Dstransgenic lines of ecotype Nossen used to generate the eaf1mutant line were described previously (Honma et al., 1993). The 35S-Acst and rbcS-Acst lines express the Actransposase under the control of either the cauliflower mosaic virus 35S or the Arabidopsis rbcS-1A promoters. TheDsALS construct carries the Arabidopsis acetolactate synthase gene that encodes resistance to the herbicide chlorsulfuron. The DsALS element resides within the untranslated leader of a kanamycin resistance gene that serves as marker for excision of Ds. The eaf1mutant was originally identified in a mutant screen of a population carrying transposed Ds elements (Honma et al., 1993; M. Honma, unpublished data). The eaf1 mutant line contained three transposed Ds insertions: Ds-1,Ds-2, and Ds-3. Ds-1 and Ds-2 are tightly linked to the eaf1 mutation, and most of the characterization described in this paper was done using a line that carried both of these insertions. Introgression of the eaf1 mutation into the Landsberg background was accomplished by crossing eaf1 plants carryingDs-1 and Ds-2 to wild-type Landsberg plants. Backcross progeny were screened using chlorsulfuron to select for the presence of either Ds-1 orDs-2, both of which are linked to theeaf1 mutation. These chlorsulfuron-resistant plants were then backcrossed four more times to the Landsberg parent. No selection for the early-flowering trait was done during the introgression, to prevent selective maintenance of additional loci in the Nossen ecotype that may condition earlier flowering. F1 plants from the fifth backcross were self-fertilized and early-flowering F2 progeny were identified. Growth Conditions Plants were grown either under sterile conditions on germination medium (Valvekens et al., 1988) supplemented with appropriate antibiotics, as described previously (Honma et al., 1993), or in soil under LD (16 h of light/8 h of dark) or SD (8 h of light/16 h of dark) conditions at 18°C to 20°C during the dark period and at 20°C to 24°C during the light period. For most of the flowering-time experiments, seeds were either allowed to imbibe in water at 4°C for 4 d (in the dark or dim light) before transfer to soil or planted directly in moist soil and cold treated. After the cold treatment, seeds were transferred to growth rooms. Light intensity was 440 μE m−2 s−1 for SD and 240 μE m−2 s−1 for LD conditions. Lighting was supplied by a 3:1 mixture of cool-white:Wide Spectrum bulbs (General Electric). Plants were grown individually in divided flats (Hummert, St. Louis, MO) at a density of 60 to 96 plants/1290 cm2 flat. In the screen used to isolate the eaf1 mutant, seedlings were grown for 2 weeks on germination medium plates, followed by transfer to soil at a density of 150 plants/1290 cm2 flat. Chlorsulfuron was a gift from DuPont. In the initial experiment to determine linkage of the Dselements to the early-flowering phenotype, seedlings were first grown on germination medium plates for 10 to 14 d before transfer to soil. Plants were grown in a growth chamber (Conviron, Winnipeg, Manitoba, Canada) with a mixture of cool-white and incandescent bulbs. The light intensity was approximately 120 μE m−2s−1 and the temperature was maintained at 22°C. In the GA experiment sterilized seeds were plated on germination medium plates with or without 10−5mGA3 and allowed to imbibe at 4°C for 2 d before growth under SD (220 μmol photons m−2s−1) conditions in a growth chamber (model CU32L, Percival Scientific, Boone, IA). After 11 d of growth the seedlings were transferred to soil and grown under SD conditions, 440 μE m−2 s−1. GA3 (100 μL of 10−5m) was applied weekly to the base of each plant and continued until the plants had flowered. For vernalization treatment, seeds were allowed to imbibe by planting in moist soil; they were grown at 4°C for 8 weeks under low-intensity SD conditions (19 μE m−2s−1) before transfer to standard SD conditions (440 μE m−2 s−1, 22°C). Untreated control seeds were allowed to imbibe for 4 d at 4°C and transferred to the SD growth rooms on the same day as the vernalized plants. Morphological Analysis Days to flowering was scored as the length of time between germination and visible appearance of the first floral bud. The number of rosette leaves was counted weekly, and cauline leaves were counted after seed set. Juvenile-stage leaves were those true leaves present in the rosette that lacked abaxial trichomes. The appearance of abaxial trichomes was monitored using a stereomicroscope. Hypocotyl elongation in response to red light was measured as described previously by Nagatani et al. (1993). Allelism Tests eaf1 was crossed to the early-flowering mutantsclf(curlyleaf),elf3, and hy3, and F1 and F2 plants were scored for days to flowering and leaf number under SD conditions. clf andphyB(hy3) are alleles in the Landsberg ecotype and elf3 is in the Columbia ecotype. The control crosseseaf1 × Landsberg, eaf1 × Columbia, and Nossen × clf, elf3, orphyB(hy3) were included. Twenty to fifty plants were scored in each experiment. Molecular Analysis DNA was isolated from leaf tissue using a modification of the method described by Dellaporta et al. (1983). Southern blotting was by reverse capillary transfer as described previously (Ausubel et al., 1988) and Southern hybridizations were carried out according to the method of Church and Gilbert (1984). DNA fragments used as probes were a 1.5-kb fragment from the 5′ end of Ac and theDs-2 genomic flanking sequence (PCR product). These probes were generated by random-prime labeling (Feinberg and Vogelstein, 1983). Genetic Mapping of Ds-2 Insertion and theeaf1 Mutation Inverse PCR was used to isolate genomic DNA sequences flanking theDs-2 insertion (Healy et al., 1993). Mapping of this genomic sequence was done using recombinant inbred lines as described previously by Osborne et al. (1995). Wild-type Nossen, Columbia, and Landsberg ecotypes were compared using restriction fragment-length polymorphism analysis to determine if the region carrying eaf1 in Nossen showed polymorphisms with respect to the Columbia or Landsberg ecotypes. Southern analysis with restriction fragment-length polymorphism clones that map to the middle of chromosome 2 (Lister and Dean, 1993) showed that the eaf1region in Nossen appears to be polymorphic compared with the same region in Columbia (data not shown). Thus, the Nossen × Columbia cross was the most likely to yield polymorphisms that could be used for mapping. A mapping population was constructed by crossing theeaf1 mutant (Nossen background) carryingDs-1 and Ds-2 to Columbia wild type, and the F1 plants were self-fertilized to generate F2 siblings. Tissue was collected for molecular analysis from 596 early-flowering plants that arose from a population of approximately 2500 F2 plants. The position of the eaf1 mutation was determined using cleaved-amplified polymorphic sequence markers positioned on chromosome 2 (Lister and Dean, 1993) (http://genome-www.stanford.edu/Arabidopsis/ww/Aug98RImaps/index.html). Progeny from plants that showed recombination events between theeaf1 mutation and the marker tested were scored for their flowering phenotype to confirm that they were homozygous for theeaf1 mutation. Germination Assays Paclobutrazol resistance was determined as described previously (Jacobsen and Olszewski, 1993) with minor modifications. For each paclobutrazol treatment, 120 seed of each line were sterilized by treatment with 0.1% Triton. After the seeds were rinsed with water, they were washed with the respective paclobutrazol solutions and allowed to imbibe in the same solutions for 4 d at 4°C. Seeds were suspended in a small volume of 0.1% agarose, plated on four stacked filter paper circles in small Petri dishes, and allowed to dry (30 seeds/plate). Paclobutrazol (1.5 mL) in 0.01% Tween was applied to the filter paper and the Petri dishes were sealed with parafilm and incubated at 22°C for 7 d under a LD photoperiod. Germination was scored under a stereomicroscope as emergence of the radicle. Paclobutrazol was a gift from Zeneca (Wilmington, DE). RESULTS Isolation of an Early-Flowering Mutant,eaf1 A mutant that flowered earlier than the wild type was identified in a Ds-mutagenized population of plants of the Nossen ecotype grown under LD conditions. This early-flowering mutant was designated eaf1. The eaf1 mutant was characterized with regard to flowering time, appearance, and overall growth and development. Under LD conditions the mutant plants flowered 2 d before the wild type and generated three fewer leaves (TableI). Under SD conditions the early-flowering phenotype became much more extreme, and eaf1flowered 20 d earlier with 27 fewer leaves than the wild type. Theeaf1 mutant flowered earlier under LD than under SD conditions, indicating that it remains responsive to changes in the photoperiod. This is in contrast to the phenotype of other flowering-time mutants, elf3, co(constans), and gi(gigantea), that flower at the same time under either SD or LD conditions and appear to be photoperiod insensitive. Table I. Days to flowering and rosette leaf no. of wild-type Nossen and the eaf1 mutant in response to photoperiod Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) LD 21.2  ± 0.5 9.0  ± 0.3 66 eaf1 LD 19.3  ± 0.3 5.8  ± 0.1 66 WT (Nossen) SD 59.2  ± 1.0 43.5  ± 1.0 63 eaf1 SD 38.9  ± 0.5 16.8  ± 0.3 69 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) LD 21.2  ± 0.5 9.0  ± 0.3 66 eaf1 LD 19.3  ± 0.3 5.8  ± 0.1 66 WT (Nossen) SD 59.2  ± 1.0 43.5  ± 1.0 63 eaf1 SD 38.9  ± 0.5 16.8  ± 0.3 69 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type (WT) controls (P < 0.05, Student's t test). Open in new tab Table I. Days to flowering and rosette leaf no. of wild-type Nossen and the eaf1 mutant in response to photoperiod Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) LD 21.2  ± 0.5 9.0  ± 0.3 66 eaf1 LD 19.3  ± 0.3 5.8  ± 0.1 66 WT (Nossen) SD 59.2  ± 1.0 43.5  ± 1.0 63 eaf1 SD 38.9  ± 0.5 16.8  ± 0.3 69 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) LD 21.2  ± 0.5 9.0  ± 0.3 66 eaf1 LD 19.3  ± 0.3 5.8  ± 0.1 66 WT (Nossen) SD 59.2  ± 1.0 43.5  ± 1.0 63 eaf1 SD 38.9  ± 0.5 16.8  ± 0.3 69 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type (WT) controls (P < 0.05, Student's t test). Open in new tab Vernalization accelerates flowering in some Arabidopsis ecotypes (Napp-Zinn, 1985), and it is possible that eaf1 has an activated vernalization response. eaf1 and Nossen wild-type plants were tested for a response to vernalization. Seedlings were vernalized under SD conditions and scored for flowering time. As shown in Table II, both Nossen wild type andeaf1 respond to vernalization, flowering earlier with fewer rosette leaves. This result suggests that response to vernalization has not been altered in eaf1 or that this response has not been saturated. Table II. Days to flowering and rosette leaf no. of wild-type Nossen and the eaf1 mutant in response to vernalization Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 54.8  ± 1.0 41.9  ± 1.6 53 eaf1 SD 41.0  ± 0.6 20.2  ± 0.3 54 WT (Nossen) vern + SD 36.2  ± 1.4 39.9  ± 1.4 63 eaf1 vern + SD 26.1  ± 1.9 15.4  ± 0.3 63 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 54.8  ± 1.0 41.9  ± 1.6 53 eaf1 SD 41.0  ± 0.6 20.2  ± 0.3 54 WT (Nossen) vern + SD 36.2  ± 1.4 39.9  ± 1.4 63 eaf1 vern + SD 26.1  ± 1.9 15.4  ± 0.3 63 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type (WT) controls (P < 0.05, Student's t test). vern, Vernalization. Open in new tab Table II. Days to flowering and rosette leaf no. of wild-type Nossen and the eaf1 mutant in response to vernalization Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 54.8  ± 1.0 41.9  ± 1.6 53 eaf1 SD 41.0  ± 0.6 20.2  ± 0.3 54 WT (Nossen) vern + SD 36.2  ± 1.4 39.9  ± 1.4 63 eaf1 vern + SD 26.1  ± 1.9 15.4  ± 0.3 63 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 54.8  ± 1.0 41.9  ± 1.6 53 eaf1 SD 41.0  ± 0.6 20.2  ± 0.3 54 WT (Nossen) vern + SD 36.2  ± 1.4 39.9  ± 1.4 63 eaf1 vern + SD 26.1  ± 1.9 15.4  ± 0.3 63 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type (WT) controls (P < 0.05, Student's t test). vern, Vernalization. Open in new tab eaf1 Is Recessive and Not Allelic to Other Early-Flowering Mutants Genetic analysis showed that eaf1 is a recessive mutation. Homozygous eaf1 plants were crossed to wild-type Nossen, and the resulting eaf1/+ F1plants from two independent crosses (a and b) were scored for flowering (Table III). Plants that were heterozygous for eaf1 flowered similarly to wild-type control plants, showing no statistical difference in terms of days to flowering. Although the eaf1/+ plants flowered at the same time as wild-type plants, the numbers of rosette leaves appeared to be slightly reduced. One possibility is that the heterozygous plants have a slightly reduced rate of leaf initiation, and if so, this would indicate that the eaf1 mutation is not completely recessive for this phenotype. Two F1 plants were self-fertilized, and the F2 progeny were scored for flowering phenotype under SD conditions. The two F2 populations segregated mutant:wild-type plants in the ratios of 1:3, indicating that eaf1 is recessive and that the early-flowering phenotype was due to a mutation at a single locus (data not shown). Table III. Days to flowering and rosette leaf no. of the eaf1 mutant in the Nossen background Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 64.0  ± 3.1 49.6  ± 3.5 24 eaf1 SD 39.9  ± 6.0 20.3  ± 1.2 11 WT × eaf1 F1 a SD 70.0  ± 2.8 44.0  ± 1.0 12 WT × eaf1F1 b SD 65.0  ± 2.42-a 39.0  ± 3.4 8 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 64.0  ± 3.1 49.6  ± 3.5 24 eaf1 SD 39.9  ± 6.0 20.3  ± 1.2 11 WT × eaf1 F1 a SD 70.0  ± 2.8 44.0  ± 1.0 12 WT × eaf1F1 b SD 65.0  ± 2.42-a 39.0  ± 3.4 8 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type (WT) controls (P < 0.05, Student's t test). F2-a Not significantly different from wild-type control (P > 0.05, Student's t test). Open in new tab Table III. Days to flowering and rosette leaf no. of the eaf1 mutant in the Nossen background Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 64.0  ± 3.1 49.6  ± 3.5 24 eaf1 SD 39.9  ± 6.0 20.3  ± 1.2 11 WT × eaf1 F1 a SD 70.0  ± 2.8 44.0  ± 1.0 12 WT × eaf1F1 b SD 65.0  ± 2.42-a 39.0  ± 3.4 8 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 64.0  ± 3.1 49.6  ± 3.5 24 eaf1 SD 39.9  ± 6.0 20.3  ± 1.2 11 WT × eaf1 F1 a SD 70.0  ± 2.8 44.0  ± 1.0 12 WT × eaf1F1 b SD 65.0  ± 2.42-a 39.0  ± 3.4 8 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type (WT) controls (P < 0.05, Student's t test). F2-a Not significantly different from wild-type control (P > 0.05, Student's t test). Open in new tab The original eaf1 mutant line carried three Dsinsertions, Ds-1, Ds-2, andDs-3. Backcross of eaf1 to wild-type Nossen generated a set of lines carrying different combinations of theDs insertions. Lines that were hemizygous forDs-1 and Ds-2 orDs-2 and Ds-3 produced 25% mutant progeny. Plants carrying only Ds-3flowered similarly to the wild type. These results indicated thatDs-2 is most closely linked to theeaf1 mutation, with Ds-1 andDs-3 not responsible for causing the early-flowering phenotype. The Ds-1 insertion is approximately 2 centimorgans from eaf1, with theDs-3 insertion loosely linked on the same chromosome (data not shown). The genomic sequence flankingDs-2 was isolated by inverse PCR (seeMethods) and used in Southern hybridization experiments (data not shown). Analysis of 50 early-flowering F2 progeny showed that all were homozygous for the Ds-2 insertion, indicating thateaf1 was <1 centimorgan from Ds-2. Two lines hemizygous for Ds-2 alone have been identified; when self-fertilized, both lines produced mutant:wild-type progeny in the ratio of 1:3 (18 mutant:60 wild type and 20 mutant:59 wild type), confirming that Ds-1 andDs-3 were not responsible for causing the early-flowering phenotype and that the eaf1 mutation was tightly linked to the Ds-2 insertion. However, meiotic mapping experiments using cleaved-amplified polymorphic sequence markers have more precisely localized theDs-2 insertion site to 0.35 ± 0.2 centimorgans away from the eaf1 mutation; thusDs-2 is tightly linked but not inserted into theeaf1 gene (W. Jin and M. Honma, unpublished data). Moreover, sequence analysis of the genomic region flankingDs-2 indicates that the element is not inserted within an open reading frame (W. Jin and M. Honma, unpublished data). Assignment of an initial map position to the eaf1 mutation was determined with the aid of the Ds-2insertion, which lies 0.35 centimorgans away. The genomic sequence flanking Ds-2 was isolated by inverse PCR (seeMethods) and this genomic sequence was mapped using a recombinant inbred population (Lister and Dean, 1993). TheDs-2 insertion resides on chromosome 2, near mi238. A mapping population was generated (as described in Methods) by crossing eaf1 to wild-type Columbia plants, and the early-flowering phenotype was mapped using cleaved-amplified polymorphic sequence markers located in the middle of chromosome 2 (Lister and Dean, 1993) (http://genome-www.stanford.edu/Arabidopsis/ww/Aug98RImaps/index.html). Our results placed the eaf1 mutation between mi139 and mi238. The early-flowering mutationsphyB(hy3), clf, and elf3map within this region, but at locations different from eaf1(Fig. 1). eaf1 lies 1.9 centimorgans south of phyB (hy3) and is >1.7 centimorgans north of clf. elf3 is 6.6 centimorgans south ofeaf1, which is close to the marker GPA1 (Zagotta et al., 1996; K.A. Hicks, T.M. Albertson, and D.R. Meeks-Wagner, personal communication). To confirm that eaf1 was not allelic tophyB (hy3), clf, or elf3, complementation tests (described in Methods) were done. eaf1 (in Nossen) was crossed to phyB(hy3) (in Landsberg), clf (in Landsberg),elf3 (in Columbia), wild-type Landsberg, or wild-type Columbia. phyB(hy3), clf, andelf3 were each crossed to wild-type Nossen. All F1 plants flowered at the same time, and after self-fertilization they produced both early-flowering and wild-type F2 progeny (data not shown). These results demonstrate that eaf1 is not allelic tophyB(hy3), clf, and elf3. Therefore, eaf1 defines a new locus on chromosome 2 that affects flowering time, in addition to the previously knownphyB(hy3), clf, and elf3loci. Fig. 1. Open in new tabDownload slide Map location of eaf1 and other early-flowering mutants on chromosome 2. Distance shown with a bracket indicates number of centimorgans between the marker and the early-flowering phenotype. Distances shown with arrows were derived from the recombinant inbred lines (Lister and Dean, 1995). Fig. 1. Open in new tabDownload slide Map location of eaf1 and other early-flowering mutants on chromosome 2. Distance shown with a bracket indicates number of centimorgans between the marker and the early-flowering phenotype. Distances shown with arrows were derived from the recombinant inbred lines (Lister and Dean, 1995). eaf1 Regulates Vegetative-Phase Transition eaf1 mutant and wild-type plants were analyzed for the appearance of abaxial trichomes, a marker associated with the shift from the juvenile to the adult phase (Chien and Sussex, 1996; Telfer et al., 1997). The first rosette leaf bearing abaxial trichomes is counted as the first adult leaf. The number of juvenile, adult, and reproductive (cauline) leaves was determined, and the results are presented in Figure 2. Abaxial trichomes first appeared on leaf 8 in mutant plants as compared with leaf 13 in wild-type plants grown under SD conditions. Under LD conditions abaxial trichomes appeared on leaf 5 in mutant plants and leaf 6 in wild-type plants. These results show that the juvenile phase has been shortened in the mutant plants when grown in either SD or LD conditions. The adult phase in the mutant was also affected, because only nine adult leaves with abaxial trichomes were produced when grown under SD, as compared with 30 adult leaves in the wild type. Fig. 2. Open in new tabDownload slide Effect of daylength on number and type of leaves produced in wild-type Nossen and eaf1 mutant plants (A) grown under LD or SD conditions. Wild-type Landsberg andtfl, spy,phyB(hy3), and eaf1 mutant plants all in the Landsberg background grown under LD (B) or SD (C) conditions. Twenty to fifty plants of each line were tested and error bars represent ±2 se. Fig. 2. Open in new tabDownload slide Effect of daylength on number and type of leaves produced in wild-type Nossen and eaf1 mutant plants (A) grown under LD or SD conditions. Wild-type Landsberg andtfl, spy,phyB(hy3), and eaf1 mutant plants all in the Landsberg background grown under LD (B) or SD (C) conditions. Twenty to fifty plants of each line were tested and error bars represent ±2 se. eaf1 Mutant Plants Resemble Phytochrome and GA Signal Transduction Mutants eaf1 mutant plants had elongated petioles, were lighter green compared with wild-type plants (Fig.3), and produced hypocotyls that were approximately 20% longer than the wild type (data not shown). The elongated petiole and pale-green phenotype has also been observed with other early-flowering mutants, phyB, spy, andelf3 (Jacobsen and Olszewski, 1993; Reed et al., 1993;Zagotta et al., 1996) and the late-flowering mutant lhy(Schaffer et al., 1998). In addition, elf3 produces long hypocotyls when grown under SD conditions (Zagotta et al., 1992), andphyB and lhy mutant seedlings produce long hypocotyls when grown under LD conditions (Koornneef et al., 1980;Schaffer et al., 1998). Fig. 3. Open in new tabDownload slide Wild-type and eaf1 mutant plants (Landsberg ecotype) grown under SD conditions for approximately 8 weeks. eaf1 plants had flowered 3 weeks before this photograph was taken and wild-type plants had not yet flowered. Fig. 3. Open in new tabDownload slide Wild-type and eaf1 mutant plants (Landsberg ecotype) grown under SD conditions for approximately 8 weeks. eaf1 plants had flowered 3 weeks before this photograph was taken and wild-type plants had not yet flowered. Although we had previously shown that eaf1 was not allelic to phyB, it is still possible that eaf1 is defective in some other phytochrome or phytochrome-related process.eaf1 mutant and Nossen wild-type seedlings were tested for inhibition of hypocotyl elongation in response to red light, a phytochrome-mediated response. Both wild-type and eaf1mutant seedlings responded by producing shorter hypocotyls at higher fluences of red light, indicating that the eaf1 mutation was not affected in this aspect of phytochrome function. (M. Honma and J. Reed, unpublished data). GA has long been known to be involved in flowering, and application of GA accelerates the transition to the adult phase and flowering in Arabidopsis (Jacobsen and Olszewski, 1993). Some of the phenotypic characteristics of eaf1 mutant plants, such as early flowering, pale color, and elongated petioles, are similar to GA-treated plants or mutants altered in GA response (Jacobsen and Olszewski, 1993). Therefore, GA levels might be elevated or GA response may have been increased in the eaf1 mutant. To test whethereaf1 is able to respond to exogenous GA, an experiment was performed by applying GA3 and measuring flowering time. If the GA signal transduction pathway has been activated such that its response is saturated, then increased levels of exogenous GA will not have an effect. Our results showed that eaf1 was still able to respond to applied GA, and treated plants exhibited more elongated petioles, were paler green (data not shown), and were earlier flowering (Table IV). This suggests that, even if GA synthesis or signaling has been activated, the response to the hormone had not been saturated. Table IV. Days to flowering and rosette leaf no. of wild-type Nossen and the eaf1 mutant in response to GA Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 60.1  ± 1.0 ND 47 eaf1 SD 37.2  ± 0.7 ND 41 WT (Nossen) GA + SD 53.9  ± 1.8 ND 40 eaf1 GA + SD 33.8  ± 1.7 ND 32 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 60.1  ± 1.0 ND 47 eaf1 SD 37.2  ± 0.7 ND 41 WT (Nossen) GA + SD 53.9  ± 1.8 ND 40 eaf1 GA + SD 33.8  ± 1.7 ND 32 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type controls (P < 0.05, Student's t test). ND, Not determined. Open in new tab Table IV. Days to flowering and rosette leaf no. of wild-type Nossen and the eaf1 mutant in response to GA Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 60.1  ± 1.0 ND 47 eaf1 SD 37.2  ± 0.7 ND 41 WT (Nossen) GA + SD 53.9  ± 1.8 ND 40 eaf1 GA + SD 33.8  ± 1.7 ND 32 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Nossen) SD 60.1  ± 1.0 ND 47 eaf1 SD 37.2  ± 0.7 ND 41 WT (Nossen) GA + SD 53.9  ± 1.8 ND 40 eaf1 GA + SD 33.8  ± 1.7 ND 32 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type controls (P < 0.05, Student's t test). ND, Not determined. Open in new tab Quantitative measurements of response to GA or GA inhibitors on soil-grown plants are extremely difficult. To address the question of whether eaf1 is altered in some aspect of GA function, seed germination in response to paclobutrazol was examined. Germination requires GA, and paclobutrazol interferes with GA biosynthesis such that germination of wild-type seeds is inhibited. Germination of wild-type Nossen and eaf1 mutant seeds was measured in the presence of varying levels of paclobutrazol (Fig.4). eaf1 showed increased resistance to paclobutrazol compared with wild-type Nossen. Between 0 and 3 μm paclobutrazol, both the wild type andeaf1 showed similar high levels of germination. Germination of wild-type Nossen dropped to below 75% on 10 μm paclobutrazol, whereas eaf1germination remained at 95%. Increasing levels of paclobutrazol showed a decreasing level of germination for the wild type, with only 15% germination at 300 μm. In contrast, germination of eaf1 seed remained high at 100 μm, and even at 300 μmpaclobutrazol, 35% of the seed still germinated. Thus, theeaf1 mutant shows increased resistance to paclobutrazol, similarly to the spy mutant (Jacobsen and Olszewski, 1993). This resistance could be the result of elevated levels of GA, such that higher levels of paclobutrazol are required to have an effect, or the result of an altered response to GA. The pale pigmentation, early-flowering, elongated petiole, and paclobutrazol-resistant phenotypes of eaf1 mutant plants are all consistent with the notion that GA levels or responses have been altered. Fig. 4. Open in new tabDownload slide Response of eaf1 to paclobutrazol. Percentage germination of Nossen wild-type and eaf1mutant seed in the presence of exogenous paclobutrazol. Open bars represent eaf1; striped bars represent wild type. Error bars represent ±1 se. Fig. 4. Open in new tabDownload slide Response of eaf1 to paclobutrazol. Percentage germination of Nossen wild-type and eaf1mutant seed in the presence of exogenous paclobutrazol. Open bars represent eaf1; striped bars represent wild type. Error bars represent ±1 se. Comparison of eaf1 and Other Early-Flowering Mutants To determine whether all early-flowering mutants show premature appearance of abaxial trichomes, flowering time and abaxial trichome formation were examined in a variety of early-flowering mutants grown under different photoperiods (Table V; Fig. 2). eaf1 was compared with the early-flowering mutantsphyB(hy3), spy, and tfl, of which alleles exist in the Landsberg ecotype. Because wild-type Landsberg and Nossen differ in flowering time and other characteristics, the eaf1 mutation was introgressed into the Landsberg background for this comparison. Plants of each mutant line were grown in either LD or SD conditions, and the appearance of abaxial trichomes and time of floral bud emergence were noted. Two experiments were done and similar results were obtained. Table V. Days to flowering and rosette leaf no. of wild-type Landsberg and various early-flowering mutants Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Landsberg) LD 19.5  ± 0.5 5.3  ± 0.2 40 tfl1-2 LD 15.9  ± 0.4 3.2  ± 0.1 40 spy-8 LD 17.5  ± 0.4 4.0  ± 0 7 phyB(hy3) LD 16.7  ± 0.5 3.1  ± 0.1 38 eaf1L LD 15.7  ± 0.4 3.9  ± 0.1 23 WT (Landsberg) SD 61.3  ± 1.3 36.5  ± 1.3 28 tfl1-2 SD 62.3  ± 1.54-a 37.3  ± 1.04-a 25 spy-8 SD 36.6  ± 0.6 14.8  ± 0.5 22 phyB(hy3) SD 55.2  ± 2.5 23.4  ± 2.7 9 eaf1L SD 31.6  ± 0.3 12.0  ± 0.3 29 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Landsberg) LD 19.5  ± 0.5 5.3  ± 0.2 40 tfl1-2 LD 15.9  ± 0.4 3.2  ± 0.1 40 spy-8 LD 17.5  ± 0.4 4.0  ± 0 7 phyB(hy3) LD 16.7  ± 0.5 3.1  ± 0.1 38 eaf1L LD 15.7  ± 0.4 3.9  ± 0.1 23 WT (Landsberg) SD 61.3  ± 1.3 36.5  ± 1.3 28 tfl1-2 SD 62.3  ± 1.54-a 37.3  ± 1.04-a 25 spy-8 SD 36.6  ± 0.6 14.8  ± 0.5 22 phyB(hy3) SD 55.2  ± 2.5 23.4  ± 2.7 9 eaf1L SD 31.6  ± 0.3 12.0  ± 0.3 29 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type controls (P < 0.05, Student's t test). F4-a Not significantly different from wild-type controls (P > 0.05, Student's t test). Open in new tab Table V. Days to flowering and rosette leaf no. of wild-type Landsberg and various early-flowering mutants Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Landsberg) LD 19.5  ± 0.5 5.3  ± 0.2 40 tfl1-2 LD 15.9  ± 0.4 3.2  ± 0.1 40 spy-8 LD 17.5  ± 0.4 4.0  ± 0 7 phyB(hy3) LD 16.7  ± 0.5 3.1  ± 0.1 38 eaf1L LD 15.7  ± 0.4 3.9  ± 0.1 23 WT (Landsberg) SD 61.3  ± 1.3 36.5  ± 1.3 28 tfl1-2 SD 62.3  ± 1.54-a 37.3  ± 1.04-a 25 spy-8 SD 36.6  ± 0.6 14.8  ± 0.5 22 phyB(hy3) SD 55.2  ± 2.5 23.4  ± 2.7 9 eaf1L SD 31.6  ± 0.3 12.0  ± 0.3 29 Genotype . Growth Condition . Days to Flowering . Leaf No. . n . WT (Landsberg) LD 19.5  ± 0.5 5.3  ± 0.2 40 tfl1-2 LD 15.9  ± 0.4 3.2  ± 0.1 40 spy-8 LD 17.5  ± 0.4 4.0  ± 0 7 phyB(hy3) LD 16.7  ± 0.5 3.1  ± 0.1 38 eaf1L LD 15.7  ± 0.4 3.9  ± 0.1 23 WT (Landsberg) SD 61.3  ± 1.3 36.5  ± 1.3 28 tfl1-2 SD 62.3  ± 1.54-a 37.3  ± 1.04-a 25 spy-8 SD 36.6  ± 0.6 14.8  ± 0.5 22 phyB(hy3) SD 55.2  ± 2.5 23.4  ± 2.7 9 eaf1L SD 31.6  ± 0.3 12.0  ± 0.3 29 Days to flowering was measured as the no. of days from germination to appearance of the floral bud. Leaf no. is the no. of rosette leaves produced before flowering. Each value represents the mean ± 2se. Unless otherwise noted, plants from within each group were significantly different from the wild-type controls (P < 0.05, Student's t test). F4-a Not significantly different from wild-type controls (P > 0.05, Student's t test). Open in new tab One set of experiments is shown in Table V and Figure 2. In the Landsberg background, eaf1 exhibited a shorter juvenile phase and flowered early in LD conditions. spy also produced fewer juvenile leaves and both eaf1 and spyproduced the normal number of adult leaves.phyB(hy3) and tfl had slightly shortened juvenile phases, had markedly reduced adult phases, and flowered early. The phenotype of tfl1-2 in the LD condition is similar to what has been previously reported for Columbia alleles of tfl (Telfer and Poethig, 1998). When grown under SD conditions, tfl appeared to be very similar to wild-type Landsberg. The reason for no observable phenotype in the SD condition is not clear, but alleles of tfl in the Landsberg ecotype have been previously reported to show a milder phenotype than those in the Columbia ecotype (Alvarez et al., 1992). Perhaps the light intensity, light quality, or temperatures used in our experiments were unable to reveal such a phenotype.phyB(hy3) flowered early under SD conditions, but the length of the adult phase appeared to be less affected than either spy oreaf1. When grown under the LD condition, the adult phase ofphyB(hy3) appeared to have been shortened compared with the wild type. spy and eaf1 behaved very similarly when grown in SD conditions, with reduced juvenile and greatly reduced adult phases and early flowering. Thus, by comparison,eaf1 is similar phenotypically to spy under both photoperiod regimes tested. DISCUSSION The EAF1 gene that controls vegetative-phase change and flowering time has been identified by mutational analysis and shown to reside on chromosome 2. Both the juvenile and adult phases ofeaf1 mutant plants are shortened, resulting in an early transition to reproductive development. eaf1 appears primarily to affect the length of the adult phase, with a less dramatic alteration of the juvenile phase. The eaf1 allele behaves as a recessive mutation, and if this mutation is due to loss-of-function of the eaf1 gene, then the EAF1 gene product may function to repress flowering by delaying adult development. Alternatively, eaf1 could be a recessive neomorph, in which case the wildtype product may not normally function to regulate flowering. Additional alleles of the eaf1 gene will provide important information regarding the role of EAF1 in control of flowering. Seed of eaf1 show increased resistance to paclobutrazol compared with Nossen wild type, a phenotype also seen with thespy mutant. Increased resistance to paclobutrazol suggests that eaf1 is involved in regulation of GA levels or response to the hormone. An increase in bioactive GAs could be the result of increased biosynthesis, decreased catabolism or inactivation, or loss of feedback regulation on the biosynthetic pathway (Chiang et al., 1995). Elongation of the inflorescence stem (bolt) after flowering is a GA-regulated process, and paclobutrazol treatment will inhibit elongation. Preliminary results indicate that eaf1 plants are more resistant to paclobutrazol than wild-type Nossen in terms of bolt elongation, demonstrating that early developmental stages such as germination and late stages such as bolting are both altered ineaf1. These results support the idea that alteration in GA metabolism or signaling in eaf1 is responsible for the early-flowering phenotype. Levels of paclobutrazol resistance observed with the Nossen wild type are higher than has been seen previously with wild-type seeds of Landsberg or Columbia ecotypes (D. Scott and M. Honma, unpublished data). Thus, the Nossen ecotype may produce more bioactive GA than other ecotypes or have an altered response to the hormone. Plants of the Nossen ecotype have leaves that are paler green with longer petioles than Landsberg or Columbia plants, phenotypes consistent with increased GA levels or GA signaling. Identification of loci that differ between Nossen and Landsberg responsible for resistance to paclobutrazol is currently in progress. GA is known to promote vegetative-phase change and flowering in a variety of plants, including Arabidopsis (Chien and Sussex, 1996;Telfer et al., 1997). The SPY, RGA, andGAI genes are negative regulators of GA signaling (Jacobsen and Olszewski, 1993; Peng et al., 1997; Silverstone et al., 1997), and loss-of-function alleles exhibit phenotypes indicating that these genes act downstream of GA biosynthesis. The spy mutant of Arabidopsis, which is altered in response to GA, exhibits phenotypic modifications similar to GA-treated plants and is able to suppress most of the ga1 phenotypic changes, including reduced germination. Although eaf1 does possess some phenotypic alterations in common with spy, it does not show increased height or reduced fertility. However, because only one allele ofeaf1 is currently available, other mutant alleles may have more severe phenotypes or eaf1 may regulate a different subset of GA-controlled functions than spy. Bothrga and spy are able to suppress the late-flowering defect of ga1 and accelerate the production of adult leaves (Silverstone et al., 1997). It is thought thatrga functions downstream of GA biosynthesis, in a pathway independent of spy (Silverstone et al., 1997). Preliminary characterization of an eaf1 ga1 double-mutant line suggests that eaf1 is not able to suppress the germination defect ofga1, similar to rga and gai mutations (H. Ledford and M. Honma, unpublished data). If eaf1 is also involved in GA response, this mutation will likely be able to suppress the late-flowering alteration of ga1. Thus, we would expect that ga1 eaf1 would flower earlier than ga1 under LD conditions. GAI also functions as a negative regulator of GA response, and GA can release this repression (Peng et al., 1997). Recently, RGA and GAI have been shown to encode proteins with similar amino acid sequences, suggesting that they may have redundant functions in GA signaling (Peng et al., 1997;Silverstone et al., 1998). The role of GAs in promotion of flowering could be the consequence of early transition to the adult phase, which then hastens transition to reproductive development. Alternatively, regulation of transition to the adult phase might be independent of transition to the reproductive phase, but with components in common, one of which may be GAs. GAs could contribute to generate a signal that promotes phase transition or may function in making the meristem more competent to respond to such factors. Thus, GAs may act by causing developmental changes that eventually result in early flowering, rather than acting directly as a floral inducer. Recently it was shown that expression of the floral meristem identity gene LEAFY is regulated in response to GA (Blázquez et al., 1998). Construction of double-mutant lines altered in eaf1 and the GA and phytochrome genes will allow study of genetic interactions between these genes. Such experiments are in progress and will indicate whether eaf1 functions within the GA or photoperiod pathways. Flowering-time mutants can be grouped into classes based on duration of juvenile and adult phases. To compare eaf1 with other early-flowering mutants, it was introgressed into the Landsberg background and the duration of juvenile and adult phases and flowering time of all genotypes compared. eaf1 appears to be most similar to the spy mutant, which shows reduced juvenile and adult phases. In contrast, the hst gene (Telfer and Poethig, 1998) appears to have a primary role during the juvenile phase.hst mutants exhibit a shortened juvenile phase and a normal-length adult phase, flower earlier than wild-type plants, and appear to be pleiotropic (Telfer and Poethig, 1998). When grown under LD conditions, we find that tfl mutant plants have a slightly shorter juvenile phase, a greatly shortened adult phase, and flower early, as has been previously reported (Shannon and Meeks-Wagner, 1991). The existence of mutants that primarily affect one phase but not the other would suggest that flowering and phase transition are separate processes, which may share common regulatory factors. eaf1 defines a new locus in Arabidopsis that represses the shift to the adult phase. Future studies ofeaf1 in conjunction with GA response and flowering-time genes will further our understanding of the complex interactions that control vegetative-phase change and reproductive development. ACKNOWLEDGMENTS We thank Dr. J. Reed (University of North Carolina, Chapel Hill) for collaboration on the hypocotyl elongation experiment, Dr. B. Osborne and C. Corr (Plant Gene Expression Center [PGEC]-U.S. Department of Agriculture [USDA], Albany, CA) for mapping theDs-2 insertion, Dr. M. Anderson (Nottingham Arabidopsis Stock Centre) for assistance with mapping ofphyB, and Dr. C. Waddell (PGEC-USDA) for collaborating on the initial Ac/Ds mutant screen. 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IBN-9509229), and the North Carolina Biotechnology Center (no. 9513 ARG-0039) to M.A.H. * Corresponding author; e-mail [email protected]; fax 1–919–613–8177. 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)

