Plant Physiology

Hughes, Jon; Lamparter, Tilman

doi: 10.1104/pp.121.4.1059pmid: 10594094

Prokaryotic systems have been important in phytochrome studies on several different levels. Bilins from cyanobacterial phycobiliproteins have allowed the production of recombinant holophytochrome and have provided insights into the attachment and functioning of the chromophore, while the recent discovery of functional phytochromes in the cyanobacteriumSynechocystis and other prokaryotes has catalyzed work in the field. Synechocystis phytochrome is useful experimentally and, by making the modular structure and potential biochemical functions of phytochromes clearer, has provided an improved focus and new viewpoints for research. Some of the earliest studies of photobiology concerned phenomena in cyanobacteria: the complementary chromatic adaptation (CCA) of photosynthetic pigments to the light environment was first described in Engelmann's laboratory in Berlin a century ago. Numerous other effects such as phototaxis, photoperiodism, cell division, and differentiation are also regulated by light in cyanobacteria. Plant plastids probably evolved from endosymbiotic cyanobacteria whose genes gradually moved to the host nucleus. There is thus every reason to expect evolutionary relationships between photoperception systems in cyanobacteria and plants. Detailed information about cyanobacterial photoreceptors was lacking, however, until genomic sequencing revealed a cyanobacterial phytochrome: ironically, the ease with which molecular methods can be used in prokaryotes has now turned the tables, with the cyanobacterial model providing a wealth of new ideas about the origins of phytochrome and its mode of action. Here we review the different ways in which cyanobacteria and other prokaryotes have contributed to research into plant photomorphogenesis and the phytochrome system (for review, seeElich and Chory, 1997; Quail, 1997a; Pepper, 1998). Phytochrome is an ubiquitous plant photoreceptor that was first characterized in the late 50s in relation to its peculiar photochromic behavior in red and far-red light (Butler et al., 1959). Phytochromes carry an open-chain tetrapyrrole (bilin) chromophore, which the apoprotein autocatalytically attaches to a conserved C residue (#380 in our alignment2) via a Schiff base (Lagarias and Lagarias, 1989). In darkness, this autoassembly produces the red-light-absorbing form Pr (λmax ≈ 660 nm). In red light this is photoisomerized to another form, Pfr, which absorbs maximally in far-red light (λmax ≈ 730 nm). In far-red light, Pfr is in turn converted back to Pr. Both forms are thermodynamically stable and can be interconverted by any number of photocycles. Because even tiny amounts of Pfr have major physiological effects, it is generally accepted that this is the active form of phytochrome, while Pr seems to be physiologically inactive. In plants, phytochromes control a variety of developmental processes such as seed germination, stem elongation, construction of the photosynthetic apparatus, chloroplast movements, shade avoidance, and photoperiodic induction of flowering. In lower plants they are also involved in sensing light direction. CYANOBACTERIA AND PLANTS CONTAIN FIVE DIFFERENT BILIN CHROMOPHORES Based on the first spectral measurements, it was correctly argued that the phytochrome chromophore might be a bilin similar to those of phycobiliproteins in cyanobacteria and red algae (Butler et al., 1959). Phycobiliproteins bear four types of bilin, namely phycocyanobilin (PCB), the chromophore of phycocyanin (PC, which is thus generally the most abundant), phycoerythrobilin (PEB), phycoviolobilin (PVB), and phycourobilin (PUB) (see Fig. 1). Apophytochromes autoassemble with PCB to form holophytochrome photoreceptors—a useful feature, as PCB can be prepared rather easily from commercially available cyanobacteria, enabling the preparation of functional phytochromes by recombinant methods (Wahleithner et al., 1991). However, in all land-plant phytochromes examined so far, another member of the family, phytochromobilin (PΦB), is the natural chromophore (Rüdiger and Thümmler, 1994). Phytochrome chromophores undergo a characteristic Z → E isomerization around the C15C16 double bond between rings C and D during Pr → Pfr conversion. PEB also assembles with apophytochrome in vitro, but the product is not photochromic because the C15-C16 bond is saturated (Li and Lagarias, 1992). The five bilins (PCB, PΦB, PVB, PEB, and PUB) above differ only in single double bonds and are derived from biliverdin, the first open-chain tetrapyrrole in this biosynthetic pathway (Fig. 1). Fig. 1. Open in new tabDownload slide Structure of heme and the natural bilins biliverdin (BV), PΦB, PCB, PEB, PVB, and PUB. Heme oxygenase converts heme to bilivirdin by cleaving between rings A and D at the positions marked. Differences in the other bilins with respect to bilivirdin are also indicated. Fig. 1. Open in new tabDownload slide Structure of heme and the natural bilins biliverdin (BV), PΦB, PCB, PEB, PVB, and PUB. Heme oxygenase converts heme to bilivirdin by cleaving between rings A and D at the positions marked. Differences in the other bilins with respect to bilivirdin are also indicated. ONLY THE FIRST GENE FOR THE BILIN SYNTHETIC PATHWAY IS KNOWN In seed plants, the enzymes for bilin synthesis are located in the plastids but are nuclear encoded. The pathway begins with the opening of the heme tetrapyrrole ring between pyrroles A and D by heme oxygenase to form the linear tetrapyrrole (bilin) biliverdin IXα. Genes for this enzyme are known from the cyanobacteriumSynechocystis PCC6803 (Cornejo et al., 1998), from the plastome of red algae, and from the genomes of Arabidopsis and several animals. In Arabidopsis heme oxygenase is encoded by the nuclear geneHY1 (Muramoto et al., 1999). The ptr116phototropic mutant of the moss Ceratodon can be rescued by exogenous biliverdin by microinjecting cells with mammalian heme oxygenase enzyme or by overexpressing HY1, showing that all three functionally complement the defective ptr116 gene (Brücker et al., 1999). The subsequent steps are not fully understood biochemically, nor have any further associated genes been described, although the Arabidopsis HY2 locus is a likely candidate. Further sequences from theSynechocystis genome must also be involved in the production of PCB and other bilins. CYANOBACTERIAL MODELS FOR PHYTOCHROME Early physiological studies indicated that cyanobacteria might harbor useful information about the phytochrome system. Action spectroscopy revealed photoreversible effects analogous to those of plant phytochrome but with the interesting distinction that, while in plants the responses are maximally induced by red light and reverted by far-red light, most photoreversible effects in cyanobacteria respond to red light (λmax ≈ 650 nm) and green light (λmax ≈ 520 nm) (Vogelman and Scheibe, 1978). The most intensely studied effect here is CCA. Unlike plants, cyanobacteria possess phycobilisome structures that funnel energy into the photosynthetic system. The principle accessory pigments involved are the blue-green (red-absorbing) PC and allophycocyanin and the red (blue-green-absorbing) phycoerythrin (PE). Some species are able to use CCA to adjust the ratio of these pigments according to environmental conditions. In green light PE dominates, whereas in red light PC dominates. In this way, the λmax of photosynthesis is shifted to the λmax of the light environment (Gaidukov, 1902). When a series of green (≈ 540 nm) and red (≈ 650 nm) pulses was given to a culture of the cyanobacterium Fremyella diplosiphon and the culture kept in darkness, the last light pulse determined the dominant accessory pigment formed (Vogelman and Scheibe, 1978). This kind of photoreversibility points to a photoreceptor with photochromic properties, an unusual spectral feature that allowed plant phytochrome to be isolated and characterized. However, this approach was less successful in cyanobacteria, principally because, unlike angiosperms, they do not etiolate. Although a photoreversible pigment showing difference maxima at 520 and 650 nm has been described (Scheibe, 1972), it was not characterized further. Later, the α-subunit of the minor phycobilisome component phycoerythrocyanin (PEC) was shown to be photochromic—but with difference maxima at 500 and 570 nm. The physiological role of PEC is unknown, although it might have a role as a photoreceptor as well as acting as a photosynthesis antenna. The PECα chromophore is PVB, which undergoes a Z → E photoisomerization analogous to phytochrome (Zhao et al., 1995). Moreover, the PECα and PCα sequences are approximately 65% identical and the molecules have similar three-dimensional structures—yet only the former is photochromic. While one might therefore suppose that the differences between these biliproteins could provide a master key to unlock the secrets of photochromicity at the atomic level, whether this key would fit phytochrome is questionable. First, there is no sequence homology between phycobiliproteins and phytochrome. Second, isolated phycobiliproteins show far stronger fluorescence than phytochrome, implying that their chromophores are much more tightly held and/or have a very different photochemistry. Third, relative to the dark state, the conformational changes in PECα are associated with a blue shift, whereas a red shift is seen with phytochrome. Interestingly, although phycobiliprotein apoproteins are generally capable of autocatalytically attaching bilin chromophores in vitro, it has been shown that specific lyases mediate the assembly in vivo, accelerating the reaction and ensuring correct bilin attachment. Phytochrome autoassembles in vitro too, but whether a discrete phytochrome bilin lyase exists is simply not known. TWO-COMPONENT SIGNALING ENTERS THE FRAY A quite different line of investigation also connects plant phytochromes with prokaryotic systems. Schneider-Poetsch et al. (1991)drew attention to a significant amphiphilic sequence similarity between the phytochrome C terminus and the transmitter module of bacterial sensory His protein kinases (HPKs). HPKs are a group of proteins responsible for the first step in the so-called two-component signal transduction pathways (see Fig. 2A) that provide the prokaryotic cell with its capacity for perception and response. Fig. 2. Open in new tabDownload slide Two-component signal transduction. A, Basic scheme. The HPK dimer is activated by conformation changes induced by stimuli perceived by the sensor module, usually N-terminal. The transmitter module of each subunit then transfers a phosphate (red dot) from ATP to a conserved His residue (circle) of the other subunit. The phosphate group is transferred to a conserved Asp residue (square) of the cognate response regulator, which is thereby activated. An autophosphatase activity returns the response regulator to its inactive state in the phosphorylation cycle (PC)—the rate of hydrolysis is sometimes regulated by interactions with the “inactive” form of the same HPK or with another molecule. B, Cph1/Rcp1 system. Pr, The ground state of the phytochrome in darkness or far-red light is the active HPK; irradiation with red light converts the molecule to Pfr, in which the HPK activity of the transmitter module is inhibited. Following autophosphorylation at H538#995, Pr transfers the phosphate to the response regulator Rcp1. The biochemical functions of Pfr and the response regulator are unknown. Fig. 2. Open in new tabDownload slide Two-component signal transduction. A, Basic scheme. The HPK dimer is activated by conformation changes induced by stimuli perceived by the sensor module, usually N-terminal. The transmitter module of each subunit then transfers a phosphate (red dot) from ATP to a conserved His residue (circle) of the other subunit. The phosphate group is transferred to a conserved Asp residue (square) of the cognate response regulator, which is thereby activated. An autophosphatase activity returns the response regulator to its inactive state in the phosphorylation cycle (PC)—the rate of hydrolysis is sometimes regulated by interactions with the “inactive” form of the same HPK or with another molecule. B, Cph1/Rcp1 system. Pr, The ground state of the phytochrome in darkness or far-red light is the active HPK; irradiation with red light converts the molecule to Pfr, in which the HPK activity of the transmitter module is inhibited. Following autophosphorylation at H538#995, Pr transfers the phosphate to the response regulator Rcp1. The biochemical functions of Pfr and the response regulator are unknown. The activity of each HPK is regulated by an associated sensory module whose conformation changes in response to an environmental signal such as an interaction with a specific ion or molecule. The HPK is a dimer and, upon sensor activation, each subunit phosphorylates the other at a conserved H target residue (H#995) within the transmitter module. The