Shimogawara, Kosuke; Wykoff, Dennis D.; Usuda, Hideaki; Grossman, Arthur R.

doi: 10.1104/pp.120.3.685pmid: 10398703

Abstract P-starved plants scavenge inorganic phosphate (Pi) by developing elevated rates of Pi uptake, synthesizing extracellular phosphatases, and secreting organic acids. To elucidate mechanisms controlling these acclimation responses in photosynthetic organisms, we characterized the responses of the green algaChlamydomonas reinhardtii to P starvation and developed screens for isolating mutants (designated psr[phosphorus-stress response]) abnormal in their responses to environmental levels of Pi. Thepsr1-1 mutant was identified in a selection for cells that survived exposure to high concentrations of radioactive Pi. psr1-2 andpsr2 were isolated as strains with aberrant levels of extracellular phosphatase activity during P-deficient or nutrient-replete growth. The psr1-1 andpsr1-2 mutants were phenotypically similar, and the lesions in these strains were recessive and allelic. They exhibited no increase in extracellular phosphatase activity or Pi uptake upon starvation. Furthermore, when placed in medium devoid of P, the psr1 strains lost photosynthetic O2evolution and stopped growing more rapidly than wild-type cells; they may not be as efficient as wild-type cells at scavenging/accessing P stores. In contrast, psr2 showed elevated extracellular phosphatase activity during growth in nutrient-replete medium, and the mutation was dominant. The mutant phenotypes and the roles of Psr1 and Psr2 in P-limitation responses are discussed. P is a nutrient that often limits plant growth in the natural environment. The primary source of P in soils is Pi, which is actively accumulated by both plants and microbes. However, most soil Pi is either covalently bonded to C molecules as Pi esters, or exists as Fe3+, Al3+, or Ca2+ salts. These Pi salts are relatively insoluble and, therefore, are not readily available for transport into microbial cells or plant roots (Halstead and McKercher, 1975). When plants or microbes are starved for P, they exhibit increased Pi uptake (McPharlin and Bieleski, 1987; Furihata et al., 1992;Shimogawara and Usuda, 1995; Muchhal et al., 1996; Jeschke et al., 1997; Schachtman et al., 1998), secrete acid and alkaline phosphatases (Lefebvre et al., 1990; Duff et al., 1991) and RNases (Loffler et al., 1993; Bariola et al., 1994; Kock et al., 1995; Dodds et al., 1996), and exude low-Mr organic acids that help mobilize stores of Pi that are present in the soil as insoluble salts (Marschner, 1995). Studies with photosynthetic organisms have also demonstrated that glycolysis and photosynthetic activities are modified during P deprivation (Brooks, 1986; Duff et al., 1989; Dietz and Heilos, 1990; Jacob and Lawlor, 1993; Theodorou and Plaxton, 1993;Plesnicar et al., 1994; Wykoff et al., 1998). The mechanisms that control the acclimation of Escherichia coli andSaccharomyces cerevisiae to P limitation have been extensively studied (Wanner, 1993; Oshima et al., 1996; Oshima, 1997). In E. coli a two-component regulatory system governs the transcription of many genes that are responsive to the P levels of the environment (Wanner, 1993). Recently, similar regulatory systems have been identified in Bacillus subtilis and the cyanobacteriumSynechococcus sp. strain PCC 7942 (Aiba et al., 1993;Hulett, 1996). In S. cerevisiae many mutants (phoseries mutants) have been isolated that have lost their ability to regulate the synthesis of extracellular phosphatases in response to P starvation (Lenburg and O'Shea, 1996; Oshima, 1997). The lesions in these mutants define genes encoding both catalytic and regulatory functions that are important for the acclimation of S. cerevisiae to P limitation. Some of these gene products, including acid and alkaline phosphatases (Pho5 and Pho8) and a high-affinity phosphate transporter (Pho84), are involved in scavenging the limiting nutrient. Others function as transcriptional regulators (Pho4 and Pho2), a cyclin (Pho80), a cyclin-dependent kinase (Pho85), and a cyclin-dependent kinase inhibitor (Pho81); these regulators coordinate limited nutrient availability with the growth and metabolism of the cell. The existence of a similar regulatory pathway inNeurospora crassa has also been established (Kang and Metzenburg, 1990, 1993; Pelleg et al., 1996). In an attempt to define mechanisms that control the acclimation of photosynthetic eukaryotes to low levels of P, we have identified mutants of Chlamydomonas reinhardtii with aberrant responses to P limitation. C. reinhardtii is a unicellular green alga that has been developed as a model organism for analyzing a number of different physiological processes in photosynthetic eukaryotes, and in particular for the dissection of photosynthesis (Harris, 1989). Many molecular techniques have been developed that allow for sophisticated molecular manipulation of this organism (Rochaix, 1995; Davies and Grossman, 1998; Shimogawara et al., 1998). To elucidate mechanisms that photosynthetic organisms use to sense and respond to P deprivation, we have characterized the responses of C. reinhardtii to P limitation (Quisel et al., 1996; Wykoff et al., 1998) and have isolated mutants of this alga that do not properly acclimate to P limitation. These mutants were selected in two different screens. The first screen involved the isolation of strains that survived high concentrations of radioactive Pi during starvation for P. The second screen identified mutants that were unable to accumulate extracellular phosphatases during P-limited growth or that were unable to completely repress the accumulation of extracellular phosphatase during nutrient-replete growth. Here we describe the physiological and genetic characteristics of these mutants and what they have revealed about the mechanisms that control the responses of C. reinhardtii to P limitation. MATERIALS AND METHODS Strains, Culture Medium, and Growth Condition Chlamydomonas reinhardtii Dangeard strains CC125 (wild type mt+), CC124 (wild type mt−), and CC425 (cw15 arg7-8 mt+) were grown in TAP (Tris-acetate-Pi) medium (Harris, 1989) or TAP medium supplemented with 50 μmol mL−1 Arg. Cells grown in Erlenmeyer flasks were agitated on a gyratory shaker (120 rpm), maintained at 27°C, and illuminated at 80 μmol photons m−2s−1 from cool-white fluorescent tubes. For P-starvation experiments Pi was eliminated from the culture medium and replaced with TA medium (1.5 mm KCl). To prepare TA solid medium, 0.5% (w/v) agarose (Agarose-I, electrophoretic grade, Dojindo, Kumamoto, Japan) was used instead of 1.2% agar, because there was Pi contamination in the latter. UV Mutagenesis and 32Pi Suicide Selection of Mutants Strain CC125 was grown to mid-logarithmic phase (2 × 106 cells mL−1), pelleted by centrifugation (4000g), and suspended in 10 mL of fresh TA medium. The cell suspension was placed in a Petri dish, exposed to UV irradiation from a germicidal UV tube (20 W, distance 50 cm, and 150 s), and then incubated in the dark for 1 d. High specific activity of 32Pi (10 μCi nmol−1) was added to the cultures of mutagenized cells to a final concentration of 10 μm to kill cells that developed an elevated capacity for Pi uptake during P limitation. The cell suspension was incubated in the light (50 μmol photons m−2 s−1) for 1 d and then placed at 4°C in the dark for 1 week. Cold treatment accelerated cell death by retarding processes involved in repairing damage caused by 32Pi accumulation. Surviving cells were spread onto solid medium containing 10 mm Pi and then screened for growth on solid medium containing high (10 mm) and low (10 μm) Pi. Strains that grew normally on high Pi but did not grow well on low Pi were further analyzed. Putative mutants were back-crossed four to five times with parental strains (CC124 and then CC125) before further characterizations. Insertional Mutagenesis and Screening for Phosphatase Mutants The plasmid pJD67, harboring the arginosuccinate lyase gene (ARG7) (Davies et al., 1994), was linearized by digestion with HindIII and transformed (Kindle, 1990) into the Arg auxotroph CC425. Mutagenized cells were screened for aberrant accumulation of extracellular phosphatases (Quisel et al., 1996) during P starvation. ARG transformants were replica plated at low density onto solid TAP medium with low (10 μm) or normal (1 mm) Pi and grown for 2 or 3 d in fluorescent light of 50 μmol photons m−2s−1. Colonies were sprayed with an aqueous solution of 10 mm X-Pi as a visual assay for phosphatase activity (Davies et al., 1994). Direct Measurement of Pi Uptake Pi uptake was measured using a procedure similar to that described for the uptake of S (Yildiz et al., 1994). The cells were stirred as a dilute suspension in the light (200 μmol photons m−2 s−1) for 2 min before the addition of 33Pi. At varying times after the addition of the radiolabeled anion, the cells were vacuum filtered onto Supor-450 membranes (pore size 0.45 μm, Gelman Sciences, Ann Arbor, MI), and the membranes were washed with 10 mL of ice-cold TAP medium containing 20 mm Pi. The radioactivity on each filter was quantified in a liquid-scintillation counter (LKB Wallac, Turku, Finland). Generation of Vegetative Diploids Vegetative diploids were constructed according to the method ofHarris (1989). NIT1 and NIT2 alleles were introduced into parental haploid strains, and diploid cells were selected for growth on solid medium containing nitrate as the sole N source (TAP-N +NO3; NH4Cl in the TAP medium was replaced by 3.5 mmKNO3). O2 Evolution Light-saturated (800 μmol photons m−2s−1) photosynthesis was measured at 27°C as O2 evolution using a Clark-type O2 electrode (Hansatech, UK) as described elsewhere (Wykoff et al., 1998). Secreted Phosphatase Activity and Periplasmic Protein Analysis Cells were washed twice with TA medium and then resuspended in appropriate medium for growth. Phosphatase activity was measured at 27°C and pH 8.5 using p-nitrophenyl phosphate as the substrate, as previously described (Quisel et al., 1996). The hydrolysis of the substrate was limited by the amount of cells added to the assay mixture and was proportional to the incubation time for at least 1 h. The back-crossed (3–5 times) mutant strains were crossed to our cell wall-less wild-type strain (cw15), and periplasmic polypeptides were isolated from the mutant, cell wall-less progeny and resolved by SDS-PAGE according to the method of Davies et al. (1994). The polypeptides were visualized by silver staining (Porro et al., 1982). RESULTS Measurement of Pi Uptake Filtration assays were used to determine the characteristics of Pi transport into C. reinhardtii cells grown under both nutrient-replete and P-starved conditions (Yildiz et al., 1994). The uptake of Pi by cells grown in nutrient-replete medium or starved for Pi for 24 h is shown as a function of the initial Pi concentration in Figure 1. TheVmax for Pi uptake increased by over 10-fold in cells starved for P (Fig. 1). The kinetic analysis of Pi uptake after nutrient-replete growth revealed two distinct kinetic components. The Km for one (low-affinity component) was approximately 10 μm, whereas that for the other (high-affinity component) was between 0.1 and 0.3 μm, as derived from the double-reciprocal plot (Fig. 1, inset). The low-affinity component comprised approximately 80% of total Pi uptake under nutrient-replete conditions. After 24 h of P starvation, all of the Pi uptake seemed to occur via the high-affinity system. Hence, P starvation of C. reinhardtiiresulted in an enhanced capacity of the cells to take up Pi and an enhanced affinity for Pi. These results suggest that more than one Pi transport system is used by C. reinhardtii and that the high-affinity system is responsible for most of the transport observed in P-starved cells. Fig. 1. Open in new tabDownload slide The velocity of Pi uptake as a function of substrate concentration. 33Pi uptake of the wild-type strain CC125 was performed as described in Methods. Cultures in the early logarithmic phase of growth were either transferred to TAP (+P, •) or TA (−P, ○) medium and allowed to continue growth for 24 h before measuring Pi uptake. The insets are double-reciprocal plots of the data, and theKm values estimated from these plots are 0.16 μm for the high-affinity component and 10 μm for the low-affinity component. chl, Chlorophyll. Fig. 1. Open in new tabDownload slide The velocity of Pi uptake as a function of substrate concentration. 33Pi uptake of the wild-type strain CC125 was performed as described in Methods. Cultures in the early logarithmic phase of growth were either transferred to TAP (+P, •) or TA (−P, ○) medium and allowed to continue growth for 24 h before measuring Pi uptake. The insets are double-reciprocal plots of the data, and theKm values estimated from these plots are 0.16 μm for the high-affinity component and 10 μm for the low-affinity component. chl, Chlorophyll. Isolation of Mutants Defective in Acclimation to P Limitation The results presented in Figure 1 and previous data showing that P-starved cells synthesize high levels of extracellular phosphatases (Lien and Knudsen, 1972; Loppes, 1976a, 1976b; Matagne et al., 1976;Patni et al., 1977; Quisel et al., 1996) suggested possible screens for isolating mutants unable to properly acclimate to P limitation. One screen was based on a preferential killing of cells that attain the capacity for elevated 32Pi transport upon exposure to P limitation. The second screen was based on a colorimetric assay to identify mutants with abnormal levels of extracellular phosphatase activity (see Methods). The latter screen is conceptually similar to a screen previously used to isolate mutants in C. reinhardtii that were unable to acclimate to S deprivation (Davies et al., 1994). From the first screen we isolated the mutantpsr1-1 (phosphorus-stressresponse), whereas the second screen yieldedpsr1-2 and psr2-1 (referred to as psr2 in the remainder of the text). Before physiological and biochemical analyses of the mutants, they were back-crossed four to five times to our wild-type strain (CC125) to ensure that the phenotypes analyzed were the result of a single lesion in a homogeneous genetic background. A qualitative analysis of extracellular phosphatase activity (based on the accumulation of a blue precipitate that forms around colonies capable of cleaving Pi from X-Pi) of the mutant and wild-type strains is presented in Figure2A. In wild-type cells and thepsr1-1 and psr1-2 mutants, little extracellular phosphatase activity was detected during growth on solid TAP medium (+P in Fig. 2A). In contrast, the psr2 and the psr1-2 psr2 double mutant accumulated relatively high levels of extracellular phosphatase activity during nutrient-replete growth. Upon P starvation (−P in Fig. 2A), wild-type cells (CC125) and psr2 accumulated high levels of extracellular phosphatase, as indicated by the intense blue halo that surrounds the cells. However, the psr1-1 andpsr1-2 mutants showed almost no extracellular phosphatase activity. The psr1-2 psr2 double mutant appeared to accumulate similar levels of extracellular phosphatase activity under both nutrient-replete and P-starvation conditions. Fig. 2. Open in new tabDownload slide Qualitative analysis of phosphatase activity secreted by wild-type cells, mutant strains, and vegetative diploids. Wild-type cells (wt),psr1-1,psr1-2, psr2, andpsr1-2 psr2 (A) and vegetative diploids of wild-type and the various mutant strains (B) were streaked onto TAP (+P) and TA (−P) solid media before spraying the plates with the colorimetric phosphatase substrate X-Pi. The plates were allowed to develop for 2 h before recording the results. The template shows the positions of the various mutants (A, right) and the different vegetative diploids (B, right) on the plates. Fig. 2. Open in new tabDownload slide Qualitative analysis of phosphatase activity secreted by wild-type cells, mutant strains, and vegetative diploids. Wild-type cells (wt),psr1-1,psr1-2, psr2, andpsr1-2 psr2 (A) and vegetative diploids of wild-type and the various mutant strains (B) were streaked onto TAP (+P) and TA (−P) solid media before spraying the plates with the colorimetric phosphatase substrate X-Pi. The plates were allowed to develop for 2 h before recording the results. The template shows the positions of the various mutants (A, right) and the different vegetative diploids (B, right) on the plates. Genetic Characterization of the Mutants Since the psr1-1 andpsr1-2 strains were both unable to secrete active phosphatase during P limitation, we constructed vegetative diploids to determine if the lesions in the strains were allelic. Vegetative diploids of wild-type cells and the individual mutants were also constructed to determine if the mutations were dominant or recessive. The phenotype of a vegetative diploid of psr1-1and psr1-2 was essentially identical, with respect to phosphatase activity, to that of the individual mutants; almost no phosphatase activity was observed when the diploid was starved for P (−P in Fig. 2B). In contrast, a vegetative diploid of the wild type and either psr1-1 orpsr1-2 had a phenotype that was identical to that of wild-type cells; P starvation led to high-level accumulation of alkaline phosphatase activity. Furthermore, a cross ofpsr1-1 to psr1-2 yielded no wild-type cells in over 400 progeny that were tested. These results clearly demonstrate that the lesions in psr1-1and psr1-2 are recessive and allelic. A vegetative diploid of psr2 and wild-type cells exhibited a phenotype that was dominant with respect to psr2. The diploid cells accumulated phosphatase activity in TAP medium even in the presence of a wild-type copy of PSR2. Like the originalpsr2 mutant, the diploid appeared to have elevated phosphatase activity when starved for P. A similar phenotype was observed for the vegetative diploid of psr1-1 psr2 (data not shown). Furthermore, the psr1 andpsr2 mutations segregated independently, indicating that they are nonallelic. Quantitative Analysis of Phosphatase Activity in the Mutant Strains A quantitative analysis of the accumulation of phosphatase activity in the medium of wild-type cells and the mutant strains during nutrient-replete and P-limited growth is presented in Figure3. Little phosphatase activity accumulated in cultures of wild-type cells grown on nutrient-replete medium (Fig. 3A). After the transfer of wild-type cells to medium devoid of P, a high level of phosphatase activity accumulated (Quisel et al., 1996); 24 to 48 h after the initiation of P deprivation, the alkaline phosphatase activity was at least 100-fold higher (Fig.3B) than in cells that were not starved. Thepsr1-1 and psr1-2 mutants showed no extracellular phosphatase activity when grown on nutrient-replete medium (Fig. 3A) or after exposure to medium devoid of P (Fig. 3B). The psr2 strain exhibited constitutive phosphatase activity when grown on nutrient-replete medium that was approximately 25% of the level observed after starvation (compare activity for psr2 in Fig. 3). Most of the extracellular phosphatase activity associated with psr2 grown in nutrient-replete medium remained in the supernatant when the cells were washed by centrifugation (data not shown). This explains why the amount of extracellular phosphatase activity was initially low after the transfer of psr2 cells to fresh medium (Fig. 3, 0 time point). The psr1-2 psr2 double mutant accumulated approximately the same amount of phosphatase activity as thepsr2 strain during growth on complete medium (Fig. 3A), however, this level did not change when the cells were transferred to medium lacking P (Fig. 3B). These results demonstrate that thepsr1 lesion prevents the induction of phosphatase activity in the psr2 strain but does not prevent constitutive, extracellular phosphatase accumulation. Fig. 3. Open in new tabDownload slide Quantitation of extracellular alkaline phosphatase activity in wild-type and mutant cultures. All strains were grown in TAP medium to early logarithmic phase, washed twice with TA medium, and then transferred to either TAP (A) or TA (B) liquid medium for 16, 25, 40, and 48 h before measuring the alkaline phosphatase activity. ◊, Wild type; ▪, psr2; ▵,psr1-1; ○,psr1-2; asterisks,psr1-2psr2. PNPP,p-Nitrophenylphosphate; chl, chlorophyll. Fig. 3. Open in new tabDownload slide Quantitation of extracellular alkaline phosphatase activity in wild-type and mutant cultures. All strains were grown in TAP medium to early logarithmic phase, washed twice with TA medium, and then transferred to either TAP (A) or TA (B) liquid medium for 16, 25, 40, and 48 h before measuring the alkaline phosphatase activity. ◊, Wild type; ▪, psr2; ▵,psr1-1; ○,psr1-2; asterisks,psr1-2psr2. PNPP,p-Nitrophenylphosphate; chl, chlorophyll. To determine if the aberrations in the psr1-1,psr1-2, and psr2 mutants were specific to P deprivation and not involved in global regulation of stress responses, we tested these strains for their ability to acclimate to S limitation. During S limitation C. reinhardtii synthesizes an extracellular arylsulfatase that can be assayed colorimetrically (Davies et al., 1994, 1996). None of the three mutant strains exhibited arylsulfatase activity before starving the cells for S, and they all accumulated normal levels of the extracellular arylsulfatase after S deprivation (D.D. Wykoff and A.R. Grossman, data not shown). Pi Transport Several tests were performed to determine if the lesions in the mutants resulted in aberrations in other responses observed in wild-type cells during P-limited growth. Initially, the mutant strains were tested for their ability to take up Pi after growth in TAP and TA media (Table I). Measurements of theVmax for Pi uptake for both the wild-type and mutant strains grown in complete medium varied from 3.11 to 6.43 pmol Pi μg−1 chlorophyll min−1. After 24 h of P starvation, the wild-type and psr2 mutant cells exhibited a 14-fold increase in the Vmax for Pi uptake. In contrast, P starvation of psr1-1 orpsr1-2 for 24 h resulted in little increase in the Vmax. The psr1-2 psr2 double mutant also exhibited little increase in theVmax for Pi uptake after starvation. Finally, wild-type cells grown in nutrient-replete medium and thepsr1 mutant strains maintained in either nutrient-replete or P-deficient medium exhibited both low- and high-affinity Pi transport (data not shown). Table I. Maximal rate of Pi uptake in wild-type and mutant strains Strain . Vmax Pi Uptake . Increase in Vmax . TAP medium 1d . TA medium 1d . pmol Pi μg−1chlorophyll min−1 -fold Wild type 6.43  (1.66) 91.1  (3.9) 14.2 psr2 6.42  (0.24) 90.8  (36) 14.1 psr1-1 3.11  (2.28) 5.38  (3.1) 1.7 psr1-2 5.32  (1.64) 7.71  (2.21) 1.5 psr1-2 psr2 4.12  (2.35) 4.50  (2.6) 1.1 Strain . Vmax Pi Uptake . Increase in Vmax . TAP medium 1d . TA medium 1d . pmol Pi μg−1chlorophyll min−1 -fold Wild type 6.43  (1.66) 91.1  (3.9) 14.2 psr2 6.42  (0.24) 90.8  (36) 14.1 psr1-1 3.11  (2.28) 5.38  (3.1) 1.7 psr1-2 5.32  (1.64) 7.71  (2.21) 1.5 psr1-2 psr2 4.12  (2.35) 4.50  (2.6) 1.1 The units below are the means of two independently grown cultures. TheVmax of Pi uptake was derived from at least three different Pi concentrations (i.e. 5, 10, and 15 μm). The values in parentheses are the difference from the mean in the same units. Open in new tab Table I. Maximal rate of Pi uptake in wild-type and mutant strains Strain . Vmax Pi Uptake . Increase in Vmax . TAP medium 1d . TA medium 1d . pmol Pi μg−1chlorophyll min−1 -fold Wild type 6.43  (1.66) 91.1  (3.9) 14.2 psr2 6.42  (0.24) 90.8  (36) 14.1 psr1-1 3.11  (2.28) 5.38  (3.1) 1.7 psr1-2 5.32  (1.64) 7.71  (2.21) 1.5 psr1-2 psr2 4.12  (2.35) 4.50  (2.6) 1.1 Strain . Vmax Pi Uptake . Increase in Vmax . TAP medium 1d . TA medium 1d . pmol Pi μg−1chlorophyll min−1 -fold Wild type 6.43  (1.66) 91.1  (3.9) 14.2 psr2 6.42  (0.24) 90.8  (36) 14.1 psr1-1 3.11  (2.28) 5.38  (3.1) 1.7 psr1-2 5.32  (1.64) 7.71  (2.21) 1.5 psr1-2 psr2 4.12  (2.35) 4.50  (2.6) 1.1 The units below are the means of two independently grown cultures. TheVmax of Pi uptake was derived from at least three different Pi concentrations (i.e. 5, 10, and 15 μm). The values in parentheses are the difference from the mean in the same units. Open in new tab Periplasmic Proteins Profiles of periplasmic polypeptides from wild-type cells,psr1-1, psr2 and the double mutantpsr1-1 psr2 grown in both complete medium and medium devoid of P are shown in Figure 4. For wild-type cells a periplasmic polypeptide of approximately 190 kD (marked by a filled arrow) accumulated as the cells grew in medium devoid of P (lanes 2 and 3). This polypeptide was previously shown to correspond to the major, derepressible extracellular phosphatase (Quisel et al., 1996). Cultures of the psr1-1mutant did not accumulate the 190-kD polypeptide upon P starvation (compare lanes 4 and 5). Similar results were observed for thepsr1-2 strain (data not shown). Hence, the 190-kD phosphatase is not synthesized, not exported, or rapidly degraded in the psr1 strains. In the psr2 mutant the 190-kD polypeptide accumulated only in the growth medium when the cells were starved for P (compare lanes 6 and 7). This suggests that the phosphatase activity that is constitutive in the psr2 strain is not a consequence of abnormal expression of the gene encoding the 190-kD species. Consistent with this interpretation is the finding that the constitutive phosphatase activity that accumulated inpsr2 cultures in nutrient-replete medium was independent of Ca2+ (data not shown), whereas the 190-kD phosphatase requires Ca2+ for activity (Quisel et al., 1996). Furthermore, the psr1-1 psr2 double mutant did not accumulate the 190-kD polypeptide upon starvation for P (compare Fig. 4, lanes 8 and 9), which is consistent with the measurements of phosphatase activity in this strain (Fig. 3). Fig. 4. Open in new tabDownload slide Profiles of periplasmic polypeptides from wild-type (wt) and the mutant strains after transfer to TAP (lanes 2, 4, 6, and 8) or TA (lanes 3, 5, 7, and 9) medium for 48 h. The samples (3 μg per lane) loaded in the different lanes are from wild type (lanes 2 and 3), psr1-1 (lanes 4 and 5), psr2 (lanes 6 and 7), andpsr1-1 psr2 (lanes 8 and 9). Lane M, Benchmark Mr markers (GIBCO-BRL). The positions of the two alkaline phosphatases are marked with arrows (black arrow for the 190-kD species and white arrow for the 70-kD species), and a cluster of other prominent polypeptides that accumulate in the medium in response to P limitation is marked with an asterisk. The polypeptides between 10 and 30 kD in the psr1strains were observed consistently throughout three independent protein isolations. Fig. 4. Open in new tabDownload slide Profiles of periplasmic polypeptides from wild-type (wt) and the mutant strains after transfer to TAP (lanes 2, 4, 6, and 8) or TA (lanes 3, 5, 7, and 9) medium for 48 h. The samples (3 μg per lane) loaded in the different lanes are from wild type (lanes 2 and 3), psr1-1 (lanes 4 and 5), psr2 (lanes 6 and 7), andpsr1-1 psr2 (lanes 8 and 9). Lane M, Benchmark Mr markers (GIBCO-BRL). The positions of the two alkaline phosphatases are marked with arrows (black arrow for the 190-kD species and white arrow for the 70-kD species), and a cluster of other prominent polypeptides that accumulate in the medium in response to P limitation is marked with an asterisk. The polypeptides between 10 and 30 kD in the psr1strains were observed consistently throughout three independent protein isolations. Quisel et al. (1996) reported the accumulation of a second extracellular alkaline phosphatase that accumulated during P limitation and migrates with an apparent molecular mass of 70 kD; the polypeptide marked with a white arrow may represent that phosphatase (Fig. 4). This polypeptide appeared to be absent in the psr1-1strain (lanes 4 and 5) but, like the 190-kD species, accumulated inpsr2 upon P deprivation (lanes 6 and 7). In thepsr1-1 psr2 double mutant a polypeptide that migrated at a position that was slightly higher than the 70-kD phosphatase accumulated in the culture medium. It is unlikely that this species is the 70-kD phosphatase, although we cannot rule out that possibility. Wild-type cells also synthesized a cluster of extracellular polypeptides with molecular masses ranging from 50 to 60 kD during P-limited growth (marked by an asterisk in Fig.4). The functions of these polypeptides, which accumulated normally inpsr2 but not in psr1-1 or thepsr1-1 psr2, are not known. Finally, low-molecular-mass polypeptides observed (in three different periplasmic protein preparations) in the medium frompsr1-1 and psr1-1 psr2cultures were not apparent in extracellular protein preparations from wild-type cells. These polypeptides may arise from increased proteolysis, aberrant processing of extracellular proteins, or increased leakage of cytoplasmic proteins in the mutant strains. Growth and Photosynthetic O2 Evolution The psr1-1 and psr1-2strains did not grow to the same extent as wild-type cells or thepsr2 mutant when exposed to conditions of P deprivation (Fig. 5). Wild-type cells and thepsr2 mutant doubled three to four times after they were placed in medium devoid of P. The psr1-1 mutant doubled only once, whereas the psr1-2 mutant doubled between one and two times after being placed in medium devoid of P. Growth characteristics of the psr1-2 psr2double mutant were similar to those of psr1-2. Fig. 5. Open in new tabDownload slide Increase in cell density after the transfer of CC125 (wt), psr1-1,psr1-2, psr2, andpsr1-2 psr2 to medium devoid of P. All of the cultures were grown to a density of 2 to 4 × 106cells mL−1, washed twice with TA medium, and adjusted to a final density of 5 × 105 cells mL−1 in TA medium. The increase in cell density was determined at 12, 24, 36, and 48 h after the transfer. The data presented here are from one experiment, but identical trends were observed in two additional experiments. Fig. 5. Open in new tabDownload slide Increase in cell density after the transfer of CC125 (wt), psr1-1,psr1-2, psr2, andpsr1-2 psr2 to medium devoid of P. All of the cultures were grown to a density of 2 to 4 × 106cells mL−1, washed twice with TA medium, and adjusted to a final density of 5 × 105 cells mL−1 in TA medium. The increase in cell density was determined at 12, 24, 36, and 48 h after the transfer. The data presented here are from one experiment, but identical trends were observed in two additional experiments. Recently, it was shown that starvation for either P or S leads to a decline in photosynthetic electron transport activity (Wykoff et al., 1998). Within 4 d of P starvation and 1 d of S starvation, O2 evolution declined by approximately 75%. This decrease reflects damage to PSII and the generation of PSII QB-nonreducing centers. Furthermore, a mutant abnormal for many responses to S deprivation dies much more rapidly than wild-type cells during S stress. This death is light dependent and appears to reflect an inability of the mutant to down-regulate photosynthetic electron transport (Davies et al., 1996). Wild-type cells and the psr2 mutant showed a similar decrease in photosynthetic O2 evolution during P deprivation. O2 evolution declined more rapidly in the psr1-1, psr1-2 (data not shown), and psr1-2 psr2 mutants than in wild-type cells (Table II), even though the viability of all of the mutant strains during P deprivation was similar to that of wild-type cells. The rapid decline in both photosynthesis and growth in the mutants suggests that they may more rapidly experience starvation when P is removed from the medium. Table II. Photosynthetic rate of wild-type and mutant strains Strain . Rate of O2 Evolution Relative to Wild-Type Unstarved Cells . TAP medium . TA medium 1d . TA medium 2d . Wild type 100%  (5.9%) 89.7%  (5.5%) 59.8%  (9.0%) psr2 97.5%  (5.0%) 77.9%  (3.8%) 59.3%  (5.0%) psr1-1 115%  (10.3%) 36.8%  (6.7%) 26.5%  (11.3%) psr1-2 psr2 112%  (10.0%) 43.6%  (7.9%) 37.3%  (3.9%) Strain . Rate of O2 Evolution Relative to Wild-Type Unstarved Cells . TAP medium . TA medium 1d . TA medium 2d . Wild type 100%  (5.9%) 89.7%  (5.5%) 59.8%  (9.0%) psr2 97.5%  (5.0%) 77.9%  (3.8%) 59.3%  (5.0%) psr1-1 115%  (10.3%) 36.8%  (6.7%) 26.5%  (11.3%) psr1-2 psr2 112%  (10.0%) 43.6%  (7.9%) 37.3%  (3.9%) The means of three independently grown cultures are indicated below with the se in parentheses. The wild-type rate of O2 evolution (100%) was 204 μmol O2mg−1 chlorophyll h−1. Open in new tab Table II. Photosynthetic rate of wild-type and mutant strains Strain . Rate of O2 Evolution Relative to Wild-Type Unstarved Cells . TAP medium . TA medium 1d . TA medium 2d . Wild type 100%  (5.9%) 89.7%  (5.5%) 59.8%  (9.0%) psr2 97.5%  (5.0%) 77.9%  (3.8%) 59.3%  (5.0%) psr1-1 115%  (10.3%) 36.8%  (6.7%) 26.5%  (11.3%) psr1-2 psr2 112%  (10.0%) 43.6%  (7.9%) 37.3%  (3.9%) Strain . Rate of O2 Evolution Relative to Wild-Type Unstarved Cells . TAP medium . TA medium 1d . TA medium 2d . Wild type 100%  (5.9%) 89.7%  (5.5%) 59.8%  (9.0%) psr2 97.5%  (5.0%) 77.9%  (3.8%) 59.3%  (5.0%) psr1-1 115%  (10.3%) 36.8%  (6.7%) 26.5%  (11.3%) psr1-2 psr2 112%  (10.0%) 43.6%  (7.9%) 37.3%  (3.9%) The means of three independently grown cultures are indicated below with the se in parentheses. The wild-type rate of O2 evolution (100%) was 204 μmol O2mg−1 chlorophyll h−1. Open in new tab DISCUSSION Little is known about the ways in which photosynthetic eukaryotes perceive and respond to P limitation. Generally, when organisms are starved for P, they synthesize both phosphatases and RNases that help them scavenge Pi from external and internal pools. Vascular plants may also increase their root-to-shoot ratio, allowing for more effective mining of Pi from the soil (Lynch, 1995), associate with mycorrhizae, which would facilitate Pi uptake (Smith and Read, 1997), and secrete organic acids, which helps mobilize stores of bound Pi in the soil (Marschner, 1995). In C. reinhardtii there are two major extracellular phosphatases that accumulate in response to Pi limitation (Quisel et al., 1996). The most abundant of these phosphatases has an apparent molecular mass of approximately 190 kD and its activity is Ca2+ dependent. At pH 9.5 this phosphatase is responsible for between 90% and 95% of the extracellular phosphatase activity in wild-type cells that are starved for P; the pH optimum for this phosphatase is 9.5 with very low activity below pH 7.0. A second extracellular phosphatase, which accounts for most of the remaining activity, has a molecular mass of approximately 70 kD and its activity is independent of Ca2+. Some mutants of C. reinhardtii with impaired phosphatase activity have been isolated (Loppes, 1978; Bachir et al., 1996), but they have not been extensively characterized. Pi uptake in C. reinhardtii is also influenced by the P status of the medium. There appear to be two different kinetic components associated with the transport of Pi into cells grown under P-replete conditions. This suggests that at least two different Pi transport systems are present in C. reinhardtii. One of these systems has a much higher affinity for Pi than the other (0.1–0.3 μm compared with approximately 10 μm). When wild-type cells are starved for P, there is an over 10-fold increase in the rate of Pi transport and only the high-affinity system is detected. Increased Pi uptake also occurs in vascular plants when Pi levels in the environment are low. The kinetics of Pi uptake by vascular plants is still controversial; most studies suggest the presence of multiple transport systems (Ullrich-Eberius et al., 1984; McPharlin and Bieleski, 1987; Nandi et al., 1987; Furihata et al., 1992), whereas others have argued for one transport system (Drew et al., 1984;Lefebvre et al., 1990; Shimogawara and Usuda, 1995). In the majority of studies there appears to be a constitutively expressed low-affinity Pi transport system and a second, high-affinity system that is derepressed during Pi-limited growth, although more than two systems may exist (Nandi et al., 1987). The high-affinity Pi transport system in plants has a Km of 3 to 7 μm, whereas the low-affinity system has aKm of 50 to 330 μm(Schachtman et al., 1998). Recently, a number of genes encoding Pi transport systems have been cloned from vascular plants (Muchhal et al., 1996; Kai et al., 1997; Leggewie et al., 1997; Mitsukawa et al., 1997; Smith et al., 1997; Daram et al., 1998; Liu et al., 1998; Okumura et al., 1998). There appear to be both differences and similarities between Pi transport in vascular plants and C. reinhardtii. First, theKm values for the low- and high-affinity transporters in C. reinhardtii are 1 order of magnitude lower than the Km values for the corresponding transporters of vascular plants. Furthermore, whereasC. reinhardtii and at least some vascular plants (Drew et al., 1984; Schmidt et al., 1992; Shimogawara and Usuda, 1995) have high-affinity Pi transport when grown in P-replete medium, the level of induction of the high-affinity transporter during P starvation is higher for C. reinhardtii (more than 10-fold) than for vascular plants (2- to 5-fold). The detection of high-affinity Pi transport in nutrient-replete C. reinhardtii cultures suggests constitutive synthesis of this transport system. It is not known whether the increase in high-affinity Pi transport that accompanies P limitation is a consequence of increased synthesis of the constitutive system or induction of a second high-affinity Pi transporter. It is also possible that under the optimal growth conditions being used, the cells have the capacity for more rapid intracellular utilization of Pi than can be supplied by the low-affinity Pi transport system. Therefore, the cells would experience Pi limitation and high-affinity transport would be partially induced. Like wild-type cells, the psr1-1 andpsr1-2 mutants have both low- and high-affinity Pi transport during nutrient-replete growth (data not shown). These results suggest that the high-affinity Pi transport activity observed in unstarved, wild-type cells is not regulated by the Psr1 polypeptide. We have used two different approaches to isolate mutants that are unable to properly acclimate to P deprivation. One approach involved a suicide selection procedure using 32Pi. Cells that synthesize elevated levels of the high-affinity Pi transport system during P starvation would rapidly incorporate radiolabeled Pi into nucleic acids and phospholipids, which would result in lethality; mutants unable to synthesize the high-affinity transport system in response to P starvation would survive longer periods of exposure to the radioisotope. The second approach exploited the finding that P-starved cells secrete extracellular phosphatases that are readily detected by spraying the colonies with the chromogenic phosphatase substrate X-Pi. Colonies that cannot synthesize extracellular phosphatases during P starvation do not develop a blue ”halo” (e.g.psr1), whereas colonies that constitutively produce high levels of extracellular phosphatase develop a blue halo when grown in nutrient-replete medium (e.g. psr2). Two mutants with a similar phenotype, psr1-1 andpsr1-2, were isolated by the different screens described above. These mutants were defective in the synthesis of extracellular phosphatases and were unable to increase the rate of Pi transport upon exposure to P limitation. Whereas thepsr1-1 allele appears to be null for both activities, the psr1-2 allele accumulates a small amount (less than 5% relative to wild-type cells) of extracellular phosphatase activity upon P starvation (Fig. 3B, 48 h). Furthermore, both strains failed to accumulate periplasmic polypeptides specifically associated with P-limited growth (Fig. 4). The finding that the phenotype of a psr1-1 psr1-2vegetative diploid was essentially identical to that of each of the haploid strains demonstrated that psr1-1 andpsr1-2 were alleles of the same gene. Additional characterizations of the mutants demonstrated a decline in photosynthetic activity and growth, after transfer to medium devoid of P, that was more rapid than in wild-type cells; the extent of growth was slightly more for psr1-2 than forpsr1-1 (again showing a slight difference in the phenotype of the two mutant alleles). The kinetics of the decline in growth and photosynthetic O2 evolution suggest that the psr1 mutants are more sensitive to P depletion than wild-type cells. This phenotype may result from the inability of these strains to access low levels of external Pi and/or to mobilize internal Pi stores. With a decreased ability to scavenge Pi, these mutants would more rapidly down-regulate metabolic processes such as photosynthetic O2 evolution, and growth would rapidly stop. Furthermore, whereas the decrease in photosynthetic activity in thepsr1 mutants occurs more rapidly than in wild-type cells, the modes by which photosynthetic electron transport is down regulated appear to be similar to that of wild-type cells (D.D. Wykoff and A.R. Grossman, data not shown). There is a decrease in linear electron transport with the major site of inhibition being PSII. The inhibition results from decreased photochemical efficiency and the accumulation of reaction centers that can perform a charge separation but that have an extremely slow rate of electron transfer between QA and QB. These PSII, QB-nonreducing centers were previously shown to be major components in the down-regulation of photosynthetic activity in wild-type cells during both P and S deprivation (Wykoff et al., 1998). Finally, whereas the mutant cells stop growing more rapidly than wild-type cells upon elimination of P from the medium, they survive long periods of P limitation, just like wild-type cells. These results suggest that the “general responses” to nutrient deprivation (Davies et al., 1996), which lead to the cessation of cell division and decreased photosynthetic activity and allow for extended survival during nutrient limitation, can still occur in the psr1strains. Based on the phenotype of the psr1 mutants, the lesions in these strains are likely to be in a regulatory gene that is needed to activate the specific but not the general responses to P deprivation. The PSR1 gene product may be directly involved in sensing the P status of the environment, or a component of the signal transduction chain that transmits the P deprivation to the transcriptional machinery of the cell. The cloning and characterization of the PSR1 gene (data not shown) suggests that Psr1 may be a transcription factor. Recently, we have also identified several mutants of C. reinhardtii that are abnormal in their responses to S limitation (Davies et al., 1996, 1999). One such mutant has been designated sac1. This mutant, unlike thepsr1 strain, is unable to control both the specific and general responses; this strain can neither regulate the synthesis of arylsulfatase (specific response) nor down-regulate photosynthetic electron transport (general response) during S starvation and, as a consequence, dies rapidly upon the imposition of S deprivation. In contrast to the psr1 mutants, the psr2 mutant shows constitutive extracellular phosphatase activity and has a dominant phenotype. 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J Cell Biochem 51 1993 47 54 Google Scholar Crossref Search ADS PubMed WorldCat 60 Wykoff DD Davies JP Melis A Grossman AR The regulation of photosynthetic electron transport during nutrient deprivation in Chlamydomonas reinhardtii. Plant Physiol 117 1998 129 139 Google Scholar Crossref Search ADS PubMed WorldCat 61 Yildiz FH Davies JP Grossman AR Characterization of a high-affinity sulfate transport system that accumulates during sulfur-limited growth of C. reinhardtii. Plant Physiol 104 1994 981 987 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the Japan-U.S. Cooperative Science Program from Japan Society of Plant Physiologists (JSPS) and the National Science Foundation (no. INT 9513 133 to H.U. and A.R.G.), the Research for the Future Program (no. JSPS-RFTF97R16001) from JSPS (to H.U.), the Asahi Glass Foundation (to K.S.), the Ministry of Education, Science, Sports and Culture, Japan (to K.S.), and the U.S. Department of Agriculture (grant no. 9302076 to A.R.G.). This is Carnegie Institution of Washington publication no. 1412. * Corresponding author; e-mail [email protected]; fax 1–650–325–6857. 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)