phosphate is then transmitted to a conserved D residue in the receiver module on the second component of the transduction system, the response regulator. This then does as its name suggests, either by activating transcription of specific genes itself or by interacting with other proteins to bring about specific physiological changes in the cell appropriate to the environmental signal. These two-component signal transduction systems seem to be the primary regulatory connections between prokaryotic metabolism and the environment. Schneider-Poetsch's suggestion that phytochrome might represent a plant sensory HPK was enhanced by the discovery of eukaryotic HPK homologs SLN1 in yeast and ETR1 in plants a year or so later, and indeed the idea that phytochrome might be a light-dependent kinase was nothing new. A seductive aspect was that it provided phytochrome with its long-sought reaction partner: This should be a response regulator homolog. Many were not convinced by the HPK/phytochrome homology, however, and there was one very large problem: although all of the functional subdomains characteristic of HPK transmitters are recognizable in phytochromes, the all-important H#995 target residue itself is poorly conserved (see alignment). It seemed that the affair was over when Boylan and Quail (1996) showed that even in phytochromes in which H#995 was conserved, it could be mutated without noticeable physiological effect. POSSIBLE PHOTORECEPTORS IN CYANOBACTERIA Work with the cyanobacterial CCA perception system, however, continued independently. Complementation methods were used to clone genes involved in the regulation of chromatic adaptation in F. diplosiphon. RcaE (Kehoe and Grossman, 1996) encodes a 74-kD polypeptide with an approximately 150-residue portion toward the N terminus showing homologies to several regions around the chromophore-binding domain of plant phytochromes, as well as C-terminal motifs typical of transmitter modules. Intriguingly, RcaE also bears two subdomains (T2L and R2L, T105#265-L129#289 and R241#436-L268#463, see alignment) showing homology to the plant ethylene receptor ETR1. Although the biochemistry has yet to be demonstrated, RcaE probably phosphorylates the response regulator RcaF, which in turn phosphorylates RcaC. The latter bears a transmitter- and two receiver-like modules, as well as a DNA-binding motif thought to mediate differential transcription of the PC and PE gene complexes. In the similar cyanobacterium Calothrix, RcaD and RcaA act as phosphorylation-dependent activators of the PC and PE gene clusters, respectively. The 155-kD conceptual gene product of PlpA (Wilde et al., 1997; sll1124 in CyanoBase) in Synechocystis PCC6803 also shows regions of similarity to phytochromes (hence the name Plp for phytochrome-like protein) and two-component modules (see alignment). Indeed, BLAST searches show that RcaE has approximately 25% amino acid identity (40% similarity) to PlpA, although the latter has a long N-terminal extension. This particular Synechocystis strain does not show CCA. It does, however, change the stochiometry between PS1 and PS2 according to the irradiance and spectral distribution of the light environment: In plpA −knockouts the balance between the photosystems is disturbed. Furthermore, in contrast to the wild type, theplpA − mutant cannot grow photoautotrophically in blue light. Although both PlpA and RcaE are clearly important in cyanobacterial photoperception, it has proven difficult to demonstrate that they are photoreceptors. The sensory function could be fulfilled by a separate molecule, as in many two-component systems. At least on the basis of homology to the phytochrome N terminus, there is little reason to expect either RcaE or PlpA gene products to be bona fide phytochromes: If one assumes that the chromophore is thioether-linked to a C residue, as in phycobiliproteins and plant phytochromes, the alignment in this region is constrained to C198#380 and C784#380 for RcaE and PlpA, respectively. The surrounding subdomain is quite different from that seen in phytochromes, where it is well conserved and changes generally lead to a complete loss of function. However, although difficulties with overexpressing PlpA andRcaE in Escherichia coli have hampered in vitro studies, it has now been reported that both do seem to be capable of attaching bilins (A. Wilde, T. Börner, D. Kehoe, and A. Grossman, unpublished data). A PROKARYOTIC PHYTOCHROME CREATES EXCITEMENT IN THE FIELD Quite separately, the entire 3.57-Mbp chromosome ofSynechocystis PCC6803 was sequenced in a singularly efficient project at the Kazusa Institute in Japan (CyanoBase:http://www.kazusa.or.jp), providing the scientific community with a wealth of valuable new data. Among the 3,168 open reading frames identified, a phytochrome-like sequence (slr0473) was recognized (Kaneko et al., 1995; Hughes et al., 1996). The N-terminal moiety showed patchy but unmistakable similarity to phytochromes, including the all-important chromophore binding region, whereas the 30-kD C-terminal moiety was clearly homologous to typical two-component transmitter modules with characteristically conserved H-, N-, G1-, F-, and G2-boxes (see alignment and Fig. 3). Fig. 3. Open in new tabDownload slide Synechocystis phytochrome (Cph1) in relation to plant phytochromes and sensory His protein kinases (HPK's). Residue numbering is that from the alignment (http://www.plantphysiol.org/cgi/content/full/121/4/1059/DC2 andhttp://www.biologie.fu-berlin.de/phytochrome/align2x.htm). The N-terminal bilin-bearing sensor module (blue) is recognizable in all phytochromes, while the C-terminal transmitter module (yellow) is common to phytochromes and most HPKs of the two-component type. The sensory modules of other HPKs (brown) are different and are sometimes carried on separate polypeptides. A PAS module (green), important for plant phytochrome signal transduction, is absent from Cph1. The chromophore attachment site, two PAS repeats, the signal transduction core (STC or Quail box), and the H-, N-, G1-, F-, and G2-boxes are highly conserved. Plant B-type phytochromes generally have an N-terminal extension, otherwise the N terminus of all phytochromes is Ser/Thr rich. Fig. 3. Open in new tabDownload slide Synechocystis phytochrome (Cph1) in relation to plant phytochromes and sensory His protein kinases (HPK's). Residue numbering is that from the alignment (http://www.plantphysiol.org/cgi/content/full/121/4/1059/DC2 andhttp://www.biologie.fu-berlin.de/phytochrome/align2x.htm). The N-terminal bilin-bearing sensor module (blue) is recognizable in all phytochromes, while the C-terminal transmitter module (yellow) is common to phytochromes and most HPKs of the two-component type. The sensory modules of other HPKs (brown) are different and are sometimes carried on separate polypeptides. A PAS module (green), important for plant phytochrome signal transduction, is absent from Cph1. The chromophore attachment site, two PAS repeats, the signal transduction core (STC or Quail box), and the H-, N-, G1-, F-, and G2-boxes are highly conserved. Plant B-type phytochromes generally have an N-terminal extension, otherwise the N terminus of all phytochromes is Ser/Thr rich. The question nevertheless remained: is it a phytochrome? The 85-kD gene product was further analyzed simultaneously by Lagarias's group at University of California-Davis and by our laboratory (Hughes et al., 1997; Lamparter et al., 1997; Yeh et al., 1997). The apoprotein overexpressed in E. coli autocatalytically attached PCB chromophore in vitro to form a blue-green photochromic pigment, clearly establishing that it encodes a bona fide cyanobacterial phytochrome, Cph1. The discovery of this phytochrome caused an immediate paradigm shift in the field. Schneider-Poetsch's suggestion that the unknown mechanism of primary signal transduction could be related to the well-established two-component system in bacteria suddenly became a very hot topic. Moreover, the utility of a prokaryotic phytochrome system in biochemical and molecular-genetic studies opens new experimental possibilities. In particular, the efficiency with which highly soluble recombinant phytochrome can be prepared from E. coli overexpressors offers fresh hope that the three-dimensional structure of this class of photoreceptors could be resolved via NMR and x-ray diffraction analysis of phytochrome crystals. Cph1 IN VITRO Cph1 has been subjected to a range of optical and biophysical studies in our laboratory (Lamparter et al., 1997) and in those of our colleagues. Recombinant apoprotein (Cph1°) is expressed remarkably efficiently in E. coli, accumulating to approximately 30% of cytosolic protein. Moreover, equipped with a C-terminal oligohistidine tag, it can be purified almost to homogeneity in a single Ni2+-affinity chromatographic step.E. coli does not support bilin synthesis, and thus autoassembly does not occur in vivo. If Cph1° is added to PCB in vitro, however, a dramatic blue to blue-green color change occurs within seconds. This results from two processes. Initially, recombinant holoprotein (Cph1*) is formed as Pr, whereby the PCB red absorbance peak at 610 nm is shifted to 658 nm as the helical form of the free bilin becomes unwound in the protein environment. Thereafter, if observed in daylight, the Pr is photoconverted to Pfr, whose absorbance peak is shifted even further to 702 nm. Both photochromic forms are quite stable in darkness. The yield of pure Cph1* is routinely about 20 mg per liter of culture. It can be concentrated to above 15 mg/mL quite easily, satisfying an important further precondition for many biophysical and physicochemical studies, including crystallization. Like other HPKs and plant phytochromes, Cph1* behaves as a dimer in solution. As one would expect for a photoreceptor, the extinction coefficient of Cph1* is very high (approximately 100 mm −1cm−1 for Pr at λmax) and the quantum conversion efficiency is about 0.16 in both directions—values similar to those for plant phytochromes. Cph1° can also be assembled with other chromophores: The PΦB adduct shows a red shift of about 15 nm for both Pr and Pfr, as seen with plant phytochromes. PEB adducts cannot photoconvert because of their missing C15C16 double bond (see Fig. 1); the quantum energy is released as fluorescence and their absorbance maximum is blue-shifted to 579 nm. Fourier-transform Raman-resonance (FTRR) and flash photolysis (Remberg et al., 1997), low-temperature fluorescence (Sineshchekov et al., 1998), and Fourier-transform IR absorbance (FTIR, H. Förstendorf and F. Siebert, unpublished data) spectroscopic methods have also been used. Despite the considerable differences in the peptide sequences, Cph1* shows remarkably similar physicochemical properties to those of B-type phytochromes. FTRR is a sensitive probe for the status of the chromophore in biliproteins, revealing in this case many similarities between the chromophores of native oat phytochrome A and the equivalent PΦB adduct of Cph1*. For both phytochromes, spectral differences between the Pr and the Pfr form reflect the Z → E isomerization of the chromophore and changes in its hydrogen bonding with the protein. Moreover, as in plant phytochromes, subtle differences between the PCB and the PΦB adduct of Cph1* can be attributed to the ring D side chain (vinyl group for PΦB versus ethyl group for PCB). FTRR also indicated different torsions around methine bridges within the chromophore and differences in chromophore/protein interactions between Cph1* and oat phytochrome. As for other phytochromes, the formation of intermediates during Pr → Pfr photoconversion of Cph1* was readily observed by flash photolysis and fast spectroscopy. The first photoproduct detected (lumi-R) of Cph1* appeared substantially more quickly than that for plant phytochromes and was followed by a novel intermediate whose kinetics were delayed almost 2-fold by2H exchange, implying that a protonation/deprotonation is involved at this point. FTIR difference spectra also indicate 2H effects, and a photoreversible pH shift (J. Hughes and J. van Thor, unpublished data) seems to confirm that proton extrusion accompanies Pfr formation. Fluorescence measurements at low temperature address the photoconversion from a different point of view. Whereas at ambient temperature phytochrome fluorescence yields are very low, these rise dramatically upon cooling; Pr → Pfr photoconversion is inhibited, although photoconversion into intermediate forms is sometimes possible. For plant PHYA at 70 K, up to 50% of the Pr can convert into lumi-R, whereas this conversion is not possible for plant PHYB. PCB and PΦB Cph1* adducts are also unable to form lumi-R at this temperature, implying that Cph1 is more related to PHYB than to PHYA photochemically. Different activation barriers for the photoreaction are thought to explain the differences between phytochrome types. The only intermediate photoproduct found after allowing the temperature to rise seemed to be rather different from the lumi-R of plant phytochromes. Cph1* IS A LIGHT-DEPENDENT HIS PROTEIN KINASE The Lagarias group (Yeh et al., 1997) analyzed the biochemistry of Cph1* regarding its apparent homology to two-component systems (see Fig. 2B). Cph1* autophosphorylates at the expected H538#995, but it was a great surprise that the active kinase was not Pfr but Pr. This flew in the face of most plant physiological data, which implied that Pfr was the active form. There was more to come, however. Unlike eukaryotes, prokaryotes often group biochemically related genes together in a single cistron, thereby keeping the job of coordinating expression simple while obligingly providing the scientist with clues to unknown biochemical associations. While biochemists had long sought the primary reaction partner(s) for plant phytochrome, the likely reaction partner for Synechocystis phytochrome was clear from the Kazusa chromosome map. Fifteen bases downstream of theCph1 stop codon begins a short open reading frame, slr0474, unmistakably coding for a 17-kD response regulator of the two-component type (Lamparter et al., 1997). Yeh et al. (1997) overexpressed this gene in yeast and demonstrated that the autophosphorylated Pr form of Cph1* promptly transmitted its phosphate to the expected Asp residue D68 of the putative response regulator. slr0474 was thus the first primary reaction partner for phytochrome to be identified and was named response regulator for cyanobacterial phytochrome, Rcp1. Here again, Pr was more active than Pfr. All of the known enzymatic activities of phytochrome (bilin ligase, His autokinase, His-Asp transphosphorylase, and, as we shall see, Ser/Thr kinase) were first described by the Lagarias laboratory. Although many response regulators are DNA-binding proteins and act as transcriptional activators, the Synechocystis model is not quite so simple. Rcp1 has no DNA-binding motifs and thus presumably acts as an intermediate phosphocarrier in a more complex relay. This is also seen in other two-component systems such as Spo and Rca. The epithet refers to the transmitter module of the kinase and the receiver module of the response regulator, whereas the transduction system as a whole can be considerably more extensive, with pathways converging and branching to form a sophisticated control network. So with what does Rcp1 interact? Unfortunately, in Synechocystis onlyCph1 and Rcp1 are co-transcribed, the flanking genes being read in the opposite direction. As the next partner cannot simply be deduced from the genome map, finding the rest of the transduction chain will prove more difficult. Cph1 IN VIVO REMAINS A MYSTERY Although Synechocystis certainly contains abundant PCB, the native chromophore of Cph1 is not known. CCA in other cyanobacteria shows well-separated maxima in the blue-green and red regions, whereas the absorbance maxima of PCB and PΦB adducts of Cph1* are poorly separated and are at longer wavelengths (see above). If a Cph1 homolog is indeed the photoreceptor for CCA, then the blue shift might result from the use of a different chromophore. Alternatively, a quite separate photoreceptor might be involved. Measuring difference spectra in extracts of Synechocystis was unsuccessful because of masking pigments, even when using PC– deletion mutants. In an attempt to overcome this, homologous recombination was used to replace the wild-type chromosomal gene with a sequence extended to provide an oligohistidine-tagged translation product similar to that in the E. coli overexpression clones, thereby allowing the photoreceptor to be purified by affinity methods. However, despite this technology, the extracts have yielded only tiny amounts of the modified native Cph1, indicating a very low expression level. Interestingly, the purified fraction showed not only the expected red/far-red difference spectrum with maxima around 650 and 700 nm, but also a red/green difference with maxima at 530 and 650 nm, close to the maxima for CCA. Whether the red/green reversible signal relates to a co-purified protein or directly to Cph1 remains to be determined (T. Lamparter, A. Wilde, and T. Hübschmann, unpublished data). Ironically, despite all the studies of Cph1 in vitro, its physiological function is unknown: What aspect of the light environment does it perceive and what response does it mediate? This gap in our knowledge is all the more surprising because the efficient homologous recombination available in Synechocystis allows knockout mutants to be created with some ease. Indeed, both Cph1− and Cph1−/Rcp1− knockouts have been generated in several labs, but an associated phenotype has yet to be found (D. Scanlan, A. Wilde, and T. Börner, unpublished data). Perhaps the effects are masked by another photoreceptor system, or the Cph1-Rcp1 pathway might lead to a physiological dead end in this particular strain. One might expect Cph1 to regulate CCA—but, unfortunately, PCC6803 and most other strains ofSynechocystis lack PE entirely and thus could not show CCA even if they wanted to. However, Synechocystis PCC6701 shows classical CCA, carrying a PE gene cluster closely homologous to that inFremyella and replacing PC with PE in blue-green light. The Cph1 homolog in CCA-active Calothrix has also now been cloned (N. Tandeau de Marsac, unpublished data). It will be interesting to see if a cph1 − knockout in one of the CCA-active types shows the Rca− phenotype. OTHER PROKARYOTIC PHYTOCHROMES HAVE ALSO BEEN IDENTIFIED Several other Synechocystis genes also show similarities to phytochromes. It seems that the true homolog ofRcaE is not PlpA (sll1124), as implied above, but, rather, is represented in Cyanobase by two pseudogenes separated by a transposon (sll11473–sll1475); in other PCC6803 cultures theRcaE homolog is intact (A. Wilde, unpublished data). sll0821 is also intriguing as it shows two regions with homology to the phytochrome chromophore subdomain, both of which bind PCB in vitro (S.-H. Wu and J.C. Lagarias, unpublished data). Even further removed from plant phytochromes are the BphP(bacterial phytochrome photoreceptor) genes recently found on the chromosomes of Deinococcus radiodurans and Pseudomonas aeruginosa. The former autoassembles with bilin chromophores in vitro to yield a phytochrome-like red/far-red light photochromic product (R. Vierstra and S. Davis, unpublished data). This result is surprising because, although a region resembling the phytochrome chromophore subdomain is apparent, residue #380 is M rather than C. While we assume that the chromophore attachment site of Cph1 is C259#380, this has yet to be demonstrated chemically. (See also “Note Added in Proof”) FROM PROKARYOTES TO PLANT PHYTOCHROME Most of the above discussion concerns phytochrome in prokaryotes, but how does that help the plant physiologist? Most importantly, it provides conceptual links. First, the alignment of Cph1 to plant phytochromes and HPKs provided a new and clearer view of phytochrome molecular architecture. Second, while the initial cloning of phytochrome was a great technical achievement in itself, the sequence did not provide us with beguiling homologies to molecules of known function. Cph1 provides a link not only to bacterial two-component systems, but also to several other eukaryotic homologs including SLN1, DHKA and DHKB, ETR1, and CKI1, all thought to take part in phosphorelay-mediated signaling. This rejuvenated the idea that phytochromes might be light-dependent protein kinases. PLANT PHYTOCHROMES POSSESS A PAS MODULE INVOLVED IN SIGNAL TRANSDUCTION Cph1 alignments revealed an additional approximately 300-residue module peculiar to plant phytochromes, placed between the sensory and transmitter modules (probably between A497#648and L498#954, see alignment and Fig. 3). The module is also missing from the Deinococcus andPseudomonas homologs. This region of the plant phytochrome sequence had already aroused interest since it contains a repeated motif (#709-#751 and #846-#888) related to the PAS3 domain family (Jones and Edgerton, 1994; Lagarias et al., 1995). It would seem that this PAS module was added to a Cph1-like progenitor early in plant evolution, perhaps even before eukaryotes arose, bringing with it a set of biochemical features probably including a new signaling mechanism. PAS domains (see Taylor and Zhulin, 1999) are found in diverse proteins throughout the living world; particularly interesting is the apparent PAS homology of the bacterial photoreceptor PYP (photoactive yellow protein) (Lagarias et al., 1995). PAS domains often bind ligands and are involved in protein-protein interactions including signal transduction. There is ambivalent evidence that the PAS repeats S599#675 to L683#766 and L685#768 to R815#901 are involved in the dimerization of phytochrome A (Edgerton and Jones, 1993; Quail, 1997b). Furthermore, random mutagenesis studies indicate that the PAS module is crucial to the plant phytochrome signaling mechanism (Quail et al., 1995). Yeast two-hybrid studies identified several phytochrome interacting factors that seem likely to bind to the PAS module. One of these is involved in phytochrome signaling in vivo, is nuclear localized, and even possesses a DNA-binding domain (Ni et al., 1998; Halliday et al., 1999), offering a remarkably—if not deceptively—simple picture of plant phytochrome action, given that newly formed Pfr migrates to the nucleus (Sakamoto and Nagatani, 1996). Although the plant phytochrome PAS module is missing from Cph1, a Hidden Markov model (http://coot.embl-heidelberg.de/SMART/) detects PAS-domain-related structures in Cph1, RcaE, and PlpA at different positions. A weak but significant similarity between HPK modules and the PAS domain has also been pointed out (Yeh and Lagarias, 1998), providing the latest twist to an unfinished story. KINASE AND KINASE-RELATED FUNCTIONS? As far as kinase function is concerned, the conceptual framework is not simple. As we have seen, athough the H538#995 target in Cph1 and its homologs in other sensory HPKs in both prokaryotes and eukaryotes is essential for autokinase and phosphorelay function, the homologous residue in plant phytochromes is neither conserved nor functional. While it is nevertheless possible that plant phytochromes could act as HPKs (for example, some PHYAs show an H-box-like [L/V][A/P]SHELQ[Q/H]AL#961-#970 motif at the PAS/transmitter module boundary) and response-regulator homologs certainly exist in plants, we emphasize that there is no evidence that any plant phytochrome functions as an HPK. Nevertheless, as we shall see, the HPK transmitter domain seems to be very much involved in signal transduction. The two-component paradigm might help us to understand plant phytochrome function independently of the prokaryotic kinase action; the H-box is by no means the best conserved of the two-component transmitter subdomains in plant phytochromes. The structures might have been retained for some purpose other than autophosphorylation and phosphotransfer. The unusual architecture of the chemotaxis HPK CheA suggests two possibilities. First, the three-dimensional structure of CheA shows a relict H-box in the conventional position, which, along with downstream residues, comprises the K290#981 to R354#1055 dimerization site (Bilwes et al., 1999). All HPKs seem to form stable dimers with submicromolar dissociation constants as a result of subunit binding in this region. Plant phytochromes are also dimers but the domains involved are uncertain (see Quail, 1997b). The CheA dimerization domain is made up of two antiparallel, highly amphiphilic α-helices with hydrophobic residues exposed on the subunit surface; PHD (http://www.embl-heidelberg.de/ predictprotein) predicts that this region in plant phytochromes is also largely helical with an amphiphilic pattern. Second, recent studies (U. Sweere and K. Harter, unpublished data) in Arabidopsis indicate that the N-terminal 100-residue fragment of phytochrome B binds the response regulator homolog ARR4, whereas the equivalent phytochrome A fragment does not. B-type phytochromes generally bear a characteristic N-terminal extension (#1–#37, see alignment) relative to other family members, so it is possible that the extension mediates the interaction. This would be analogous to the unconventional H-target subdomain at the N terminus of CheA, although little sequence homology is apparent and there is no evidence that the phytochrome is involved in a phosphotransfer. The system seems to connect to a two-component system involved in hormone signaling. Missing a conserved H#995-target residue in plant phytochromes, it was suggested that the perfectly conserved Y#991 nearby might have taken over the acceptor function (Schneider-Poetsch et al., 1991). Tyr and Ser/Thr protein kinases (YPKs and S/TPKs, respectively) work differently from HPKs; after autophosphorylation, rather than donating their own single phosphates, they phosphorylate their substrates with phosphate groups from free ATP4. There are, however, precedents for