Tanaka, Ryouichi; Oster, Ulrike; Kruse, Elisabeth; Rüdiger, Wolfhart; Grimm, Bernhard

doi: 10.1104/pp.120.3.695pmid: 10398704

Abstract The enzyme geranylgeranyl reductase (CHL P) catalyzes the reduction of geranylgeranyl diphosphate to phytyl diphosphate. We identified a tobacco (Nicotiana tabacum) cDNA sequence encoding a 52-kD precursor protein homologous to the Arabidopsis and bacterial CHL P. The effects of deficient CHL P activity on chlorophyll (Chl) and tocopherol contents were studied in transgenic plants expressing antisense CHL P RNA. Transformants with gradually reduced Chl P expression showed a delayed growth rate and a pale or variegated phenotype. Transformants grown in high (500 μmol m−2 s−1; HL) and low (70 μmol photon m−2 s−1; LL) light displayed a similar degree of reduced tocopherol content during leaf development, although growth of wild-type plants in HL conditions led to up to a 2-fold increase in tocopherol content. The total Chl content was more rapidly reduced during HL than LL conditions. Up to 58% of the Chl content was esterified with geranylgeraniol instead of phytol under LL conditions. Our results indicate that CHL P provides phytol for both tocopherol and Chl synthesis. The transformants are a valuable model with which to investigate the adaptation of plants with modified tocopherol levels against deleterious environmental conditions. The main constituents of the photosynthetic apparatus are Chls, carotenoids, the plastid-encoded apoproteins of the core complex of the reaction centers, and the nuclear-encoded light-harvesting Chl-binding proteins of the antenna complexes. Control of pigment metabolism and expression of pigment-binding proteins ensure a synchronous synthesis of all components in stoichiometric amounts to prevent pigment or protein degradation or photooxidative deterioration. Chl consists of two moieties, chlorophyllide and phytol, which are formed from the precursor molecules 5-aminolevulinate and isopentenyl diphosphate, respectively, in the two different pathways of tetrapyrrole and isoprenoid biosynthesis. Most of the previous investigations of Chl biosynthesis have emphasized the tetrapyrrolic pathway, which is entirely located in plastids and converts Glu to Chl. Most of the genes involved in tetrapyrrole biosynthesis have been characterized previously (Chadwick and Ackrill, 1994; von Wettstein et al., 1995; Porra, 1997; Rüdiger, 1997; Grimm, 1998). The branched isoprenoid pathway is rather complex and comprises enzymatic steps in at least two compartments. The cytosolic isoprenoid-synthesizing pathway proceeds from acetyl-CoA via mevalonate in the plant cytoplasm, leading, for example, to sterol compounds. Incorporation studies of labeled early precursors indicated a mevalonate-independent pathway in plastids for phytol biosynthesis, for which 1-deoxy-xylulose-5-P is an intermediate (Rohmer et al., 1993;Lichtenthaler et al., 1997). The exact localization of the pathways for carotenoids and other end products was hampered to a certain extent because isopentenyl diphosphate, which is the common intermediate of both pathways, is transferred through the plastid envelope (Kreuz and Kleinig, 1984; Gray, 1987). Four molecules of isopentenyl diphosphate are subsequently joined to form the C20-intermediate GGPP, which is then allocated to the synthesis of various end products such as carotenoids, quinones, Chl, or tocopherol. The hydrogenation of GGPP is catalyzed by a CHL P (EC 1.3.1.-). The enzyme Chl synthase links tetrapyrrole and isoprenoid metabolism by esterifying chlorophyllide with GGPP or PhyPP. In etiolated plants Chl synthase esterifies chlorophyllide preferentially with GGPP to form ChlaGG. The recombinant Chl synthase that is encoded in the G4 gene of Arabidopsis (Gaubier et al., 1995) and overexpressed in Escherichia coli also gives preference to GGPP relative to PhyPP (Oster and Rüdiger, 1997). The NADPH-dependent hydrogenation to the phytol chain of Chl is observed after prenylation of chlorophyllide (Schoch et al., 1977; Benz et al., 1980). Conversely, Chl synthase assayed from green plants favors PhyPP for Chla synthesis rather than GGPP (Soll et al., 1983;Rüdiger, 1987). PhyPP is the preferential substrate of two overexpressed proteins, the Chl synthase derived from thechlG gene of Synechocystis and the bacteriochlorophyll synthase encoded by the Rhodobacter bchGgene (Oster et al., 1997). PhyPP is also an obligatory precursor for the synthesis of tocopherol (Soll and Schultz, 1981), and is directed into the tocopherol-synthesizing pathway by condensation with homogentisate derived from the shikimate pathway. Several methylations and a cyclization step of the quinol intermediate leads to α-tocopherol, the major form of the vitamin E fraction (Soll, 1987). Tocopherol prevents the Chl-photosensitized oxidation of thylakoid components, especially when plants are subjected to environmental stress, mainly by quenching activated singlet oxygen or by scavenging oxygen radicals (Fryer, 1992). A bchP gene has been previously detected as part of the 46-kb photosynthetic gene cluster of Rhodobacter capsulatus(Marrs, 1981; Zsebo and Hearst, 1984) encoding almost all constituents required for photosynthesis. Insertional mutagenesis of thebchP gene resulted in a mutant that produced bacteriochlorophyll esterified with geranylgeraniol instead of phytol (Bollivar et al., 1994). The bchP gene encodes CHL P. Electron transfer and energy transfer from the light-harvesting complex to the reaction center were apparently not affected in the R. capsulatus mutant. Nevertheless, the growth rate under photosynthetic conditions was severely reduced in the mutant (Bollivar et al., 1994). The authors suggested a reduced stability of the pigment-protein complexes, if bacteriochlorophyll is esterified with geranylgeraniol. Other reasons for a reduced growth rate could not be ruled out. Nevertheless, all normal functions were restored when the homologous chlP gene from Synechocystis sp. PCC 6803 was expressed in the Rhodobacter bchP-deficient mutant (Addlesee et al., 1996). We isolated the tobacco Chl P sequence encoding CHL P and examined its metabolic function as well as its expression in transgenic tobacco plants. CHL P is located at the branch point toward Chl and tocopherol. Our aim was to improve the understanding of the molecular and physiological effects of reduced synthesis of PhyPP on the controlled distribution of substrate for Chl and tocopherol synthesis under different light conditions. MATERIALS AND METHODS Plant Growth and Harvest Wild-type (Nicotiana tabacum var Samsun NN) and transgenic tobacco plants were cultivated in growth chambers with a 16-h light/8-h dark cycle at 25°C. The light intensities were 500 μmol m−2 s−1 (HL) or 70 μmol m−2 s−1 (LL). Primary transformants were used for analysis. Leaves were harvested from 8-week-old plants grown in growth chambers, frozen in liquid nitrogen, and analyzed immediately, freeze-dried, or stored at −80°C before analysis. Isolation and Analysis of a cDNA Clone Encoding CHL P Protein A tobacco cDNA library (Nicotiana tabacum SR1, Stratagene) in a Lambda ZAP II cDNA library was screened using the expressed sequence tag clone 4D9T7P (accession no. T04791) from Arabidopsis obtained from the Arabidopsis Biological Research Center (Ohio State University, Columbus). The sequence is homologous to that of the bchP gene of Rhodobacter capsulatus (Young et al., 1989) and of Arabidopsis Chl P. The cDNA sequences were analyzed with the PCGENE program (Intelligenetics, Mountain View, CA). Alignment of peptide sequences was done with the Clustal W program. Construction of a Chl P Antisense Gene and Plant Transformation The full-length cDNA sequence was cut out of the vector with the restriction enzymes KpnI and XbaI and ligated into the same restriction sites of the plant binary vector BinAR (Höfgen and Willmitzer, 1992), a pBIB derivative containing the cauliflower mosaic virus 35S promoter. The transformation of tobacco leaf discs was mediated byAgrobacterium tumefaciens, as described by Horsch et al.(1985). The insertion of copies of the transgene was confirmed by kanamycin resistance of regenerated explants and by genomic Southern hybridization or PCR amplification using a Chl P-specific probe and oligonucleotide primers. RNA Analysis Total RNA was isolated by the acid-phenol extraction method (Chomczinski and Sacchi, 1987). Aliquots of 10 μg of RNA were blotted onto nylon membranes (Hybond N, Amersham) and hybridized with [32P]dCTP using the nick-translation method. Hybridized filters were exposed to radiographic film (Kodak) or to phosphor-imaging plates (Fuji Film, Tokyo) and analyzed (STORM 960, Molecular Dynamics, Krefeld, Germany). Equal loading of samples was controlled by rehybidizing the RNA filter with a cDNA probe for 18S rRNA (Thompson et al., 1994). Antiserum Preparation and Western Analysis Two oligonucleotide primers were designed to amplify the coding sequences of Chl P: Csyn1, 5′cgc cat ggg ccg caa tct tcg tgt tgc ggt 3′; Csyn 2, 5′gca gat ctg tcc att tcc ctt ctt agt gca 3′. The PCR fragment was cloned into the NcoI and BglI sites of the expression vector pQE 60 (Qiagen, Hilden, Germany). The subcloned tobacco Chl P sequence continues behind the initiation codon of the expression plasmid with the nucleotide at position 148 of the cDNA clone (accession no. AJ007789). Overexpression of recombinant CHL P protein was performed in Escherichia coli XL-1 Blue or SG 13009 (Stratagene). The protein was purified by metal chelate affinity chromatography and used for immunization of rabbits. Antiserum was collected after triple injection of the antigen. Plant material (100 mg) was ground under liquid nitrogen, suspended in 1 mL of solubilization buffer (56 mmNa2CO3, 56 mmDTT, 2% SDS, 12% Suc, and 2 mm EDTA), and denatured for 15 min at 70°C. The soluble protein fraction was quantified and 10-μg protein aliquots were analyzed by western blot with the anti-CHL P antiserum using an immunoblotting kit (ECL, Amersham). Analysis of Chl Leaf tissue (100 mg fresh weight) was pulverized in liquid nitrogen with a mortar and pestle and extracted twice with 400 μL of acetone:water (3:1, v/v). The liquid phases were collected in a 2-mL test tube. The extraction was repeated three times until the pellet was colorless. The combined acetone extracts were cleared by centrifugation and mixed with 500 μL of n-hexane. The hexane phase was separated and the acetone phase again extracted with the same volume ofn-hexane. A small aliquot of the combined hexane phases was washed with water until it was free of acetone.A660 and A642were determined after a suitable dilution (normally 1:10) in a spectrophotometer (model 8451A, Hewlett-Packard) and used for calculation of the total contents of Chl a and baccording to the method of French (1960). The rest of the combined hexane phases was acidified with three drops of concentrated HCl. The color of the solution changed from green to brown, indicating the formation of pheophytin from Chl. The hexane phase was then repeatedly washed with water until the aqueous phase reached a pH of ≥5.5. The last traces of water were removed from the hexane phase by freezing at −20°C. Hexane was then removed by evaporation and the precipitate was dissolved in 500 μL of acetone. Twenty microliters of the 1:10 diluted samples was applied for HPLC analysis (model 480, Gynkothek, Ramsey, NJ). Pigments were separated on a column (4 × 250 mm) filled with RP18 (Gromsil 120, Grom Analytic, Herrenberg, Germany) at 1.2 mL/min with the following gradient consisting of 60% acetone (solvent A) and 100% acetone (solvent B): 75% A/25% B for 2 min, followed by 45% A/55% B for 2 min, 30% A/70% B for 11 min, and 100% B for 8 min. Pigments were detected by a UV-visible light detector (model SP5V, Shimadzu, Columbia, MD) at 410 nm and by a fluorescence detector (model RF551, Shimadzu) at 665 nmem and 425 nmex. The peak areas indicated the ratios of ChlaGG to Chl aPhyand Chl bGG to Chl bPhy. Analysis of Tocopherol Tocopherol was measured independently in both laboratories by two methods. In the first, leaf material was pulverized in liquid nitrogen and lyophilized. A precisely weighted 5-mg aliquot of the dry powder was extracted four times in a precooled mortar with 350 μL each of dioxane:n-hexane (1:1, v/v), and the combined supernatants were cleared by centrifugation and evaporated. The residue was dissolved in 100 μL of dioxane:n-hexane (3:97, v/v). For each analysis, 20 μL of this solution was analyzed by HPLC (model 300C, Gynkothek) using a column (4.6 × 250 mm) filled with Nucleosil 50 (5 μm) at a flow rate of 1.5 mL/min with dioxane:n-hexane (3:97, v/v). Tocopherol was quantified at 295 nmex and 325 nmem using a fluorescence detector (model RF1001, Shimadzu). In the second method, tocopherol was extracted from frozen leaf powder with acetone containing 10 μm KOH and separated on a HPLC system equipped with a C18 column (3.9 × 150 mm, Nova-Pak, Waters) with a gradient of solvent A (30% methanol, and 10% 0.1 m ammonium acetate, pH 5.2) and solvent B (100% methanol) as follows: a linear gradient from 6% A/94% B at 0 min to 1% A/99% B at 10 min until 23 min with the same ratio of solutions A and B. Standards for α, β/γ, and δ tocopherol and for α, β/γ, and δ tocotrienol were purchased from Merck (Darmstadt, Germany) and used to quantify and qualify the tocopherol forms eluted by our HPLC program. Analysis of Carotenoids Carotenoids were extracted from 100 mg of leaf powder with acetone containing 10 μm KOH and separated by HPLC with a linear gradient beginning with 100% eluate A (86.7% acetonitrile, 9.6% methanol, and 3.6% 0.1 m Tris-HCl, pH 8.0) to 100% eluate B at 15 min (80% methanol and 20% hexane) on a 5-μm column (Lichrosphere 100 RP-18, Merck, Darmstadt, Germany) and monitored by a photodiode array detector (model 996, Waters) at a flow rate of 1 mL/min. Carotenoid standards were purchased from Roth (Karlsruhe, Germany). RESULTS Tobacco Chl P cDNA Sequence Encoding CHL P and the Expression of Recombinant Protein A full-length cDNA clone encoding CHL P was identified. The cDNA sequence is composed of 1510 nucleotides without the poly(A+) chain and is deposited in the database under accession no. AJ007789. Nucleotides 1 through 1392 encode a 52-kD protein consisting of 464 amino acid residues. The deduced peptide sequence shows similarity to the Mesembryanthemum crystallinum CHL P (accession no. AF069318) (82% identical amino acid residues), to the Arabidopsis CHL P sequence (accession no.Y14044) (81% identical amino acid residues) (Keller et al., 1998), to the Synechocystis sp PCC 6803 ChlP (accession no. Q55087) (67%) (Addlesee et al., 1996), and to the R. capsulatuscounterpart (34%) (Zsebo and Hearst, 1984; Bollivar et al., 1994). In contrast to the bacterial peptides, the three plant sequences of CHL P contain amino-terminal extensions that resemble plastid transit sequences. The overall similarity among the five sequences was 29.1%. The coding region of a truncated CHL P peptide (amino acid residues 50–464) was fused in frame behind the initiation codon into anE. coli expression vector. The beginning of the open reading frame codes for the Met-Gly-Arg-Asn-Leu of the recombinant CHL P. The recombinant protein was insoluble in aqueous solution. The His-tagged protein was dissolved in 8 m urea and was purified by metal chelate affinity chromatography as recommended by manufacturer's instructions. The purified protein of an approximate molecular mass of 47 kD was injected into rabbits for immunization. Phenotypical Differences between Transgenic Plants Expressing Antisense-Oriented Chl P Genes and Control Plants The full-length Chl P-cDNA sequence was inserted in inverse orientation between the cauliflower mosaic virus 35Spromoter and the 3′ termination sequence of the octopine synthase gene of the binary plant vector BinAR. The antisense gene construct was introduced into the tobacco genome by A. tumefaciens-mediated transformation. Approximately 100 different transgenic lines were generated and analyzed for the insertion of copies of the transgene. The transformants were generally characterized by a growth rate slower than or similar to that of control plants and a gradually reduced Chl content compared with the wild type. Most of the transgenic lines displayed a uniform low pigmentation, and some of them had yellow areas along the leaf veins or different variegation patterns (Fig.1). The green pigmentation was generally more reduced in older than in younger leaves of the same transformant. The transformants grown in the greenhouse or in the growth chamber under controlled conditions did not show any necrotic leaf lesions that could be generated by accumulating photosensitizing tetrapyrrole intermediates. Fig. 1. Open in new tabDownload slide Primary transformant 6 with Chl Pantisense genes (PL24–6) and a wild-type tobacco plant (SNN). Plants were grown for 8 weeks under greenhouse conditions in an average light intensity of 300 μmol m−2 s−1. Fig. 1. Open in new tabDownload slide Primary transformant 6 with Chl Pantisense genes (PL24–6) and a wild-type tobacco plant (SNN). Plants were grown for 8 weeks under greenhouse conditions in an average light intensity of 300 μmol m−2 s−1. Metabolic Effects of Reduced Expression of Chl P in Transgenic Plants At first, all transformants were phenomenologically and biochemically evaluated to select a few transgenic lines for further detailed analysis. The primary data were obtained with plants grown in the greenhouse under ambient conditions. The plants were exposed to diurnal changes in temperature (15°C–24°C) and light intensity (up to 800 μm m−2s−1) before analysis. TableI illustrates the gradual variation of the inhibitory effects on Chl and tocopherol contents by Chl P antisense RNA expression among a representative set of transformants (lines 6, 10, 20, 21, 24, and 47) and wild-type plants. These lines represent a broad range of gradually increasing transgenic phenotypes; they retained these characteristics in the course of the studies over almost 2 years and were characterized by a progressive reduction in growth rate and pigmentation. Their progenies of the T1 and T2 generation showed the same typical deficiency symptoms or were more severely impaired in growth and pigmentation. Table I. Comparative analysis of Chl P antisense RNA-expressing primary transformants and control plants Plant . ChlGG . Total Chl . ChlPhy . Tocopherol . % % WT WT 0 100 100 100  6 53 32 15 14 10 56 40 18 19 20 7 59 55 52 21 4 90 87 99 24 30 99 70 91 47 25 37 28 18 Plant . ChlGG . Total Chl . ChlPhy . Tocopherol . % % WT WT 0 100 100 100  6 53 32 15 14 10 56 40 18 19 20 7 59 55 52 21 4 90 87 99 24 30 99 70 91 47 25 37 28 18 The fifth leaf of each plant was harvested and analyzed. Total Chl, ChlGG, ChlPhy, and tocopherol were determined by HPLC and are given as percentage of the wild-type (WT) levels (except for ChlGG, which is given as percentage of the total Chl in the respective transformant). Open in new tab Table I. Comparative analysis of Chl P antisense RNA-expressing primary transformants and control plants Plant . ChlGG . Total Chl . ChlPhy . Tocopherol . % % WT WT 0 100 100 100  6 53 32 15 14 10 56 40 18 19 20 7 59 55 52 21 4 90 87 99 24 30 99 70 91 47 25 37 28 18 Plant . ChlGG . Total Chl . ChlPhy . Tocopherol . % % WT WT 0 100 100 100  6 53 32 15 14 10 56 40 18 19 20 7 59 55 52 21 4 90 87 99 24 30 99 70 91 47 25 37 28 18 The fifth leaf of each plant was harvested and analyzed. Total Chl, ChlGG, ChlPhy, and tocopherol were determined by HPLC and are given as percentage of the wild-type (WT) levels (except for ChlGG, which is given as percentage of the total Chl in the respective transformant). Open in new tab The pale-green leaves of some transgenic plants correlated with the reduced total Chl content. The severely affected lines 6, 47, and 10 accumulated only 32%, 37%, and 40%, respectively, of the wild-type Chl content. The transformants also contained a lower tocopherol content. Lines 6, 47, and 10 also showed the strongest deficiency of tocopherol (14%, 18%, and 19% of the control values, respectively). As indicated, other transgenic lines displayed only minor decreases in Chl and tocopherol contents. Tocotrienol could be an expected product if GGPP were also used for the synthesis of tocopherol. The HPLC elution program allowed the separation of tocopherol and tocotrienol derivatives, but tocotrienol was not detected in extracts of the transformants. The esterification of Chl with various alcohols was analyzed by HPLC. To avoid allomerization and oxidation, we removed the central Mg2+ and analyzed the corresponding pheophytins. A typical chromatogram is shown in Figure2. In the control samples, pheophytinsa and b (Phe αPhy and PhebPhy) were detectable as the only Chl derivatives. In the samples of the transformants, three new peaks were identified by co-chromatography with authentic samples: the new compound Phe a′Phy and Phe α and Pheb esterified with geranylgeraniol (PheaGG and PhebGG). The identity of the pigments was confirmed by the absorption and the fluorescence emission spectra of the single peaks and by comparison with the authentic compounds. We could not detect in any of the transformants the pigments containing intermediate alcohols between geranylgeraniol and phytol. This observation of the steady-state content in transgenic plants differs from the results of the in vitro assays with recombinant Arabidopsis CHL P (Keller et al., 1998). The small peaks between those of the identified pigments do not indicate Chl derivatives, but they exhibited only diffuse fluorescence and absorption spectra (Fig. 2). Fig. 2. Open in new tabDownload slide HPLC chromatogram of pheophytins derived from Chls of tobacco leaves. A, Wild-type tobacco; B, tobacco transformed with the Chl P gene in antisense orientation. Labeled peaks are: Phe aPhy (1); Phea′Phy (1a); PhebPhy (1a); PheaGG (3); and PhebGG (4). Fig. 2. Open in new tabDownload slide HPLC chromatogram of pheophytins derived from Chls of tobacco leaves. A, Wild-type tobacco; B, tobacco transformed with the Chl P gene in antisense orientation. Labeled peaks are: Phe aPhy (1); Phea′Phy (1a); PhebPhy (1a); PheaGG (3); and PhebGG (4). The percentages of Chl aGG, ChlbGG, Chl aPhy, and Chl bPhy were determined by quantification of the peak areas. As expected, the wild-type plants contained only phytylated Chl. Whereas line 21, which had wild-type-like characteristics, contained only 4% ChlGG, the strongly reduced amounts of Chl of lines 10 and 6 consisted of 56% and 53% ChlGG, respectively (Table I). If the phytylated portion of Chl was related to the reduced amount of total Chl in the transformants, the values for ChlPhy and α-tocopherol were about equal (TableI), indicating that antisense inhibition of Chl P expression affects the pathways leading to ChlPhy and to α-tocopherol to the same extent. These initial data of six representative transformants reflect the variation of inhibitory effects by Chl P antisense gene expression in transgenic tobacco plants. The percentage of ChlGG is apparently a suitable indicator for the extent of CHL P inhibition and was used to substantiate the primary examinations of the antisense inhibition upon different physiological conditions. We grew plants at a constant temperature of 22°C in light intensities of 70 and 500 μmm−2 s−1 and designated the conditions as LL and HL growth, respectively. All plants exhibited the distinct properties of either LL- and HL-exposed plants apart from their characteristic transgenic phenotype. The LL-grown plants displayed smaller and paler leaves as well as extended internodia. HL-grown plants looked more vigorous and robust and contained more spacious leaves. To test a possible developmental regulation, we analyzed leaves 3 to 10 (counting from the top). Wild-type leaves contained exclusively ChlPhy. In contrast, ChlGG was detected in all transformants under conditions of irradiation and leaf development (TableII). In leaves 1 to 3, ChlGG was not detectable in any transformant. The percentage of ChlGG increased generally from leaves 5 to 7 (Table II) and did not increase further in leaves 8 to 13 (data not shown). Therefore, the data indicate an increasing effect of the antisense inhibition with leaf development. Furthermore, the percentage of ChlGG was generally higher in transgenic plants grown under LL conditions than in the same plants cultivated under HL conditions. Under HL conditions only the three transformants, 6, 10, and 47, showed significant accumulation of ChlGG in the analyzed leaves (Table II). Table II. Percentage of ChlGG in leaves 5 and 7 of tobacco plants containing Chl P antisense genes and control plants Plant . HL-5 . HL-7 . LL-5 . LL-7 . Chl a . Chl b . Chl a . Chlb . Chl a . Chl b . Chla . Chl b . % WT 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0  6 9.4 4.1 31.6 23.0 49.9 47.7 54.9 54.1 10 0.0 0.0 12.0 7.8 39.3 36.3 57.0 57.9 20 0.5 0.5 0.7 0.7 7.2 7.0 13.0 12.7 21 0.0 0.0 1.6 2.0 2.8 2.7 9.4 9.5 24 0.0 0.0 0.0 0.0 29.9 32.3 29.1 35.0 47 9.1 5.7 11.1 7.5 25.2 22.0 22.5 29.4 Plant . HL-5 . HL-7 . LL-5 . LL-7 . Chl a . Chl b . Chl a . Chlb . Chl a . Chl b . Chla . Chl b . % WT 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0  6 9.4 4.1 31.6 23.0 49.9 47.7 54.9 54.1 10 0.0 0.0 12.0 7.8 39.3 36.3 57.0 57.9 20 0.5 0.5 0.7 0.7 7.2 7.0 13.0 12.7 21 0.0 0.0 1.6 2.0 2.8 2.7 9.4 9.5 24 0.0 0.0 0.0 0.0 29.9 32.3 29.1 35.0 47 9.1 5.7 11.1 7.5 25.2 22.0 22.5 29.4 Plants were grown under HL and LL conditions, and leaf extracts were analyzed by HPLC and spectrometry. Presented are the percentages of ChlaGG and Chl bGG based on total Chl in the respective transgenic or control plant (ChlGG + ChlPhy = 100%). Open in new tab Table II. Percentage of ChlGG in leaves 5 and 7 of tobacco plants containing Chl P antisense genes and control plants Plant . HL-5 . HL-7 . LL-5 . LL-7 . Chl a . Chl b . Chl a . Chlb . Chl a . Chl b . Chla . Chl b . % WT 