protein kinases showing a different substrate specificity from that implied by their primary structure. Moreover, immunological methods suggest a light-regulated Y phosphorylation of oat PHYA (Sommer et al., 1996). Although the residue involved is not known, motifs around Y#319 and Y#1055 in various phytochromes resemble the phosphotyrosine-binding site of SH2 domains. While this appears to be the sum of current evidence for phytochrome YPK function, the possible relationship between two-component systems and YPK/Ras GTPase signaling (Stock and Lukat, 1991) should encourage a careful search for related mechanisms in the case of phytochrome. PLANT PHYTOCHROME IS A DIFFERENT KIND OF KINASE Even before the sequence of oat PHYA was published, Quail and co-workers drew attention to its peculiarly S/T-rich N terminus as a possible kinase substrate. This was perhaps born of necessity as it was the only feature of the sequence that hinted at a function. While phytochrome N-terminal sequences are not well conserved, S and T residues predominate—also in Cph1. Indeed, S8#45in oat PHYA is phosphorylated; however, the physiological significance of this is unclear, as the level of phosphorylation is similar for Pr and Pfr (Lapko et al., 1997). Interestingly, mutation of the N-terminal Ser residues in PHYA increases rather than decreases its physiological potency in transgenic plants, so the phospho-Ser modification might serve to attenuate phytochrome action, analogously to arrestin-mediated quenching of rhodopsin (Elich and Chory, 1997). In the case of phytochrome it is uncertain whether this is an autophosphorylation event or whether a separate kinase is involved. The idea that phytochrome might be a S/T kinase bothered biochemists for many years, but recent evidence using recombinant systems in vitro and in vivo indicates that plant phytochromes can indeed autophosphorylate S/T residues and phosphorylate other proteins, including Rcp1, in a light-dependent manner (Yeh and Lagarias, 1998;Fankhauser et al., 1999; Lapko et al., 1999). Major differences from the Cph1 system should be made clear, however. First, plant Pfr becomes more strongly labeled than Pr, implying that the assembled sensory module in its ground state represses the autokinase activity—the opposite of Cph1*. Second, although plant phytochrome phosphorylated the Rcp1 response regulator in vitro, the target was not the D68 used by Cph1*. As histones too were effective substrates, the relevance of this observation might be called into question. However, it seems that the phytochrome kinase substrate PKS1 is phosphorylated by Pfr both in vitro and in vivo, with overexpression leading to repression of phytochrome action. Third, rather than H538#995, one or more unknown S/T residue(s) in plant phytochrome are autophosphorylated. S599#675 of oat PHYA—within the PAS module but N-terminal of the first repeat—shows Pfr-enhanced phosphorylation in vivo and would thus seem to be an obvious candidate. However, it is not conserved and the S599K#675mutant still autophosphorylates and phosphorylates PKS1—although light regulation is lost. In relation to domain function it is interesting to note that, while it is probably not a functional HPK, the plant phytochrome transmitter module alone is sufficient for PSK1 binding. Three-dimensional structural studies with kinases and their allies are already advanced (for example, Bilwes et al., 1999), providing useful background information regarding possible functions in phytochrome. Most of the residues directly responsible for ATP binding in HPKs and gyrases (GXG#1166-#1168, GLGL#1196-#1199 and G#1213) are well conserved in phytochromes–interestingly deviant are N#1118 and D#1168. We look forward to the day when phytochrome will contribute to studies of kinase function in general. Pr VERSUS Pfr For the plant physiologist, perhaps the most intriguing aspect of Cph1 is that Pr is the active kinase, while in plants Pr is thought to be inactive, Pfr being the “active form of phytochrome”. The red-light-induced formation of tiny amounts of Pfr from the Pr pool in the cytoplasm of imbibed seeds or dark-grown seedlings leads to the profound physiological changes associated with germination or de-etiolation. Physiological responses do correlate quite well with the Pfr concentration of PHYA in etiolated tissues, although in green tissues this is less certain because spectroscopic measurements are hampered by strong chlorophyll fluorescence and the approximately 100-fold lower amounts of phytochrome. Genetic studies seem to have settled the issue, however, as phy −mutants phenocopy Pr. There is also an attitude problem. For the physicist, photoreceptors are in their ground state in darkness and are excited by light—thus the ground state of Cph1 is Pr and the excited state is Pfr. But for the Cph1 biochemist, the active kinase is Pr—mirroring the behavior of the Rhizobium HPK oxygen sensor, FixL, in which ligand binding represses kinase activity. For the biologist, on the other hand, prolonged darkness is equivalent to starvation for a photosynthetic organism—“ground state” is hardly an appropriate description. As discussed above, there is no reason to suppose that plant phytochromes act as HPKs—even if that was the original function of Pr. On the other hand, it seems now that plant phytochromes act as Pfr-active S/T protein kinases. This leaves a question open: What is the biochemical function of Pfr in cyanobacteria? Once again, the two-component paradigm provides possible answers. Many HPKs are known to be bi-functional, phosphorylating or promoting de-phosphorylation of the response regulator according to their conformation as determined by the sensor module. Both activities are important as they allow the transduction system to differentiate rather than integrate the input signals from the sensor module, a principle that also applies to eukaryotic G-protein-coupled signaling, as in the rhodopsin/arrestin system. The photochromic nature of Cph1 offers the attractive possibility that the excited Pfr form might play the opposite role to that of Pr, promoting the de-phosphorylation of Rcp1. Of course, cyanobacterial Pfr might have a quite separate biochemical activity or it may simply be inactive. Whatever the role of Pfr in Cph1 is, it might be retained in plant phytochromes. CONCLUSIONS AND OUTLOOK As we have discussed, photochromic detection systems and chromophores associated with the cyanobacterial phycobilisome have made crucial contributions to the study of phytochrome at the molecular and conceptual levels. The unexpected discovery of phytochromes in other prokaryotes both answers and poses many questions. Phytochrome apparently appeared before eukaryotes, evolving over vast tracts of time and under changing selection pressures to glean and then transmit pertinent information about the light environment to allow the organism to respond appropriately. Perhaps most surprising, then, is the clarity of the homologies between prokaryotic and plant phytochromes. The origin of the PAS module involved in plant phytochrome signaling can also be traced to prokaryotes. Thus, a variety of prokaryotic models are acting catalytically in studies of phytochrome, the active sites being the modes of signal transduction and the photochromic mechanism itself. The reaction products should prove most interesting. ACKNOWLEDGMENTS We thank Annegret Wilde and Thomas Hübschmann (Humboldt University, Berlin), Richard Vierstra and Seth Davis (University of Wisconsin, Madison), David Kehoe (University of Indiana, Bloomington), Uta Sweere (University of Freiburg, Germany), Jasper van Thor (University of Amsterdam), Pill-Soon Song (University of Nebraska, Lincoln), Clark Lagarias (University of California, Davis), Peter Quail (Plant Gene Expression Center, Albany, CA), David Scanlan (Warwick University, UK) and Harald Förstendorf (University of Freiburg) for helpful discussions and for providing data prior to publication. 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Biochim Biophys Acta 1228 1995 235 243 Google Scholar Crossref Search ADS WorldCat NOTE ADDED IN PROOF Since this review was submitted, Jiang et al. (Z.Y. Jiang, L.R. Swen, B.G. Rushing, S. Devanathan, G. Tollin, C.E. Bauer [1999] Science 285: 406–409) have reported a photoreceptor, Ppr, in the purple photosynthetic bacterium Rhodospirillum centenum showing homology to HPKs and phytochromes but, like Cph1, missing the PAS module. An N-terminal extension resembles PYP and, like PYP, the apoprotein binds p-hydrocinnamic acid. The reconstituted holoprotein is a functional HPK whose autokinase activity is inhibited by blue light. Also, BphP from Deinococcus has now been shown to attach PCB at H260#381 — adjacent to C#380 (M in PphP), the traditional binding site (S. Davis and R. Vierstra, unpublished data). Interestingly, H#381 is conserved in all phytochromes (see alignment). Author notes 1 This work was supported by grants from the Deutsche Forschungsgemeinschaft. 2 Residues numbered with a # refer to an alignment available athttp://www.plantphysiol.org/cgi/content/full/121/4/1059/DC2 and http://www.biologie.fu-berlin.de/phytochrome/align2x.htm. Other numbering refers to the specific gene product involved (start codon = 1). 3 Originally the PAS domain referred to the entire region including both repeats, but as it now seems that the repeated region in PAS can also appear alone, we refer here to a “single-copy” PAS domain. 4 This is significant in signal transduction. Although an HPK activated by its sensor module might be able to carry out autophosphorylation and phosphotransfer many times, amplifying the initial signal (gain > 1), the rest of the prokaryotic phosphorelay does not amplify (gain < 1). Eukaryotes commonly employ Y and S/T protein kinases in a cascade, providing strong amplification (gain ≫ 1). Interestingly, the eukaryotic two-component HPK systems SLN1 and ETR1 both connect to such cascades. * Corresponding author; e-mail [email protected]; fax 49–30–838–4357. 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)

Kosegarten, Harald U.; Hoffmann, Bernd; Mengel, Konrad

doi: 10.1104/pp.121.4.1069pmid: 10594095

Abstract It has been hypothesized that under NO3 − nutrition a high apoplastic pH in leaves depresses Fe3+ reductase activity and thus the subsequent Fe2+ transport across the plasmalemma, inducing Fe chlorosis. The apoplastic pH in young green leaves of sunflower (Helianthus annuus L.) was measured by fluorescence ratio after xylem sap infiltration. It was shown that NO3 − nutrition significantly increased apoplastic pH at distinct interveinal sites (pH ≥ 6.3) and was confined to about 10% of the whole interveinal leaf apoplast. These apoplastic pH increases presumably derive from NO3 −/proton cotransport and are supposed to be related to growing cells of a young leaf; they were not found in the case of sole NH4 + or NH4NO3 nutrition. Complementary to pH measurements, the formation of Fe2+-ferrozine from Fe3+-citrate was monitored in the xylem apoplast of intact leaves in the presence of buffers at different xylem apoplastic pH by means of image analysis. This analysis revealed that Fe3+reduction increased with decreasing apoplastic pH, with the highest rates at around pH 5.0. In analogy to the monitoring of Fe3+ reduction in the leaf xylem, we suggest that under alkaline nutritional conditions at interveinal microsites of increased apoplastic pH, Fe3+ reduction is depressed, inducing leaf chlorosis. The apoplastic pH in the xylem vessels remained low in the still-green veins of leaves with intercostal chlorosis. Various investigations have shown that leaves may show Fe-deficiency symptoms even with leaf Fe concentrations higher than in green leaves (e.g. Carter, 1980; Mengel and Malissiovas, 1981; Sahu et al., 1987). Aktas and Van Egmond (1979) reported that chlorosis increased with elevated NO3 − supply. The chlorosis-inducing effect of NO3 − was also found byMengel and Geurtzen (1988) and could be reversed by switching from NO3 − to NH4 + without any external supply of Fe. Hoffmann et al. (1992) were the first to report a relationship between leaf apoplastic pH and the form of N nutrition. With NH4 + supply the leaf apoplastic pH was low, while NO3 − resulted in high apoplastic pH. Mengel et al. (1994) and Kosegarten and English (1994)found an inverse relationship between the chlorophyll concentration and leaf apoplastic pH. NO3 −was thought to be taken up into the cell via a NO3 −/H+cotransport (Ullrich, 1992; Crawford and Glass, 1998), and the perfusion of excised leaves with NO3 − resulted in microsites with an apoplastic pH of around 7.0 (Hoffmann and Kosegarten, 1995). These findings suggested that high leaf apoplastic pH restricts cellular Fe acquisition (Mengel, 1995), and this conclusion was corroborated by the observation that spraying leaves with dilute acids resulted in a re-greening of chlorotic leaves (Sahu et al., 1987;Tagliavini et al., 1995). Sahu et al. (1987) found that spraying