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0  6 9.4 4.1 31.6 23.0 49.9 47.7 54.9 54.1 10 0.0 0.0 12.0 7.8 39.3 36.3 57.0 57.9 20 0.5 0.5 0.7 0.7 7.2 7.0 13.0 12.7 21 0.0 0.0 1.6 2.0 2.8 2.7 9.4 9.5 24 0.0 0.0 0.0 0.0 29.9 32.3 29.1 35.0 47 9.1 5.7 11.1 7.5 25.2 22.0 22.5 29.4 Plant . HL-5 . HL-7 . LL-5 . LL-7 . Chl a . Chl b . Chl a . Chlb . Chl a . Chl b . Chla . Chl b . % WT 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0  6 9.4 4.1 31.6 23.0 49.9 47.7 54.9 54.1 10 0.0 0.0 12.0 7.8 39.3 36.3 57.0 57.9 20 0.5 0.5 0.7 0.7 7.2 7.0 13.0 12.7 21 0.0 0.0 1.6 2.0 2.8 2.7 9.4 9.5 24 0.0 0.0 0.0 0.0 29.9 32.3 29.1 35.0 47 9.1 5.7 11.1 7.5 25.2 22.0 22.5 29.4 Plants were grown under HL and LL conditions, and leaf extracts were analyzed by HPLC and spectrometry. Presented are the percentages of ChlaGG and Chl bGG based on total Chl in the respective transgenic or control plant (ChlGG + ChlPhy = 100%). Open in new tab No significant difference was found between the portion of ChlaGG and Chl bGGin transformants grown under LL conditions. In HL-cultivated transgenic plants 6, 10, and 47, the degree of Chl aGGwas higher than that of Chl bGG (Table II). As expected, the total Chl a/b ratio was higher in HL-grown wild-type and transgenic plants than in LL-grown plants. In LL-grown plants the Chl a/b ratio did not differ between the geranylgeranylated and the phytylated Chl molecules. Under HL conditions transformants contained a lower degree of ChlbGG than ChlaGG, resulting in a higher Chla/b ratio for the geranylgeranylated molecules (5.2 in leaf 5 of transformant 47, 6.3 in leaf 5 of transformant 6, and of 7.5 in leaf 7 of transformant 10). Chl P Expression in CHL P-Deficient Transgenic Plants under Two Different Light Intensities We extended our comparative biochemical and genetic analysis of transgenic plants to dependency on light intensity to substantiate the primary examinations of the effects of reduced CHL P contents in leaf 3, 5, and 7 of the transgenic plants. We chose lines 6, 10, and 20 for further analysis because they represented transgenic plants with significant macroscopic modifications. The Chl P RNA levels remained constant and the CHL P protein levels did not vary much during the development of leaves 3, 5, and 7 of control plants under LL and HL conditions. In the transformants the steady-state Chl Ptranscript and CHL P protein levels progressively decreased with age (Fig. 3). Steady-state Chl Ptranscript levels were generally lower in LL-grown transformants than in those grown under HL conditions and were more rapidly diminished during leaf development. The CHL P protein levels correlated with RNA content during leaf development under identical conditions. The more rapidly descending levels of the CHL P protein under LL conditions might reflect its lower stability because of fewer reduction equivalents and less substrate in these transformants. Fig. 3. Open in new tabDownload slide Expression of CHL P in transgenic (6, 10, and 20) and wild-type tobacco plants (SNN) was determined by northern and western analysis of HL- and LL-grown plants. Total RNA was isolated from leaves 3, 5, and 7 (counted from the top of each plant). Ten micrograms of RNA was loaded per lane, separated on a 1% formaldehyde-agarose gel, and hybridized to a tobacco Chl P cDNA probe. A cDNA probe for 18S rRNA was subsequently hybridized to the RNA on the same filter. Equal amounts of protein extracted from leaf 3, 5, and 7 were loaded on a SDS-polyacrylamide gel. After transfer to a nitrocellulose filter, immunodetection was performed with antiserum raised against recombinant CHL P. Fig. 3. Open in new tabDownload slide Expression of CHL P in transgenic (6, 10, and 20) and wild-type tobacco plants (SNN) was determined by northern and western analysis of HL- and LL-grown plants. Total RNA was isolated from leaves 3, 5, and 7 (counted from the top of each plant). Ten micrograms of RNA was loaded per lane, separated on a 1% formaldehyde-agarose gel, and hybridized to a tobacco Chl P cDNA probe. A cDNA probe for 18S rRNA was subsequently hybridized to the RNA on the same filter. Equal amounts of protein extracted from leaf 3, 5, and 7 were loaded on a SDS-polyacrylamide gel. After transfer to a nitrocellulose filter, immunodetection was performed with antiserum raised against recombinant CHL P. Chl, Tocopherol, and Carotenoid Contents Are Developmentally Controlled in Transgenic Plants under LL and HL Conditions Chl accumulation during leaf development was determined in leaves 3 to 10 of HL- and LL-grown transgenic and control plants (Fig.4). Since there was no significant difference between Chl a and b under most physiological conditions (see Table II), we continued to determine only total Chl. Eight-week-old control plants reached the maximum Chl level upon LL exposure in leaf 5 (1.36 nmol/g fresh weight) and under HL exposure in the same leaf (1.85 nmol/g fresh weight). The transient increase of total Chl contents up to leaf 5 and its subsequent decrease in the transgenic lines followed the wild-type pattern. Under LL conditions the Chl content was lower only in analyzed leaves of transgenic lines 6 and 10 (maximal 30% and 40% Chl, respectively, less than the wild-type leaves). All HL-grown transformants contained significantly reduced Chl contents compared with wild-type values, particularly if older leaves were compared (e.g. 70%, 48%, and 26% less Chl in leaf 10 of transformants 6, 10, and 20, respectively, than in wild-type plants; Fig. 4). Fig. 4. Open in new tabDownload slide Contents of total Chl (C and D) and α-tocopherol (A and B) in extracts of leaf 3, 5, 7, and 10 from LL-grown (A and C) and HL-grown (B and D) transgenic tobacco plants with reduced CHL P expression (lines 6, 10, and 20) and control plants (SNN). White bars, Leaf 3; right-hatched bars, leaf 5; left-hatched bars, leaf 7; and cross-hatched bars, leaf 10. fw, Fresh weight. Fig. 4. Open in new tabDownload slide Contents of total Chl (C and D) and α-tocopherol (A and B) in extracts of leaf 3, 5, 7, and 10 from LL-grown (A and C) and HL-grown (B and D) transgenic tobacco plants with reduced CHL P expression (lines 6, 10, and 20) and control plants (SNN). White bars, Leaf 3; right-hatched bars, leaf 5; left-hatched bars, leaf 7; and cross-hatched bars, leaf 10. fw, Fresh weight. Tocopherol content was analyzed in HL- and LL-grown control and transgenic plants (Fig. 4). Evaluation of the tocopherol content in premature leaves of control plant and of lines 20, 10, and 6 revealed that under HL conditions the plants contained 110%, 83%, 139%, and 47%, respectively, more tocopherol than under LL growth conditions. In wild-type plants the tocopherol content increased maximally up to leaf 7 under both light intensities, remained constant during plant development, and decreased progressively during senescence (data not shown). Tocopherol content was progressively reduced from the young to the older leaves in lines 6 and 10 and only slightly lower in line 20 after a similar profile during leaf development, compared with wild-type plants. The degree of lowered tocopherol levels was different in each transformant, but the relative decrease in tocopherol contents was very similar in each respective line under conditions of either LL or HL growth (Fig. 4). In leaf 10 of lines 6 and 10, tocopherol yielded approximately 20% of the wild-type contents under both growth conditions and in line 20 the yield was approximately 75%. Comparison of the decline in Chl and tocopherol in the transgenic relative to the wild-type plants during leaf development revealed that the relative tocopherol contents were diminished to a similar extent under both light intensities and that the relative amount of total Chl was more intensively reduced with age in HL-grown than in LL-grown plants of the same transgenic lines. The total carotenoid contents during leaf development of LL-and HL-grown transformants paralleled approximately the total Chl content. Accumulation of the carotenoid species neoxanthin, violaxanthin, lutein, and β-carotene is depicted in TableIII. The amounts of zeaxanthin and antheraxanthin could not be determined by our HPLC program because ChlaGG was eluted from the HPLC column simultaneously with zeaxanthin and Chl bGGwith antheraxanthin. All carotenoid species accumulated to lower amounts in the transgenic plants grown under HL or LL conditions than in control plants. However, the reduction of lutein and β-carotene content was more pronounced in the transgenic plants relative to wild-type plants than the reduction of violaxanthin and neoxanthin. Comparing the contents of lutein and β-carotene during developmental growth of the transgenic lines, it is noticeable that both carotenoids were more diminished under HL than under LL growth conditions. Table III. Quantitative analysis of carotenoids in leaves 3, 5, 7, and 10 of control and selected transgenic lines 6, 10, and 20 Carotenoid . LL . HL . SNN . 6 . 10 . 20 . SNN . 6 . 10 . 20 . μg g−1 fresh wt Neoxanthin  Leaf    3 18.5  (0.64) 13.2  (1.38) 12.8  (1.55) 15.7  (0.43) 20.6  (1.03) 18.3  (1.83) 19.0  (3.99) 13.4  (1.48)    5 28.5  (0.98) 25.4  (0.20) 22.2  (0.65) 28.1  (3.81) 29.4  (1.48) 27.0  (2.10) 26.7  (1.79) 20.8  (1.87)    7 28.0  (3.83) 28.1  (0.20) 28.2  (1.24) 26.0  (0.65) 27.8  (3.23) 27.4  (2.23) 33.8  (5.17) 21.7  (2.43)   10 17.3  (1.47) 26.5  (0.59) 20.6  (3.24) 16.9  (2.03) 25.2  (1.79) 15.7  (4.72) 31.3  (8.06) 18.4  (2.85) Violaxanthin  Leaf    3 28.3  (0.85) 21.0  (1.38) 20.4  (0.69) 24.9  (0.64) 33.0  (1.47) 30.7  (5.50) 32.5  (6.86) 25.5  (2.23)    5 34.6  (1.57) 29.7  (0.59) 24.7  (0.65) 35.4  (4.98) 45.9  (1.03) 41.1  (1.31) 39.6  (4.26) 38.2  (4.10)    7 27.8  (3.43) 24.0  (1.18) 22.2  (1.22) 26.6  (1.29) 38.5  (4.25) 30.1  (4.19) 35.6  (6.55) 35.2  (4.15)   10 15.9  (1.77) 17.3  (0.39) 13.5  (1.01) 15.8  (2.29) 29.4  (3.40) 13.2  (3.28) 22.4  (7.08) 21.9  (5.55) Lutein  Leaf    3 89.1  (6.77) 59.8  (7.42) 59.6  (5.26) 77.4  (1.24) 124.8  (4.75) 87.9  (6.11) 109.8  (24.86) 80.7  (9.22)    5 132.3  (8.30) 103.5  (3.93) 87.9  (4.17) 129.8  (20.06) 161.0  (7.24) 105.0  (16.16) 130.1  (12.97) 114.0  (11.54)    7 121.0  (20.73) 81.0  (7.21) 78.9  (2.52) 111.4  (2.13) 142.9  (20.11) 83.0  (13.68) 129.3  (26.90) 113.1  (13.80)   10 75.6  (5.40) 62.4  (3.06) 47.8  (7.43) 64.7  (7.43) 127.7  (9.84) 39.2  (7.42) 89.7  (29.04) 87.1  (9.10) β-Carotene  Leaf    3 52.7  (2.93) 38.5  (3.16) 32.5  (2.08) 49.0  (9.03) 76.7  (2.06) 48.0  (4.21) 60.3  (12.51) 51.2  (4.79)    5 69.2  (2.53) 51.9  (2.37) 46.5  (3.03) 71.8  (11.10) 93.7  (4.03) 53.3  (9.68) 68.7  (5.96) 70.0  (6.38)    7 62.4  (9.45) 32.8  (4.74) 31.5  (2.41) 59.1  (2.10) 83.9  (11.15) 35.8  (8.21) 59.1  (14.33) 66.2  (7.08)   10 36.8  (3.24) 18.3  (1.89) 15.7  (3.24) 31.4  (3.23) 72.8  (6.34) 11.9  (2.63) 30.5  (12.79) 46.1  (5.85) Carotenoid . LL . HL . SNN . 6 . 10 . 20 . SNN . 6 . 10 . 20 . μg g−1 fresh wt Neoxanthin  Leaf    3 18.5  (0.64) 13.2  (1.38) 12.8  (1.55) 15.7  (0.43) 20.6  (1.03) 18.3  (1.83) 19.0  (3.99) 13.4  (1.48)    5 28.5  (0.98) 25.4  (0.20) 22.2  (0.65) 28.1  (3.81) 29.4  (1.48) 27.0  (2.10) 26.7  (1.79) 20.8  (1.87)    7 28.0  (3.83) 28.1  (0.20) 28.2  (1.24) 26.0  (0.65) 27.8  (3.23) 27.4  (2.23) 33.8  (5.17) 21.7  (2.43)   10 17.3  (1.47) 26.5  (0.59) 20.6  (3.24) 16.9  (2.03) 25.2  (1.79) 15.7  (4.72) 31.3  (8.06) 18.4  (2.85) Violaxanthin  Leaf    3 28.3  (0.85) 21.0  (1.38) 20.4  (0.69) 24.9  (0.64) 33.0  (1.47) 30.7  (5.50) 32.5  (6.86) 25.5  (2.23)    5 34.6  (1.57) 29.7  (0.59) 24.7  (0.65) 35.4  (4.98) 45.9  (1.03) 41.1  (1.31) 39.6  (4.26) 38.2  (4.10)    7 27.8  (3.43) 24.0  (1.18) 22.2  (1.22) 26.6  (1.29) 38.5  (4.25) 30.1  (4.19) 35.6  (6.55) 35.2  (4.15)   10 15.9  (1.77) 17.3  (0.39) 13.5  (1.01) 15.8  (2.29) 29.4  (3.40) 13.2  (3.28) 22.4  (7.08) 21.9  (5.55) Lutein  Leaf    3 89.1  (6.77) 59.8  (7.42) 59.6  (5.26) 77.4  (1.24) 124.8  (4.75) 87.9  (6.11) 109.8  (24.86) 80.7  (9.22)    5 132.3  (8.30) 103.5  (3.93) 87.9  (4.17) 129.8  (20.06) 161.0  (7.24) 105.0  (16.16) 130.1  (12.97) 114.0  (11.54)    7 121.0  (20.73) 81.0  (7.21) 78.9  (2.52) 111.4  (2.13) 142.9  (20.11) 83.0  (13.68) 129.3  (26.90) 113.1  (13.80)   10 75.6  (5.40) 62.4  (3.06) 47.8  (7.43) 64.7  (7.43) 127.7  (9.84) 39.2  (7.42) 89.7  (29.04) 87.1  (9.10) β-Carotene  Leaf    3 52.7  (2.93) 38.5  (3.16) 32.5  (2.08) 49.0  (9.03) 76.7  (2.06) 48.0  (4.21) 60.3  (12.51) 51.2  (4.79)    5 69.2  (2.53) 51.9  (2.37) 46.5  (3.03) 71.8  (11.10) 93.7  (4.03) 53.3  (9.68) 68.7  (5.96) 70.0  (6.38)    7 62.4  (9.45) 32.8  (4.74) 31.5  (2.41) 59.1  (2.10) 83.9  (11.15) 35.8  (8.21) 59.1  (14.33) 66.2  (7.08)   10 36.8  (3.24) 18.3  (1.89) 15.7  (3.24) 31.4  (3.23) 72.8  (6.34) 11.9  (2.63) 30.5  (12.79) 46.1  (5.85) Extracts were prepared as for tocopherol determination and subjected to HPLC as described in Methods. Compounds were identified and quantified with the help of authentic standards. Values in parentheses are sd. Open in new tab Table III. Quantitative analysis of carotenoids in leaves 3, 5, 7, and 10 of control and selected transgenic lines 6, 10, and 20 Carotenoid . LL . HL . SNN . 6 . 10 . 20 . SNN . 6 . 10 . 20 . μg g−1 fresh wt Neoxanthin  Leaf    3 18.5  (0.64) 13.2  (1.38) 12.8  (1.55) 15.7  (0.43) 20.6  (1.03) 18.3  (1.83) 19.0  (3.99) 13.4  (1.48)    5 28.5  (0.98) 25.4  (0.20) 22.2  (0.65) 28.1  (3.81) 29.4  (1.48) 27.0  (2.10) 26.7  (1.79) 20.8  (1.87)    7 28.0  (3.83) 28.1  (0.20) 28.2  (1.24) 26.0  (0.65) 27.8  (3.23) 27.4  (2.23) 33.8  (5.17) 21.7  (2.43)   10 17.3  (1.47) 26.5  (0.59) 20.6  (3.24) 16.9  (2.03) 25.2  (1.79) 15.7  (4.72) 31.3  (8.06) 18.4  (2.85) Violaxanthin  Leaf    3 28.3  (0.85) 21.0  (1.38) 20.4  (0.69) 24.9  (0.64) 33.0  (1.47) 30.7  (5.50) 32.5  (6.86) 25.5  (2.23)    5 34.6  (1.57) 29.7  (0.59) 24.7  (0.65) 35.4  (4.98) 45.9  (1.03) 41.1  (1.31) 39.6  (4.26) 38.2  (4.10)    7 27.8  (3.43) 24.0  (1.18) 22.2  (1.22) 26.6  (1.29) 38.5  (4.25) 30.1  (4.19) 35.6  (6.55) 35.2  (4.15)   10 15.9  (1.77) 17.3  (0.39) 13.5  (1.01) 15.8  (2.29) 29.4  (3.40) 13.2  (3.28) 22.4  (7.08) 21.9  (5.55) Lutein  Leaf    3 89.1  (6.77) 59.8  (7.42) 59.6  (5.26) 77.4  (1.24) 124.8  (4.75) 87.9  (6.11) 109.8  (24.86) 80.7  (9.22)    5 132.3  (8.30) 103.5  (3.93) 87.9  (4.17) 129.8  (20.06) 161.0  (7.24) 105.0  (16.16) 130.1  (12.97) 114.0  (11.54)    7 121.0  (20.73) 81.0  (7.21) 78.9  (2.52) 111.4  (2.13) 142.9  (20.11) 83.0  (13.68) 129.3  (26.90) 113.1  (13.80)   10 75.6  (5.40) 62.4  (3.06) 47.8  (7.43) 64.7  (7.43) 127.7  (9.84) 39.2  (7.42) 89.7  (29.04) 87.1  (9.10) β-Carotene  Leaf    3 52.7  (2.93) 38.5  (3.16) 32.5  (2.08) 49.0  (9.03) 76.7  (2.06) 48.0  (4.21) 60.3  (12.51) 51.2  (4.79)    5 69.2  (2.53) 51.9  (2.37) 46.5  (3.03) 71.8  (11.10) 93.7  (4.03) 53.3  (9.68) 68.7  (5.96) 70.0  (6.38)    7 62.4  (9.45) 32.8  (4.74) 31.5  (2.41) 59.1  (2.10) 83.9  (11.15) 35.8  (8.21) 59.1  (14.33) 66.2  (7.08)   10 36.8  (3.24) 18.3  (1.89) 15.7  (3.24) 31.4  (3.23) 72.8  (6.34) 11.9  (2.63) 30.5  (12.79) 46.1  (5.85) Carotenoid . LL . HL . SNN . 6 . 10 . 20 . SNN . 6 . 10 . 20 . μg g−1 fresh wt Neoxanthin  Leaf    3 18.5  (0.64) 13.2  (1.38) 12.8  (1.55) 15.7  (0.43) 20.6  (1.03) 18.3  (1.83) 19.0  (3.99) 13.4  (1.48)    5 28.5  (0.98) 25.4  (0.20) 22.2  (0.65) 28.1  (3.81) 29.4  (1.48) 27.0  (2.10) 26.7  (1.79) 20.8  (1.87)    7 28.0  (3.83) 28.1  (0.20) 28.2  (1.24) 26.0  (0.65) 27.8  (3.23) 27.4  (2.23) 33.8  (5.17) 21.7  (2.43)   10 17.3  (1.47) 26.5  (0.59) 20.6  (3.24) 16.9  (2.03) 25.2  (1.79) 15.7  (4.72) 31.3  (8.06) 18.4  (2.85) Violaxanthin  Leaf    3 28.3  (0.85) 21.0  (1.38) 20.4  (0.69) 24.9  (0.64) 33.0  (1.47) 30.7  (5.50) 32.5  (6.86) 25.5  (2.23)    5 34.6  (1.57) 29.7  (0.59) 24.7  (0.65) 35.4  (4.98) 45.9  (1.03) 41.1  (1.31) 39.6  (4.26) 38.2  (4.10)    7 27.8  (3.43) 24.0  (1.18) 22.2  (1.22) 26.6  (1.29) 38.5  (4.25) 30.1  (4.19) 35.6  (6.55) 35.2  (4.15)   10 15.9  (1.77) 17.3  (0.39) 13.5  (1.01) 15.8  (2.29) 29.4  (3.40) 13.2  (3.28) 22.4  (7.08) 21.9  (5.55) Lutein  Leaf    3 89.1  (6.77) 59.8  (7.42) 59.6  (5.26) 77.4  (1.24) 124.8  (4.75) 87.9  (6.11) 109.8  (24.86) 80.7  (9.22)    5 132.3  (8.30) 103.5  (3.93) 87.9  (4.17) 129.8  (20.06) 161.0  (7.24) 105.0  (16.16) 130.1  (12.97) 114.0  (11.54)    7 121.0  (20.73) 81.0  (7.21) 78.9  (2.52) 111.4  (2.13) 142.9  (20.11) 83.0  (13.68) 129.3  (26.90) 113.1  (13.80)   10 75.6  (5.40) 62.4  (3.06) 47.8  (7.43) 64.7  (7.43) 127.7  (9.84) 39.2  (7.42) 89.7  (29.04) 87.1  (9.10) β-Carotene  Leaf    3 52.7  (2.93) 38.5  (3.16) 32.5  (2.08) 49.0  (9.03) 76.7  (2.06) 48.0  (4.21) 60.3  (12.51) 51.2  (4.79)    5 69.2  (2.53) 51.9  (2.37) 46.5  (3.03) 71.8  (11.10) 93.7  (4.03) 53.3  (9.68) 68.7  (5.96) 70.0  (6.38)    7 62.4  (9.45) 32.8  (4.74) 31.5  (2.41) 59.1  (2.10) 83.9  (11.15) 35.8  (8.21) 59.1  (14.33) 66.2  (7.08)   10 36.8  (3.24) 18.3  (1.89) 15.7  (3.24) 31.4  (3.23) 72.8  (6.34) 11.9  (2.63) 30.5  (12.79) 46.1  (5.85) Extracts were prepared as for tocopherol determination and subjected to HPLC as described in Methods. Compounds were identified and quantified with the help of authentic standards. Values in parentheses are sd. Open in new tab DISCUSSION The plastidal metabolite GGPP is an intermediate in several biosynthetic pathways. It can be channeled into the Chl pathway by reduction to PhyPP and final esterification with chlorophyllide or vice versa by the initial prenylation of chlorophyllide and a subsequent reduction of ChlGG (Fig.5). GGPP can also be directed into tocopherol synthesis when PhyPP is condensed with homogentisate. CHL P catalyzes the reduction of GGPP and ChlGG in vitro (Keller et al., 1998). It is not known if the enzyme accepts both substrates with the same specificity. Moreover, it remains to be seen if the same enzyme serves simultaneously the Chl- and the tocopherol-synthesizing pathways in planta. Fig. 5. Open in new tabDownload slide Scheme of the branched pathway starting from GGPP to α-tocopherol, γ-tocopherol, or ChlPhy. CHL P, Chl synthase, and an isoprenyl transferase are indicated. CHL P uses GGPP and ChlGG as substrates and directs PhyPP to the tocopherol- and the Chl-synthesizing pathway. Fig. 5. Open in new tabDownload slide Scheme of the branched pathway starting from GGPP to α-tocopherol, γ-tocopherol, or ChlPhy. CHL P, Chl synthase, and an isoprenyl transferase are indicated. CHL P uses GGPP and ChlGG as substrates and directs PhyPP to the tocopherol- and the Chl-synthesizing pathway. We identified the tobacco Chl P cDNA sequence and introduced a binary vector harboring an antisense Chl P gene into tobacco to reduce specifically the enzyme activity of CHL P. This transgenic approach not only addresses the question of whether CHL P functions for two pathways but also enables a prediction on the control mechanism that distributes the enzymatic product of CHL P toward Chl or tocopherol synthesis. The antisense inhibition of the Chl P expression led to an increasing proportion of ChlGG in the total amount of Chl and to lower tocopherol contents during leaf development of the analyzed transformants (Table II). The deficiency of tocopherol was found synchronously with the increasing amount of ChlGG (Table I). Therefore, the involvement of CHL P in the formation of Chl and tocopherol can be described as two equivalent functions. Our results obtained with the Chl Pantisense plants are in agreement with the assumption of hydrogenation of GGPP to PhyPP and of ChlGG to ChlPhy and with the second function of CHL P protein in contributing to the formation of tocopherol. The recombinant Arabidopsis CHL P protein expressed in E. coli was shown to be active in both the reduction of GGPP and of ChlGG (Keller et al., 1998). The R. capsulatus bchP insertion mutant fails in the hydrogenation step from geranylgeraniol to phytol, the esterifying alcohol of bacteriochlorophyll (Bollivar et al., 1994). Under increased light intensities, wild-type plants and the analyzed transformants accumulated more Chl (between approximately 35% in leaves of wild-type plants and 12% in those of line 6) and up to 2 times more tocopherol (Fig. 4). Since the wild-type contents of CHL P seemed to be similar under LL and HL conditions in leaves of the same age, it is suggested that CHL P expression does not normally limit the supply of precursor for Chl and tocopherol during plant development. However, as was apparent from the analysis of the transgenic lines, significant reduction of the amount of CHL P by antisense RNA synthesis affects the levels of both biomolecules in a light and developmentally dependent manner. It is remarkable that transformants with diminished amounts of CHL P not only contained reduced contents of phytylated Chl but also contained less total Chl than wild-type plants. It is assumed either that ChlGG is not associated as stably as ChlPhy in the photosynthetic pigment-binding proteins, resulting in a faster Chl breakdown (analogous to the assumption of labile bacteriochlorophyllGG-protein complexes; Bollivar et al., 1994), or that the reduced synthesis of isoprenoid and Chl precursors is indirectly caused by the antisense inhibition of CHL P expression. The percentage of phytylated Chl was generally higher in HL transformants than in the same LL-grown transgenic lines, which could be explained by several factors. First, if Chl synthase activity were higher under HL than under LL conditions, more PhyPP but also more accumulating GGPP could be channeled into the Chl-synthesizing pathway. ChlGG competes with GGPP for the residual hydrogenation activity of CHL P, leading to more ChlPhy. Second, an increasing pool of NADPH or other reducing biomolecules due to the stimulated photosynthesis in HL-exposed leaves could activate CHL P, which should lead to more ChlPhy and α-tocopherol. However, it seems more likely that the low ChlGG levels in HL-grown transgenic plants result from photooxidation of ChlGG so that ChlPhyremains preferentially. It is unknown whether HL conditions cause destabilization of pigments, especially of ChlGGin the Chl P antisense plants. Chl a and b were phytylated to the same extent in most transformants grown under LL conditions. Some transformants exposed to HL conditions showed more ChlaGG than ChlbGG, which is indicative for a preferential supply of Chl for the reaction center core complex under increasing light intensities. This would mean that Chl a exhibits a faster turnover than Chl b in HL-exposed transformants, most likely in the reaction center of the photosystems and to lesser extent in the antenna complexes. Approximately 2 times more tocopherol accumulated upon increased light intensity in wild-type and transgenic plants compared with those exposed to lower light intensities. It has been shown previously that HL exposure modulates the tocopherol content in photosynthetic membranes (Lichtenthaler, 1979). The antioxidant function of tocopherol is expected to scavenge photogenerated singlet oxygen or other organic radicals in photosynthetic membranes. Exposure to enhanced light intensities increases the risks of photodynamic damage. An increased requirement of tocopherol under HL conditions is most likely adjusted by stimulation of its synthesis in response to light intensities. The analysis of the transgenic lines with reduced CHL P expression did not provide evidence that HL exposure would lead to preferential channelling of PhyPP under limiting synthesis of this precursor into the tocopherol-synthesizing pathway. However, the consequences of insufficient levels of tocopherol became apparent in the transgenic plants with reduced CHL P content. The transformants grew slower and showed a bleached phenotype. Lower CHL P activity could result in accumulation of GGPP, which can also be directed into carotenoid synthesis. However, the analyzed carotenoid levels were lower in the transformants than in the wild-type plants and almost paralleled the reduction in Chl content. The assembly of ChlGG instead of ChlPhywith the pigment-binding proteins does not significantly modify the composition of carotenoids in the pigment-protein complexes. Our observation reflects the synchronized need for both pigment fractions and the pigment-binding proteins to stabilize the photosynthetic complexes and is consistent with observations of the parallel loss of pigments and pigment-binding proteins in mutants with deficiencies in Chl or carotenoid synthesis (Plumley and Schmidt, 1995). In conclusion, we demonstrate here, for the first time to our knowledge, that deficiency of CHL P simultaneously affects two pathways, leading to a decline in tocopherol and phytylated Chl contents and in the level of total Chl molecules. Lack of tocopherol could destabilize the thylakoid membrane and negatively influence the photosynthetic machinery. Analysis of the photoprotective functions of tocopherol and other antioxidants under various environmental conditions and for photosynthesis in the Chl P antisense plants is currently being performed. Transgenic plants with reduced tocopherol contents seem to be an appropriate model with which to investigate how the transformants prevent the deleterious photo effects and if and how they compensate for the tocopherol deficit. ACKNOWLEDGMENTS We thank P. Linow, E. Fraust, and I. Blos for excellent technical assistance. Abbreviations: Chl chlorophyll CHL P geranylgeranyl reductase GGPP geranylgeranyl diphosphate HL high light LL low light PhyPP phytyl diphosphate LITERATURE CITED 1 Addlesee HA Gibson LCD Jensen PE Hunter CN Cloning sequencing and functional assignment of the chlorophyll biosynthesis gene chlP of Synechocystis sp PCC 6803. FEBS Lett 389 1996 126 130 Google Scholar Crossref Search ADS PubMed WorldCat 2 Benz J Wolf C Rüdiger W Chlorophyll biosynthesis: hydrogenation of geranylgeraniol. 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SFB 184), Bonn, Germany. 2 Present address: The Institute of Low Temperature Science, Hokkaido University, N19 W8, Sapporo 060–0819, Japan. * Corresponding author; e-mail [email protected]; fax 49–39482–5139. 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)

Watt, Michelle; Evans, John R.

doi: 10.1104/pp.120.3.705pmid: 10398705

Abstract White lupin (Lupinus albus L.) was grown in hydroponic culture with 1 μm phosphorus to enable the development of proteoid roots to be observed in conjunction with organic acid exudation. Discrete regions of closely spaced, determinate secondary laterals (proteoid rootlets) emerged in near synchrony on the same plant. One day after reaching their final length (4 mm), citrate exudation occurred over a 3-d pulse. The rate of exudation varied diurnally, with maximal rates during the photoperiod. At the onset of citrate efflux, rootlets had exhausted their apical meristems and had differentiated root hairs and vascular tissues along their lengths. Neither in vitro phosphoenolpyruvate carboxylase nor citrate synthase activity was correlated with the rate of citrate exudation. We suggest that an unidentified transport process, presumably at the plasma membrane, regulates citrate efflux. Growth with elevated (700 μL L−1) atmospheric [CO2] promoted earlier onset of rootlet determinacy by 1 d, resulting in shorter rootlets and citrate export beginning 1 d earlier as a 2-d diurnal pulse. Citrate was the dominant organic acid exported, and neither the rate of exudation per unit length of root nor the composition of exudate was altered by atmospheric [CO2]. Proteoid roots develop on a range of plant species, including white lupin (Lupinus albus L.), adapted to environments with poorly available phosphorus (Gardner et al., 1981; Dinkelaker et al., 1995). First described in 1960 in a study of the Proteaceae (Purnell, 1960), proteoid roots have discrete clusters of closely spaced laterals (rootlets) along their lengths that greatly increase the surface area for nutrient uptake compared with the proteoid root axis (Lamont et al., 1984). These clusters of rootlets release exudates, notably organic acids that can solubilize phosphorus by chelating the metal ions that immobilize it (Gardner et al., 1983; Gerke et al., 1994). The organic acids exported from white lupin proteoid roots can include citrate, malate, succinate, and fumarate (Gardner et al., 1983;Dinkelaker et al., 1989; Johnson et al., 1994; Keerthisinghe et al., 1998) and can account for a substantial portion of the total plant carbon. For example, Dinkelaker et al. (1989) showed that citrate exported from white lupin growing in a calcareous soil was 23% of the plant dry weight at 13 weeks, and Johnson et al. (1996b) measured malate, succinate, and citrate totaling 12% of the plant dry weight at 3 weeks. The metabolism linked to the synthesis and efflux of organic acids from proteoid roots has been studied recently in white lupin (Johnson et al., 1994, 1996a, 1996b). Johnson et al. (1996a) showed that approximately 30% of the carbons released from the roots as malate or citrate were fixed within the proteoid roots by the enzyme PEPC (EC 4.1.1.31). The increased activity and expression of PEPC and malate dehydrogenase (EC 1.1.1.37) coincided with the start of organic acid efflux and was considered to be part of an altered metabolism within proteoid roots that was required for the synthesis of the exported organic acids (Johnson et al., 1996b). The morphology and anatomy of proteoid rootlets was reported previously for some members of the Proteaceae (for summary, see Dinkelaker et al., 1995; Skene et al., 1996, 1998a, 1998b). These studies showed that a cluster of rootlets starts as many meristematic primordia that subsequently mature into determinate rootlets with no apical meristem and root hairs around their tips. In Hakea obliqua, approximately 5 d is required for the rootlets to reach their final determinate length (Dell et al., 1980). To our knowledge, there are no published studies of the anatomy of white lupin proteoid rootlets, although macrographs of white lupin rootlets growing in calcareous soil show root hairs extending around their tips (Dinkelaker et al., 1989), suggesting that the rootlets also reach a determinate stage. There is evidence that the efflux of exudates, including organic acids, occurs when rootlets are young and that this efflux is transient (Dinkelaker et al., 1995; Neumann et al., 1995; Keerthisinghe et al., 1998). Keerthisinghe and co-workers (1998) collected exudates along a proteoid root axis of white lupin and found that most of the efflux occurred in the youngest portion of the root axis, where the rootlets were young, and that the in vitro activities of PEPC and malate dehydrogenase were not strictly correlated with citrate fluxes from the same portion of root. The transition from primordial tissue to fully differentiated, determinate root tissue during rootlet growth suggests that enzymatic changes associated with development could be confused with those associated with exudation and, as suggested by Dinkelaker et al. (1995), elucidation of the mechanisms related to exudation requires a detailed time-course study of rootlets of different ages. To our knowledge, there have been no such studies linking anatomical changes with biochemical changes and the efflux of organic acids on the proteoid rootlets of any species. In the present study, we applied the root-incubation chamber used byKeerthisinghe et al. (1998) to study proteoid rootlets of white lupin from the time of emergence through d 8. Our first objective was to resolve the developmental and metabolic stages associated with efflux of organic acids by performing a detailed time-course study of rootlet anatomy and biochemistry for plants grown with low phosphorus and ambient atmospheric [CO2]. Increases in atmospheric [CO2] can alter root growth and turnover (for review, see Rogers et al., 1994; for example, see Fitter et al., 1996) and can increase the amount of carbon-containing compounds exported to the rhizosphere (Paterson et al., 1997). However, there have been few studies measuring the quantity and quality of exudates on a per-root basis under ambient and elevated [CO2] (Sadowsky and Schortemeyer, 1997), particularly those exudates that function in nutrient acquisition, such as organic acids. If such exudates are increased under elevated atmospheric CO2, they may confer an advantage to those species with these processes (Gifford et al., 1996; DeLucia et al., 1997). Because white lupin exports a large amount of carbon as citrate from proteoid lateral roots with defined, determinate development, they are an ideal system with which to study the effects of atmospheric [CO2] on root and exudate processes. Our second objective was to investigate the effect of atmospheric [CO2] on proteoid root growth, development, and efflux. MATERIALS AND METHODS Plant Growth White lupin (Lupinus albus L. cv Kiev mutant) was grown in ambient (350 μL L−1) or elevated (700 μL L−1) atmospheric CO2 in climate-controlled growth cabinets with a 12-h photoperiod, 600 μmol m−2 s−1 light at leaf level, 70% RH, and 15°C/22°C night/day temperatures. Seeds were germinated in damp sand and at d 6 (2 d after cotyledon emergence) were transferred to black, 22-L hydroponics tanks. Four seedlings per tank were supported by removable foam discs that fit into the lid. Each tank contained a solution of 1 μmKH2PO4, 0.25 mm CaCl2, 0.7 mm KNO3, 0.25 mm MgSO4, 11 μmH3BO3, 2 μm MnSO4, 0.35 μm ZnSO4, 0.2 μm CuSO4, and 6 μm ferric EDTA and was adjusted daily to pH 6.0 (Keerthisinghe et al., 1998). Each day, phosphate levels were assayed using Malachite green dye and replenished to 1 μm (Irving and McLaughlin, 1990; Keerthisinghe et al., 1998). The complete solution in the tanks was changed weekly or biweekly. Preliminary experiments indicated that nitrate had been depleted by less than 20% between solution changes. The nutrient solution was aerated continuously from rings with small perforations supported at the bottom of the tanks. Scoring Proteoid Root Development The working definition of a proteoid root was a primary lateral root with defined clusters of more than 10 secondary lateral roots (proteoid rootlets) per centimeter (Figs.1 and 3; Johnson et al., 1996b). The emergence of these clusters of proteoid rootlets was scored daily for 3 to 4 weeks by removing a plant from the hydroponics tank, spreading the root system in a shallow dish of water, and counting clusters that had recently emerged from the root cortex (rootlets 0.5–1.0 mm long). Fig. 1. Open in new tabDownload slide Root system of an 18-d-old white lupin plant germinated on sand for 6 d and then transferred to nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. All of the primary basal laterals have become proteoid roots and the first cluster of secondary laterals (proteoid rootlets) has fully emerged. The basal laterals are longer and thicker than the thinner, shorter, acropetal primary laterals. Scale bar = 1 cm. Fig. 1. Open in new tabDownload slide Root system of an 18-d-old white lupin plant germinated on sand for 6 d and then transferred to nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. All of the primary basal laterals have become proteoid roots and the first cluster of secondary laterals (proteoid rootlets) has fully emerged. The basal laterals are longer and thicker than the thinner, shorter, acropetal primary laterals. Scale bar = 1 cm. Fig. 3. Open in new tabDownload slide Clusters of rootlets along proteoid roots of white lupin grown in nutrient solution with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. A cluster is defined as a length of root with more than 10 rootlets per centimeter that have emerged in near synchrony. A, Proteoid root of a 32-d-old plant. Clusters of emerged rootlets are numbered. Studies relating development with citrate efflux were done on the fourth cluster of rootlets. Arrow indicates the sixth cluster. B, Developmental series of a proteoid root cluster. Rootlets emerged in near synchrony on d 1 and developed to similar final lengths of approximately 4 mm on d 4.   Fig. 3. Open in new tabDownload slide Clusters of rootlets along proteoid roots of white lupin grown in nutrient solution with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. A cluster is defined as a length of root with more than 10 rootlets per centimeter that have emerged in near synchrony. A, Proteoid root of a 32-d-old plant. Clusters of emerged rootlets are numbered. Studies relating development with citrate efflux were done on the fourth cluster of rootlets. Arrow indicates the sixth cluster. B, Developmental series of a proteoid root cluster. Rootlets emerged in near synchrony on d 1 and developed to similar final lengths of approximately 4 mm on d 4.   In Situ Collection of Root Exudates Exudates were collected from incubated clusters of proteoid rootlets attached to the plant in their growing environments (Ryan et al., 1993; Keerthisinghe et al., 1998, see figure 1 therein). Plants were transferred to tanks partially filled with nutrient solution and the proteoid roots were supported on trays in the tanks. A Perspex resin incubation ring, 2 cm in diameter and 1.2 cm in height, with two small notches to fit over the axis of the lateral root, was sealed around a cluster of developing rootlets with silicon grease. Nutrient solution (2 mL) was placed around the isolated cluster and the tank was filled to cover the rest of the root system. In all experiments the solution in the ring was replaced every 6, 12, or 18 h with fresh nutrient solution. Keerthisinghe et al. (1998) reported that degradation of citrate did not occur in the incubation rings, so no precautions were taken to prevent the breakdown of organic acids during exudate collection in the experiments reported here. Any breakdown would have resulted in an underestimation of exported organic acids. Once collected from the rings, the solutions with the exudates were immediately frozen and stored at −20°C until organic acid analysis. Exudates were collected from the fourth cluster of proteoid rootlets developing on the basal laterals on both ambient- and elevated-[CO2]-grown plants when the plants were 26 d old (Figs. 2 and3). In one experiment plants were grown with either ambient or elevated [CO2], and exudates were collected continuously for 6 to 8 d, from the time that the rootlets emerged from the cortex and were 0.5 to 1 mm long (Fig. 3B). The cluster was photographed daily in the incubation ring for correlation of rootlet growth with exudate efflux. Rootlet length was measured directly from photographs using a digitizing tablet. Eight rootlets per cluster and 12 clusters from four plants per CO2 treatment were measured. Fig. 2. Open in new tabDownload slide Emergence of clusters of proteoid rootlets scored daily on a white lupin plant grown in nutrient solution with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. Newly formed clusters emerged simultaneously in discrete pulses on all of the proteoid roots of the root system. The plot is of one representative plant; eight other plants exhibited similar patterns of cluster emergence. The primary laterals emerged when seedlings were 8 d old. Fig. 2. Open in new tabDownload slide Emergence of clusters of proteoid rootlets scored daily on a white lupin plant grown in nutrient solution with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. Newly formed clusters emerged simultaneously in discrete pulses on all of the proteoid roots of the root system. The plot is of one representative plant; eight other plants exhibited similar patterns of cluster emergence. The primary laterals emerged when seedlings were 8 d old. In the second experiment plants were grown with ambient [CO2] only. After the developmental time course shown in Figure 3B, exudates were collected for 24 h, and then the cluster was harvested. A sample of two to three rootlets plus approximately 1 mm of the adjoining main axis was excised from each harvested cluster and immediately fixed in glutaraldehyde on ice for anatomical studies. The remaining tissue was immediately frozen in liquid nitrogen for enzymatic studies. Analysis of Exudates Enzyme Assay for Citrate Glycylglycine buffer (100 mm with 0.2 mmZnCl2, pH 7.9), 0.3 mm NADH, 600 kilounits L−1 lactate dehydrogenase (EC 1.1.2.3), and 600 kilounits L−1 malate dehydrogenase were combined with 1 mL of thawed exudate, mixed thoroughly, and read at 340 nm in a spectrophotometer. Citrate lyase (EC 4.1.3.6) was added (final activity in cuvette 40 kilounits L−1), and the absorbance was monitored until it stabilized. The decrease in absorbance was proportional and stoichiometric to the amount of citrate present in the reaction (Möllering, 1985; Keerthisinghe et al., 1998). HPLC for Organic Acids Exudates were thawed and 1 mL was passed through a 0.45-μm pore-size filter with a syringe. The filtrate was evaporated to dryness in a freeze drier (Speed Vac, Savant Instruments, Holbrook, NY), redissolved with 40 to 100 μL of 13 mmH2SO4, and spun at 13,000 rpm for 5 min. Twenty-five microliters of the supernatant was injected into an HPLC (model 1090M, Hewlett-Packard) that was fitted with an ion-exclusion column (300 × 7.8 mm; HPX-87H Aminex, Bio-Rad) and an organic acid guard column (Bio-Rad). The mobile phase was 13 mm H2SO4 run at 0.5 mL min−1 at 60°C. The acids were detected at 210 nm with a photodiode-array UV detector. Standards for oxalic acid, citric acid, α-ketoglutaric acid, malic acid, succinic acid, pyruvic acid, and fumaric acid were made in distilled water at concentrations within the ranges found in the exudates. As with the samples, the standards were filtered, evaporated, suspended in 13 mm H2SO4, centrifuged, and then run through the HPLC system individually or as a mixture. Linear standard curves were generated for each acid from the areas under the peaks corresponding to different concentrations. The standard curves were used to quantify acids in the exudates. To verify running conditions, a mixture of the standard acids was run before each batch of samples was analyzed. Specific Enzyme Activities and Protein Content Each root sample was frozen, transferred to a chilled, 2-mL glass homogenizer, and ground on ice for 1 to 2 min in 250 μL of freshly prepared grinding solution (50 mm Hepes, 5 mmMgCl2, 1 mm EDTA, 1 mmEGTA, 0.1% Triton X-100, 10% glycerol, 0.5 mm PMSF dissolved in isopropanol, 5 mm DTT, 2 mmbenzamidine, and 2 mm ε-amino-n-caproic acid). The ground sample was divided into three aliquots, snap-frozen in liquid nitrogen, and stored at −80°C until analysis. Just before analysis, samples were thawed on ice and centrifuged at 13,000 rpm for 10 min at 4°C; the supernatant was maintained on ice. PEPC activity in the tissue supernatant was measured by monitoring the oxidation of NADH at 340 nm in a spectrophotometer (Vance et al., 1983). The tissue supernatant (40–80 μL) was initially incubated for 10 min at 25°C in 100 mm Bicine, pH 8.0, with 5 mm MgCl2, 10 mmNaHCO3, 0.16 mm NADH, and 60 units of malate dehydrogenase. Next, 15 μL of 100 mm PEP was added and the decrease in absorbance per unit of time was measured. Citrate synthase (EC 4.1.3.7) activity was measured by following the rate of 3-acetylpyridine adenine dinucleotide reduction at 365 nm in a spectrophotometer (Stitt, 1983). Tissue supernatant (60–100 μL) was incubated with 100 mm triethanolamine, pH 8.5, 3.5 mm malate, 0.3 mm 3-acetylpyridine adenine dinucleotide, and 30 units of malate dehydrogenase at 25°C until the absorbance had stabilized (approximately 15 min). Next, 15 μL of 10 mm acetyl-CoA was added and the increase in absorbance per unit of time was measured. Protein in the tissue supernatant was estimated using the Coomassie Plus protein assay reagent kit (Pierce). Tissue Preservation and Staining for Anatomy Rootlets were fixed in 3% glutaraldehyde in 25 mm potassium-phosphate buffer, pH 6.8, on ice overnight, rinsed four times for 15 min each in buffer, postfixed in 1% osmium tetroxide for 2 h, and rinsed three times for 15 min each in buffer. The tissue was taken through a gradual dehydration series from 4% to 100% ethanol over 2 d on ice and then at room temperature and slowly infiltrated with Spurr's resin starting with 2.5% in ethanol and reaching 100% in 2 d. The Spurr's resin was replaced daily for 5 d and then the resin-embedded tissue was polymerized at 70°C overnight. Transverse and longitudinal sections (2–3 μm thick) of resin-embedded material were cut with a glass knife, transferred to drops of water on gelatin-coated glass slides, and dried on a hot plate for 1 h. Sections were first stained with 1% toluidine blue in borate, pH 11.0, and viewed with bright-field optics to show general anatomy and development. To visualize phloem differentiation, the resin was etched from the sections with sodium ethoxide (1–2 min), rinsed with 70% ethanol, and then rinsed for 1 min in running tap water. The sections were then stained with Schiff's reagent for 4 min to reduce background wall autofluorescence, rinsed for 5 min in running tap water, and stained with 0.05% aniline blue in 67 mmpotassium-phosphate buffer, pH 8.6, for 2 h. Sections were mounted in fresh aniline blue and viewed with UV fluorescence optics (Axioplan microscope, Zeiss). The callose deposited in the walls and plates of the sieve tubes fluoresces bright blue/green and could therefore be distinguished from lignin and suberin autofluorescence (O'Brien and McCully, 1981). RESULTS Root System Architecture and Proteoid Root Morphology We found in white lupin a taproot system with approximately 20 thick, indeterminate, basal primary laterals in addition to the thinner, shorter, acropetal primary laterals (Fig. 1). All of the basal primary laterals became proteoid roots, developing distinct clusters of closely spaced, determinate, secondary laterals or proteoid rootlets. Few of the thinner, acropetal laterals became proteoid roots as the plant aged. The clusters of proteoid rootlets emerged in near synchrony on all of the proteoid roots of the root system, regardless of root length (Fig. 2). The emergence of the first cluster of rootlets was predictable, occurring 7 d after the emergence of the basal laterals when the plant was approximately 2 weeks old. This first cluster often appeared on 7 to 15 basal primary laterals; the second and third clusters recruited more basal laterals. By the third cluster, a plateau was reached in the number of primary laterals becoming proteoid roots (Fig. 2). An example of a basal primary lateral from a 32-d-old plant is shown in Figure 3A. Five root clusters are shown, with a sixth cluster just emerging from the cortex. The length and spacing of the root axis of each cluster varied; for the earlier clusters the morphology was more variable. For this study, all data were collected from the fourth cluster. A time course of rootlet development of this cluster is shown in Figure 3B. Within a cluster, almost all rootlets emerged from the cortex in near synchrony; they grew and developed at similar rates, with an occasional rootlet reaching 2 to 3 times the length of other rootlets. Effect of Atmospheric [CO2] on Proteoid Rootlet Elongation and Efflux of Organic Acids Rootlet length was measured on clusters restrained in the incubation rings. Rootlets grew rapidly for the first 2 d, reaching their final 4-mm length 4 d after emergence from the cortex in plants grown with ambient [CO2] (Fig.4). Elevated atmospheric [CO2] resulted in the rootlets stopping growth after only 3 d, reaching only a 2.5-mm final length. Fig. 4. Open in new tabDownload slide Lengths of proteoid rootlets of white lupin grown in nutrient culture with 1 μm phosphorus. Rootlets grown with 700 μL L−1 atmospheric [CO2] (•) reached a shorter final length 1 d earlier than rootlets grown with 350 μL L−1 atmospheric CO2 (♦). Each point represents the mean ± se of 12 roots from four plants per CO2 treatment. Fig. 4. Open in new tabDownload slide Lengths of proteoid rootlets of white lupin grown in nutrient culture with 1 μm phosphorus. Rootlets grown with 700 μL L−1 atmospheric [CO2] (•) reached a shorter final length 1 d earlier than rootlets grown with 350 μL L−1 atmospheric CO2 (♦). Each point represents the mean ± se of 12 roots from four plants per CO2 treatment. At both CO2 concentrations, the onset of citrate efflux occurred after the rootlets stopped elongating; onset was 1 d earlier in plants grown with elevated atmospheric [CO2] (Fig. 5). Citrate efflux continued for 2 d in the elevated-[CO2]-grown plants and for 3 d in the ambient-[CO2]-grown plants. Peaks in efflux occurred during the day under both CO2treatments, and the peak efflux rate per unit of length of proteoid root was not altered by atmospheric [CO2]. Citrate was the dominant organic acid exuded from the roots, being 60-fold higher than malate at peak efflux of citrate (TableI). Other organic acids were detected at very low concentrations and were unaffected by the atmospheric [CO2]. Fig. 5. Open in new tabDownload slide Citrate efflux from developing clusters of proteoid roots of white lupin grown in nutrient culture with 1 μm phosphorus. Shading indicates dark periods. Each point represents the mean ± se of 12 roots from four plants per CO2 treatment. A, Plants grown with 350 μL L−1 atmospheric [CO2]. Citrate efflux began 4 d after rootlet emergence, when rootlets had stopped elongating, and lasted 3 d, with strong peaks during light periods. B, Plants grown with 700 μL L−1 atmospheric [CO2]. Citrate efflux began 3 d after rootlet emergence when rootlets had stopped elongating and lasted 2 d with strong peaks during light periods. Fig. 5. Open in new tabDownload slide Citrate efflux from developing clusters of proteoid roots of white lupin grown in nutrient culture with 1 μm phosphorus. Shading indicates dark periods. Each point represents the mean ± se of 12 roots from four plants per CO2 treatment. A, Plants grown with 350 μL L−1 atmospheric [CO2]. Citrate efflux began 4 d after rootlet emergence, when rootlets had stopped elongating, and lasted 3 d, with strong peaks during light periods. B, Plants grown with 700 μL L−1 atmospheric [CO2]. Citrate efflux began 3 d after rootlet emergence when rootlets had stopped elongating and lasted 2 d with strong peaks during light periods. Table I. Rates of efflux of organic acids detected around proteoid roots before, during, and after peaks in efflux activity Organic Acid . Efflux Rate . Ambient [CO2] . Elevated [CO2] . Prepeak . Peak . Postpeak . Prepeak . Peak . Postpeak . nmol min−1 m−1 Enzymatic analysis  Citrate 0.27  ± 0.22 23  ± 7 ND-a 0.19  ± 0.11 19  ± 3 ND HPLC analysis  Citrate ND 33  ± 10 0.29  ± 0.29 ND 23  ± 4 0.1  ± 0.04  Oxalate 1.2  ± 0.2 0.019  ± 0.019 0.3  ± 0.03 0.8  ± 0.15 0.7  ± 0.31 0.23  ± 0.02  α-Ketoglutarate 0.082  ± 0.044 0.055  ± 0.055 0.11  ± 0.09 0.07  ± 0.04 0.17  ± 0.11 0.004  ± 0.004  Malate 1.4  ± 0.78 0.55  ± 0.25 0.34  ± 0.12 0.35  ± 0.19 0.43  ± 0.16 0.13  ± 0.04  Succinate 0.069  ± 0.069 0.042  ± 0.042 ND 0.83  ± 0.48 0.87  ± 0.42 0.17  ± 0.12  Pyruvate 0.81  ± 0.52 ND 1.18  ± 0.8 0.49  ± 0.3 0.043  ± 0.043 0.05  ± 0.05  Fumarate 0.12  ± 0.12 ND 0.12  ± 0.05 0.31  ± 0.2 0.043  ± 0.036 0.09  ± 0.07  Total-b 3.6  ± 1.3 34  ± 10 2.3  ± 0.68 2.86  ± 0.57 26  ± 4 0.75  ± 0.09 Organic Acid . Efflux Rate . Ambient [CO2] . Elevated [CO2] . Prepeak . Peak . Postpeak . Prepeak . Peak . Postpeak . nmol min−1 m−1 Enzymatic analysis  Citrate 0.27  ± 0.22 23  ± 7 ND-a 0.19  ± 0.11 19  ± 3 ND HPLC analysis  Citrate ND 33  ± 10 0.29  ± 0.29 ND 23  ± 4 0.1  ± 0.04  Oxalate 1.2  ± 0.2 0.019  ± 0.019 0.3  ± 0.03 0.8  ± 0.15 0.7  ± 0.31 0.23  ± 0.02  α-Ketoglutarate 0.082  ± 0.044 0.055  ± 0.055 0.11  ± 0.09 0.07  ± 0.04 0.17  ± 0.11 0.004  ± 0.004  Malate 1.4  ± 0.78 0.55  ± 0.25 0.34  ± 0.12 0.35  ± 0.19 0.43  ± 0.16 0.13  ± 0.04  Succinate 0.069  ± 0.069 0.042  ± 0.042 ND 0.83  ± 0.48 0.87  ± 0.42 0.17  ± 0.12  Pyruvate 0.81  ± 0.52 ND 1.18  ± 0.8 0.49  ± 0.3 0.043  ± 0.043 0.05  ± 0.05  Fumarate 0.12  ± 0.12 ND 0.12  ± 0.05 0.31  ± 0.2 0.043  ± 0.036 0.09  ± 0.07  Total-b 3.6  ± 1.3 34  ± 10 2.3  ± 0.68 2.86  ± 0.57 26  ± 4 0.75  ± 0.09 Plants were grown with ambient (350 μL L−1) or elevated (700 μL L−1) atmospheric [CO2], and exudates were collected in situ with incubation rings on roots for 1 week. Each value represents the mean ± se of exudates from one root from each of three to six plants. F0-a ND, Not detected. F0-b Sum of all organic acids detected by HPLC. Open in new tab Table I. Rates of efflux of organic acids detected around proteoid roots before, during, and after peaks in efflux activity Organic Acid . Efflux Rate . Ambient [CO2] . Elevated [CO2] . Prepeak . Peak . Postpeak . Prepeak . Peak . Postpeak . nmol min−1 m−1 Enzymatic analysis  Citrate 0.27  ± 0.22 23  ± 7 ND-a 0.19  ± 0.11 19  ± 3 ND HPLC analysis  Citrate ND 33  ± 10 0.29  ± 0.29 ND 23  ± 4 0.1  ± 0.04  Oxalate 1.2  ± 0.2 0.019  ± 0.019 0.3  ± 0.03 0.8  ± 0.15 0.7  ± 0.31 0.23  ± 0.02  α-Ketoglutarate 0.082  ± 0.044 0.055  ± 0.055 0.11  ± 0.09 0.07  ± 0.04 0.17  ± 0.11 0.004  ± 0.004  Malate 1.4  ± 0.78 0.55  ± 0.25 0.34  ± 0.12 0.35  ± 0.19 0.43  ± 0.16 0.13  ± 0.04  Succinate 0.069  ± 0.069 0.042  ± 0.042 ND 0.83  ± 0.48 0.87  ± 0.42 0.17  ± 0.12  Pyruvate 0.81  ± 0.52 ND 1.18  ± 0.8 0.49  ± 0.3 0.043  ± 0.043 0.05  ± 0.05  Fumarate 0.12  ± 0.12 ND 0.12  ± 0.05 0.31  ± 0.2 0.043  ± 0.036 0.09  ± 0.07  Total-b 3.6  ± 1.3 34  ± 10 2.3  ± 0.68 2.86  ± 0.57 26  ± 4 0.75  ± 0.09 Organic Acid . Efflux Rate . Ambient [CO2] . Elevated [CO2] . Prepeak . Peak . Postpeak . Prepeak . Peak . Postpeak . nmol min−1 m−1 Enzymatic analysis  Citrate 0.27  ± 0.22 23  ± 7 ND-a 0.19  ± 0.11 19  ± 3 ND HPLC analysis  Citrate ND 33  ± 10 0.29  ± 0.29 ND 23  ± 4 0.1  ± 0.04  Oxalate 1.2  ± 0.2 0.019  ± 0.019 0.3  ± 0.03 0.8  ± 0.15 0.7  ± 0.31 0.23  ± 0.02  α-Ketoglutarate 0.082  ± 0.044 0.055  ± 0.055 0.11  ± 0.09 0.07  ± 0.04 0.17  ± 0.11 0.004  ± 0.004  Malate 1.4  ± 0.78 0.55  ± 0.25 0.34  ± 0.12 0.35  ± 0.19 0.43  ± 0.16 0.13  ± 0.04  Succinate 0.069  ± 0.069 0.042  ± 0.042 ND 0.83  ± 0.48 0.87  ± 0.42 0.17  ± 0.12  Pyruvate 0.81  ± 0.52 ND 1.18  ± 0.8 0.49  ± 0.3 0.043  ± 0.043 0.05  ± 0.05  Fumarate 0.12  ± 0.12 ND 0.12  ± 0.05 0.31  ± 0.2 0.043  ± 0.036 0.09  ± 0.07  Total-b 3.6  ± 1.3 34  ± 10 2.3  ± 0.68 2.86  ± 0.57 26  ± 4 0.75  ± 0.09 Plants were grown with ambient (350 μL L−1) or elevated (700 μL L−1) atmospheric [CO2], and exudates were collected in situ with incubation rings on roots for 1 week. Each value represents the mean ± se of exudates from one root from each of three to six plants. F0-a ND, Not detected. F0-b Sum of all organic acids detected by HPLC. Open in new tab Rootlet Development and Anatomy The development and anatomy of the proteoid rootlets are illustrated in Figures 6 and7 and summarized in TableII. One day after emergence from the proteoid root, the rootlets were almost entirely meristematic (Figs. 6A and 7F); only the cells of the rootlet cortex within the proteoid root had started to vacuolate and elongate. The phloem sieve tubes were visible at approximately six cell lengths from the phloem of the stele of the proteoid root axis, and extended over only 25% of the rootlet length. The root cap adhered tightly to the rootlet tip; at this stage no root hairs had developed. On d 2, many rootlet cells were developing vacuoles and elongating, whereas cells toward the tip of the stele and cortex still had high amounts of cytoplasm (Fig. 7G). Root hairs were beginning to develop; root-cap cells were sloughing from the epidermis and could be seen along the length of the rootlet; and callose in the phloem sieve tubes had differentiated along 60% of the rootlet length (Fig. 7, G and L). Fig. 6. Open in new tabDownload slide Cross-sections (3 μm thick) through the tips of resin-embedded proteoid rootlets of white lupin grown in nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. A, Rootlet 1 d after emergence from the cortex of the proteoid root. All of the cells shown are meristematic except for the root-cap cells, which are tightly anchored to the epidermis. Magnification = ×280. B, Rootlet 4 d after emergence from the cortex. The apical meristem is no longer present and all cells in the rootlet have vacuolated and differentiated. The epidermis has differentiated root hairs to the tip of the rootlet, and the root-cap cells are loosely anchored. Magnification = ×300. C, Rootlet 8 d after emergence from the cortex. The stele has two xylem and two phloem poles and is surrounded by an endodermis with a Casparian band. Sections from d 1 and 4 were stained with toluidine blue and viewed with bright-field optics. The section from d 8 was etched after sectioning to remove the resin, stained with Schiff's reagent, mounted in aniline blue, and viewed with fluorescence optics. Black arrowheads, Root hairs; white arrowheads, Casparian bands; white arrow, phloem sieve tube; c, root cap; x, xylem vessel. Magnification = ×1000. Fig. 6. Open in new tabDownload slide Cross-sections (3 μm thick) through the tips of resin-embedded proteoid rootlets of white lupin grown in nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. A, Rootlet 1 d after emergence from the cortex of the proteoid root. All of the cells shown are meristematic except for the root-cap cells, which are tightly anchored to the epidermis. Magnification = ×280. B, Rootlet 4 d after emergence from the cortex. The apical meristem is no longer present and all cells in the rootlet have vacuolated and differentiated. The epidermis has differentiated root hairs to the tip of the rootlet, and the root-cap cells are loosely anchored. Magnification = ×300. C, Rootlet 8 d after emergence from the cortex. The stele has two xylem and two phloem poles and is surrounded by an endodermis with a Casparian band. Sections from d 1 and 4 were stained with toluidine blue and viewed with bright-field optics. The section from d 8 was etched after sectioning to remove the resin, stained with Schiff's reagent, mounted in aniline blue, and viewed with fluorescence optics. Black arrowheads, Root hairs; white arrowheads, Casparian bands; white arrow, phloem sieve tube; c, root cap; x, xylem vessel. Magnification = ×1000. Fig. 7. Open in new tabDownload slide Development from 1 d after emergence from the cortex of proteoid rootlets from white lupin grown in nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. Left, Whole mounts of glutaraldehyde- and osmium-fixed rootlets embedded in resin. Center, Longitudinal sections (3 μm thick) through rootlets shown at left stained with toluidine blue (F, G, and I) or left unstained and viewed with differential image contrast bright-field optics (H and J). Right, Longitudinal sections (3 μm thick) etched to remove resin, stained with Schiff's reagent followed by aniline blue, and viewed with UV fluorescence optics. Arrows, Points of visible callose deposition in the phloem; x, xylem secondary wall; arrowheads, root hairs; c, root cap. At d 1, rootlets are almost entirely meristematic. At d 2, most of the cells are developing vacuoles, root hairs have started to develop at the base, and the phloem callose extends to 75% of the rootlet length. At d 3, the apical meristem is completely exhausted, and root hairs and vascular tissues have differentiated along the length of the rootlet. Days 4 and 6 are similar to d 3. Refer to text for more details. Rootlet diameter was approximately 225 μm. Magnification: A to E, ×60; F and G, ×100; H and M, ×180; I, J, L, and O, ×170; K, ×150; and N, ×425. The root tip in L, J, and O is pointing up. Fig. 7. Open in new tabDownload slide Development from 1 d after emergence from the cortex of proteoid rootlets from white lupin grown in nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. Left, Whole mounts of glutaraldehyde- and osmium-fixed rootlets embedded in resin. Center, Longitudinal sections (3 μm thick) through rootlets shown at left stained with toluidine blue (F, G, and I) or left unstained and viewed with differential image contrast bright-field optics (H and J). Right, Longitudinal sections (3 μm thick) etched to remove resin, stained with Schiff's reagent followed by aniline blue, and viewed with UV fluorescence optics. Arrows, Points of visible callose deposition in the phloem; x, xylem secondary wall; arrowheads, root hairs; c, root cap. At d 1, rootlets are almost entirely meristematic. At d 2, most of the cells are developing vacuoles, root hairs have started to develop at the base, and the phloem callose extends to 75% of the rootlet length. At d 3, the apical meristem is completely exhausted, and root hairs and vascular tissues have differentiated along the length of the rootlet. Days 4 and 6 are similar to d 3. Refer to text for more details. Rootlet diameter was approximately 225 μm. Magnification: A to E, ×60; F and G, ×100; H and M, ×180; I, J, L, and O, ×170; K, ×150; and N, ×425. The root tip in L, J, and O is pointing up. Table II. Development and citrate efflux of proteoid rootlets, ranging from none detected (−) to maximum (+++), in plants grown with ambient [CO2] Rootlet Age . Development . Citrate Efflux . Meristematic cells . Phloem and xylem . Root hairs . d 1 +++ + − − 2 ++ ++ + − 3 − +++ ++ − 4 − +++ +++ ++ 5 − +++ +++ +++ 6 − +++ +++ ++ 7-8 − +++ +++ − Rootlet Age . Development . Citrate Efflux . Meristematic cells . Phloem and xylem . Root hairs . d 1 +++ + − − 2 ++ ++ + − 3 − +++ ++ − 4 − +++ +++ ++ 5 − +++ +++ +++ 6 − +++ +++ ++ 7-8 − +++ +++ − Open in new tab Table II. Development and citrate efflux of proteoid rootlets, ranging from none detected (−) to maximum (+++), in plants grown with ambient [CO2] Rootlet Age . Development . Citrate Efflux . Meristematic cells . Phloem and xylem . Root hairs . d 1 +++ + − − 2 ++ ++ + − 3 − +++ ++ − 4 − +++ +++ ++ 5 − +++ +++ +++ 6 − +++ +++ ++ 7-8 − +++ +++ − Rootlet Age . Development . Citrate Efflux . Meristematic cells . Phloem and xylem . Root hairs . d 1 +++ + − − 2 ++ ++ + − 3 − +++ ++ − 4 − +++ +++ ++ 5 − +++ +++ +++ 6 − +++ +++ ++ 7-8 − +++ +++ − Open in new tab By d 3, rootlets were approaching their final length (Fig. 4), all cells had vacuolated, and the apical meristem was no longer present. Root hairs were long and dense toward the base of the rootlet, and epidermal cells around the tip were just developing hairs (Fig. 7, C and H). The distribution of root-hair growth along the rootlet varied, however, with some rootlets showing very dense proliferation toward the tip and others toward the base. The phloem sieve tubes and protoxylem secondary walls were visible to within five cell lengths of the rootlet tip, indicating that the stele had differentiated along more than 95% of the rootlet length (Fig. 7H). By d 4, citrate efflux had started (Fig. 5A) and root hairs continued to expand (Fig. 7D). The root cap was loosely anchored and sloughed clumps of root-cap cells were often associated with root hairs (Figs.6B and 7I). Rootlet development from d 5 to 8 was similar to that seen at d 4, although root hairs continued to develop until d 6. A completely differentiated rootlet had sloughed root-cap cells and an epidermal layer with root hairs had differentiated around the tip of the rootlet. Four layers of cortical cells (the innermost being an endodermal layer with a suberized Casparian band) are shown in Figure 6C, and a stele occupying approximately 10% of the rootlet (in cross-section) is shown in Figure 6B. The stele had two phloem poles and two protoxylem poles (with spiral secondary wall thickenings; Figs.6C and 7H). The dry weight of clusters of rootlets increased between d 1 and 5, plateaued, and then dropped at d 8 (Fig.8A). Fresh weight increased abruptly on d 3 and 4 because of root-hair development and vacuolation of cells (Fig.8B); as a consequence, the soluble protein content per unit of fresh weight fell sharply (Fig. 8C). The soluble protein content per unit of root length increased steadily to a peak on d 3 before declining again (Fig. 9C). Fig. 8. Open in new tabDownload slide Dry weight (A), fresh weight (B), and soluble protein content (C) of proteoid root clusters of white lupin grown in nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. Each point represents the mean ± se of one root from each of four to six plants. Fig. 8. Open in new tabDownload slide Dry weight (A), fresh weight (B), and soluble protein content (C) of proteoid root clusters of white lupin grown in nutrient culture with 1 μm phosphorus and 350 μL L−1 atmospheric [CO2]. Each point represents the mean ± se of one root from each of four to six plants. Fig. 9. Open in new tabDownload slide In vitro PEPC (A) and citrate synthase activities (B) per unit of soluble protein of developing proteoid root clusters from white lupin grown in nutrient culture with 1 μmphosphorus and 350 μL L−1 atmospheric [CO2]. C, Soluble protein peaks 3 d after rootlet emergence. Each point represents the mean ± se of one root from each of four to six plants. Fig. 9. Open in new tabDownload slide In vitro PEPC (A) and citrate synthase activities (B) per unit of soluble protein of developing proteoid root clusters from white lupin grown in nutrient culture with 1 μmphosphorus and 350 μL L−1 atmospheric [CO2]. C, Soluble protein peaks 3 d after rootlet emergence. Each point represents the mean ± se of one root from each of four to six plants. Enzyme Activities and Citrate Efflux In vitro PEPC activity per unit of protein peaked 3 d after rootlet emergence, doubling in activity between d 1 and 3 before decreasing to 25% of peak activity by d 8 (Fig. 9A). Citrate synthase activity per unit of protein declined by 50% on d 1, after which it plateaued for 5 d before declining again (Fig. 9B). Both PEPC and citrate synthase had maximal activity per unit length of proteoid root 3 d after rootlet emergence, preceding the onset of citrate efflux by 1 d and the peak of efflux by 2 d (Fig.10). PEPC and citrate synthase had approximately 4- and 3-fold increases in activity, respectively, between d 1 and 3. Citrate efflux from the developmental time-course samples that were harvested daily lasted 3 d, with efflux peaks during photoperiods (Fig. 10C). This length of time was similar to that of the efflux from developing roots maintained in the incubation rings for 8 d (Fig. 5A); however, efflux from roots collected from rings in place for just 24 h was twice that of the roots maintained in rings for 8 d. The total amount of citrate exudation was equivalent to 10% of the proteoid-root dry weight. In vitro activities of PEPC and citrate synthase were well in excess of citrate-exudation rates at all times. Fig. 10. Open in new tabDownload slide In vitro PEPC activity (A), citrate synthase activity (B), and citrate efflux (C) per unit length of white lupin proteoid root grown in nutrient culture with 1 μmphosphorus and 350 μL L−1 atmospheric [CO2]. PEPC and citrate synthase activities per unit length of main root peaked 3 d after rootlet emergence, 1 d before onset of citrate efflux and 2 d before the peak in citrate efflux on d 5. Each point represents the mean ± se of one root from each of four to six plants. Fig. 10. Open in new tabDownload slide In vitro PEPC activity (A), citrate synthase activity (B), and citrate efflux (C) per unit length of white lupin proteoid root grown in nutrient culture with 1 μmphosphorus and 350 μL L−1 atmospheric [CO2]. PEPC and citrate synthase activities per unit length of main root peaked 3 d after rootlet emergence, 1 d before onset of citrate efflux and 2 d before the peak in citrate efflux on d 5. Each point represents the mean ± se of one root from each of four to six plants. DISCUSSION Synchronous Development of Proteoid Roots A striking finding of this study was the predictable and synchronous development of clusters of rootlets on the proteoid roots in white lupin grown in solution culture (Fig. 2). This implicates a central signaling cascade for development of clusters of rootlets. Studies showing that the internal phosphorus status of the plant can determine proteoid root development in white lupin (Marschner et al., 1987; Keerthisinghe et al., 1998) and wax myrtle (Louis et al., 1990) support a central signaling mechanism. The signal may start in the shoot and radiate down to the roots, reaching the basal laterals first. It is possible that the signaling process is delivered to the roots in pulses related to the plant phosphorus status, if so, the frequency and duration would determine the spacing and length of each cluster of rootlets along the proteoid root axis. As the severity of phosphorus stress grew, signaling would intensify, thus increasing the proportion of the root system covered with proteoid rootlets (Keerthisinghe et al., 1998). The signals involved in proteoid root development probably include auxin. Gilbert et al. (1997) were able to use auxin to induce the production of proteoid rootlets in white lupin when phosphorus was supplied at a level that normally suppresses their development, and they could suppress proteoid rootlet formation in minus-phosphorus treatments by supplying auxin-transport inhibitors to the nutrient solution. Auxin plays a role in lateral root initiation in other species (Thimann, 1936; Wightman et al., 1980); in the Arabidopsis mutant, superroot is responsible for a phenotype that is very similar to a proteoid root (Boerjan et al., 1995). The rootlets that developed within clusters along a proteoid root reached a similar, determinate length, although an occasional rootlet extended two to three times the length of the other rootlets (Fig. 3A). Determinacy has been reported in all species that form proteoid roots (Dinkelaker et al., 1995) but has also been observed in roots from other types of species (Varney and McCully, 1991; Dubrovsky, 1997). The average final length of a rootlet varies within and among proteoid species; recently Skene et al. (1998a) showed that rootlets ofGrevillea robusta were shorter when they developed in hydroponics compared with development in vermiculite. To our knowledge, we are the first to report that determinacy varies with environmental treatment ([CO2]) in a common rooting medium (Fig. 4), indicating that root determinacy is under internal control. Controls for root determinacy are unknown, but the switch from indeterminacy to determinacy in stem nodules has been linked to the environment and the presence of the hormone ethylene (Fernández-López et al., 1998). Although auxin is responsible for initiating lateral root growth, continued exposure to auxins can inhibit elongation of laterals (Thimann, 1936). In addition, cytokinins have been shown to suppress lateral root formation (Wightman et al., 1980). The effects of cytokinins or ethylene on proteoid root development are not yet known. Rootlet Development and Citrate Efflux The anatomy of white lupin proteoid rootlets is similar in most respects to that of members of the Proteaceae, in which the mature rootlets have a differentiated apex, root hairs to the tip, a loosely anchored root cap, an endodermis with a Casparian band, and a diarch stele (for review, see Dinkelaker et al., 1995; Skene et al., 1998b). Root-hair development in white lupin differs from that of G. robusta (Skene et al., 1996, 1998a) and Hakea obliqua(Dell et al., 1980). In G. robusta, root hairs were produced only after the rootlet had reached its final length; they then developed back from the tip. In H. obliqua, few hairs were produced in water culture, whereas they were produced extensively in soil. Citrate efflux began shortly after the rootlets reached their final length. By that stage, phloem and xylem tissues had differentiated to the tips of the rootlets, enabling the import of photosynthates and the export of nutrients mobilized by the citrate exudation. Root hairs started to develop 2 d after emergence from the cortex, which was also 2 d before the onset of citrate efflux. This suggests that the presence of root hairs does not alone determine organic acid export. Citrate export lasted only 2 to 3 d and then stopped. Neumann et al. (1995) also reported transient release of organic acids from a proteoid root cluster of Hakea undulata, although they did not document root development. Keerthisinghe et al. (1998) found that citrate efflux was maximal 1 to 3 cm from the tip and was only one-tenth that rate from either the 0- to 1-cm or the 5- to 9-cm region. The protein content of the rootlets peaked 3 d after emergence and then declined during citrate efflux (Fig. 9C). The links among determinacy, longevity, metabolism related to senescence, and efflux of organic acids remain to be investigated. Some plants export organic anions from their roots upon exposure to Al; the organic acids chelate the Al, conferring tolerance. The fact that white lupin can export large amounts of citrate, an excellent chelator of Al, suggests that it may be tolerant to soils with high levels of available Al. However, citrate is not exported to the rhizosphere until the rootlets are fully mature, 2 to 4 d past the time that the rootlets are dividing and meristematic and are thought to be most susceptible to Al damage (Ryan et al., 1993). If the emerging rootlets are not protected from damage by exposure to Al, they will be hindered in their ability to access phosphorus when they are mature, imposing a double stress in very acidic soils. To confer Al tolerance, organic acid efflux from roots should coincide with the time that they are meristematic. Enzyme Activities and Citrate Efflux We did not find a correlation between PEPC activity and the magnitude of citrate efflux, because the peak in in vitro PEPC activity preceded the onset of citrate efflux by 1 d and preceded peak efflux by 2 d (Fig. 10). Furthermore, in vitro citrate synthase activity per unit of protein did not increase during citrate efflux, although citrate was the dominant organic acid exported. Our results contrast with those of Johnson et al. (1994, 1996b), who suggested that increases in PEPC activity per unit of protein, as well as mRNA expression and abundance, coincide with organic acid efflux from the roots, and thus provide necessary carbons for the anapleurotic functioning of the tricarboxylic acid cycle during exudation. In the studies by Johnson et al. (1994, 1996b), enzyme activities were not measured on the same tissues from which exudates were collected; exudates were collected from the entire root system by flushing the pot with nutrient solution every 2 d, whereas enzyme activities were measured on root portions pooled from many plants. Although PEPC activity is clearly involved in citrate synthesis, it does not appear to determine the rate, onset, or duration of citrate exudation.Keerthisinghe et al. (1998) made concomitant measurements of citrate efflux and PEPC activity from 2-cm regions along a proteoid root axis and found that rates of citrate efflux did not vary proportionally with PEPC activity, which is in agreement with our findings. In vitro activities of both PEPC and citrate synthase exceeded the rates of citrate exudation at every stage; however, these activities may not reflect the actual rates realized in vivo. The white lupin PEPC may be regulated by phosphorylation, similar to the regulation of PEPC in other species (for review, see Vidal and Chollet, 1997). Neumann et al. (1999) measured similar citrate concentrations (20 μmol g−1 fresh weight) in juvenile, mature, and senescent proteoid root segments of white lupin, but significant citrate exudation was observed only in mature proteoid roots. Therefore, PEPC activity and tissue citrate concentration must not control the rate of citrate exudation. We have made preliminary measurements of respiration rates of whole pieces of proteoid roots that greatly exceed the peak rate of exudation (4 μmol O2 g−1 dry weight min−1 for 0.025 μmol citrate g−1 dry weight min−1), suggesting that the flux through the TCA cycle is unlikely to limit the rate of citrate exudation. Neumann et al. (1999) measured similar respiration rates by mature and senescent proteoid root segments of white lupin, despite significant citrate exudation by mature but not senescent proteoid roots. Most of the respiration in hydroponically grown, mature proteoid roots is therefore involved in other metabolic processes such as ion uptake and amino acid synthesis (Jeschke and Pate, 1995; Johnson et al., 1996b) and maintenance rather than in citrate synthesis for exudation. In the present study, there was a strong effect of day and night on citrate efflux, with peaks in efflux occurring during the light periods. To our knowledge, this is the first time a diurnal rhythm has been reported for organic acids exported from roots. A diurnal rhythm has been reported for the export of phytosiderophores from graminaceous roots in response to Fe deficiency (Ma and Nomoto, 1996). The highest tissue concentrations and effluxes of phytosiderophores also occur in the light, but it is unclear whether the mechanisms are directly linked to light or temperature (Ma and Nomoto, 1996). Our data strongly suggest that citrate export is not simply a result of PEPC activity supplying the tricarboxylic acid cycle. Another key step must be regulating the export of citrate from the roots. In a study byRyan et al. (1995), neither malate dehydrogenase nor PEPC showed enhanced specific activities associated with malate efflux from wheat root tips. The same investigators showed that efflux of malate is likely to be linked to the activity of an anion transporter that was detected on the plasma membrane of the root tip cells (Ryan et al., 1997). The transport mechanism for organic acid efflux from proteoid roots is not known, although Dinkelaker et al. (1989) concluded that citrate is excreted concomitantly with protons. It seems likely that an anion transporter in the plasma membrane is synthesized and becomes active as rootlet elongation ceases. The transporter is then inactivated after 2 to 3 d, stopping further exudation. This transporter could be located in specific cells, but whether those are in the cortex, the epidermis, or elsewhere is unknown. Incubation of proteoid roots in solutions of anion-channel blockers reduced citrate exudation by 50% (Neumann et al., 1999), supporting the existence of an anion channel. Effect of Elevated Atmospheric [CO2] Growth under elevated atmospheric [CO2] altered the development of root clusters. Rootlets were shorter, reaching their final length and beginning citrate exudation 1 d earlier than ambient-[CO2]-grown plants (Figs.4 and 5). It is unclear how atmospheric [CO2] alters rootlet determinacy. Atmospheric CO2 can enhance lateral root turnover in some species and in certain soil environments (Pregitzer et al., 1995; Berntson and Bazzaz, 1996; Fitter et al., 1996) and may do so in proteoid rootlets by promoting exhaustion of the apical meristem and shortening growing time. By advancing the onset of determinacy and the onset of citrate efflux, residency time of a proteoid root cluster in a patch of soil may be decreased. In the present study, elevated atmospheric [CO2] did not change the rate of citrate efflux or the composition of background organic acids per unit of length of proteoid root, although it did shorten the period over which this efflux occurred (Fig. 5). Unfortunately, the present experimental technique was not able to provide an integrated picture of efflux for a whole plant such as can be gained by the method of Johnson et al. (1996b). There is a paucity of data regarding measurements of root exudation in the literature, but there is a great deal of speculation. The data shown in Figure 5 suggest that an increased efflux from roots under elevated atmospheric [CO2] would occur if growth of the basal lateral roots were enhanced. However, given that the exudation pulse was shorter, this is by no means certain. The white lupin system illustrates the transient nature of exudation, its dependence on a specific stage of root development and time of day, and, even at peaks in efflux, the variability among samples from similar atmospheric CO2 environments. Whipps (1985) showed that carbon export per unit of length of maize root was unaltered by ambient CO2 treatment. Similarly, Norby et al. (1987) concluded that there was no consistent effect of elevated [CO2] on root exudation from pine seedlings, either per unit mass of fine root or as a percentage of photosynthate. Gifford et al. (1996), measuring citrate efflux per unit of dry weight of Danthonia root tips, were unable to detect a significant difference among atmospheric CO2treatments, although exudation was consistently greater at elevated [CO2]. DeLucia et al. (1997) detected an increase in oxalic acid in the rhizosphere of pine seedlings in response to increased atmospheric [CO2]; however, these investigators did not measure efflux per unit of root length and did not differentiate between root-derived and hyphae-derived exudates. Using stable-isotope techniques, Hungate et al. (1997) showed that the total labile carbon pool, including root exudation and respiration, was larger in grassland growing with elevated atmospheric [CO2]. In conclusion, with the limited amount of data available, it is not possible to conclude that elevated atmospheric [CO2] alters the rate of carbon exudation from a given length of root. However, if root production were to be enhanced by elevated atmospheric [CO2], then overall exudation could also be enhanced. ACKNOWLEDGMENTS We are grateful to Gamini Keerthisinghe (Commonwealth Scientific and Industrial Research Organization Plant Industry, Canberra, Australia) for help with establishing the hydroponics and exudate collection system and to Charlie Hocart (Research School of Biological Sciences, Australian National University) for help with the HPLC analyses. Abbreviation: PEPC PEP carboxylase LITERATURE CITED 1 Berntson GM Bazzaz FA The allometry of root production and loss in seedlings of Acer rubrum (Aceraceae) and Betula papyrifera (Betulaceae): implications for root dynamics in elevated CO2. Am J Bot 83 1996 608 616 Google Scholar Crossref Search ADS WorldCat 2 Boerjan W Cervera MT Delarue M Beeckman T Dewitte W Bellini C Caboche M Van Onckelen H Van Montagu M Inzé D superroot, a recessive mutation in Arabidopsis, confers auxin overproduction. 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Physiol Plant 49 1980 304 314 Google Scholar Crossref Search ADS WorldCat Author notes 1 This study was funded in part by an Overseas Postgraduate Award to M.W. from the Australian Government. * Corresponding author; e-mail [email protected]; fax 61–2–6249–4919. 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)