caused a 2-fold increase in yield; interestingly, the same yield increase was found by treating the plants with Fe-EDDHA. Apoplastic pH has been shown to be related to plasmalemma proton pumps (Petzold and Dahse, 1988; Hoffmann et al., 1992) and spraying chlorotic leaves with fusicoccin resulted in a lowering of leaf apoplastic pH (Hoffmann et al., 1992) and in leaf re-greening (Mengel and Geurtzen, 1988). Based on these observations, Mengel (1995) hypothesized that high pH in the leaf apoplast hampers the reduction of Fe3+-citrate; reduction of Fe3+ is the prerequisite for the transport of Fe2+ across the plasmalemma (Chaney et al., 1972;Fox et al., 1996). Recently, a Fe2+ transporter has been identified in yeast (Eide et al., 1996). High pH in the root medium depressed the reduction of Fe3+ complexes (Romera et al., 1991). The investigations of Römheld and Marschner (1983), Toulon et al. (1992), and Susin et al. (1996) have shown that the reduction of Fe3+ in the apoplast of intact roots occurred at low pH. Various researchers (e.g.Brüggemann and Moog, 1989) working with membrane vesicles from barley roots found a pH optimum of Fe3+ reduction at pH 6.8; others (e.g. Holden et al., 1991), using vesicles from tomato roots, found an optimum of pH 6.5 for the reduction of Fe3+. These high pH optima, however, presumably relate to the cytosolic side of the plasma membrane-located Fe3+ reductase and were also found for vesicles from mesophyll cells (Brüggemann et al., 1993; Rombola et al., 1999). The pH optimum for the apoplastic domain of the Fe3+ reductase appeared to be lower (Mengel, 1995). If this apoplastic condition is not met, substantial amounts of Fe remain in the apoplast and are not transported into the symplasm, where it is required for cellular processes. The main objective of this study was to test the pH dependence of Fe3+ reduction in the leaf apoplast. Also, apoplastic pH measurements were carried out with excised leaves fed via the petiole with xylem sap obtained from plants grown on NO3 −, NO3 −/HCO3 −, NH4 +, or NH4NO3 as a control. It was possible to display by use of microscope image analysis apoplastic pH at the cellular level and apoplastic Fe3+reduction in intact leaf tissue. Since Fe chlorosis is a symptom of young leaves, measurements were carried out with young green leaves before leaf chlorosis occurred. MATERIALS AND METHODS Chemicals 2′,7′-Bis-(2-carboxyethyl)-5(and-6)-carboxyfluorescein- dextran and rhodamine were purchased from Molecular Probes (Eugene, OR). All other chemicals were from Sigma Chemical (St. Louis). Plant Growth Sunflower (Helianthus annuus L. cv Solostar) seeds were soaked with 0.5 mm CaSO4(24 h) and then germinated under darkness in a humid chamber at 25°C for 2 d. Plants were cultivated at 25°C during the day (15 h) and at 20°C during the night (9 h) in nutrient solution for 14 d. The control plants were grown for 14 d with NH4NO3. The plants of the other treatments were cultivated for 12 d in NH4NO3 and then transferred for another 2 d to two different N forms, NH4Cl and Ca(NO3)2, and in one treatment to Ca(NO3)2 and KHCO3. The total N concentration in each treatment was 6 mm, and the HCO3 − concentration was 10 mm. In a further treatment plants were cultivated for 9 d in 3 mmNH4NO3, then transferred for 2 d in solution without N, and then cultivated for a further 3 d with 1 mm NH4Cl. The Fe concentration in all series was 1 μm. The basic nutrient solution was as described by Kosegarten et al. (1998). Collection and Analysis of the Xylem Sap Xylem sap was obtained by sampling the exudation sap as described by Van Beusichem et al. (1988). Collection was carried out 4 h after the onset of the photoperiod, and the xylem sap was sampled for 60 min after plants were decapitated about 3 cm above the root. The sap from the first 5 min was discarded. Analyses of pH and of N compounds (NO3 −, NH4 +, and amino acids) were carried out on fresh samples. The pH measurements were conducted with an electrode (U402 M3/S7/60, Ingold, Mettler Toledo, Steinbach, Germany). NO3 − was analyzed by means of a continuous flow analyzer (Technicon Autoanalyzer II, Bran and Luebbe, Hamburg, Germany). NH4 + and amino acids were determined with an amino acid analyzer (Biotronic LC 3000, Eppendorf, Maintal, Germany). The method was modified according to Moore and Stein (1954). Samples were centrifuged at 15,000g for 15 min at 4°C, and a 20-μL aliquot of the supernatant was taken and isolated by a cation-exchange column (Eppendorf Biotronic, TS 01044P). Separation was carried out in a buffer system (Eppendorf Biotronic, Typ H1) at increasing pH at a flow rate of 0.2 mL min−1. Each sample comprised the xylem sap from four plants (xylem sap pH) and 24 plants (N compounds), respectively. Xylem sap pH was measured in 20 samples (n = 20). Apoplastic pH Measurements in Intact Sunflower Leaves Measurements of leaf apoplastic pH were carried out according to the method of Hoffmann and Kosegarten (1995), working with young leaves with a leaf area of about 800 mm2. Apoplastic pH was measured after infiltration of fresh xylem sap obtained from the different nutritional N sources into excised leaves. If not noted otherwise, the apoplastic pH was monitored in the interveinal area at the leaf base. Apoplastic pH measurements were conducted with (a) a fluorescence photometer (LS 50, Perkin-Elmer Applied Biosystems, Foster City, CA) at the tissue level on leaf areas of 9 mm2 at three different sites per leaf (in each treatment five leaves were analyzed; n = 15), and (b) under a fluorescence microscope (Axiotron/UV-fluorescence microscope, Carl Zeiss, Jena, Germany) at the cellular level. The basic configuration of microscope analysis was as described by Hoffmann and Kosegarten (1995). Excitation light between 450 and 490 nm was specified with a monochromator (bandwidth 15 nm). A measuring diaphragm of 30 × 150 μm was positioned on various cell areas (hair cells, stomata, epidermal cells, and xylem vessels). The illumination field diaphragm was about 20% larger than the measuring diaphragm. Apoplastic pH of xylem vessels (first to fourth order) and hair cells was examined at nine positions per leaf blade; in each treatment three leaves were examined (n = 27). To investigate apoplastic pH distribution in the intercostal leaf area, 20 cell complexes consisting of three to five epidermal and stomatal cells on leaf areas of 50 mm2 per leaf at the base were examined; in each treatment monitoring of apoplastic pH was conducted with five leaves (n = 100). Apoplastic pH gradients were also measured by microscope image analysis as described byHoffmann and Kosegarten (1995). A back-illuminated integrating CCD camera (Princeton Applied Research, Trenton, NJ) was used to improve the signal-to-noise ratio. To monitor the apoplasic pH of green and chlorotic areas of leaves with intercostal chlorosis, leaves were only perfused with 0.1 mm MgCl2, 0.1 mmCaCl2, and 2 mm KCl. The apoplastic pH was monitored with a fluorescence microscope (Axiotron/UV-fluorescence microscope, Carl Zeiss) in the chlorotic intercostal area (epidermal and stomatal cells) and in the green xylem vessels (mid-rib and first order veins). Green leaves were also examined for comparison. Ten positions per leaf blade were examined and pH measurements were conducted with three leaves (n = 30). Measurement of Fe3+ Reduction in Relation to Apoplastic pH in Intact Sunflower Leaves Fe3+ reduction in relation to apoplastic pH was examined in the xylem vessels (first order according to Canny, 1990) by microscope image analysis. Youngest leaves were excised, ferrozine (1 mm) was preloaded into the leaf for 24 h, and then for a further 6 h, 80 μmFeCl3 and 80 μm citrate were perfused in the presence of various buffers: 100 mm2-(N-morpholino)-ethanesulfonic acid (MES)/KOH, pH 4.0 to 6.5, and 100 mm 4-(2-hydroxyethyl)-1-piperazine 2-ethanesulfonic acid (HEPES)/KOH, pH 7.0 to 8.0. Ferrozine specifically complexes Fe2+ and exhibits an absorption maximum at 560 nm (Stookey, 1970). At 720 nm the Fe2+-ferrozine complex shows no absorption (data not shown). The principle of the measurement is based on monitoring the light transmission at 560 nm in the apoplast area of the xylem vessel. To compensate for differences in leaf absorption, light transmission was also measured at 720 nm. By calculating the ratio of light transmission at 720 nm and at 560 nm, a specific measure for the Fe2+-ferrozine complex in the xylem vessel was obtained. The light transmission ratio was calculated on frames of 512 × 512 pixels captured by a standard CCD camera (XC57CE, Sony, Tokyo). The resulting ratio values were displayed in pseudocolor. Figure 1 shows the light transmission at 720 nm (A) and at 560 nm (B) of a control leaf without ferrozine perfusion. The yellow pseudocolor in the xylem vessel corresponds to the maximal light transmission ratio (C). The histogram of Figure 1D shows the distribution of pixel gray values (0–255) with a maximum at a gray level of 101.3 ± 3.9 (n = 9), representing the maximum of the light transmission ratio in the xylem vessel (yellow pseudocolor). Fig. 1. Open in new tabDownload slide Light transmission at 720 nm (A) and 560 nm (B) and the maximal light transmission ratio (720/560 nm; C) in the xylem vessel of a control leaf (without ferrozine) of sunflower. The maximal light transmission ratio (720/560 nm) in the xylem vessel (without Fe-ferrozine complex) is displayed by yellow pseudocolor (C). Scale, 240 μm. The histogram (D) shows the distribution of the pixel intensity in the ratio picture (C) of the xylem vessel. The maximal light transmission ratio shows a pixel value of 101.3 ± 3.9 (n = 9). Fig. 1. Open in new tabDownload slide Light transmission at 720 nm (A) and 560 nm (B) and the maximal light transmission ratio (720/560 nm; C) in the xylem vessel of a control leaf (without ferrozine) of sunflower. The maximal light transmission ratio (720/560 nm) in the xylem vessel (without Fe-ferrozine complex) is displayed by yellow pseudocolor (C). Scale, 240 μm. The histogram (D) shows the distribution of the pixel intensity in the ratio picture (C) of the xylem vessel. The maximal light transmission ratio shows a pixel value of 101.3 ± 3.9 (n = 9). The apoplastic pH of first-order xylem vessels after buffer infiltration was measured in separate experiments using fluorescein isothiocyanate-dextran and 2′,7′-bis-(2-carboxyethyl)-5(and-6)-carboxyfluorescein-dextran (100 μm each). The different light transmission ratios after the various pH treatments in relation to the formation of Fe2+-ferrozine are shown as different distributions of the pixel intensity in the ratio picture of the xylem vessels (see histograms in Fig. 6). A high pixel value represents a reduction in the light transmission ratio. Changes in gray levels were expressed in pseudocolor. Light transmission at each apoplastic pH value was measured in nine different areas of first-order xylem vessels per leaf (n = 9), and each pH treatment consisted of three leaves (n = 27). In the xylem vessel of the unbuffered leaf (with a xylem apoplastic pH ≤ 5.0; Table V), the light transmission ratio was minimal due to high formation of the Fe2+-ferrozine complex. The minimal light transmission ratio shows a maximum at a gray level of 141.2 ± 4.1 (n = 27; not shown). After subtracting the minimal pixel value of the control leaf (100.5 ± 4.2; n = 27), the maximal rate of Fe3+ reduction was obtained and was defined as 100% (Table V). For each pH treatment the corresponding percentage of Fe3+ reduction was calculated (Table V). To check for variations in dye loading, the pH-independent fluorescent dye rhodamine (100 μm) was also perfused into leaves. Excitation of rhodamine was conducted at 560 nm and emission was observed at 580 nm, cutting off reflected excitation light by use of a long-pass filter (OG 570, Schott, Mainz, Germany). Statistical Treatment Significant differences between the control and the other nutritional treatments were calculated for xylem sap pH and for leaf apoplastic pH by use of the t test (Köhler et al., 1984; Table I). Table I. Xylem sap pH (n = 20; ±sd) and apoplast pH (n = 15; ±sd) in the intercostal area of young sunflower leaves after infiltration with xylem sap of the different nutritional N sources Nutritional N Source . Xylem Sap pH . Apoplast pH . Light . Dark (5 h) . NH4NO3 (3 mm)1-a 5.12 ± 0.08 5.36 ± 0.06 5.47 ± 0.07 NH4 + (6 mm) 5.14 ± 0.07 5.36 ± 0.06 5.43 ± 0.07 NH4 + (1 mm) 5.31 ± 0.02** 5.41 ± 0.09 Not determined NO3 − (6 mm) 5.45 ± 0.05*** 5.50 ± 0.05** 5.62 ± 0.07* NO3 −/HCO3 − (6 mm) 5.53 ± 0.04*** 5.51 ± 0.07* 5.60 ± 0.08* Nutritional N Source . Xylem Sap pH . Apoplast pH . Light . Dark (5 h) . NH4NO3 (3 mm)1-a 5.12 ± 0.08 5.36 ± 0.06 5.47 ± 0.07 NH4 + (6 mm) 5.14 ± 0.07 5.36 ± 0.06 5.43 ± 0.07 NH4 + (1 mm) 5.31 ± 0.02** 5.41 ± 0.09 Not determined NO3 − (6 mm) 5.45 ± 0.05*** 5.50 ± 0.05** 5.62 ± 0.07* NO3 −/HCO3 − (6 mm) 5.53 ± 0.04*** 5.51 ± 0.07* 5.60 ± 0.08* Significant differences between the