Lancien, Muriel; Ferrario-Méry, Sylvie; Roux, Yvette; Bismuth, Evelyne; Masclaux, Céline; Hirel, Bertrand; Gadal, Pierre; Hodges, Michael

doi: 10.1104/pp.120.3.717pmid: 10398706

Abstract Mitochondrial NAD-dependent (IDH) and cytosolic NADP-dependent isocitrate dehydrogenases have been considered as candidates for the production of 2-oxoglutarate required by the glutamine synthetase/glutamate synthase cycle. The increase in IDH transcripts in leaf and root tissues, induced by nitrate or NH4+ resupply to short-term N-starved tobacco (Nicotiana tabacum) plants, suggested that this enzyme could play such a role. The leaf and root steady-state mRNA levels of citrate synthase, acotinase, IDH, and glutamine synthetase were found to respond similarly to nitrate, whereas those for cytosolic NADP-dependent isocitrate dehydrogenase and fumarase responded differently. This apparent coordination occurred only at the mRNA level, since activity and protein levels of certain corresponding enzymes were not altered. Roots and leaves were not affected to the same extent either by N starvation or nitrate addition, the roots showing smaller changes in N metabolite levels. After nitrate resupply, these organs showed different response kinetics with respect to mRNA and N metabolite levels, suggesting that under such conditions nitrate assimilation was preferentially carried out in the roots. The differential effects appeared to reflect the C/N status after N starvation, the response kinetics being associated with the nitrate assimilatory capacity of each organ, signaled either by nitrate status or by metabolite(s) associated with its metabolism. Nitrate and NH4+assimilation depends on C metabolism for energy, reducing power, and C skeletons. An important site for the production of critical organic acids, and perhaps ATP and reductant especially in nongreen tissues, is the mitochondria. In higher plants the major NH4+ assimilatory pathway is carried out in the plastids by the concerted action of GS and GOGAT. The GS/GOGAT cycle requires the input of an organic acid in the form of 2-OG to produce Glu. However, the presence of several different 2-OG-producing enzymes, as well as their isoenzymatic forms, means that this substrate is potentially synthesized in a number of subcellular compartments, including mitochondria, cytosol, and plastids. This has led to a poor understanding of how 2-OG is provided and how its provision is regulated for NH4+assimilation. In the literature it is often proposed that the 2-OG for the GS/GOGAT cycle is produced by either an isocitrate dehydrogenase (Gálvez and Gadal, 1995) or an Asp amino acid transferase (Lam et al., 1996). In the first case, there is a net synthesis of Glu, whereas in the second case, there is a production of Asp via an interconversion of C and amino compounds. Two isocitrate dehydrogenase enzymes can be distinguished that differ in their cofactor specificity: IDH (Lancien et al., 1998), a Krebs cycle enzyme that is restricted to the mitochondria, and ICDH. Several ICDH isoenzymes have been shown to be present in the plant, located in different subcellular compartments: the cytosol (Gálvez et al., 1996), chloroplasts (Gálvez et al., 1994), mitochondria (Gálvez et al., 1998), and peroxisomes (Yamazaki and Tolbert, 1970). Two hypotheses have been considered for the isocitrate dehydrogenase-dependent origin of 2-OG for N assimilation. Miflin and Lea (1980) suggested that this essential keto-acid is produced in the mitochondria by the IDH. However, according to Chen and Gadal (1990) citrate synthesized in the mitochondria is transferred to the cytosol where it is decarboxylated to 2-OG via the action of cytosolic ACO and ICDH (ICDH1). As yet, both hypotheses are poorly supported experimentally, but several observations have argued in favor of an ICDH1 origin. These include (a) a 2- to 3-fold higher export rate of citrate over 2-OG by isolated spinach mitochondria oxidizing malate (Hanning and Heldt, 1993), (b) an increase in ICDH1 transcript levels in NR mutants grown on high nitrate and the subsequent accumulation of 2-OG because of reduced nitrate assimilation (Scheible et al., 1997), and (c) an increase in ICDH1 transcript levels in detached potato leaves fed with nitrate (Fieuw et al., 1995). On the other hand, an attempt to show such a role for cytosolic ICDH using an antisense transgenic plant strategy has shown that a dramatic reduction of ICDH1 activity to less than 5% of the control plant level did not affect either plant growth and development or nitrogen metabolism under normal growth conditions (Kruse et al., 1998). To explain the lack of phenotype, these authors suggested that under such conditions either there was a contribution from other ICDH isoenzymatic activities or that the remaining ICDH1 activity was sufficient and that the rate of 2-OG synthesis was a nonlimiting step for N assimilation. Of course, it is possible that ICDH1 is not involved in the main pathway for 2-OG production for NH4+ assimilation. The interaction between C and N metabolisms in higher plant cells is governed by many regulatory factors. This need to coordinate C and N metabolisms is reflected by the complex interplay between signals involving C metabolism, such as Suc and light, and those associated with N metabolism, such as nitrate (Crawford, 1995; Lam et al., 1996). It has become well established that nitrate (or N-containing compounds derived from nitrate metabolism) increases the expression of several genes encoding enzymes involved in N metabolism: NR (Pouteau et al., 1989), nitrate reductase, and (Rastogi et al., 1993), GS and GOGAT (Redinbaugh and Campbell, 1993). Recently, it has been shown that nitrate (Scheible et al., 1997) and/or sugars (Morcuende et al., 1998) also lead to a stimulation of organic acid biosynthesis. The initial aim of this work was to investigate the role of an I(C)DH in 2-OG supply for amino acid synthesis during nitrate assimilation. The effect of N supplies (nitrate or ammonium) on I(C)DH transcript levels when supplied to short-term N-starved tobacco plants was carried out. It was found that only IDH was affected by both N sources in roots and leaves. The effect of nitrate resupply was chosen to see if the IDH changes were coordinated, at the transcript level, with other enzymes involved in N assimilation and 2-OG production. In this way it was found that nitrate supply led to a differential but coordinated response of several genes involved in N and C metabolisms in both roots and leaves. The analysis of metabolite levels was carried out to determine the signals involved in the observed simultaneous expression and to explain the observed differences between roots and leaves. MATERIALS AND METHODS Plants and Growth Conditions Tobacco (Nicotiana tabacum L. cv Xanthi) was grown aeroponically in a greenhouse under a 16-h day/8-h night photoperiod, with a supplementation by white fluorescent light to provide a PAR of 200 μmol photons m−2s−1. The day/night temperature was 22°C/18°C. The plants were supplied every 3 d during 2 weeks with nutrient solution A containing the Murashige and Skoog macroelements, microelements, and Fe-EDTA (Murashige and Skoog, 1962). Then, the plants were treated with N-deficient solution B (solution A without mineral N source, K was supplied in the form of 10 mm KCl) for 4 d. Finally, 2 h after the beginning of the photoperiod, the plants were supplied with a solution consisting of solution B either with 10 mmpotassium nitrate or 10 mm ammonium chloride added. Deveined leaves and roots were harvested at different times after the resupply of the N-containing solution. Plant tissues were fixed in liquid N2 and conserved at −80°C. Isolation of RNA and Northern Analyses Total RNA was extracted according to the method of Chirgwin et al. (1979) and Harpster et al. (1986). Northern analyses were carried out according to the standard procedures as described previously (Sambrook et al., 1989) using 20 μg of deveined leaf or root total RNA. DNA probes were generated, using the DNA fragments described in TableI, by random priming with the Nonaprimer kit (Pharmacia) and [32P]dCTP (Amersham). Specific 3′-noncoding region fragments were generated by PCR using Taq polymerase (Appligene, Illkirch, France) and the following oligonucleotide pairs: TGTCTGGGCAGACAAGAGG/TTGTAATTACGGACCTC and ATCCTGTAGCACAGAA/AGGATAGATACGCTA, for  ICDH1 and mtICDH, respectively. Hybridization and wash conditions were as described previously (Gálvez et al., 1996) with final washes from 1× SSC and 0.2% SDS to 0.2× SSC and 0.2% SDS, depending on the origin of the probe. Table I. DNA probes used for northern analyses Name . Description . Source (Reference) . IDHa 656-bpEcoRI-HindIII fragment Tobacco (Lancien et al., 1998) ICDH1 142-bp PCR fragment Tobacco (Gálvez et al., 1996) mtICDH 186-bp PCR fragment Tobacco (Gálvez et al., 1998) CS 400-bp EcoRI fragment Tobacco (Landschütze et al., 1995) ACO 2,000-bpNotI-XhoI fragment Tobacco (M. Surpili and B. Müller-Röber, unpublished data) FUM 18,000-bpBamHI fragment Potato (Nast and Müller-Röber, 1996) GS2 1,800-bp EcoRI fragment Tobacco (Becker et al., 1992) GS1 200-bp HindIII-EcoRI fragment Tobacco (Dubois et al., 1996) NR 1,600-bpEcoRI fragment Tobacco, nia2 (Calza et al., 1987) Name . Description . Source (Reference) . IDHa 656-bpEcoRI-HindIII fragment Tobacco (Lancien et al., 1998) ICDH1 142-bp PCR fragment Tobacco (Gálvez et al., 1996) mtICDH 186-bp PCR fragment Tobacco (Gálvez et al., 1998) CS 400-bp EcoRI fragment Tobacco (Landschütze et al., 1995) ACO 2,000-bpNotI-XhoI fragment Tobacco (M. Surpili and B. Müller-Röber, unpublished data) FUM 18,000-bpBamHI fragment Potato (Nast and Müller-Röber, 1996) GS2 1,800-bp EcoRI fragment Tobacco (Becker et al., 1992) GS1 200-bp HindIII-EcoRI fragment Tobacco (Dubois et al., 1996) NR 1,600-bpEcoRI fragment Tobacco, nia2 (Calza et al., 1987) Open in new tab Table I. DNA probes used for northern analyses Name . Description . Source (Reference) . IDHa 656-bpEcoRI-HindIII fragment Tobacco (Lancien et al., 1998) ICDH1 142-bp PCR fragment Tobacco (Gálvez et al., 1996) mtICDH 186-bp PCR fragment Tobacco (Gálvez et al., 1998) CS 400-bp EcoRI fragment Tobacco (Landschütze et al., 1995) ACO 2,000-bpNotI-XhoI fragment Tobacco (M. Surpili and B. Müller-Röber, unpublished data) FUM 18,000-bpBamHI fragment Potato (Nast and Müller-Röber, 1996) GS2 1,800-bp EcoRI fragment Tobacco (Becker et al., 1992) GS1 200-bp HindIII-EcoRI fragment Tobacco (Dubois et al., 1996) NR 1,600-bpEcoRI fragment Tobacco, nia2 (Calza et al., 1987) Name . Description . Source (Reference) . IDHa 656-bpEcoRI-HindIII fragment Tobacco (Lancien et al., 1998) ICDH1 142-bp PCR fragment Tobacco (Gálvez et al., 1996) mtICDH 186-bp PCR fragment Tobacco (Gálvez et al., 1998) CS 400-bp EcoRI fragment Tobacco (Landschütze et al., 1995) ACO 2,000-bpNotI-XhoI fragment Tobacco (M. Surpili and B. Müller-Röber, unpublished data) FUM 18,000-bpBamHI fragment Potato (Nast and Müller-Röber, 1996) GS2 1,800-bp EcoRI fragment Tobacco (Becker et al., 1992) GS1 200-bp HindIII-EcoRI fragment Tobacco (Dubois et al., 1996) NR 1,600-bpEcoRI fragment Tobacco, nia2 (Calza et al., 1987) Open in new tab Enzyme Activities Plant material was initially ground to a fine powder in liquid N2 using a mortar and pestle and then further ground in the appropriate extraction buffer with respect to the enzyme activity to be measured. After centrifugation at 20,000g, the supernatant gave the crude extract, which was used for subsequent enzyme activity measurements. ICDH activity was measured spectrophotometrically as the reduction of NADP at 340 nm, as described by Gálvez et al. (1996). Total GS was measured as the synthetase activity as described previously (O'Neal and Joy, 1973). GOGAT (Fd/NADH) activity was measured as described previously (Suzuki and Gadal, 1984) using Fd from spinach leaves or NADH. NR activity measurements were carried out in the presence of 5 mm EDTA according to the method ofFerrario-Méry et al. (1997). Metabolite Measurements Carbohydrates and organic acids were extracted with 1m perchloric acid and amino acids were extracted with 3% sulfosalycilic acid, both from dry matter of roots and de-veined leaves. Suc, Fru, and Glc were measured in the soluble fraction using the Boerhinger Mannheim Suc/d-Glc/d-Fru test kit, following the manufacturer's instructions. The nitrate content was measured as described by Cataldo et al. (1975), and the ammonium content was measured as described by Rochat and Boutin (1989). Total amino acids were quantified by the method of Rosen (1957). Separation and analysis of amino acids were done by ion-exchange chromatography as in Rochat and Boutin (1989). Citrate and 2-OG were assayed as described previously (Bergmeyer, 1965). SDS-PAGE and Western Analysis Crude protein extracts (50 μg) were separated on 12% SDS-polyacrylamide gels according to the method of Laemmli (1970). For western analyses, protein transfer onto nitrocellulose membranes and immunodetection were performed as in Gálvez et al. (1996), using a 500-fold dilution of the 33% ammonium sulfate precipitate of the rabbit antiserum containing IDHa polyclonal antibodies (Lancien et al., 1998). RESULTS Effect of N Supply on Isocitrate Dehydrogenase Steady-State mRNA Levels If a specific isocitrate dehydrogenase produces the 2-OG for NH4+ assimilation, it is possible that the addition of either nitrate or NH4+ to N-starved plants could affect the steady-state mRNA level of this enzyme. Therefore, mRNA levels of IDH and ICDH, taken at various times from roots and deveined leaves after the resupply of either 10 mm nitrate or 10 mm ammonium to aeroponically grown 4-d N-starved tobacco plants, were analyzed by northern blot. Figure1 shows the effect of the above-mentioned treatments on the mRNA levels of IDH, ICDH1, and mtICDH. To distinguish the two different ICDH mRNAs, specific 3′-noncoding region probes were used as described in Methods. To investigate IDH expression a NtIDHa probe was used, which encodes the catalytic IDH subunit shown to be necessary for IDH activity (Lancien et al., 1998). In the experiment shown in Figure 1, nitrate supply led to a significant increase in IDHa mRNA steady-state levels only in the roots (similar changes have also been reported for IDHb and IDHc regulatory subunit mRNA; Lancien et al., 1998). This was seen in the roots and deveined leaves when NH4+ was given to the plants, although the response was slower in the roots when compared with nitrate addition. In contrast, nitrate did not affect ICDH1 and mtICDH mRNA levels (Fig. 1), whereas the addition of NH4+ gave rise to an increase in ICDH1 (only in the roots) and mtICDH (only in the leaves) transcript levels. Since only the IDH steady-state mRNA level was affected by the addition of the two different inorganic N supplies, IDH appeared to be a good candidate for the production of 2-OG with respect to N assimilation. Fig. 1. Open in new tabDownload slide Effect of nitrate and ammonium supply on isocitrate dehydrogenase transcripts in tobacco. Deveined leaves and roots were harvested after a 4-d N starvation (0 h) and after the addition of a N source (lanes 15, 30, and 96 h). mRNA was probed for IDH, ICDH1, and mtICDH. Twenty micrograms of total mRNA was loaded in each lane, which was checked by ethidium bromide staining. Fig. 1. Open in new tabDownload slide Effect of nitrate and ammonium supply on isocitrate dehydrogenase transcripts in tobacco. Deveined leaves and roots were harvested after a 4-d N starvation (0 h) and after the addition of a N source (lanes 15, 30, and 96 h). mRNA was probed for IDH, ICDH1, and mtICDH. Twenty micrograms of total mRNA was loaded in each lane, which was checked by ethidium bromide staining. If there is a major pathway leading to C skeleton production for N assimilation, the genes associated with this pathway could respond in a similar manner and be coordinated with N assimilation genes. Therefore, it was decided to investigate further the effect of nitrate resupply to the N-starved plants to see if similar changes to those observed for the IDH could be seen for enzymes associated with N assimilation and organic acid synthesis. Differential but Coordinated Expression of Some Genes Encoding Enzymes Involved in C or N Metabolisms between Roots and Leaves during Nitrate Resupply The effect of nitrate resupply to 4-d N-starved tobacco plants was investigated in roots (Fig. 2) and deveined leaves (Fig. 3) by northern analyses using DNA probes for N-assimilatory enzymes (NR and GS), certain Krebs cycle and putative organic acid pathway enzymes (IDH, CS, ACO, FUM, and ICDH1). Fig. 2. Open in new tabDownload slide Changes in steady-state transcript abundance in roots after nitrate resupply to 4-d N-starved tobacco plants. Northern analyses were performed at 0 h (after N starvation), and 3, 8, 23, and 32 h after nitrate resupply. An ethidium bromide (EtBr)-stained gel is presented to show loading in each lane. mRNAs were hybridized with probes for CS, ACO, IDH, FUM, ICDH1, NR, and GS1. Twenty micrograms of total mRNA was loaded in each lane. Fig. 2. Open in new tabDownload slide Changes in steady-state transcript abundance in roots after nitrate resupply to 4-d N-starved tobacco plants. Northern analyses were performed at 0 h (after N starvation), and 3, 8, 23, and 32 h after nitrate resupply. An ethidium bromide (EtBr)-stained gel is presented to show loading in each lane. mRNAs were hybridized with probes for CS, ACO, IDH, FUM, ICDH1, NR, and GS1. Twenty micrograms of total mRNA was loaded in each lane. Fig. 3. Open in new tabDownload slide Changes in leaf steady-state transcript abundance in deveined leaves after nitrate resupply to 4-d-starved tobacco plants. Northern analyses were performed at 0 h (after N starvation), and 3, 8, 23, and 32 h after nitrate resupply. An ethidium bromide (EtBr)-stained gel is presented to show loading in each lane. mRNAs were hybridized with probes for CS, ACO, IDH, FUM, ICDH1, NR, and GS2 and GS1. Twenty micrograms of total mRNA was loaded in each lane. Fig. 3. Open in new tabDownload slide Changes in leaf steady-state transcript abundance in deveined leaves after nitrate resupply to 4-d-starved tobacco plants. Northern analyses were performed at 0 h (after N starvation), and 3, 8, 23, and 32 h after nitrate resupply. An ethidium bromide (EtBr)-stained gel is presented to show loading in each lane. mRNAs were hybridized with probes for CS, ACO, IDH, FUM, ICDH1, NR, and GS2 and GS1. Twenty micrograms of total mRNA was loaded in each lane. As expected, nitrate given to the N-starved plants induced a rapid and transient increase of the NR steady-state mRNA level in both roots and leaves. These changes were considered as a reference for a nitrate-induced gene response. In the roots (Fig. 2), this response was also seen for GS1, encoding a cytosolic GS (Dubois et al., 1996), and the enzymes involved in the first decarboxylative step of the Krebs cycle: CS, ACO, and IDH. However, in the leaves (Fig. 3), the transcript levels of these enzymes, as well as GS2, appeared to be coordinated but showed slower response kinetics with respect to NR. This coordinated response was not observed for either the FUM or the ICDH1 (Figs. 2 and 3). These results show that there was a different response to nitrate supply between the roots and the leaves of N-starved tobacco plants affecting the steady-state mRNA levels of specific enzymes involved in N and C metabolisms. Although the responses were characterized by different kinetics between the two organs, there appeared to be a simultaneous response between the IDH and several other genes that could suggest a role for this enzyme in organic acid supply to N assimilation. Therefore, it seemed that a regulatory control could occur to coordinate C skeleton partitioning for amino acid biosynthesis via the nitrate assimilatory pathway. To establish a possible correlation between gene expression and plant metabolic status of the different organs, an analysis of certain metabolite levels and enzymatic capacities was undertaken. The Effect of N Starvation on Plant Metabolism It is interesting to note that N starvation already affected the roots and the leaves to different extents with respect to N- and C-containing metabolites. As expected, the 4-d treatment caused a drastic decrease in stored endogenous nitrate in both leaves and roots (Fig. 4), leading to undetectable levels in the leaves, whereas a small amount remained in the roots. In general, leaves showed a larger decrease in the content of measured amino acids than roots. This was particularly pronounced for Gln, Asp, Asn, and Ser (compare leaves and roots in Fig.5). On the other hand, the roots showed a greater increase in sugar levels, especially Suc and Fru (Fig.6). N starvation also resulted in a differential response with respect to the 2-OG pool, since it increased in the leaves but decreased in the roots (Fig. 6). Fig. 4. Open in new tabDownload slide The effect of nitrate resupply on inorganic N (NH4+ and NO3−), amino acid content, and NR activity. Deveined leaves and roots were harvested from 4-d N-starved plants (0 h) and after nitrate resupply (3, 8, 23, 32 h), as shown by the shaded histograms. Control, nitrate-fed plants are shown by the nonshaded histograms. Maximal NR activities (nmol−1 h−1 μg−1 protein [prot]) were measured in the presence of EDTA. Values are the means of two independent experiments for nitrate supply experiments, whereas the controls are a single experimental series. Data from each individual experiment are the averages of material taken from four independent plants. DW, Dry weight. Fig. 4. Open in new tabDownload slide The effect of nitrate resupply on inorganic N (NH4+ and NO3−), amino acid content, and NR activity. Deveined leaves and roots were harvested from 4-d N-starved plants (0 h) and after nitrate resupply (3, 8, 23, 32 h), as shown by the shaded histograms. Control, nitrate-fed plants are shown by the nonshaded histograms. Maximal NR activities (nmol−1 h−1 μg−1 protein [prot]) were measured in the presence of EDTA. Values are the means of two independent experiments for nitrate supply experiments, whereas the controls are a single experimental series. Data from each individual experiment are the averages of material taken from four independent plants. DW, Dry weight. Fig. 5. Open in new tabDownload slide The effect of nitrate addition on the pool of major amino acids, Gln, Glu, Asp, Asn, Gly, and Ser in deveined leaves and roots. Results are shown relative to dry weight (DW). A calculation relative to total amino acid content did not change the observed amino acid variations. N-starved/nitrate-resupplied plants are shown by the shaded histograms; control nitrate-fed plants are depicted by the nonshaded histograms. Values are the means of two independent experiments for nitrate supply experiments, whereas the controls are single experimental series. Data from each individual experiment are the averages of material taken from four independent plants. Fig. 5. Open in new tabDownload slide The effect of nitrate addition on the pool of major amino acids, Gln, Glu, Asp, Asn, Gly, and Ser in deveined leaves and roots. Results are shown relative to dry weight (DW). A calculation relative to total amino acid content did not change the observed amino acid variations. N-starved/nitrate-resupplied plants are shown by the shaded histograms; control nitrate-fed plants are depicted by the nonshaded histograms. Values are the means of two independent experiments for nitrate supply experiments, whereas the controls are single experimental series. Data from each individual experiment are the averages of material taken from four independent plants. Fig. 6. Open in new tabDownload slide The effect of nitrate addition on the pool of organic acids (citrate and 2-OG) and sugars (Glc, Fru, and Suc) in deveined leaves and roots. N-starved/nitrate-resupplied plants are shown by the shaded histograms; control nitrate-fed plants are depicted by the nonshaded histograms. Values are the means of two independent experiments for nitrate supply experiments, whereas the controls are single experiments. Data from each individual experiment are the averages of material taken from four independent plants. DW, Dry weight. Fig. 6. Open in new tabDownload slide The effect of nitrate addition on the pool of organic acids (citrate and 2-OG) and sugars (Glc, Fru, and Suc) in deveined leaves and roots. N-starved/nitrate-resupplied plants are shown by the shaded histograms; control nitrate-fed plants are depicted by the nonshaded histograms. Values are the means of two independent experiments for nitrate supply experiments, whereas the controls are single experiments. Data from each individual experiment are the averages of material taken from four independent plants. DW, Dry weight. In both roots and leaves, N starvation led to a decrease in NR activity (Fig. 4) but no significant changes were seen in either the in vitro total GS and ICDH activities (Fig. 7, compare + and 0). Fig. 7. Open in new tabDownload slide The effect of nitrate addition on GS, Fd-dependent GOGAT (GOGAT), and ICDH activities in root (A) and deveined leaf (B) extracts. Values are the average of two independent experiments. + corresponds to activities from control plants grown on nitrate and harvested at 0 h. Data from each individual experiment are the average of material taken from four independent plants. Fig. 7. Open in new tabDownload slide The effect of nitrate addition on GS, Fd-dependent GOGAT (GOGAT), and ICDH activities in root (A) and deveined leaf (B) extracts. Values are the average of two independent experiments. + corresponds to activities from control plants grown on nitrate and harvested at 0 h. Data from each individual experiment are the average of material taken from four independent plants. A Differential Effect of Nitrate Supply on N Metabolism between Leaves and Roots Nitrate resupply after N starvation led to a rapid accumulation of nitrate in the roots, which was mirrored by an increase in NR activity (Fig. 4). The induced changes allowed the roots to attain the control plant levels during the first 8 h after nitrate supply. These changes were accompanied by similar rapid increases in measured amino acid levels, including Glu, Gln, and Asp (Fig. 5). Gln reached and exceeded the control level 8 h after resupply. No significant changes were observed in GS and Fd-GOGAT activities (Fig. 7A); however, there was an increase in NADH-GOGAT activity from 10 to 390 nmol NADH oxidized h−1mg−1 protein, reaching 30% of the total GOGAT capacity 32 h after nitrate addition. In the leaves the differential kinetic response seen at the mRNA level with respect to the roots was also observed at the N metabolism level. Nitrate supply led to a much slower accumulation of nitrate, which like the NR activity, increased to reach the control plant level only 23 h after resupply (Fig. 4). Similar kinetics were also observed for the NH4+ content (Fig. 4) and to a lesser extent for the Gln content, both of which returned to the control plant levels (Fig. 5). However, the other measured amino acids did not increase significantly, staying close to the low N-starved levels (Fig. 5). As in the roots, the in vitro GS/GOGAT cycle capacity was not modified (Fig. 7B) and an NADH-GOGAT activity was induced to give an activity of 164 nmol NADPH oxidized h−1 mg−1 protein 32 h after the nitrate addition (10% of the total leaf GOGAT activity). A Differential Effect of Nitrate Supply on C Metabolism between Leaves and Roots The resupply of N-starved plants with nitrate also led to a differential response between the roots and the leaves with respect to organic acid levels and to a lesser extent with respect to sugar content (Fig. 6). After nitrate supply, the citrate content slowly increased, whereas 2-OG levels decreased in the leaf tissues, both reaching the control plant levels. However, in the roots citrate and 2-OG contents decreased to levels lower than the control plants. Nitrate addition also led to a decrease in Glc and Fru levels in both roots and leaves; in all cases the level remained higher than in the control plants. A differential effect between the roots and leaves was observed for the Suc content that stayed at the nonstarved level in the leaves but remained higher than the control level in the roots. IDH protein content (Fig. 8) and total ICDH activity (Fig. 7) were unchanged after nitrate resupply, staying at the control plant level in both leaves and roots. Fig. 8. Open in new tabDownload slide The effect of nitrate resupply on IDH protein content in roots (A) and leaves (B). Antibodies raised against recombinant IDHa protein were used to detect IDH in 50 μg of crude protein extracts. Control, Nitrate-grown plant extracts. Resupply corresponds to 4-d N-starved plants re-fed with 10 mmnitrate. Fig. 8. Open in new tabDownload slide The effect of nitrate resupply on IDH protein content in roots (A) and leaves (B). Antibodies raised against recombinant IDHa protein were used to detect IDH in 50 μg of crude protein extracts. Control, Nitrate-grown plant extracts. Resupply corresponds to 4-d N-starved plants re-fed with 10 mmnitrate. DISCUSSION 2-OG Production for N Assimilation The hypothesis that cytosolic ICDH plays an important role in the production of C skeletons for NH4+ assimilation (Chen and Gadal, 1990) has recently been supported indirectly by several observations. These include an increase in ICDH1 transcript levels and 2-OG levels by the addition of nitrate to NR tobacco mutants (Scheible et al., 1997) and increased ICDH1 transcript levels in detached potato leaves fed with nitrate (Fieuw et al., 1995). However, in the first case (Scheible et al., 1997), the ICDH activity was not measured, and in the second example, the changes in transcript levels could not be correlated with the measured ICDH activity (Fieuw et al., 1995). In this work the addition of 10 mm nitrate to 4-d N-starved tobacco plants did not give rise to significant increases in leaf ICDH1 transcript levels (Figs. 1 and 2), although root ICDH1 mRNA occasionally showed a slow increase (Fig. 2). However, in both organs total ICDH activity did not change after nitrate resupply (Fig. 7). The apparent contradiction with the above-mentioned works could be explained by the different experimental models/conditions used. The comparison of results obtained using slow-growing NR mutants containing high nitrate and low Gln levels (Scheibel et al., 1997), nutrient-fed detached leaves (Fieuw et al., 1995), and N-starved plants containing low nitrate and low Gln levels is probably imprudent. However, in our case it is not possible to say that ICDH1 does not play a role in N metabolism, because it was also seen that our N starvation and N resupply conditions did not alter the in vitro GS and GOGAT enzymatic capacities (Fig. 7). An absence of any modification in the GS/GOGAT cycle capacity could be explained by the fact that nitrate assimilation only produces a small fraction of the total NH4+ assimilated by a leaf (Keys et al., 1978). Although it appeared that photorespiration was reduced by N starvation (as judged by the changes in Gly, Ser, and NH4+ levels after the treatment), it was possible that sufficient activity remained to justify the stability of leaf GS/GOGAT capacity. The absence of an increase in ICDH1 activity could also reflect the fact that 2-OG production is a nonlimiting step for the GS/GOGAT cycle, as suggested by the absence of a phenotype in antisense potato (Kruse et al., 1998) and tobacco (S. Gálvez and M. Hodges, unpublished observations) severely inhibited in ICDH1 activity. It has already been well documented that nitrate addition can induce an increase in NR, nitrite reductase, GS, and Fd-GOGAT transcript levels (see the introduction). On the other hand, NH4+ has been shown to decrease NR levels (Hoff et al., 1994) and increase only the transcript levels of enzymes involved in its assimilation (e.g. GS; Hirel et al., 1987). Therefore, it is interesting that the addition of either nitrate or NH4+ to N-starved tobacco plants led to an increase in IDH transcript levels, both in roots and leaves. Although, NH4+ did produce an increase in mtICDH and ICDH1 mRNA levels in the leaves and roots, respectively. These observations concerning IDH could reflect a role in 2-OG production for N assimilation. However, as with total GS activity, the increase in IDH mRNA levels was not mirrored by an increase in IDH protein levels, as judged by western analyses (Fig. 8). This shows that the nitrate-induced effect, presumably at the transcriptional level, does not necessarily lead to a similar response at the protein/activity level, perhaps reflecting a control at the translational level. Further indications for a possible role of IDH in nitrate assimilation are seen by the coordinated changes in steady-state transcript levels between IDH, NR and GS1 (in the roots), and GS2 (in the leaves). This could be because of the important role of Krebs cycle activity in general cell functioning, in which case it might be predicted that all Krebs cycle genes would be modified. However, this was not observed because only CS, mitochondrial ACO, and IDH mRNA levels were equally affected, whereas FUM transcripts were unaffected by the addition of nitrate, this occurring in both the roots and the leaves. From these results, it is clear that nitrate addition exerts a control on certain Krebs cycle enzyme genes in a manner that suggests the induction of an organic acid production pathway for N assimilation. Recently, using NR mutants, PEP carboxylase, and CS mRNA levels and organic acid content were shown to increase when nitrate accumulates (Scheible et al., 1997), and it was proposed that nitrate induced organic acid production for nitrate assimilation via the ICDH1 pathway. However, in this work IDH was not investigated. Differential Response between the Roots and Leaves The experimental conditions used in this work provoked a differential response between the roots and the leaves both during the N-starvation period and after the addition of nitrate. First, the withdrawal of N supply led to less-pronounced changes in N-containing metabolite levels in the roots than in the leaves; this was seen for nitrate, NH4+, and amino acid contents (Figs. 4 and 5). It seemed that leaf N metabolism was down-regulated and export of N-containing compounds to the roots was up-regulated by the N stress. As a consequence, such changes would favor root growth under “N-limiting” conditions. It has already been shown that a moderate N deficiency stimulates root growth (Agren and Ingestad, 1987). It has been suggested that under such conditions root growth would be selectively stimulated by a regulation of shoot/root allocation and that roots would recruit more amino acids, which have been shown to cycle between the leaves and the roots (Cooper and Clarkson, 1989). Under our experimental conditions the root contained higher sugar levels both after and during the N-starvation period. Such changes could reflect the requirement of an increased Krebs cycle activity to sustain cell metabolic functions, including organic acid synthesis for N-metabolism. The decrease in the root 2-OG level could be explained by an increased demand for C skeletons for N assimilation. On the other hand, the increase in leaf 2-OG levels after N starvation might reflect the observed decrease in nitrate assimilatory capacity. Therefore, the level of 2-OG seems to reflect the changes in N assimilatory capacity and C/N status in the different organs. Perhaps, the 2-OG content, alone or in association with a N compound like the Gln, allows the cell to sense the C/N status and signal the need to regulate/coordinate C and N metabolic pathways. Such a signaling pathway is well documented in bacteria in which Gln and 2-OG levels regulate the so-called two-component system (Jiang et al., 1997). The addition of nitrate to the N-starved plants also produced a differential effect on the roots and leaves. This could be explained by a preferential assimilation of the newly supplied nitrate in the roots. Under normal growth conditions, nitrate is mainly exported to the leaves to be assimilated but it appears that under our conditions the transport of nitrate from the roots could be either inhibited or drastically reduced. The proposed stimulation of root nitrate assimilation is suggested by the observed rapid increase in NR activity and amino acid levels (especially Gln) and the decrease in sugar and 2-OG levels. It is possible that during this time the root exports Gln to the leaves, thus explaining why the leaf Gln content increased, whereas there was no significant change in nitrate accumulation, NR activity, and the content of the other measured amino acids. The changes in leaf metabolite levels after nitrate addition suggest a much slower up-regulation in leaf N metabolism with respect to that measured in the root. On the whole, the kinetics of mRNA accumulation and the up-regulation of nitrate assimilation capacity seem to be similar. Therefore, in both organs the effect on the observed transcript levels (except for leaf NR) could be explained by a similar signaling pathway involving either nitrate directly or a nitrate assimilation derived metabolite. In the roots nitrate accumulated and appeared to be metabolized more rapidly than in the leaves. However, since IDH transcript levels were found also to respond to NH4+ supply in both roots and leaves, it could be that a downstream signal linked to nitrate assimilation might be involved to control the observed coordination. In the leaves only the NR was rapidly affected by the addition of nitrate (mRNA and enzymatic capacity). This could be because of a specific signaling pathway between the roots and the leaves. NR is known to be responsive to cytokinins (Vincentz et al., 1993), and recent advances have identified a related action of nitrate and cytokinins in the metabolic cross-talk between roots and leaves. A response regulator homologous to the bacterial two-component system has been isolated in maize leaves and has been assumed to be involved in the early response and signal transduction pathway involved in the inorganic N supply transduction, mediated by cytokinins (Sakakibara et al., 1998). Perhaps, such a signaling pathway could have triggered the rapid leaf NR response. 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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)

Verhoeven, Amy S.; Adams, William W.; Demmig-Adams, Barbara; Croce, Roberta; Bassi, Roberto

doi: 10.1104/pp.120.3.727pmid: 10398707

Abstract The distribution of xanthophyll cycle pigments (violaxanthin plus antheraxanthin plus zeaxanthin [VAZ]) among photosynthetic pigment-protein complexes was examined in Vinca majorbefore, during, and subsequent to a photoinhibitory treatment at low temperature. Four pigment-protein complexes were isolated: the core of photosystem (PS) II, the major light-harvesting complex (LHC) protein of PSII (LHCII), the minor light-harvesting proteins (CPs) of PSII (CP29, CP26, and CP24), and PSI with its LHC proteins (PSI-LHCI). In isolated thylakoids 80% of VAZ was bound to protein independently of the de-epoxidation state and was found in all complexes. Plants grown outside in natural sunlight had higher levels of VAZ (expressed per chlorophyll), compared with plants grown in low light in the laboratory, and the additional VAZ was mainly bound to the major LHCII complex, apparently in an acid-labile site. The extent of de-epoxidation of VAZ in high light and the rate of reconversion of Z plus A to V following 2.5 h of recovery were greatest in the free-pigment fraction and varied among the pigment-protein complexes. Photoinhibition caused increases in VAZ, particularly in low-light-acclimated leaves. The data suggest that the photoinhibitory treatment caused an enrichment in VAZ bound to the minor CPs caused by de novo synthesis of the pigments and/or a redistribution of VAZ from the major LHCII complex. Photoinhibition refers to a condition in which a persistent decrease in the efficiency of photosynthetic energy conversion in leaves is observed. Photoinhibition occurs in the field in plants exposed to conditions of high light in combination with environmental stress, such as cold temperatures, but can also be induced by exposure of shade-acclimated leaves to high light (Krause, 1994; Osmond, 1994). Under various photoinhibitory conditions large quantities of the xanthophyll cycle pigments Z and A have been found to be retained in leaves for extended periods after darkening (Demmig et al., 1988; Adams et al., 1995; Verhoeven et al., 1996; Demmig-Adams et al., 1998). The xanthophyll cycle pigments Z and A are formed from V when light is excessive, and they are involved in a photoprotective process whereby excess absorbed excitation energy is dissipated thermally in the light-harvesting antennae of PSII (Demmig-Adams and Adams, 1996;Eskling et al., 1997; Gilmore, 1997). The retention of Z plus A in photoinhibited leaves often correlates closely with sustained low PSII efficiencies measured as theFv/Fm (Adams et al., 1995; Verhoeven et al., 1996; Demmig-Adams et al., 1998, and refs. therein). Such correlations have led to the suggestion that Z plus A may be engaged for thermal energy dissipation under these conditions and may therefore be involved in the reduced PSII efficiencies observed. To influence Chl fluorescence yield xanthophylls must be localized in close proximity to the pigment-protein complexes of the thylakoid membrane, and knowledge of their precise organization is important to understand the mechanism of (Z plus A)-dependent energy dissipation. Several studies have demonstrated that the xanthophyll cycle pigments (VAZ) are associated with all light-harvesting components, including LHCI (Thayer and Björkman, 1992; Lee and Thornber, 1995). Among LHCII components, VAZ has been reported to be enriched in the minor CPs (CP29, CP26, and CP24) relative to the major LHC of PSII, LHCII (Bassi et al., 1993; Ruban et al., 1994; Lee and Thornber, 1995; Goss et al., 1997), suggesting an important role for the minor CPs in photoprotection (Bassi et al., 1993; Gilmore, 1997). Upon illumination, V is apparently converted to Z in all complexes (Ruban et al., 1994;Lee and Thornber, 1995; Phillip and Young, 1995; Färber et al., 1997; Zhu et al., 1997), with the degree of epoxidation varying among the different LHCs (Ruban et al., 1994; Croce et al., 1996a). Although several studies have been conducted to examine the distribution of the xanthophyll cycle pigments among pigment-protein complexes isolated from unstressed leaves, very little is know about their distribution under photoinhibitory conditions when the rate of Z epoxidation is severely slowed following leaf darkening. The major goal of this study was to assess whether changes occur in the levels or distribution of VAZ in the pigment-protein complexes when plants are treated with such photoinhibitory high light, as well as whether differences exist in the rate at which Z and A are reconverted to V on the different pigment-proteins during the slow recovery from photoinhibition. MATERIALS AND METHODS Plant Material and Photoinhibitory Treatments Vinca major var variegata Loud. plants were acquired at a local greenhouse in northern Italy on two occasions, resulting in plants that were acclimated to two different growth conditions. One set of plants was grown at a low light intensity (approximately 50 μmol photons m−2s−1) provided by a combination of HQE fluorescent tubes and R80-Natura (Osram, Munich, Germany) incandescent bulbs. Plants were grown in soil, watered every 2 d, and received Nitsch's nutrient solution (Nitsch and Nitsch, 1969) once per week. The temperature was 22°C/28°C, night/day, with 80% RH. Plants were acclimated to low-light conditions for at least 6 weeks prior to treatment. A second set of plants was grown outside in northern Italy in September 1998, when the maximum light intensity at midday was approximately 1900 μmol photons m−2 s−1. The temperature in the last 6 weeks prior to experimentation was 18°C to 24°C (night)/26°C to 34°C (day). Plants were watered and received nutrients daily. For the photoinhibitory treatment, plants were exposed to continuous light and chilling temperatures (480 μmol photons m−2 s−1 at 15°C for the plants grown in GCs and 430 μmol photons m−2 s−1 at 10°C for the plants grown OD in full sunlight) until photoinhibition was observed, which was measured by moving a leaf into darkness and measuring Fv/Fmafter 30 min. Very slowly relaxingFv/Fm was achieved after 48 h of exposure to continuous light in plants grown in GCs and after 122 h of exposure to continuous light in plants grown OD. Three sets of leaves were harvested for thylakoid isolation: controls harvested after 12 h of darkness before each treatment, leaves harvested directly following high-light treatment, and leaves harvested after an additional 2.5 h of recovery in darkness at room temperature. In each case, all of the plant leaves were harvested for thylakoid isolation. Thylakoid Isolation Thylakoid membranes were isolated as described previously by Bassi et al. (1988) except the grinding buffer consisted of 0.1 mTricine, pH 7.8, 0.4 m sorbitol, 0.5% nonfat dried milk, 0.2 mm PMSF, 5 mmε-amino-n-caproic acid, and 1 mmbenzamidine; the washing buffer consisted of 25 mm Hepes/KOH, pH 7.5, and 10 mm EDTA; and the resuspension buffer consisted of 10 mm Hepes/KOH, pH 7.5, 1 mm EDTA, and 50% (v/v) glycerol. Chl Fluorescence Fluorescence measurements were performed on intact leaves under the respective growth PPFD conditions with a portable fluorometer (PAM-2000, Walz, Effeltrich, Germany). Fluorescence measurements and calculations were performed as described previously (Demmig-Adams and Adams, 1996; Demmig-Adams et al., 1996). Solubilization and Fractionation of Thylakoids and Identification of Pigment-Protein Complexes Thylakoids were resuspended in 2 mg Chl/mL and solubilized by adding an equal volume of 2.8% dodecyl maltoside in water. The sample was then vortexed for 20 s and put on ice for 1 min. The solubilized sample was spun for 2 min at 15,000g and 4°C and rapidly loaded onto a 0.1 to 1 m Suc gradient containing 10 mm Hepes, pH 7.6, and 0.06% dodecyl maltoside. The gradient was spun in a Beckman SW41 rotor at 39,000 rpm for 27 h at 4°C. Individual green fractions were harvested with a syringe. Pigment-protein complexes were identified using analytical SDS-PAGE, spectroscopy, and pigment analysis, as described previously for several different species (Di Paolo et al., 1990; Santini et al., 1994; Kilian et al., 1997). Electrophoresis and Immunoblotting Analytical SDS-PAGE was performed with gradient gels (10%–16% acrylamide) using the Tris/Tricine buffer system of Schägger and von Jagow (1987). Alternatively, a Tris-sulfate buffer system was used (Bassi et al., 1985a). Preparative IEF was performed as previously described (Dainese et al., 1990). Nondenaturing green gel electrophoresis was done according to the method of Knoetzel and Simpson (1991). For immunoblot assays, samples were separated by the Tris-sulfate gel system and transferred to a nitrocellulose filter (Millipore). The filters were incubated with antibodies and detected with alkaline phosphatase coupled to anti-rabbit IgG (Sigma). Analysis of Pigments Total pigments were extracted with 80% acetone. Analysis of the extracts by HPLC was as described previously (Gilmore and Yamamoto, 1991). In this study the determination of the Chl-to-carotenoid ratio in pigment proteins was of crucial importance. The results of HPLC analysis were therefore verified by fitting the absorption spectra of ethanolic extracts of pigment proteins with the spectra of pure pigments in ethanol (Connelly et al., 1997). Spectra (350–750 nm) were recorded using a DW2000 spectrophotometer in the split-beam mode (Aminco, Silver Spring, MD). Individual pigments were purified by HPLC, dried under a vacuum, and resuspended in 96% ethanol. Curve fitting was obtained by using a nonlinear least-squares fitting code (Origin, MicroCal Software, Northampton, MA). Verification of Specificity of Pigment Binding To verify the specificity of xanthophyll binding to the different pigment-protein complexes, an experiment was undertaken in which excess xanthophylls were added to solubilized thylakoids before their fractionation. No change in pigment composition of the protein complexes was observed after fractionation, suggesting that no nonspecific binding of the xanthophylls was caused by the solubilization conditions. DEAE Chromatography of the PSII Core The Suc-gradient fraction containing the PSII core was purified using DEAE chromatography as described previously (Giuffra et al., 1996). RESULTS Characterization of Pigment Content and Distribution prior to Photoinhibitory Treatment The pigment content of V. major varvariegata Loud. was examined in control conditions from both isolated thylakoids and whole-leaf extracts (TableI). In control conditions the differences in carotenoid levels, relative to Chl a, from plants grown in the GC relative to those grown OD involved predominantly the fraction of VAZ (Table I). In both isolated thylakoids and leaf extracts from plants grown OD, the content of V and VAZ was approximately twice that of the plants grown in GCs. A comparison of the pigment composition of whole-leaf extracts versus isolated thylakoids demonstrates that the ratios of total carotenoid and of VAZ to Chl were not significantly changed following thylakoid isolation. Table I. Pigment composition from both thylakoids and leaves of V. major grown either in GCs or OD under natural sunlight Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . mol/100 mol Chl a Chlb 39.8 (38.5) 39.1 (38.9) 37.1 (38.4) 39.1 (34.4) 40.1 (42.0) 38.4 (35.2) Neoxanthin 5.5 (5.6) 5.0 (4.8) 5.3 (5.7) 5.9 (5.9) 6.3 (5.9) 6.1 (5.6) Lutein 18.0 (18.3) 21.0 (23.6) 19.4 (19.6) 18.8 (18.8) 21.8 (21.7) 20.1 (19.6) β-Carotene 6.2 5.8 5.8 6.4 6.8 6.4 V 4.5 (4.4) 1.8 (2.4) 2.2 (2.3) 9.1 (9.6) 2.9 (3.5) 4.3 (4.6) A 0 (0) 1.2 (1.4) 1.3 (1.3) 0 (0) 2.6 (2.9) 2.5 (3.2) Z 0 (0) 4.2 (5.2) 3.5 (3.6) 0 (0) 5.4 (5.3) 3.2 (3.8) VAZ 4.5 (4.4) 7.2 (9.0) 7.0 (7.2) 9.1 (9.6) 10.9 (11.8) 10.1 (11.7) (Z+A)/(VAZ) 0 (0) 0.75 (0.73) 0.69 (0.68) 0 (0) 0.74 (0.70) 0.56 (0.60) Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . mol/100 mol Chl a Chlb 39.8 (38.5) 39.1 (38.9) 37.1 (38.4) 39.1 (34.4) 40.1 (42.0) 38.4 (35.2) Neoxanthin 5.5 (5.6) 5.0 (4.8) 5.3 (5.7) 5.9 (5.9) 6.3 (5.9) 6.1 (5.6) Lutein 18.0 (18.3) 21.0 (23.6) 19.4 (19.6) 18.8 (18.8) 21.8 (21.7) 20.1 (19.6) β-Carotene 6.2 5.8 5.8 6.4 6.8 6.4 V 4.5 (4.4) 1.8 (2.4) 2.2 (2.3) 9.1 (9.6) 2.9 (3.5) 4.3 (4.6) A 0 (0) 1.2 (1.4) 1.3 (1.3) 0 (0) 2.6 (2.9) 2.5 (3.2) Z 0 (0) 4.2 (5.2) 3.5 (3.6) 0 (0) 5.4 (5.3) 3.2 (3.8) VAZ 4.5 (4.4) 7.2 (9.0) 7.0 (7.2) 9.1 (9.6) 10.9 (11.8) 10.1 (11.7) (Z+A)/(VAZ) 0 (0) 0.75 (0.73) 0.69 (0.68) 0 (0) 0.74 (0.70) 0.56 (0.60) Leaves were collected following 12 h of darkness (control), at the end of the photoinhibitory treatment (stress), or following 2.5 h of recovery at low light (recovery). The leaf data are in parentheses, except β-carotene for which the leaf data are not available. SeeMethods for growth conditions and stress treatments. Open in new tab Table I. Pigment composition from both thylakoids and leaves of V. major grown either in GCs or OD under natural sunlight Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . mol/100 mol Chl a Chlb 39.8 (38.5) 39.1 (38.9) 37.1 (38.4) 39.1 (34.4) 40.1 (42.0) 38.4 (35.2) Neoxanthin 5.5 (5.6) 5.0 (4.8) 5.3 (5.7) 5.9 (5.9) 6.3 (5.9) 6.1 (5.6) Lutein 18.0 (18.3) 21.0 (23.6) 19.4 (19.6) 18.8 (18.8) 21.8 (21.7) 20.1 (19.6) β-Carotene 6.2 5.8 5.8 6.4 6.8 6.4 V 4.5 (4.4) 1.8 (2.4) 2.2 (2.3) 9.1 (9.6) 2.9 (3.5) 4.3 (4.6) A 0 (0) 1.2 (1.4) 1.3 (1.3) 0 (0) 2.6 (2.9) 2.5 (3.2) Z 0 (0) 4.2 (5.2) 3.5 (3.6) 0 (0) 5.4 (5.3) 3.2 (3.8) VAZ 4.5 (4.4) 7.2 (9.0) 7.0 (7.2) 9.1 (9.6) 10.9 (11.8) 10.1 (11.7) (Z+A)/(VAZ) 0 (0) 0.75 (0.73) 0.69 (0.68) 0 (0) 0.74 (0.70) 0.56 (0.60) Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . mol/100 mol Chl a Chlb 39.8 (38.5) 39.1 (38.9) 37.1 (38.4) 39.1 (34.4) 40.1 (42.0) 38.4 (35.2) Neoxanthin 5.5 (5.6) 5.0 (4.8) 5.3 (5.7) 5.9 (5.9) 6.3 (5.9) 6.1 (5.6) Lutein 18.0 (18.3) 21.0 (23.6) 19.4 (19.6) 18.8 (18.8) 21.8 (21.7) 20.1 (19.6) β-Carotene 6.2 5.8 5.8 6.4 6.8 6.4 V 4.5 (4.4) 1.8 (2.4) 2.2 (2.3) 9.1 (9.6) 2.9 (3.5) 4.3 (4.6) A 0 (0) 1.2 (1.4) 1.3 (1.3) 0 (0) 2.6 (2.9) 2.5 (3.2) Z 0 (0) 4.2 (5.2) 3.5 (3.6) 0 (0) 5.4 (5.3) 3.2 (3.8) VAZ 4.5 (4.4) 7.2 (9.0) 7.0 (7.2) 9.1 (9.6) 10.9 (11.8) 10.1 (11.7) (Z+A)/(VAZ) 0 (0) 0.75 (0.73) 0.69 (0.68) 0 (0) 0.74 (0.70) 0.56 (0.60) Leaves were collected following 12 h of darkness (control), at the end of the photoinhibitory treatment (stress), or following 2.5 h of recovery at low light (recovery). The leaf data are in parentheses, except β-carotene for which the leaf data are not available. SeeMethods for growth conditions and stress treatments. Open in new tab Distribution of VAZ among Pigment Proteins In a first approach to examining the location of pigments within the protein complexes, thylakoid membranes were solubilized with dodecyl maltoside, and Suc-gradient fractionation was performed, followed by analysis of the pigment and protein content of the fractions (Fig. 1; TablesII andIII). Five major fractions were obtained. Fig. 1. Open in new tabDownload slide Fully denaturing Tris-sulfate SDS-PAGE of the fractions obtained from Suc-gradient ultracentrifugation of V. major thylakoids collected from dark-adapted (at least 12 h) leaves of GC plants. Fractions from stressed and recovered thylakoids, as well as the fractions obtained from plants grown OD, were very similar and are therefore not depicted. Whole thylakoids (T) were loaded onto the gel in addition to the four fractions (F2–F5) collected; fraction 2 contained the LHCII monomer (B), the minor Chl proteins CP29 (A), CP26 (comigrating with LHCII; B), and CP24 (C); fraction 3 contained the LHCII trimer (D); fraction 4 contained the PSII core with the D1/D2 heterodimer (E), CP47 (F), CP43 (G), and D1/D2 monomer (H); fraction 5 contained the PSI core (I) and LHCI (J). Fraction 1 contained the free pigments (Table III) and is not depicted here. Fig. 1. Open in new tabDownload slide Fully denaturing Tris-sulfate SDS-PAGE of the fractions obtained from Suc-gradient ultracentrifugation of V. major thylakoids collected from dark-adapted (at least 12 h) leaves of GC plants. Fractions from stressed and recovered thylakoids, as well as the fractions obtained from plants grown OD, were very similar and are therefore not depicted. Whole thylakoids (T) were loaded onto the gel in addition to the four fractions (F2–F5) collected; fraction 2 contained the LHCII monomer (B), the minor Chl proteins CP29 (A), CP26 (comigrating with LHCII; B), and CP24 (C); fraction 3 contained the LHCII trimer (D); fraction 4 contained the PSII core with the D1/D2 heterodimer (E), CP47 (F), CP43 (G), and D1/D2 monomer (H); fraction 5 contained the PSI core (I) and LHCI (J). Fraction 1 contained the free pigments (Table III) and is not depicted here. Table II. Pigment composition of the free-pigment fractions (fraction 1) after Suc-gradient ultracentrifugation of solubilized thylakoid membranes Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . % in free pigment band % in free pigment band Chl a 1.4 1.4 1.1 0.7 1.3 1.4 Chl b 0.8 0.6 0.4 0.3 0.4 0.5 Neoxanthin 4.2 2.2 2.0 4.4 2.5 2.9 Lutein 9.3 11.9 8.7 5.2 8.6 9.2 β-Carotene 0 0 0 4.7 4.0 6.5 V 19.5 6.9 8.7 12.4 8.0 13.0 A 16.6 24.8 20.8 0 18.3 18.7 Z 1.3 23.3 16.3 0 16.3 13.4 VAZ 18.4 19.0 14.6 12.4 14.4 14.5 (Z+A)/(VAZ) 0.02 0.90 0.81 0.00 0.82 0.59 Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . % in free pigment band % in free pigment band Chl a 1.4 1.4 1.1 0.7 1.3 1.4 Chl b 0.8 0.6 0.4 0.3 0.4 0.5 Neoxanthin 4.2 2.2 2.0 4.4 2.5 2.9 Lutein 9.3 11.9 8.7 5.2 8.6 9.2 β-Carotene 0 0 0 4.7 4.0 6.5 V 19.5 6.9 8.7 12.4 8.0 13.0 A 16.6 24.8 20.8 0 18.3 18.7 Z 1.3 23.3 16.3 0 16.3 13.4 VAZ 18.4 19.0 14.6 12.4 14.4 14.5 (Z+A)/(VAZ) 0.02 0.90 0.81 0.00 0.82 0.59 Open in new tab Table II. Pigment composition of the free-pigment fractions (fraction 1) after Suc-gradient ultracentrifugation of solubilized thylakoid membranes Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . % in free pigment band % in free pigment band Chl a 1.4 1.4 1.1 0.7 1.3 1.4 Chl b 0.8 0.6 0.4 0.3 0.4 0.5 Neoxanthin 4.2 2.2 2.0 4.4 2.5 2.9 Lutein 9.3 11.9 8.7 5.2 8.6 9.2 β-Carotene 0 0 0 4.7 4.0 6.5 V 19.5 6.9 8.7 12.4 8.0 13.0 A 16.6 24.8 20.8 0 18.3 18.7 Z 1.3 23.3 16.3 0 16.3 13.4 VAZ 18.4 19.0 14.6 12.4 14.4 14.5 (Z+A)/(VAZ) 0.02 0.90 0.81 0.00 0.82 0.59 Pigment . V. major from GCs . V. major from OD . Control . Stress . Recovery . Control . Stress . Recovery . % in free pigment band % in free pigment band Chl a 1.4 1.4 1.1 0.7 1.3 1.4 Chl b 0.8 0.6 0.4 0.3 0.4 0.5 Neoxanthin 4.2 2.2 2.0 4.4 2.5 2.9 Lutein 9.3 11.9 8.7 5.2 8.6 9.2 β-Carotene 0 0 0 4.7 4.0 6.5 V 19.5 6.9 8.7 12.4 8.0 13.0 A 16.6 24.8 20.8 0 18.3 18.7 Z 1.3 23.3 16.3 0 16.3 13.4 VAZ 18.4 19.0 14.6 12.4 14.4 14.5 (Z+A)/(VAZ) 0.02 0.90 0.81 0.00 0.82 0.59 Open in new tab Table III. Pigment composition of the fractions collected from Suc-gradient ultracentrifugation of solubilized thylakoids from V. major grown in GCs or OD under natural sunlight Pigment and Treatment . Minor CPs + LHCII Monomer (Fraction 2) . LHCII Trimer (Fraction 3) . PSII Core (Fraction 4) . PSI-LHCI (Fraction 5) . GC . OD . GC . OD . GC . OD . GC . OD . mol/100 mol Chl a Chlb 68.0 ± 1.4  67.8 ± 0.3  78.8 ± 2.0  74.8 ± 2.7  5.5 ± 1.3 2.8 ± 0.8 11.5 ± 0.5  11.5 ± 0.2  Neoxanthin 10.6 ± 0.7  10.0 ± 0.7  12.7 ± 0.6  11.3 ± 0.9  0.6 ± 0.5 0.7 ± 0.3 0 0.3 ± 0.1 Lutein 28.8 ± 0.4  27.6 ± 1.3  34.8 ± 0.5  31.4 ± 1.7  4.7 ± 0.5 3.1 ± 0.4 6.5 ± 0.3 5.7 ± 0.6 β-Carotene 0 0.4 ± 0.3 0 0 17.0 ± 0.1  12.0 ± 2.9  9.9 ± 0.7 13.6 ± 0.5  V  Control 4.8 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 2.1 3.2 0.6 2.0 1.3 0.3 2.0 1.9  Recovery 2.5 5.0 1.0 3.3 1.0 0.7 2.3 2.4 A  Control 0.2 0 0 0 0 0 0 0  Stress 1.1 2.1 0.9 1.9 0.7 0.6 0.5 0.8  Recovery 1.4 3.0 1.0 1.9 0.9 0.8 0.6 0.8 Z  Control 0.7 0 0 0 0 0 0 0  Stress 5.2 5.9 2.8 4.8 1.2 1.2 1.3 1.3  Recovery 4.8 4.3 2.7 2.5 1.0 0.9 1.3 1.7 VAZ  Control 5.7 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 8.5 11.2 4.3 8.8 3.2 2.2 3.8 4.0  Recovery 8.7 12.3 4.7 7.6 2.8 2.4 4.2 4.8 (Z+A)/(VAZ)  Control 0.15 0 0 0 0 0 0 0  Stress 0.75 0.68 0.86 0.74 0.61 0.82 0.48 0.47  Recovery 0.72 0.55 0.78 0.53 0.66 0.69 0.44 0.46 Pigment and Treatment . Minor CPs + LHCII Monomer (Fraction 2) . LHCII Trimer (Fraction 3) . PSII Core (Fraction 4) . PSI-LHCI (Fraction 5) . GC . OD . GC . OD . GC . OD . GC . OD . mol/100 mol Chl a Chlb 68.0 ± 1.4  67.8 ± 0.3  78.8 ± 2.0  74.8 ± 2.7  5.5 ± 1.3 2.8 ± 0.8 11.5 ± 0.5  11.5 ± 0.2  Neoxanthin 10.6 ± 0.7  10.0 ± 0.7  12.7 ± 0.6  11.3 ± 0.9  0.6 ± 0.5 0.7 ± 0.3 0 0.3 ± 0.1 Lutein 28.8 ± 0.4  27.6 ± 1.3  34.8 ± 0.5  31.4 ± 1.7  4.7 ± 0.5 3.1 ± 0.4 6.5 ± 0.3 5.7 ± 0.6 β-Carotene 0 0.4 ± 0.3 0 0 17.0 ± 0.1  12.0 ± 2.9  9.9 ± 0.7 13.6 ± 0.5  V  Control 4.8 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 2.1 3.2 0.6 2.0 1.3 0.3 2.0 1.9  Recovery 2.5 5.0 1.0 3.3 1.0 0.7 2.3 2.4 A  Control 0.2 0 0 0 0 0 0 0  Stress 1.1 2.1 0.9 1.9 0.7 0.6 0.5 0.8  Recovery 1.4 3.0 1.0 1.9 0.9 0.8 0.6 0.8 Z  Control 0.7 0 0 0 0 0 0 0  Stress 5.2 5.9 2.8 4.8 1.2 1.2 1.3 1.3  Recovery 4.8 4.3 2.7 2.5 1.0 0.9 1.3 1.7 VAZ  Control 5.7 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 8.5 11.2 4.3 8.8 3.2 2.2 3.8 4.0  Recovery 8.7 12.3 4.7 7.6 2.8 2.4 4.2 4.8 (Z+A)/(VAZ)  Control 0.15 0 0 0 0 0 0 0  Stress 0.75 0.68 0.86 0.74 0.61 0.82 0.48 0.47  Recovery 0.72 0.55 0.78 0.53 0.66 0.69 0.44 0.46 The pigments Chl b, neoxanthin, lutein, and β-carotene are means ± sd of the control, stress (48 h of continuous light at 15°C for leaves from GC plants and 122 h of continuous light at 10°C for leaves from plants grown OD), and recovery (2.5 h in low light at 22°C) treatments. Open in new tab Table III. Pigment composition of the fractions collected from Suc-gradient ultracentrifugation of solubilized thylakoids from V. major grown in GCs or OD under natural sunlight Pigment and Treatment . Minor CPs + LHCII Monomer (Fraction 2) . LHCII Trimer (Fraction 3) . PSII Core (Fraction 4) . PSI-LHCI (Fraction 5) . GC . OD . GC . OD . GC . OD . GC . OD . mol/100 mol Chl a Chlb 68.0 ± 1.4  67.8 ± 0.3  78.8 ± 2.0  74.8 ± 2.7  5.5 ± 1.3 2.8 ± 0.8 11.5 ± 0.5  11.5 ± 0.2  Neoxanthin 10.6 ± 0.7  10.0 ± 0.7  12.7 ± 0.6  11.3 ± 0.9  0.6 ± 0.5 0.7 ± 0.3 0 0.3 ± 0.1 Lutein 28.8 ± 0.4  27.6 ± 1.3  34.8 ± 0.5  31.4 ± 1.7  4.7 ± 0.5 3.1 ± 0.4 6.5 ± 0.3 5.7 ± 0.6 β-Carotene 0 0.4 ± 0.3 0 0 17.0 ± 0.1  12.0 ± 2.9  9.9 ± 0.7 13.6 ± 0.5  V  Control 4.8 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 2.1 3.2 0.6 2.0 1.3 0.3 2.0 1.9  Recovery 2.5 5.0 1.0 3.3 1.0 0.7 2.3 2.4 A  Control 0.2 0 0 0 0 0 0 0  Stress 1.1 2.1 0.9 1.9 0.7 0.6 0.5 0.8  Recovery 1.4 3.0 1.0 1.9 0.9 0.8 0.6 0.8 Z  Control 0.7 0 0 0 0 0 0 0  Stress 5.2 5.9 2.8 4.8 1.2 1.2 1.3 1.3  Recovery 4.8 4.3 2.7 2.5 1.0 0.9 1.3 1.7 VAZ  Control 5.7 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 8.5 11.2 4.3 8.8 3.2 2.2 3.8 4.0  Recovery 8.7 12.3 4.7 7.6 2.8 2.4 4.2 4.8 (Z+A)/(VAZ)  Control 0.15 0 0 0 0 0 0 0  Stress 0.75 0.68 0.86 0.74 0.61 0.82 0.48 0.47  Recovery 0.72 0.55 0.78 0.53 0.66 0.69 0.44 0.46 Pigment and Treatment . Minor CPs + LHCII Monomer (Fraction 2) . LHCII Trimer (Fraction 3) . PSII Core (Fraction 4) . PSI-LHCI (Fraction 5) . GC . OD . GC . OD . GC . OD . GC . OD . mol/100 mol Chl a Chlb 68.0 ± 1.4  67.8 ± 0.3  78.8 ± 2.0  74.8 ± 2.7  5.5 ± 1.3 2.8 ± 0.8 11.5 ± 0.5  11.5 ± 0.2  Neoxanthin 10.6 ± 0.7  10.0 ± 0.7  12.7 ± 0.6  11.3 ± 0.9  0.6 ± 0.5 0.7 ± 0.3 0 0.3 ± 0.1 Lutein 28.8 ± 0.4  27.6 ± 1.3  34.8 ± 0.5  31.4 ± 1.7  4.7 ± 0.5 3.1 ± 0.4 6.5 ± 0.3 5.7 ± 0.6 β-Carotene 0 0.4 ± 0.3 0 0 17.0 ± 0.1  12.0 ± 2.9  9.9 ± 0.7 13.6 ± 0.5  V  Control 4.8 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 2.1 3.2 0.6 2.0 1.3 0.3 2.0 1.9  Recovery 2.5 5.0 1.0 3.3 1.0 0.7 2.3 2.4 A  Control 0.2 0 0 0 0 0 0 0  Stress 1.1 2.1 0.9 1.9 0.7 0.6 0.5 0.8  Recovery 1.4 3.0 1.0 1.9 0.9 0.8 0.6 0.8 Z  Control 0.7 0 0 0 0 0 0 0  Stress 5.2 5.9 2.8 4.8 1.2 1.2 1.3 1.3  Recovery 4.8 4.3 2.7 2.5 1.0 0.9 1.3 1.7 VAZ  Control 5.7 9.1 3.4 10.4 1.7 1.4 3.2 3.6  Stress 8.5 11.2 4.3 8.8 3.2 2.2 3.8 4.0  Recovery 8.7 12.3 4.7 7.6 2.8 2.4 4.2 4.8 (Z+A)/(VAZ)  Control 0.15 0 0 0 0 0 0 0  Stress 0.75 0.68 0.86 0.74 0.61 0.82 0.48 0.47  Recovery 0.72 0.55 0.78 0.53 0.66 0.69 0.44 0.46 The pigments Chl b, neoxanthin, lutein, and β-carotene are means ± sd of the control, stress (48 h of continuous light at 15°C for leaves from GC plants and 122 h of continuous light at 10°C for leaves from plants grown OD), and recovery (2.5 h in low light at 22°C) treatments. Open in new tab Near the top of the gradient a distinct yellow band was evident (fraction 1). This fraction contained free pigments, and V was the major component; only very low levels of Chl were present (Table II). Quantification of pigments in this fraction, relative to the fractions containing protein, showed that 80% to 88% of V in isolated thylakoids was bound to proteins (Table II). This fraction was not examined for proteins using SDS-PAGE; however, in a parallel experiment solubilized thylakoids from all treatments were applied to a nondenaturing deriphat gel and then to SDS-PAGE gels in the second dimension. No proteins were visible in the free-pigment fraction in any of the treatments when the gels were stained with Coomassie Blue. The second fraction from the top (fraction 2) was heterogeneous, containing LHCII monomer and the minor Chl proteins (CP24, CP26, and CP29) of PSII; the fraction migrating below it (fraction 3) contained pure LHCII trimer. The higher LHCII content in fraction 2 (derived from monomerization of trimeric LHCII) compared with findings from previous reports (Bassi and Dainese, 1992) was due to the relatively high detergent concentration used here to ensure fractionation of relatively pure PSI-LHCI and PSII core complexes. The composition of the xanthophyll cycle pigments in fractions 2 and 3 were strikingly different in fractions isolated from plants grown OD in the sun versus low-light GCs: the V content was 3 times higher in the LHCII trimer fraction (fraction 3) from plants grown OD (Table III). Although a 40% increase in V content was also observed in fraction 2 (minor CPs), this increase was possibly due entirely to the LHCII monomers present in this fraction (Fig. 1; Table III). The PSII core fraction (fraction 4), migrating below LHCII, was also relatively pure (the Chl a/b ratio was >20). The bottom fraction (fraction 5) contained pure PSI-LHCI with a Chl a/b ratio of about 9 and was the only gradient fraction that contained PSI-LHCI components. VAZ was present in all of the pigment-protein fractions, although in very different amounts. The major V-binding complexes were PSI-LHCI, which bound 40% of Chl a, 12% to 15% of Chl b,and 24% to 26% of V in both sets of control plants, and PSII LHCs (minor CPs plus LHCII), which together bound 50% of Chl a, 86% of Chl b, and 49% of V in GC plants; the corresponding values in plants grown OD were 39%, 83%, and 59%. Effect of Photoinhibitory Treatment V. major plants were subjected to treatments of continuous light (48 or 122 h for plants grown in GCs and OD, respectively) and chilling temperatures (15°C and 10°C for plants grown in GCs and OD, respectively) to induce photoinhibition, after which plants were allowed to recover at room temperature in darkness for 2.5 h. Plants were monitored during the treatment to ensure that photoinhibitory conditions (i.e. persistent reductions inFv/Fm) were achieved. The different treatment conditions reflect different requirements necessary to photoinhibit the plants, with the plants acclimated to the lower-light environment becoming photoinhibited much more rapidly than the plants grown in full sunlight. Fluorescence parameters ascertained at different times during the experimental treatment are depicted in TableIV. At the end of the stress treatment (after continuous high light and chilling temperatures) PSII efficiency at the actual degree of reaction center closure was quite low in both sets of plants. Increases in the efficiency of open PSII units (Fv/Fm) upon leaf darkening were very slow (Table IV), i.e. sustained decreases in PSII efficiency (photoinhibition) were present. Table IV. PSII efficiency of V. major grown in GCs or OD under natural sunlight V. major . Efficiency of PSII Units . GCs  Control (5) 0.72  ± 0.03 (D)  Stress (7) 0.04  ± 0.04 (L)  1.5 h of recovery (12) 0.33  ± 0.12 (D)  2.5 h of recovery (19) 0.37  ± 0.12 (D) OD  Control (11) 0.77  ± 0.02 (D)  Stress (13) 0.15  ± 0.08 (L)  1 h of recovery (14) 0.38  ± 0.08 (D)  2.5 h of recovery (22) 0.47  ± 0.13 (D) V. major . Efficiency of PSII Units . GCs  Control (5) 0.72  ± 0.03 (D)  Stress (7) 0.04  ± 0.04 (L)  1.5 h of recovery (12) 0.33  ± 0.12 (D)  2.5 h of recovery (19) 0.37  ± 0.12 (D) OD  Control (11) 0.77  ± 0.02 (D)  Stress (13) 0.15  ± 0.08 (L)  1 h of recovery (14) 0.38  ± 0.08 (D)  2.5 h of recovery (22) 0.47  ± 0.13 (D) Measurements were taken after 12 h of darkness (control), at the end of the high-light treatment (stress), and during recovery in darkness at room temperature. PSII efficiency is either that of open PSII units in darkness (Fv/Fm; D) or that at the actual degree of closure in the light ([Fm′ −F]/Fm′, whereFm′ is the maximal fluorescence measured in the light and F is the actual fluorescence measured in the light; L). The number of leaves sampled at each time is indicated in parentheses. Data are means ± sd. Open in new tab Table IV. PSII efficiency of V. major grown in GCs or OD under natural sunlight V. major . Efficiency of PSII Units . GCs  Control (5) 0.72  ± 0.03 (D)  Stress (7) 0.04  ± 0.04 (L)  1.5 h of recovery (12) 0.33  ± 0.12 (D)  2.5 h of recovery (19) 0.37  ± 0.12 (D) OD  Control (11) 0.77  ± 0.02 (D)  Stress (13) 0.15  ± 0.08 (L)  1 h of recovery (14) 0.38  ± 0.08 (D)  2.5 h of recovery (22) 0.47  ± 0.13 (D) V. major . Efficiency of PSII Units . GCs  Control (5) 0.72  ± 0.03 (D)  Stress (7) 0.04  ± 0.04 (L)  1.5 h of recovery (12) 0.33  ± 0.12 (D)  2.5 h of recovery (19) 0.37  ± 0.12 (D) OD  Control (11) 0.77  ± 0.02 (D)  Stress (13) 0.15  ± 0.08 (L)  1 h of recovery (14) 0.38  ± 0.08 (D)  2.5 h of recovery (22) 0.47  ± 0.13 (D) Measurements were taken after 12 h of darkness (control), at the end of the high-light treatment (stress), and during recovery in darkness at room temperature. PSII efficiency is either that of open PSII units in darkness (Fv/Fm; D) or that at the actual degree of closure in the light ([Fm′ −F]/Fm′, whereFm′ is the maximal fluorescence measured in the light and F is the actual fluorescence measured in the light; L). The number of leaves sampled at each time is indicated in parentheses. Data are means ± sd. Open in new tab The photoinhibitory (stress) treatment induced changes in pigment composition measured in extracts from both isolated thylakoids and whole leaves (Table I). At the end of the high-light treatment VAZ content (expressed per total Chl a) had increased in both sets of plants, with the increase being more pronounced in the plants grown in GCs (an increase of 53% versus 20%, calculated from the thylakoid data). A comparison of the pigment content measured from isolated thylakoids versus leaf extracts in the stressed plants showed that the ratios of carotenoids to Chl a were fairly consistent in both sets of data. A possible exception is the VAZ content of the GC plants subjected to photoinhibitory treatment, in which an approximate 20% decrease in VAZ occurred following thylakoid isolation. The de-epoxidation state of VAZ at the end of the stress treatment was high in both sets of thylakoids. The reconversion of Z plus A to V was somewhat less in the thylakoids from GC plants versus the plants grown OD after 2.5 h in darkness (the change in [Z plus A]/[VAZ] upon recovery was −0.06 versus −0.18 in thylakoids from the GC versus OD, respectively), which correlated with a slower increase inFv/Fm in the GC plants (Table IV). Thylakoids from leaves collected before and after the photoinhibitory treatment were analyzed for the presence of ELIPs. A possible role of ELIPs as xanthophyll-binding proteins, expressed when leaves were exposed to excessive irradiance, has been discussed (Adamska, 1997). Western blots, using double labeling with an antibody raised against fully denatured LHCII (which also weakly recognizes ELIPs) and the maize anti-ELIPs, indicated no induction of ELIPs following the photoinhibitory treatment. A faint band at approximately 17 kD (recognized by both antibodies) was apparent both before and after photoinhibition. In a control experiment with maize,cold stress induced the appearance of a 17-kD band reactive to polyclonal antibodies directed against an ELIPs epitope. However, because of the unusually high degree of species specificity of the ELIP antibody (especially the lack of cross-reactivity between monocot and dicot antibody to ELIP; B. Andersson, I. Adamska, and K. Kloppstech, unpublished observations and personal communication), it is likely that the maize antibody did not cross-react in V. major. Suc-Gradient Fractionation of Thylakoid Membranes upon Stress