control and the other treatments at three different levels are denoted by: *, 5% level; **, 1% level;***, 0.1% level. F1-a Control. Open in new tab Table I. Xylem sap pH (n = 20; ±sd) and apoplast pH (n = 15; ±sd) in the intercostal area of young sunflower leaves after infiltration with xylem sap of the different nutritional N sources Nutritional N Source . Xylem Sap pH . Apoplast pH . Light . Dark (5 h) . NH4NO3 (3 mm)1-a 5.12 ± 0.08 5.36 ± 0.06 5.47 ± 0.07 NH4 + (6 mm) 5.14 ± 0.07 5.36 ± 0.06 5.43 ± 0.07 NH4 + (1 mm) 5.31 ± 0.02** 5.41 ± 0.09 Not determined NO3 − (6 mm) 5.45 ± 0.05*** 5.50 ± 0.05** 5.62 ± 0.07* NO3 −/HCO3 − (6 mm) 5.53 ± 0.04*** 5.51 ± 0.07* 5.60 ± 0.08* Nutritional N Source . Xylem Sap pH . Apoplast pH . Light . Dark (5 h) . NH4NO3 (3 mm)1-a 5.12 ± 0.08 5.36 ± 0.06 5.47 ± 0.07 NH4 + (6 mm) 5.14 ± 0.07 5.36 ± 0.06 5.43 ± 0.07 NH4 + (1 mm) 5.31 ± 0.02** 5.41 ± 0.09 Not determined NO3 − (6 mm) 5.45 ± 0.05*** 5.50 ± 0.05** 5.62 ± 0.07* NO3 −/HCO3 − (6 mm) 5.53 ± 0.04*** 5.51 ± 0.07* 5.60 ± 0.08* Significant differences between the control and the other treatments at three different levels are denoted by: *, 5% level; **, 1% level;***, 0.1% level. F1-a Control. Open in new tab RESULTS Effects of N Form and HCO3 − on Xylem Sap pH and Leaf Apoplastic pH Table I shows that under alkaline conditions in the nutrient solution (NO3 − and NO3 −/HCO3 −), both xylem sap pH and apoplastic pH significantly increased compared with the NH4NO3 treatment (control). Addition of HCO3 − had no influence on apoplastic pH compared with the NO3 − treatment. Moreover, when plants were exclusively fed with NH4 + (at both 1 and 6 mm), the apoplastic pH decreased. In darkness, the apoplastic pH increased by about 0.1 pH unit in all treatments. The apoplastic pH values shown in Table I are mean values at the leaf tissue level (9 mm2) from the intercostal area at the leaf base and were recorded by use of fluorescence photometry. Thus, with this experimental approach, the mean pH response in the apoplast of several hundred cells and also of various cell types, i.e. the apoplastic pH of leaf epidermal, stomatal, and hair cells, was measured. Since such apoplastic pH values at the tissue level may average out more pronounced pH changes at the cellular level, the local apoplastic pH of the various cell types was recorded by fluorescence microscopy (Fig. 2; Table II) combined with digital image processing (Fig. 3). These approaches revealed distinctly different apoplastic pH values at various microsites at the cellular level in the leaf. Fig. 2. Open in new tabDownload slide Relative frequency of apoplast pH (n = 100) of epidermal and stomatal cells in the intercostal area at the leaf base in relation to different N nutrition and light/dark changes. A, 1 mmNH4 +; B, 3 mmNH4NO3; C, 6 mmNO3 −/10 mmHCO3 −. Dark period, 5 h. White bars, Light; black bars, dark. Fig. 2. Open in new tabDownload slide Relative frequency of apoplast pH (n = 100) of epidermal and stomatal cells in the intercostal area at the leaf base in relation to different N nutrition and light/dark changes. A, 1 mmNH4 +; B, 3 mmNH4NO3; C, 6 mmNO3 −/10 mmHCO3 −. Dark period, 5 h. White bars, Light; black bars, dark. Table II. Effect of different N forms (NH4NO3, NO3 − in the presence of HCO3 −) on apoplast pH in xylem vessels (first to fourth order according to Canny, 1990) and of hair cells (n = 27; ±sd) of young sunflower leaves after infiltration with xylem sap of the different nutritional N sources Apoplast pH . Light . Dark . NH4NO3 . NO3 −/HCO3 − . NH4NO3 . NO3 −/HCO3 − . Xylem vessel 5.14 ± 0.24 5.16 ± 0.24 5.21 ± 0.10 5.22 ± 0.14 Hair cell 5.61 ± 0.23 5.72 ± 0.22 5.80 ± 0.15 5.82 ± 0.23 Apoplast pH . Light . Dark . NH4NO3 . NO3 −/HCO3 − . NH4NO3 . NO3 −/HCO3 − . Xylem vessel 5.14 ± 0.24 5.16 ± 0.24 5.21 ± 0.10 5.22 ± 0.14 Hair cell 5.61 ± 0.23 5.72 ± 0.22 5.80 ± 0.15 5.82 ± 0.23 Open in new tab Table II. Effect of different N forms (NH4NO3, NO3 − in the presence of HCO3 −) on apoplast pH in xylem vessels (first to fourth order according to Canny, 1990) and of hair cells (n = 27; ±sd) of young sunflower leaves after infiltration with xylem sap of the different nutritional N sources Apoplast pH . Light . Dark . NH4NO3 . NO3 −/HCO3 − . NH4NO3 . NO3 −/HCO3 − . Xylem vessel 5.14 ± 0.24 5.16 ± 0.24 5.21 ± 0.10 5.22 ± 0.14 Hair cell 5.61 ± 0.23 5.72 ± 0.22 5.80 ± 0.15 5.82 ± 0.23 Apoplast pH . Light . Dark . NH4NO3 . NO3 −/HCO3 − . NH4NO3 . NO3 −/HCO3 − . Xylem vessel 5.14 ± 0.24 5.16 ± 0.24 5.21 ± 0.10 5.22 ± 0.14 Hair cell 5.61 ± 0.23 5.72 ± 0.22 5.80 ± 0.15 5.82 ± 0.23 Open in new tab Fig. 3. Open in new tabDownload slide Apoplast pH (A) and fluorescence intensity after excitation at 490 nm (B) of the upper cell layer of a sunflower leaf after 5 h of darkness as examined by microscope image analysis. Plants were cultivated with 6 mm NO3 −/10 mmHCO3 −. The light-blue pseudocolor in the left picture corresponds to a pH of around 5.7, the green pseudocolor to a pH of around 6.5, and the yellow pseudocolor to a pH of around 7.0. The fluorescence intensity is high in the xylem vessels and in the stomatal region, as shown by the red, yellow, and light-blue pseudocolors (B). Scale, 80 μm. Fig. 3. Open in new tabDownload slide Apoplast pH (A) and fluorescence intensity after excitation at 490 nm (B) of the upper cell layer of a sunflower leaf after 5 h of darkness as examined by microscope image analysis. Plants were cultivated with 6 mm NO3 −/10 mmHCO3 −. The light-blue pseudocolor in the left picture corresponds to a pH of around 5.7, the green pseudocolor to a pH of around 6.5, and the yellow pseudocolor to a pH of around 7.0. The fluorescence intensity is high in the xylem vessels and in the stomatal region, as shown by the red, yellow, and light-blue pseudocolors (B). Scale, 80 μm. Figure 2 shows the frequency distribution of apoplastic pH in complexes comprising about three to five leaf epidermal and stomatal cells in the interveinal leaf area. Independent of N form and the addition of HCO3 −, about 50% of the apoplastic pH values in these complexes was between pH 5.0 and 5.5; about 20% to 30% was between pH 5.5 and 6.0; and 10% to 20% of the apoplastic pH was ≤ 5.0. In darkness, the frequency distribution of apoplastic pH shifted to values between 5.5 and 6.0 (70%). Only under alkaline nutritional conditions was about 10% of the apoplastic pH ≥ 6.3 (Fig. 2C), which was not different between light and dark. Such leaf cell complexes with high apoplastic pH levels are indicated by green and yellow color (arrows) in Figure 3A. The light-blue color represents a mean pH of about 5.7. The restriction of high apoplastic alkalization to small complexes of epidermal and stomatal cells (about 10% of the leaf apoplast) under alkaline nutritional conditions explains the small overall pH increase (0.15 pH unit) at the tissue level (hundreds of leaf cells) compared with the NH4NO3 treatment (Table I). Figure 3B shows the distribution of the dye fluorescence intensity at 490 nm with highest intensities around the stomatal apoplast and in the xylem area. High fluorescence intensities are caused by the high optical pathlength of the xylem vessels and by dye enrichment around the stomatal apoplast at high transpiration rates, and can be eliminated by the use of the fluorescence ratio technique (see Hoffmann and Kosegarten, 1995). The apoplastic pH of the hair cells and of the xylem vessels is shown in Table II. Apoplastic pH at these microsites was affected by neither the N form nor HCO3 −. Apoplastic pH of the hair cells was considerably higher (0.5 pH unit) than that of the xylem. Table III shows the contribution of various N compounds in the xylem sap being infiltrated into the leaf. The concentration of NO3 −was higher in treatments of alkaline nutrition (NO3 − and NO3 −/HCO3 −) than in the NH4NO3treatment (control); it was lowest when plants were fed exclusively with NH4 +. The reverse was true for the concentration of NH4 +, Gln, and Asn in the xylem sap under the applied nutritional conditions. At 1 mmNH4 + in the nutrient solution, the NH4 +concentration in the xylem sap was not much different from that found in alkaline nutritional treatments. However, the Gln concentration in the xylem sap increased 7-fold over that under alkaline conditions. Table III. Concentration (mm) of various N compounds in the xylem sap of sunflower, as depending on the nutritional N source Nutritional N Source . NO3 − . NH4 + . Glu . Asp . Other Amino Acids . NH4NO3 (3 mm)3-a 20.08 0.64 3.42 0.76 0.47 NH4 + (6 mm) 1.49 0.92 7.49 1.17 1.10 NH4 + (1 mm) 0.69 0.30 11.88 0.74 1.61 NO3 − (6 mm) 30.45 0.32 1.39 0.38 0.24 NO3 −/HCO3 − (6 mm) 26.92 0.16 1.54 0.34 0.42 Nutritional N Source . NO3 − . NH4 + . Glu . Asp . Other Amino Acids . NH4NO3 (3 mm)3-a 20.08 0.64 3.42 0.76 0.47 NH4 + (6 mm) 1.49 0.92 7.49 1.17 1.10 NH4 + (1 mm) 0.69 0.30 11.88 0.74 1.61 NO3 − (6 mm) 30.45 0.32 1.39 0.38 0.24 NO3 −/HCO3 − (6 mm) 26.92 0.16 1.54 0.34 0.42 F3-a Control. Open in new tab Table III. Concentration (mm) of various N compounds in the xylem sap of sunflower, as depending on the nutritional N source Nutritional N Source . NO3 − . NH4 + . Glu . Asp . Other Amino Acids . NH4NO3 (3 mm)3-a 20.08 0.64 3.42 0.76 0.47 NH4 + (6 mm) 1.49 0.92 7.49 1.17 1.10 NH4 + (1 mm) 0.69 0.30 11.88 0.74 1.61 NO3 − (6 mm) 30.45 0.32 1.39 0.38 0.24 NO3 −/HCO3 − (6 mm) 26.92 0.16 1.54 0.34 0.42 Nutritional N Source . NO3 − . NH4 + . Glu . Asp . Other Amino Acids . NH4NO3 (3 mm)3-a 20.08 0.64 3.42 0.76 0.47 NH4 + (6 mm) 1.49 0.92 7.49 1.17 1.10 NH4 + (1 mm) 0.69 0.30 11.88 0.74 1.61 NO3 − (6 mm) 30.45 0.32 1.39 0.38 0.24 NO3 −/HCO3 − (6 mm) 26.92 0.16 1.54 0.34 0.42 F3-a Control. Open in new tab Apoplastic pH in Green and Intercostal Chlorotic Leaves To investigate the influence of chlorosis on leaf apoplastic pH, the intercostal area and xylem vessels of leaves with intercostal chlorosis (arrows; Fig. 4) and of green control leaves were analyzed. No pH differences were found in the xylem vessels of the mid-rib and the first-order veins for green and intercostal chlorotic leaves. The apoplastic pH in the intercostal region of the chlorotic leaf was remarkably higher (about 0.5 pH unit) than that monitored in the green vessels and that in the intercostal area of the green leaf (Table IV). Fig. 4. Open in new tabDownload slide Intercostal chlorosis in young sunflower leaves. The apoplast pH was measured with a fluorescence microscope in the yellow intercostal area and the green xylem vessels (arrows). Fig. 4. Open in new tabDownload slide Intercostal chlorosis in young sunflower leaves. The apoplast pH was measured with a fluorescence microscope in the yellow intercostal area and the green xylem vessels (arrows). Table IV. Apoplast pH in the intercostal area and of xylem vessels (mid-rib and first order) in the tip of green leaves and of leaves with intercostal chlorosis (n = 30; ±sd) of H. annuus Apoplast pH . Green Leaf . Leaf with Intercostal Chlorosis . Mid-Rib 4.78 ± 0.10 4.68 ± 0.24 Xylem vessel (first order) 4.51 ± 0.22 4.72 ± 0.21 Intercostal area 4.46 ± 0.18 5.30 ± 0.14 Apoplast pH . Green Leaf . Leaf with Intercostal Chlorosis . Mid-Rib 4.78 ± 0.10 4.68 ± 0.24 Xylem vessel (first order) 4.51 ± 0.22 4.72 ± 0.21 Intercostal area 4.46 ± 0.18 5.30 ± 0.14 In the leaf with intercostal chlorosis the apoplast pH was measured in the area of green leaf veins and in the chlorotic intercostal area. Apoplast pH was monitored in the light after perfusion of 0.1 mm MgCl2, 0.1 mm CaCl2, and 2 mm KCl. Open in new tab Table IV. Apoplast pH in the intercostal area and of xylem vessels (mid-rib and first order) in the tip of green leaves and of leaves with intercostal chlorosis (n = 30; ±sd) of H. annuus Apoplast pH . Green Leaf . Leaf with Intercostal Chlorosis . Mid-Rib 4.78 ± 0.10 4.68 ± 0.24 Xylem vessel (first order) 4.51 ± 0.22 4.72 ± 0.21 Intercostal area 4.46 ± 0.18 5.30 ± 0.14 Apoplast pH . Green Leaf . Leaf with Intercostal Chlorosis . Mid-Rib 4.78 ± 0.10 4.68 ± 0.24 Xylem vessel (first order) 4.51 ± 0.22 4.72 ± 0.21 Intercostal area 4.46 ± 0.18 5.30 ± 0.14 In the leaf with intercostal chlorosis the apoplast pH was measured in the area of green leaf veins and in the chlorotic intercostal area. Apoplast pH was monitored in the light after perfusion of 0.1 mm MgCl2, 0.1 mm CaCl2, and 2 mm KCl. Open in new tab Fe3+ Reduction in Relation to Leaf Apoplastic pH Various pH-buffered solutions were infiltrated into excised leaves, and both xylem apoplastic pH and Fe3+reduction were measured; the latter by the formation of the Fe2+-ferrozine complex in the xylem vessels (first order veins; Table V). In Figure 5the light transmission of different leaves in the region of the xylem vessel after infiltration of Fe3+-citrate and ferrozine at a low (pH 5.4; Fig. 5, A and B) and a high (pH 7.7; Fig.5, C and D) xylem apoplastic pH is shown. After both pH treatments the light transmission at 720 nm in the xylem vessels (Fig. 5, A and C) was similar and comparable to the control leaf without ferrozine (Fig. 1A). Small differences were due to differences in leaf absorption. At 560 nm and high apoplastic pH (pH 7.7; Fig. 5D), light transmission in the xylem vessels was high and was similar to that in the control leaf (Fig. 1B). This finding shows that Fe2+-ferrozine formation at pH 7.7 was negligible. In contrast, at low pH levels (pH 5.4), light transmission at 560 nm was much reduced, as shown by fewer whitish strands in this picture (Fig. 5B). Therefore, at low pH, Fe3+ reduction took place and the Fe2+-ferrozine complex was formed. Table V. Fe3+ reduction in intact leaves of sunflower in relation to apoplast pH of xylem vessels (first order) Buffer (pH) . Xylem Apoplast pH . Fe3+ Reduction . Rhodamine Fluorescence Intensity . % Leaf without buffer ≤5.0 (Light) 100 2,690.1 ± 202.2 100 mm MES (pH 5.0) 5.44 ± 0.07 98 ± 5 2,651.8 ± 208.4 100 mm MES (pH 6.0) 5.92 ± 0.43 78 ± 15 n.d.5-a 100 mm HEPES (pH 7.25) 6.86 ± 0.62 54 ± 20 n.d. 100 mm HEPES (pH 8.0) 7.71 ± 0.41 22 ± 11 2,821.3 ± 177.5 Buffer (pH) . Xylem Apoplast pH . Fe3+ Reduction . Rhodamine Fluorescence Intensity . % Leaf without buffer ≤5.0 (Light) 100 2,690.1 ± 202.2 100 mm MES (pH 5.0) 5.44 ± 0.07 98 ± 5 2,651.8 ± 208.4 100 mm MES (pH 6.0) 5.92 ± 0.43 78 ± 15 n.d.5-a 100 mm HEPES (pH 7.25) 6.86 ± 0.62 54 ± 20 n.d. 100 mm HEPES (pH 8.0) 7.71 ± 0.41 22 ± 11 2,821.3 ± 177.5 Fe3+ reduction and apoplast pH was monitored after infiltration of various buffer solutions to the xylem vessels (n = 27; ±sd). The pH-insensitive dye rhodamine was infiltrated into leaves and used as an internal standard to check for variations in dye loading. F5-a n.d., Not determined. Open in new tab Table V. Fe3+ reduction in intact leaves of sunflower in relation to apoplast pH of xylem vessels (first order) Buffer (pH) . Xylem Apoplast pH . Fe3+ Reduction . Rhodamine Fluorescence Intensity . % Leaf without buffer ≤5.0 (Light) 100 2,690.1 ± 202.2 100 mm MES (pH 5.0) 5.44 ± 0.07 98 ± 5 2,651.8 ± 208.4 100 mm MES (pH 6.0) 5.92 ± 0.43 78 ± 15 n.d.5-a 100 mm HEPES (pH 7.25) 6.86 ± 0.62 54 ± 20 n.d. 100 mm HEPES (pH 8.0) 7.71 ± 0.41 22 ± 11 2,821.3 ± 177.5 Buffer (pH) . Xylem Apoplast pH . Fe3+ Reduction . Rhodamine Fluorescence Intensity . % Leaf without buffer ≤5.0 (Light) 100 2,690.1 ± 202.2 100 mm MES (pH 5.0) 5.44 ± 0.07 98 ± 5 2,651.8 ± 208.4 100 mm MES (pH 6.0) 5.92 ± 0.43 78 ± 15 n.d.5-a 100 mm HEPES (pH 7.25) 6.86 ± 0.62 54 ± 20 n.d. 100 mm HEPES (pH 8.0) 7.71 ± 0.41 22 ± 11 2,821.3 ± 177.5 Fe3+ reduction and apoplast pH was monitored after infiltration of various buffer solutions to the xylem vessels (n = 27; ±sd). The pH-insensitive dye rhodamine was infiltrated into leaves and used as an internal standard to check for variations in dye loading. F5-a n.d., Not determined. Open in new tab Fig. 5. Open in new tabDownload slide Light transmission in the xylem vessels at pH 5.4 (A, 720 nm; B, 560 nm) and at pH 7.7 (C, 720 nm; D, 560 nm) after infiltration of Fe3+-citrate and ferrozine at different apoplastic pH levels. Scale, 240 μm. Fig. 5. Open in new tabDownload slide Light transmission in the xylem vessels at pH 5.4 (A, 720 nm; B, 560 nm) and at pH 7.7 (C, 720 nm; D, 560 nm) after infiltration of Fe3+-citrate and ferrozine at different apoplastic pH levels. Scale, 240 μm. The images in Figure 6 show the light transmission ratio (720/560 nm) after infiltration of Fe3+-citrate and ferrozine at high and low xylem apoplastic pH. At pH 7.7 (Fig. 6A) the light transmission ratio in the xylem vessel was high, as indicated by yellow pseudocolor. The blue pseudocolor of the ratio picture in the xylem vessel of Figure 6B indicates a low light transmission ratio at low apoplastic pH (pH 5.4). The degree of light transmission ratio at various apoplastic pH levels corresponded to the degree of Fe2+-ferrozine formation and therefore to the capacity of Fe3+reduction. Table V summarizes the percentage data of the mean light transmission ratio of the Fe2+-ferrozine complex in the xylem vessels under various pH conditions. The capacity of Fe3+ reduction decreased with increasing apoplastic pH. Formation of the Fe2+-ferrozine complex, and therefore Fe3+ reduction, was the same at an apoplastic pH ≤ 5.0 (leaf without buffer) and at pH 5.4 (leaf with 100 mm MES, pH 5.0; TableV); therefore, the effect of buffer infiltration appeared negligible. To check for variations in dye loading, the fluorescence intensity of the pH-independent dye rhodamine was monitored as a direct measure of dye concentration inside the xylem vessels. Table V shows no difference in the fluorescence intensity of rhodamine between the pH treatments; therefore, for ferrozine infiltration variations in dye loading could be excluded as well. Fig. 6. Open in new tabDownload slide Light transmission ratio (720/560 nm) in the xylem vessels (A and B) after infiltration of Fe3+-citrate and ferrozine at different apoplastic pH levels. A, pH 7.7; yellow pseudocolor represents high light transmission ratio. B, pH 5.4; light-blue pseudocolor shows low light transmission ratio. Scale, 240 μm. The distribution of the pixel intensity in the histogram is shown at pH 7.7 (C) and at pH 5.4 (D), with a maximum at 106.6 ± 3.4 (n = 9) and at 139.7 ± 5.8 (n = 9), respectively. A shift to higher pixel values indicates a reduction in the light transmission ratio. Fig. 6. Open in new tabDownload slide Light transmission ratio (720/560 nm) in the xylem vessels (A and B) after infiltration of Fe3+-citrate and ferrozine at different apoplastic pH levels. A, pH 7.7; yellow pseudocolor represents high light transmission ratio. B, pH 5.4; light-blue pseudocolor shows low light transmission ratio. Scale, 240 μm. The distribution of the pixel intensity in the histogram is shown at pH 7.7 (C) and at pH 5.4 (D), with a maximum at 106.6 ± 3.4 (n = 9) and at 139.7 ± 5.8 (n = 9), respectively. A shift to higher pixel values indicates a reduction in the light transmission ratio. DISCUSSION Apoplastic pH of Young Green Leaves under Alkaline Conditions Fe chlorosis occurs mainly on calcareous soils, where NO3 − is the exclusive N form in the soil solution due to increased nitrification (Darrah et al., 1986) and NH3 volatilization (Paramasivam and Alva, 1997). As shown in Table I, the N form clearly influenced xylem sap pH; the highest pH values were observed with NO3 − nutrition in solution culture. Interestingly, the presence of HCO3 − (as in the soil solution of calcareous soils) did not influence xylem sap pH (Table I). Presumably, proton pumps adjacent to the xylem (Canny, 1987; Wilson et al., 1988) are efficient enough to neutralize HCO3 −. In addition, the low partial pressure in the xylem (Zimmermann et al., 1993) should favor the formation of CO2 from HCO3 −. Feeding young excised leaves with the xylem sap obtained from various N treatments resulted in a substantial apoplastic pH increase at microsites (pH ≥ 6.3) in the intercostal leaf area only in plants that had received exclusively NO3 − from the nutrient solution (Figs. 2C and 3A). Leaf apoplastic alkalization upon NO3 − nutrition was not homogenous in the intercostal leaf area, when inspected at the cellular level. About 10% of the leaf apoplast showed elevated pH levels ≥ 6.3 (Fig. 2C) at distinct apoplastic microsites on complexes of stomatal and epidermal cells. The section shown in Figure 3 with an area of 500 × 500 μm2 comprises about 100 cells. With microscope imaging only the upper cell layer could be analyzed and showed a number of epidermal and stomatal cells, as indicated by the green and yellow color (arrows), with pH levels ≥ 6.3 (Fig. 3A). The apoplast of underlying mesophyll cells may also show these increased pH levels, but this has to be proven by use of confocal microscopy. Such microsites of high apoplastic pH were not found in the case of NH4 +and NH4NO3 supply (Fig. 2, A and B). The apoplastic pH values shown in Table I are mean data at the tissue level of several hundred leaf cells and show a significantly higher apoplastic pH of only 0.15 pH units under alkaline nutritional conditions (Table I) compared with the control (NH4NO3 treatment). From this observation it is clear that at the tissue level apoplastic pH measurements average out more pronounced apoplastic pH increases at the cellular level (Figs. 2C and 3A). Therefore, the small pH increases at the tissue level (Table I) do not reflect the real physiological, site-specific apoplastic pH response of young leaves exposed to alkaline nutritional conditions. Such microsites of high apoplastic pH are distributed throughout the leaf blade (not shown), and we speculate that they are related to growing sites of a young leaf where high NO3 − uptake rates occur. This means that these sites need N for protein synthesis, as well as NO3 − for osmotic reasons in expanding cells (McIntyre, 1997). According to the composition of N compounds in the xylem sap (Table III), when NO3 − is the sole N source, it may also provide N for protein synthesis. Like N demand, Fe demand in the growing cells is high, in particular for the synthesis of ribonucleotide reductase (Reichard, 1993) and for chlorophyll synthesis (Terry and Abadia, 1986). This assumption is in line with the observation of Kosegarten et al. (1998) that in sunflowers fed with NO3 − the development of leaf primordia was inhibited in contrast to the treatment with NH4NO3 nutrition. It is well known from the work of Maksymowych (1973) that the entire blade of a dicotyledonous leaf is involved in growth. Accordingly, we have conducted a frequency study at the leaf base related to a leaf area of 50 mm2 (Fig. 2). Interestingly, in older leaves apoplastic alkalization induced by NO3 − nutrition was not observed (not shown), and this may be the reason why mature leaves are not sensitive to Fe chlorosis. In mature leaves, growth processes have been completed and, unlike young leaves, have a low demand for NO3 − (Van Egmond and Breteler, 1972). In addition, mature leaves show high net photosynthetic rates (Turgeon and Webb, 1975), presumably providing enough energy for the plasmalemma H+ pump and therefore may efficiently regulate leaf apoplastic pH. The process of apoplastic alkalization supposedly resulted from the removal of protons from the apoplast upon proton cotransport of NO3 − (Ullrich, 1992;Crawford and Glass, 1998) into the adjacent cells. NO3 − typically is the main inorganic N form transported to the leaf (Pate, 1973; Van Beusichem et al., 1988) and, presumably, at microsites of the meristematic and rapidly expanding leaf cells, high NO3 − uptake rates necessary for the growth of a young leaf occur. As evident from TableIII, the NO3 −concentration in the xylem sap was high in all treatments with NO3 − in the nutrient solution. In the treatment with NH4NO3 and NH4 +, however, the NH4 + concentration in the xylem sap was relatively high (Table III). Therefore, in these treatments NH4 + also may play an important role in N nutrition of leaf cells. Even at a concentration of 1 mmNH4 + in the nutrient solution (Table III), reflecting the concentration of most agricultural soil solutions (Wolt, 1994), a concentration of 0.3 mmNH4 + and a low NO3 − concentration (0.69 mm) were found in the xylem sap. In the leaf apoplast ofBrassica napus grown on sandy soil, Husted and Schjoerring (1995) found NH4 +concentrations up to 0.8 mm and reported high uptake rates for NH4 +, which according to Nielsen and Schjoerring (1998), may be related to a transporter with channel-like properties. NH4 + uptake depolarizes the membrane potential (Herrmann and Felle, 1995) and stimulates the H+-ATPase, which results in a low apoplastic pH (Kosegarten et al., 1999). Until now, very little information has been available concerning NH4 + transport from the leaf apoplast into the symplasm. Ninnemann et al. (1994) isolated and characterized the AMT1 gene for a high-affinity NH4 + transporter in leaves of Arabidopsis. In addition to NH4 +, Gln was a major N compound in the xylem sap upon treatment with NH4NO3; with exclusive NH4 + supply, Gln was even the dominating N compound in the xylem sap (Table III). Uptake systems for amino acids in leaves have been identified by Van Bel et al. (1986)and uptake found to occur presumably via proton cotransport (Li and Bush, 1990; Williams et al., 1990). Amino acids are protonated at the apoplastic pH level, and therefore uptake into the mesophyll cell may remove fewer protons from the apoplast than with NO3 −. If NH4 + and/or amino acids contribute to the N nutrition of leaf mesophyll cells from the apoplast, the apoplastic pH may be lowered and high apoplastic pH levels at microsites would not prevail (Fig. 2, A and B). An increase of apoplastic pH here was observed between different cell types according to the following sequence: xylem vessel (Table II) < the main portion of epidermal and stomatal cells (Fig. 2) < hair cells (Table II). The observed cell-specific differences in apoplastic pH may result from a differential abundance of H+ pumps in the different leaf cells (Bouche-Pillon