and Recovery During photoinhibitory stress the relative percentage of VAZ in the free-pigment fraction increased only slightly in thylakoids isolated from both sets of plants (Table II). However, the percentage of total Z plus A present in the free-pigment fraction (after stress) was higher than that of V under nonstressed conditions, indicating a relative enrichment of Z plus A in the free-pigment fraction following the stress treatment. Increases in bound VAZ (relative to Chla) following high-light treatment were apparent in all of the fractions, except the LHCII trimer in high-light-acclimated samples in which some decrease in bound VAZ was observed (Table III). The de-epoxidation of bound VAZ was generally higher in samples from GC plants compared with plants grown OD, except in the PSII core fraction and the fraction bound to PSI-LHCI. Within a treatment, (Z plus A)/(VAZ) was generally similar in all fractions except PSI-LHCI, which had a lower conversion state (approximately 0.46 in PSI-LHCI compared with the 0.61–0.86 in the other fractions; Table III). The free-pigment fraction had the highest degree of de-epoxidation following photoinhibitory stress (0.90 in the GC plants and 0.82 in the leaves of plants grown OD; Table II). The extent of reconversion of bound Z plus A to V following 2.5 h of darkness was consistently less in GC plants relative to plants grown OD, but the relative differences among proteins in the extent of reconversion were the same. The greatest extent of reconversion was in the free-pigment fraction (Δ[Z plus A]/[VAZ] of −0.09 and −0.23 in fractions from the GC and OD plants, respectively], with the LHCII trimer fraction exhibiting only slightly less reconversion (Δ[Z plus A]/[VAZ] of −0.08 and −0.21). The fractions containing the PSII core and the LHCII monomer/minor CPs showed approximately one-half the extent of reconversion (Δ[Z plus A]/[VAZ] of −0.05 and −0.03 in the core and minor CPs of the GC leaves and of −0.13 in both the core and minor CPs of the leaves of plants grown OD). The PSI-LHCI fraction showed the least reconversion in the plants grown OD (−0.01), whereas in the GC plants the reconversion was similar to the core and minor CPs (−0.04). Analysis of the PSII Core-Containing Fraction Immunoblot analysis of the PSII core-containing fraction, isolated from the plants grown OD, indicated that LHCII was the principal contaminant and that minor CPs or PSI-LHCI were not present. To quantify the level of contamination high amounts of the core fraction (protein equivalent to 10 μg of Chl) in addition to known amounts of pure LHCII trimer were subjected to SDS-PAGE. Densitometric analysis of the Coomassie Blue-stained gels revealed the level of contamination at a maximum of 7% LHCII. Calculations of expected pigment content (moles per 100 moles of Chl a) assuming pure PSII core and 7% contamination with LHCII were compared with the actual data in TableV. Although LHCII contamination accounted for all of the Chl b, neoxanthin, and lutein present, there was a greater concentration of VAZ in the core than could be accounted for by LHCII contamination, suggesting that some VAZ may be bound to the PSII core. Table V. Analysis of pigments in the fraction containing the PSII core (fraction 4) Pigment . PSII Core Suc Gradient (Fraction 4) . Calculated Value if 100% PSII Core . Calculated Values if 7% Contamination with LHCII . Actual/Calculated . Repurified Core with DEAE . mol/100 mol Chl a Chlb 2.7  ± 0.8 0 2.8 0.92 1.7  ± 0.3 Neoxanthin 0.8  ± 0.3 0 0.5 1.53 0 Lutein 3.1  ± 0.4 2.5 3.55 0.86 1.0  ± 0.4 β-Carotene 12.1  ± 2.9 8.9 8.55 1.41 13.4  ± 0.9 VAZ 2.0  ± 0.5 0 0.18 11.17 0.3  ± 0.2 Pigment . PSII Core Suc Gradient (Fraction 4) . Calculated Value if 100% PSII Core . Calculated Values if 7% Contamination with LHCII . Actual/Calculated . Repurified Core with DEAE . mol/100 mol Chl a Chlb 2.7  ± 0.8 0 2.8 0.92 1.7  ± 0.3 Neoxanthin 0.8  ± 0.3 0 0.5 1.53 0 Lutein 3.1  ± 0.4 2.5 3.55 0.86 1.0  ± 0.4 β-Carotene 12.1  ± 2.9 8.9 8.55 1.41 13.4  ± 0.9 VAZ 2.0  ± 0.5 0 0.18 11.17 0.3  ± 0.2 Comparison of the core fraction after Suc-gradient ultracentrifugation with values calculated based on previously published results for pure PSII core and core contaminated with 7% LHCII, in addition to pigment content after purification with DEAE-Fractogel chromatography. The data are means ± sd of the three treatments. Calculations were based on data presented by Yamamoto and Bassi (1996), except the VAZ content per LCHII was based on data from Table III. Open in new tab Table V. Analysis of pigments in the fraction containing the PSII core (fraction 4) Pigment . PSII Core Suc Gradient (Fraction 4) . Calculated Value if 100% PSII Core . Calculated Values if 7% Contamination with LHCII . Actual/Calculated . Repurified Core with DEAE . mol/100 mol Chl a Chlb 2.7  ± 0.8 0 2.8 0.92 1.7  ± 0.3 Neoxanthin 0.8  ± 0.3 0 0.5 1.53 0 Lutein 3.1  ± 0.4 2.5 3.55 0.86 1.0  ± 0.4 β-Carotene 12.1  ± 2.9 8.9 8.55 1.41 13.4  ± 0.9 VAZ 2.0  ± 0.5 0 0.18 11.17 0.3  ± 0.2 Pigment . PSII Core Suc Gradient (Fraction 4) . Calculated Value if 100% PSII Core . Calculated Values if 7% Contamination with LHCII . Actual/Calculated . Repurified Core with DEAE . mol/100 mol Chl a Chlb 2.7  ± 0.8 0 2.8 0.92 1.7  ± 0.3 Neoxanthin 0.8  ± 0.3 0 0.5 1.53 0 Lutein 3.1  ± 0.4 2.5 3.55 0.86 1.0  ± 0.4 β-Carotene 12.1  ± 2.9 8.9 8.55 1.41 13.4  ± 0.9 VAZ 2.0  ± 0.5 0 0.18 11.17 0.3  ± 0.2 Comparison of the core fraction after Suc-gradient ultracentrifugation with values calculated based on previously published results for pure PSII core and core contaminated with 7% LHCII, in addition to pigment content after purification with DEAE-Fractogel chromatography. The data are means ± sd of the three treatments. Calculations were based on data presented by Yamamoto and Bassi (1996), except the VAZ content per LCHII was based on data from Table III. Open in new tab The PSII core fraction was subjected to further purification using DEAE-Fractogel (EM Science, Gibbstown, NJ) chromatography (TableV). Immunoblot analysis of the purified core verified that after purification only trace amounts of LHCII were present (data not shown). After the core was repurified with DEAE, there were decreases in all of the xanthophylls in addition to Chl b(Table V). The only small amount of VAZ still present (0.3 mol/100 mol Chl a) may indicate that VAZ was bound loosely to the core, because it was largely removed upon DEAE chromatography. Flat-Bed IEF Fractionation of the LHCII Monomer and Minor CP-Containing Fraction from High-LightAcclimated Leaves The Suc-gradient fraction from the high-light OD samples containing LHCII monomer and the minor Chl proteins was subjected to flat-bed IEF. Fractions from IEF were applied to glycerol gradients (15%–40% glycerol) and ultracentrifuged, and two bands were collected in each case. Bands were analyzed by SDS-PAGE, and pigment analysis was performed (Fig. 2; TableVI). Although fully purified minor CPs were not obtained, the minor CPs were separated from LHCII, except for some contamination of the minor CPs by LHCII in the control sample (Fig. 2). Fig. 2. Open in new tabDownload slide Fully denaturing Tris-Tricine SDS-PAGE of the bands (bands 1–8) collected following glycerol gradients of IEF fractions from separation of LHCII monomer and the minor Chl proteins of samples obtained from V. major plants grown OD. The three gels depict samples from the control, stress, and recovery sets of experiments, as indicated. Thylakoids were loaded as a standard (CT, ST, and RT). CP29 (A), CP26 (B), LHCII (C), and CP24 (D) are indicated. Fig. 2. Open in new tabDownload slide Fully denaturing Tris-Tricine SDS-PAGE of the bands (bands 1–8) collected following glycerol gradients of IEF fractions from separation of LHCII monomer and the minor Chl proteins of samples obtained from V. major plants grown OD. The three gels depict samples from the control, stress, and recovery sets of experiments, as indicated. Thylakoids were loaded as a standard (CT, ST, and RT). CP29 (A), CP26 (B), LHCII (C), and CP24 (D) are indicated. Table VI. Average pigment composition of combined fractions (±sd) from flat-bed IEF of the Suc-gradient fraction containing LHCII and the minor Chl proteins from the plants grown OD Pigment . LHCII Combined Fractions . Minor CP Combined Fractions . Control (C1, C3, C5) . Stress (S1, S4, ND) . Recovery (R1, R4, ND) . Control (C6, C7) . Stress (S6, S8, S9) . Recovery (R6, R8, R9) . mol/100 mol Chl a Chlb 80.4  ± 2.6 82.8  ± 5.2 80.1  ± 6.7 54.4 54.5  ± 4.5 53.4  ± 4.0 Neoxanthin 11.9  ± 0.1 11.8  ± 1.1 11.8  ± 1.1 11.25 9.24  ± 1.2 9.29  ± 1.8 Lutein 27.1  ± 1.6 27.5  ± 2.3 28.1  ± 2.0 25.5 22.2  ± 4.1 20.9  ± 3.1 V 2.8  ± 0.6 1.2  ± 0.6 1.8  ± 0.3 11.1 8.3  ± 2.1 8.2  ± 1.1 A 0 1.0  ± 0.4 1.4  ± 0.6 0 3.5  ± 1.4 3.1  ± 0.4 Z 0 2.2  ± 0.7 2.1  ± 0.3 0 8.3  ± 2.8 5.9  ± 0.8 VAZ 2.8  ± 0.6 4.3  ± 1.7 5.4  ± 0.9 11.1 20.1  ± 5.9 17.2  ± 0.9 (Z+A)/(VAZ) 0 0.70  ± 0.06 0.62  ± 0.07 0 0.54  ± 0.07 0.48  ± 0.04 Pigment . LHCII Combined Fractions . Minor CP Combined Fractions . Control (C1, C3, C5) . Stress (S1, S4, ND) . Recovery (R1, R4, ND) . Control (C6, C7) . Stress (S6, S8, S9) . Recovery (R6, R8, R9) . mol/100 mol Chl a Chlb 80.4  ± 2.6 82.8  ± 5.2 80.1  ± 6.7 54.4 54.5  ± 4.5 53.4  ± 4.0 Neoxanthin 11.9  ± 0.1 11.8  ± 1.1 11.8  ± 1.1 11.25 9.24  ± 1.2 9.29  ± 1.8 Lutein 27.1  ± 1.6 27.5  ± 2.3 28.1  ± 2.0 25.5 22.2  ± 4.1 20.9  ± 3.1 V 2.8  ± 0.6 1.2  ± 0.6 1.8  ± 0.3 11.1 8.3  ± 2.1 8.2  ± 1.1 A 0 1.0  ± 0.4 1.4  ± 0.6 0 3.5  ± 1.4 3.1  ± 0.4 Z 0 2.2  ± 0.7 2.1  ± 0.3 0 8.3  ± 2.8 5.9  ± 0.8 VAZ 2.8  ± 0.6 4.3  ± 1.7 5.4  ± 0.9 11.1 20.1  ± 5.9 17.2  ± 0.9 (Z+A)/(VAZ) 0 0.70  ± 0.06 0.62  ± 0.07 0 0.54  ± 0.07 0.48  ± 0.04 The fractions combined are indicated below each treatment and correspond to lanes from the Tris-Tricine gels depicted in Figure 3. ND, The fraction is not depicted in Figure 3. Open in new tab Table VI. Average pigment composition of combined fractions (±sd) from flat-bed IEF of the Suc-gradient fraction containing LHCII and the minor Chl proteins from the plants grown OD Pigment . LHCII Combined Fractions . Minor CP Combined Fractions . Control (C1, C3, C5) . Stress (S1, S4, ND) . Recovery (R1, R4, ND) . Control (C6, C7) . Stress (S6, S8, S9) . Recovery (R6, R8, R9) . mol/100 mol Chl a Chlb 80.4  ± 2.6 82.8  ± 5.2 80.1  ± 6.7 54.4 54.5  ± 4.5 53.4  ± 4.0 Neoxanthin 11.9  ± 0.1 11.8  ± 1.1 11.8  ± 1.1 11.25 9.24  ± 1.2 9.29  ± 1.8 Lutein 27.1  ± 1.6 27.5  ± 2.3 28.1  ± 2.0 25.5 22.2  ± 4.1 20.9  ± 3.1 V 2.8  ± 0.6 1.2  ± 0.6 1.8  ± 0.3 11.1 8.3  ± 2.1 8.2  ± 1.1 A 0 1.0  ± 0.4 1.4  ± 0.6 0 3.5  ± 1.4 3.1  ± 0.4 Z 0 2.2  ± 0.7 2.1  ± 0.3 0 8.3  ± 2.8 5.9  ± 0.8 VAZ 2.8  ± 0.6 4.3  ± 1.7 5.4  ± 0.9 11.1 20.1  ± 5.9 17.2  ± 0.9 (Z+A)/(VAZ) 0 0.70  ± 0.06 0.62  ± 0.07 0 0.54  ± 0.07 0.48  ± 0.04 Pigment . LHCII Combined Fractions . Minor CP Combined Fractions . Control (C1, C3, C5) . Stress (S1, S4, ND) . Recovery (R1, R4, ND) . Control (C6, C7) . Stress (S6, S8, S9) . Recovery (R6, R8, R9) . mol/100 mol Chl a Chlb 80.4  ± 2.6 82.8  ± 5.2 80.1  ± 6.7 54.4 54.5  ± 4.5 53.4  ± 4.0 Neoxanthin 11.9  ± 0.1 11.8  ± 1.1 11.8  ± 1.1 11.25 9.24  ± 1.2 9.29  ± 1.8 Lutein 27.1  ± 1.6 27.5  ± 2.3 28.1  ± 2.0 25.5 22.2  ± 4.1 20.9  ± 3.1 V 2.8  ± 0.6 1.2  ± 0.6 1.8  ± 0.3 11.1 8.3  ± 2.1 8.2  ± 1.1 A 0 1.0  ± 0.4 1.4  ± 0.6 0 3.5  ± 1.4 3.1  ± 0.4 Z 0 2.2  ± 0.7 2.1  ± 0.3 0 8.3  ± 2.8 5.9  ± 0.8 VAZ 2.8  ± 0.6 4.3  ± 1.7 5.4  ± 0.9 11.1 20.1  ± 5.9 17.2  ± 0.9 (Z+A)/(VAZ) 0 0.70  ± 0.06 0.62  ± 0.07 0 0.54  ± 0.07 0.48  ± 0.04 The fractions combined are indicated below each treatment and correspond to lanes from the Tris-Tricine gels depicted in Figure 3. ND, The fraction is not depicted in Figure 3. Open in new tab The content of VAZ bound to LHCII following IEF (Table VI) was significantly lower than that bound to the LHCII trimer following Suc-gradient fractionation (Table II), which suggests that a portion of the VAZ bound to LHCII was stripped off during IEF (possibly due to the acidic pI of the LHCII bands that migrated between pH 3.5 and 4.5). The amount of VAZ bound to the LHCII fraction obtained after IEF was similar to that in the Suc-gradient fraction of the LHCII trimer from the GC plants (2.8–5.4 versus 3.4–4.7 mol VAZ/100 mol Chla in the IEF fraction versus the LHCII trimer fraction of GC plants, respectively). The photoinhibitory treatment induced a considerable increase in VAZ bound to the minor CPs (11–20 mol VAZ/100 mol Chl a; TableVI), as well as an increase in the VAZ still bound to LHCII (2.8–4.3 mol VAZ/100 mol Chl a), in contrast to the apparent decrease in total (tightly plus loosely bound) VAZ associated with LHCII trimers from plants grown OD (Table II). At the end of the stress treatment the conversion state of the xanthophyll cycle was higher in the LHCII versus the minor CP fraction (Table VI), whereas the extent of reconversion of Z plus A to V upon 2.5-h recovery was similar in both fractions (−0.08 versus −0.06 in LHCII and the minor CPs, respectively). When the VAZ pool that was apparently loosely bound to LHCII (present in the LHCII trimer fraction following Suc-gradient fractionation but not present in the LHCII monomer following IEF) was examined, the quantity of loosely bound VAZ had decreased after photoinhibitory treatment (7.6–4.3 mol VAZ/100 mol Chl a), whereas there was a corresponding increase in the VAZ bound to the minor CPs (TablesII and VI). This may indicate a redistribution of loosely bound VAZ (particularly the Z plus A formed) from LHCII to the minor CPs during the photoinhibitory treatment. The presumably loosely bound VAZ had a higher degree of de-epoxidation relative to VAZ that was more tightly bound to LHCII (0.82 versus 0.70, respectively) and reconverted to a greater extent following recovery (−0.45 versus −0.08, respectively), suggesting that this loosely bound VAZ was more accessible to the xanthophyll cycle enzymes than the more tightly bound VAZ. Nondenaturing Green Gel of the Minor CP-Containing Fraction IEF fractions containing the minor CPs were solubilized with 1.9% dodecyl maltoside and run on a nondenaturing green gel. Each of the fractions resulted in two green bands, which were excised, eluted from the gel, and analyzed. Although pure CPs were not obtained, SDS-PAGE analysis indicated that there was significant enrichment in CP26 or CP29 in two of the bands excised from both the stress samples and the recovery samples. Results of the densitometric analysis of the Coomassie Blue-stained gels, indicating the relative quantities of the proteins, in addition to the xanthophyll cycle pigment content for each fraction, is presented in Figure 3. A relative enrichment in V correlated with CP29, whereas enrichment with Z correlated with CP26. This suggests that the conversion state of the xanthophyll cycle was not uniform among the minor CPs and that the pigments bound to CP29 de-epoxidized to a lesser extent compared with either CP26 or the LHCIIs. Fig. 3. Open in new tabDownload slide A, Densitometric analysis of Coomassie Blue-stained gels of bands from nondenaturing green gels of the minor CPs and LHCII monomer from samples obtained from plants grown OD (two bands were apparent in each lane). A, Relative percentages of CP29, LHCII, and CP26 in each band. B, The contents of the xanthophyll cycle pigments analyzed for each band. Fig. 3. Open in new tabDownload slide A, Densitometric analysis of Coomassie Blue-stained gels of bands from nondenaturing green gels of the minor CPs and LHCII monomer from samples obtained from plants grown OD (two bands were apparent in each lane). A, Relative percentages of CP29, LHCII, and CP26 in each band. B, The contents of the xanthophyll cycle pigments analyzed for each band. DISCUSSION The results of this study demonstrate that at least 80% of the xanthophyll cycle pigments (VAZ) that were present in isolated thylakoids were bound to protein independently of the de-epoxidation state (Table II). These data are consistent with those of Thayer and Björkman (1992) who found that 14% to 24% of the VAZ pool was in the pigment front following nondenaturing deriphat electrophoresis of solubilized thylakoids. The comparison of pigment content of extracts from both whole leaves and isolated thylakoids demonstrates a high degree of pigment conservation during the thylakoid isolation procedure (Table I). Of the VAZ found in the free-pigment fractions, the de-epoxidation state was higher in the samples collected during the stress treatment than for protein-bound VAZ, and upon 2.5 h of recovery this portion was reconverted to V to a greater extent than the bound VAZ. These data may indicate that loosely bound and/or free VAZ is the most accessible to the enzymes responsible for their interconversions. In addition, because the leaf VAZ content increased during stress treatment (Table I), there may be relatively more Z in this fraction because of the presence of newly synthesized pigment. An important, novel finding of the present study is that, unlike any other pigment component, the levels of VAZ associated with a given PSII protein fraction are variable and can be altered by environmental factors such as growth conditions (e.g. sun-exposed plants grown OD versus GC plants grown at low PPFD) and/or photoinhibitory treatment. Our data suggest that an increasing demand for thermal energy dissipation results in increased levels of Z plus A bound to additional sites on, or associated with, PSII proteins. It is an attractive possibility that these additional sites may be additional, loose binding sites on given PSII proteins. But it cannot be excluded at this time that stress-induced additional proteins with binding sites for Z plus A may become associated with PSII proteins. Whereas such a possibility has been suggested for ELIPs (that can bind Chl and lutein;Adamska, 1997), no actual evidence has been obtained that ELIPs can in fact bind Z plus A. Moreover, the possibility that, at least in isolated extracts, ELIPs may associate with other proteins particularly easily should also be considered (V. Ebbert, B. Demmig-Adams, and W. Adams, unpublished observations). The distribution of the VAZ that was bound to protein was as follows. LHCII In leaves acclimated to different environmental conditions, the fraction containing LHCII exhibited an altered VAZ content. In TableVII the xanthophyll content per LHCII polypeptide was computed assuming 12 Chl a+bbound (Dainese and Bassi, 1991; Kühlbrandt et al., 1994) and using the data from Table III and VI. Thus, VAZ per polypeptide was greater in plants grown OD versus V. major grown in GCs. The additional VAZ found in the plants grown OD was mostly loosely bound, because it was removed upon IEF. In addition, there were 2 luteins and 0.8 neoxanthin per LHCII polypeptide, both of which were not altered by growth conditions or stress treatment (Tables II, VI, and VII). These data therefore indicate approximately 3.5 carotenoids per LHCII monomer in the bands from Suc gradients (Table II) and close to 3 carotenoids per LHCII following IEF (Tables VI and VII), which is in agreement with the value of 3 xanthophylls per LHCII polypeptide consistently found for highly purified LHCII (Bassi et al., 1993; Ruban et al., 1994). We propose that LHCII possesses an additional xanthophyll-binding site (in addition to the three usually found) that binds additional VAZ in leaves acclimated to high light. Since loosely bound VAZ was found to be associated with the LHCII complex, it could be hypothesized that the VAZ found in the free-pigment fraction actually derives from LHCII, which would account for 0.2 mol VAZ per mol LHCII in the case of the sample from the plants grown OD. With this complement each LHCII would bind 3.7 xanthophyll molecules of which approximately 1 would bind to the new binding site postulated above. Table VII. Xanthophyll content of LHCII as calculated on the basis of 12 Chl (a+b) per polypeptide Pigment . Control . Stress . Recovery . GC . OD . GC . OD . GC . OD . mol/mol polypeptide Chla+b 12 12 12 12 12 12 Neoxanthin 0.85 0.78 (0.79) 0.85 0.78 (0.78) 0.85 0.78 (0.78) Lutein 2.3 2.2 (1.8) 2.3 2.2 (1.8) 2.3 2.2 (1.9) VAZ 0.23 0.71 (0.18) 0.28 0.60 (0.28) 0.31 0.52 (0.36) Total xanthophylls 3.38 3.64 (2.77) 3.43 3.58 (2.86) 3.46 3.50 (3.04) Pigment . Control . Stress . Recovery . GC . OD . GC . OD . GC . OD . mol/mol polypeptide Chla+b 12 12 12 12 12 12 Neoxanthin 0.85 0.78 (0.79) 0.85 0.78 (0.78) 0.85 0.78 (0.78) Lutein 2.3 2.2 (1.8) 2.3 2.2 (1.8) 2.3 2.2 (1.9) VAZ 0.23 0.71 (0.18) 0.28 0.60 (0.28) 0.31 0.52 (0.36) Total xanthophylls 3.38 3.64 (2.77) 3.43 3.58 (2.86) 3.46 3.50 (3.04) Values in parentheses refer to xanthophyll content following acid treatment during IEF. Values are from V. major grown in GCs versus OD. Open in new tab Table VII. Xanthophyll content of LHCII as calculated on the basis of 12 Chl (a+b) per polypeptide Pigment . Control . Stress . Recovery . GC . OD . GC . OD . GC . OD . mol/mol polypeptide Chla+b 12 12 12 12 12 12 Neoxanthin 0.85 0.78 (0.79) 0.85 0.78 (0.78) 0.85 0.78 (0.78) Lutein 2.3 2.2 (1.8) 2.3 2.2 (1.8) 2.3 2.2 (1.9) VAZ 0.23 0.71 (0.18) 0.28 0.60 (0.28) 0.31 0.52 (0.36) Total xanthophylls 3.38 3.64 (2.77) 3.43 3.58 (2.86) 3.46 3.50 (3.04) Pigment . Control . Stress . Recovery . GC . OD . GC . OD . GC . OD . mol/mol polypeptide Chla+b 12 12 12 12 12 12 Neoxanthin 0.85 0.78 (0.79) 0.85 0.78 (0.78) 0.85 0.78 (0.78) Lutein 2.3 2.2 (1.8) 2.3 2.2 (1.8) 2.3 2.2 (1.9) VAZ 0.23 0.71 (0.18) 0.28 0.60 (0.28) 0.31 0.52 (0.36) Total xanthophylls 3.38 3.64 (2.77) 3.43 3.58 (2.86) 3.46 3.50 (3.04) Values in parentheses refer to xanthophyll content following acid treatment during IEF. Values are from V. major grown in GCs versus OD. Open in new tab These data have important implications with regard to the distribution of xanthophyll cycle pigments among CPs. It was previously proposed that most of the VAZ was bound to the minor CPs, whereas a minor fraction was LHCII bound in maize and spinach (Bassi et al., 1993;Ruban et al., 1994). On the basis of Suc-gradient ultracentrifugation data (Table II), VAZ can be attributed to the different CPs (TableVIII), with the conclusion that a significant portion of VAZ is actually bound to the LHCII fraction, although in a low-affinity site. Moreover, upon acclimation to different environmental conditions, the distribution of V undergoes a dramatic change. In GC-grown plants, 28% of VAZ was bound to the minor CP fraction and 24% was bound to the LHCII fraction, whereas in thylakoids from plants grown OD only 11% of VAZ was bound to minor CPs and 48% was bound to LHCII. If it is assumed that the VAZ in the free-pigment band was stripped off of LHCII (see above), then LHCII would bind 60% of VAZ in the plants grown OD and 42% in V. major grown in GCs at low PPFD. Table VIII. Distribution of Chl and VAZ in the Chl-binding complexes Complex . V. major from OD . V. major from GCs . Chl . VAZ . Chl . VAZ . % Free pigment 0.6 12.4 1.2 17.6 PSII minor CPs 12.1 11 10.2 27.3 LHCII 37.3 48 43.6 22.6 PSII core 16 4.6 13 7.5 PSI-LHCI 34 24 32 25 Complex . V. major from OD . V. major from GCs . Chl . VAZ . Chl . VAZ . % Free pigment 0.6 12.4 1.2 17.6 PSII minor CPs 12.1 11 10.2 27.3 LHCII 37.3 48 43.6 22.6 PSII core 16 4.6 13 7.5 PSI-LHCI 34 24 32 25 Values are expressed as the percentages of total pigment found in each complex. Open in new tab Table VIII. Distribution of Chl and VAZ in the Chl-binding complexes Complex . V. major from OD . V. major from GCs . Chl . VAZ . Chl . VAZ . % Free pigment 0.6 12.4 1.2 17.6 PSII minor CPs 12.1 11 10.2 27.3 LHCII 37.3 48 43.6 22.6 PSII core 16 4.6 13 7.5 PSI-LHCI 34 24 32 25 Complex . V. major from OD . V. major from GCs . Chl . VAZ . Chl . VAZ . % Free pigment 0.6 12.4 1.2 17.6 PSII minor CPs 12.1 11 10.2 27.3 LHCII 37.3 48 43.6 22.6 PSII core 16 4.6 13 7.5 PSI-LHCI 34 24 32 25 Values are expressed as the percentages of total pigment found in each complex. Open in new tab Upon photoinhibitory treatment, the loosely bound fraction of VAZ decreased (0.53–0.30 VAZ per polypeptide), whereas the more tightly bound fraction increased slightly (0.2–0.3 VAZ per polypeptide). This may be indicative of a redistribution of xanthophyll cycle pigments under light stress, which differs from their localization in darkened leaves. It is possible that the V-binding site of LHCII has a low affinity for Z. The presence of such a V-binding but not Z-binding site is supported by the recent finding that the aba-3 mutant of Arabidopsis, lacking epoxy-xanthophylls, contained one less xanthophyll molecule per LHCII polypeptide (Connelly et al., 1997). It has previously been proposed that upon de-epoxidation Z and A are released from LHCII and become free in the membrane lipids (Krupa et al., 1987;Tardy and Havaux, 1997), where they may modify membrane fluidity (Gruszecky and Strzalka, 1991; Tardy and Havaux, 1997). Although our findings are consistent with some release of VAZ from LHCII, the amount of VAZ recovered in the free-pigment band does not change significantly following de-epoxidation (Table II). The constancy of free VAZ in spite of its apparent release from LHCII suggests that newly formed Z becomes bound to one or more pigment-binding protein(s). Minor CPs are the best candidate, since (a) their VAZ content increased upon de-epoxidation and (b) their degree of de-epoxidation was less than that of the free-pigment fraction, which is consistent with the hypothesis that they bind Z that has been de-epoxidized in the free-lipid phase. Consistent with this, high degrees of de-epoxidation have been found in chlorina mutants of barley (Dainese et al., 1992), which are enriched in xanthophylls found in the free-pigment fraction (Bassi et al., 1985a). Minor Chl Proteins The pigment-binding properties of the minor Chl proteins (CP29, CP26, and CP24) were relatively constant during purification, and total VAZ content of the minor CP fraction from darkened control leaves did not seem to differ in response to the acclimation to low- versus high-growth PPFD. Assuming approximately equal proportions of the three minor CPs (CP29, CP26, and CP24), the pigment composition indicated by these data are in close agreement with data from highly purified proteins (Bassi et al., 1993; Ruban et al., 1994). Considering 6 Chla per polypeptide, there were an average of 3.2 Chlb, 1.3 lutein, 0.5 neoxanthin, and between 0.7 and 1.2 VAZ per polypeptide. There were 2.5 carotenoids per polypeptide in the control leaves and a considerable increase in the VAZ bound to the minor CP fraction during stress, resulting in 3.2 carotenoids per polypeptide during stress. It is therefore possible that 3 carotenoid-binding sites are present in the minor CPs, one-third of which is only partially occupied in control leaves. Whereas the existence of a variably occupied site was also postulated for LHCII (in control GC plants versus those grown OD in full sun), in the case of minor CPs bound VAZ increased only during stress treatment rather than as an acclimation response, thus implying that the CP site has a higher affinity for Z than for V. Thus, affinities for Z versus V appear to be opposite for these newly postulated sites on the minor CPs versus LHCII. The present results confirm that the degree of de-epoxidation is not the same among the different minor CPs. Although CP29 bound considerable amounts of V (Bassi et al., 1993; Giuffra et al., 1996), this pigment could be de-epoxidized to a much lower extent compared with CP26 (Fig. 3; it was not possible to assess the de-epoxidation state of CP24). Low xanthophyll de-epoxidation in CP29 following high-light treatment was previously reported (Ruban et al., 1994; Croce et al., 1996a). These results are in contrast to those of Färber et al. (1997), who found that, when spinach was treated with 3 h of photoinhibition, there was no increased binding of VAZ to any of the pigment-protein complexes and no redistribution of pigments. This difference may be due to the very different experimental conditions used: a 3-h photoinhibitory treatment versus a 48- or 122-h treatment used in this study. PSII Core Suc-gradient fractionation yielded PSII core fractions with a maximum of 7% contamination by LHCII. Assuming 56 Chls per PSII core, between 1 and 1.8 xanthophylls per PSII core were detected in samples from GC plants and 0.8 to 1.3 xanthophylls per PSII core were detected in samples from plants grown OD in full sunlight. The LHCII contamination accounted for less than 0.1 xanthophyll molecule per PSII core (Table V), suggesting that 1 to 2 VAZ molecules may be bound to the PSII core. This was not previously recognized in a study of maize leaves (Bassi et al., 1993) in which a multistep purification procedure was used, which would remove any loosely bound pigment. A loose binding site for VAZ in the PSII core is suggested by the results of ion-exchange chromatography; when the complex was bound to the column and the column was washed extensively, the xanthophyll content decreased to 0.17 per PSII core. The presence of VAZ in PSII core fractions has also been reported in barley (Lee and Thornber, 1995) and lettuce (Phillip and Young, 1995). There was some increase in VAZ present in the PSII core fraction following the stress treatment (Table III). Binding of Z to the PSII core upon photoinhibition was postulated previously (Jahns and Miehe, 1996; Färber et al., 1997). When expressed per polypeptide, the increase in apparent VAZ content was actually similar in magnitude for the PSII cores and minor CPs (an increase of 0.8 or 0.5 VAZ per PSII core complex in leaves from GCs or OD, respectively, versus an increase of 0.5 VAZ per minor CP in leaves from plants grown OD). However, it should be noted that the absolute content of VAZ detected in the PSII core was very small (Table II), thus precluding any firm conclusions. Again, the association of stress-induced proteins with the PSII core fraction, leading to the binding of additional VAZ, cannot be excluded. PSI-LHCI We confirm the presence of VAZ in PSI-LHCI (Thayer and Björkman, 1992; Lee and Thornber, 1995; Zhu et al., 1997). However, the VAZ bound to PSI-LHCI was de-epoxidized to a lesser extent upon photoinhibitory treatment than that bound to the PSII complexes, and the extent of reconversion following 2.5 h recovery was also lower in PSI-LHCI. The presence of photoconvertible VAZ in PSI-LHCI raises the question of whether thermal energy-dissipation mechanisms similar to those observed in PSII take place in PSI. This seemed unlikely previously, since PSI was believed to be a deep trap, which would make energy dissipation in LHCI inefficient (in decreasing Chla excited state concentrations at the level of the PSI reaction center). More recently, however, it was recognized that PSI, like PSII, is a shallow trap and that, therefore, PSI and its antenna are essentially equilibrated (Croce et al., 1996b), implying that thermal dissipation in the LHCI antenna could be an efficient regulatory mechanism for Chl a excitation in PSI. This could provide protection from the adverse effects of excess excitation on PSI as well (Terashima et al., 1994; Tjus et al., 1998). CONCLUSIONS In this study we have shown that xanthophyll cycle pigments undergo dynamic changes not only in their epoxidation state but also in their association with CPs. Upon acclimation to growth OD in full sunlight, the V content of control thylakoids was greatly increased. This additional V was bound to the major LHCII fraction. Following photoinhibitory treatment, the VAZ content of the LHCII fraction decreased and the minor CP fraction of PSII (CP29, CP26, and CP24), and to a lesser extent the PSII core complex fraction, bound increased amounts of Z plus A. This is interpreted in terms of the presence of an additional, low-affinity V-binding site in LHCII, in equilibrium with V free in the lipid phase, and of an additional high-affinity Z-binding site in the minor CPs. The higher degree of de-epoxidation in the free-pigment fraction suggests that the preferred substrate for V de-epoxidase is the pigment free in the lipid phase, which, upon conversion, becomes bound to minor CPs and PSII core CPs. ACKNOWLEDGMENTS We gratefully acknowledge Paolo Pesaresi for his assistance in performing some of the experimental techniques. Abbreviations: A antheraxanthin Chl chlorophyll CP Chl-binding protein ELIP early-light-inducible protein Fv/Fm ratio of variable to maximal Chl fluorescence GC growth chamber LHCI or II light-harvesting complex I or II OD outdoors V violaxanthin VAZ V plus A plus Z Z zeaxanthin LITERATURE CITED 1 Adams WW III Demmig-Adams B Verhoeven AS Barker DH 'Photoinhibition' during winter stress: involvement of sustained xanthophyll cycle-dependent energy dissipation. Aust J Plant Physiol 22 1995 261 276 Google Scholar OpenURL Placeholder Text WorldCat 2 Adamska I ELIPs: light-induced stress proteins. 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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)