et al., 1994; Michelet and Boutry, 1995) and/or from differential removal of protons from the respective apoplast space because of differential N uptake of the various cells. In all nutritional treatments, the pH in the apoplast of the mesophyll, the xylem, and the hair cells was higher in darkness than in light (TablesI and II; Fig. 2, B and C). This finding indicates that apoplastic pH is influenced by the prevailing metabolic condition and is in line with previous results of Hoffmann and Kosegarten (1995). Mengel and Malissiovas (1982) have shown that net proton excretion of roots of intact vine trees was higher during the day than at night. Fe3+ Reduction in Relation to Apoplastic pH in Young Green Leaves Growing tissues need a continuous Fe supply (Brown, 1978), and the anatomy of the growing leaf tissue is complex (Taylor, 1997). It is of interest whether growing tissues receive Fe from the xylem and/or from the phloem. According to U.W. Stephan (personal communication) and in contrast to their earlier findings (Stephan and Scholz, 1993), Fe in the phloem sap is mainly transported in the form of a Fe3+ complex, presumably bound to a small peptide. In our study we used young leaves of about 800 mm2. At that developmental stage, showing high transpiration rates (not shown), the supply of Fe to the expanding leaf should proceed mainly via the xylem. Here, Fe is translocated in form of Fe3+-citrate (e.g. Tiffin, 1966; Clark et al., 1973). This Fe3+ complex presumably needs to be reduced before passing the plasmalemma (Chaney et al., 1972; Fox et al., 1996). Fe3+ reductase activity in the leaf has been evidenced (Brüggemann et al., 1993; De la Guardia and Alcantara, 1996) and has been suggested as the prerequisite for Fe uptake into the growing leaf cell (Crowley et al., 1991; Mengel, 1995). In this context it is of interest that the mesophyll tissue of young green leaves showed minute areas with a high apoplastic pH exclusively with NO3 −supply (Figs. 2C and 3A). As mentioned previously, we suggest that NO3 − is taken up with high rates at these microsites of high apoplastic pH, and that these microsites comprise meristematic and rapidly expanding cells, where NO3 − is used for protein synthesis and as a major osmoticum (McIntyre, 1997). Such cells also require Fe for the synthesis of ribonucleotide reductase (Reichard, 1993) and for chlorophyll synthesis (Terry and Abadia, 1986). If at such sites the activity of the Fe3+ reductase is restricted because of a high pH at the apoplastic domain of the reductase, intracellular Fe deficiency will occur, with a concurrent reduction in leaf growth (Mengel and Malissiovas, 1981; Kosegarten et al., 1998) and hampered chlorophyll synthesis (Terry and Abadia, 1986). In our experiments, Fe3+ reduction was measured by the formation of a Fe2+-ferrozine complex in the leaf xylem after infiltration of Fe3+-citrate in the presence of various buffers and was determined by means of assessing the light transmission in the xylem vessel (Figs. 1, 5, and6). As shown in Table V, Fe3+ reduction rates clearly declined upon increase of apoplastic pH in the xylem. Our measure for Fe3+ reduction is a relative one and the most important conclusion that can be drawn from our data is that the xylem of intact leaves shows a pH-dependent Fe3+ reduction, with maximal rates at apoplastic pH 5.0 and lower. With increasing xylem apoplastic pH, Fe3+ reduction concomitantly decreased; e.g. at pH 7.7 the reducing power was only 22% of that found at apoplastic pH 5.0 (Table V). To our knowledge, until now no relationship between Fe3+ reduction power and apoplastic pH in intact leaves has been described. The maximal rates of Fe3+ reduction in the leaf apoplast at apoplastic pH 5.0 and lower compare well with that in intact roots of B. napus (Toulon et al., 1992) and of Beta vulgaris (Susin et al., 1996). Toulon et al. (1992) found the highest reduction rate at pH 4.0 in the outer solution. Taking into account the maximal H+-buffer capacity of cell walls at around pKa 5 (Sentenac and Grignon, 1981), an apoplastic pH of around 5.0 (Felle, 1998; Kosegarten et al., 1999) with maximal Fe3+ reduction rates in the root apoplast is realistic. Because of experimental difficulties in monitoring Fe3+ reduction at the apoplastic side of intact leaf mesophyll (e.g. insensitivity of absorbance measurement at low optical pathlength), Fe3+ reduction was recorded in the leaf xylem. Also, the xylem is a part of the apoplast that is separated by the plasmalemma from the leaf cells surrounding the xylem vessels. It is assumed that also these plasma membranes are equipped with Fe3+ reductases and, in analogy to the monitoring of reduced formation of the Fe2+-ferrozine complex in the xylem vessels at high xylem apoplastic pH in the presence of HEPES buffer (Table V), we suggest that at apoplastic microsites of the interveinal leaf area with high apoplastic pH ≥ 6.3 (Figs. 2C and 3A) under alkaline nutritional conditions, Fe3+ reduction is clearly decreased. At about pH 7.0 in the xylem apoplast, Fe3+ reduction was reduced by about 50% (TableV) and at microsites with pH ≥ 7.0 (Fig. 2C), the rate of Fe3+ reduction should be even lower (Table V). Such an analogous conclusion is justified, because a similar pH-dependent pattern between Fe3+ reduction and outer solution pH was found in intact roots with maximal rates of Fe3+ reduction at low pH and a concomitant decrease with increasing pH (Toulon et al., 1992; Susin et al., 1996). Using the experimental approach of microscope imaging, we monitored, for the first time to our knowledge, Fe3+reduction at the apoplastic side in intact leaf tissue. It is interesting that here a similar apoplastic pH dependency of Fe3+ reduction prevails, as is the case for intact roots. With this experimental setup, a nonenzymatic, spontaneous Fe3+ reduction (e.g. by ascorbate) cannot be excluded. However, the pH-dependent response, as shown in Table V, is pronounced and therefore strongly indicates an enzymatic mechanism of Fe3+ reduction. Phenomenon of Intercostal Chlorosis The influence of chlorosis on leaf apoplast pH was investigated and the results are shown in Table IV. The striking pH difference between the apoplast of the green leaf veins (pH 4.5–4.7) and the chlorotic intercostal area (pH 5.3) presumably reflects the overall lower energetic status of chlorotic intercostal leaf regions. In contrast to the apoplastic pH measurements in young leaves before leaf chlorosis occurred (Fig. 2), the apoplastic pH of leaves with intercostal chlorosis was measured at the cellular level in the absence of NO3 − or other N forms (Table IV). Therefore, the apoplastic pH in the intercostal area of the chlorotic and, in particular, in the green control leaves, was relatively low and no microsites with high apoplastic pH were measured. Supplying chlorotic leaves with NO3 − may cause the apoplastic pH to increase at sites of high N demand. Since growth of chlorotic leaves is restricted, the need for NO3 − is also reduced and, in particular, in fully developed chlorotic leaves high apoplastic pH levels restricting Fe3+reduction may not necessarily prevail. Compared with the chlorotic intercostal area, the apoplast pH of the green leaf veins (Fig. 4) was particularly low (Table IV), presumably because of: (a) the influx of xylem liquid, which had a relatively low pH (Table I), and (b) the efficient pH regulation via H+-ATPase at the site of xylem vessels (Michelet and Boutry, 1995). NO3 −nutrition did not increase the xylem apoplastic pH compared with the NH4NO3 treatment, and light/dark changes had only a minor effect (Table II). Therefore, Fe3+ reduction in the area of green veins of interchlorotic leaves is presumably still optimal for continuous Fe supply of the neighboring cells adjacent to the xylem vessels, and this may be the reason that during leaf yellowing the tissue around the leaf xylem remains green (Fig. 4). Several studies have shown that upon NO3 − nutrition, leaf chlorosis will be induced (e.g. Aktas and Van Egmond, 1979; Mengel and Geurtzen, 1988). As the xylem liquid enters the intercostal area of young, still green leaves under alkaline nutritional conditions, high apoplastic pH levels prevailed at microsites (Figs. 2C and 3A) over the whole leaf blade, presumably related to the growing sites due to increased uptake of NO3 −via proton cotransport. When averaged across the leaf, these substantial apoplastic pH changes were limited to about 10% of the whole interveinal leaf apoplast in this study, and were shown to be a small overall apoplastic pH change (Table I). However, the high apoplastic pH at these interveinal microsites may depress Fe3+ reductase activity by about 50% (Table V). Such a restriction is not small, in particular because at these growing microsites, different reactions such as DNA synthesis (Reichard, 1993) and chlorophyll synthesis (Terry and Abadia, 1986) compete for Fe. We therefore suggest that the uptake of Fe2+ may be depressed at these interveinal microsites and may be sufficient to induce leaf yellowing and growth retardation under alkaline conditions (Kosegarten et al., 1998). From these argumentations it is clear that future research is needed to clarify the induction of leaf yellowing and to investigate apoplastic pH throughout the leaf chlorosis process. Interestingly, leaf yellowing of young green leaves is a slowly continuing process that starts at minute areas over the whole leaf surface and not simultaneously at all interveinal sites. This observation fits with the distribution of high apoplastic pH in the leaf at interveinal microsites (Figs. 2C and 3A) and with our hypothesis that at these sites of high apoplastic pH, with with Fe3+ reduction is inhibited, which may induce leaf yellowing. Since apoplastic pH is a dynamic rather than a static parameter (see Hoffmann and Kosegarten, 1995) future studies would be of particular interest to correlate leaf paling with apoplastic pH throughout the process of leaf chlorosis at the cellular level to understand the complex nature of leaf yellowing. Also, a Fe3+-sensitive fluorochrome that can be loaded into the leaf apoplast would realize measurements of Fe3+ reduction at interveinal microsites. In the present study, apoplastic pH during leaf yellowing and apoplastic pH compared with Fe3+ reduction in yellowing leaves were not examined. It is quite possible that, in contrast to young green leaves, in growing but yellowing leaves microsites with high apoplastic pH may be increased, because at lower photosynthetic rates plasmalemma H+-ATPase activity may be restricted. Due to low photosynthetic rates of chlorotic leaves (Kosegarten et al., 1998), the amount of reducing equivalents may also be restricted and therefore Fe3+ reduction as well. OUTLOOK Plants grown on calcareous soils suffer from a physiological Fe deficiency, and a substantial amount of Fe is presumably trapped in the apoplast of leaves and roots. The supply of Fe has to overcome two critical steps: (a) the high pH in the leaf apoplast, which hampers Fe3+-citrate reduction (Table V), and (b) the high pH in the root apoplast (Toulon et al., 1992; Kosegarten et al., 1999), which may hamper Fe3+-siderophore reduction. The pH dependence of Fe3+ reduction in the root apoplast remains to be proven. ACKNOWLEDGMENTS We thank Dr. F. Grolig (Botanik, Philipps Universität, Marburg, Germany) for critically reading the manuscript. LITERATURE CITED 1 Aktas M Van Egmond M Effect of nitrate nutrition on Fe utilization by an Fe-efficient and an Fe-inefficient soybean cultivar. 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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)

Fowler, Thomas J.; Bernhardt, Christine; Tierney, Mary L.

doi: 10.1104/pp.121.4.1081pmid: 10594096

Abstract We have characterized the molecular organization and expression of four proline-rich protein genes from Arabidopsis (AtPRPs). These genes predict two classes of cell wall proteins based on DNA sequence identity, repetitive motifs, and domain organization. AtPRP1 and AtPRP3encode proteins containing an N-terminal PRP-like domain followed by a C-terminal domain that is biased toward P, T, Y, and K.AtPRP2 and AtPRP4 represent a second, novel group of PRP genes that encode two-domain proteins containing a non-repetitive N-terminal domain followed by a PRP-like region rich in P, V, K, and C. Northern hybridization analysis indicated that AtPRP1 and AtPRP3 are exclusively expressed in roots, while transcripts encoding AtPRP2 and AtPRP4 were most abundant in aerial organs of the plant. Histochemical analyses of promoter/β-glucuronidase fusions localized AtPRP3 expression to regions of the root containing root hairs. AtPRP2 and AtPRP4expression was detected in expanding leaves, stems, flowers, and siliques. In addition, AtPRP4 expression was detected in stipules and during the early stages of lateral root formation. These studies support a model for involvement of PRPs in specifying cell-type-specific wall structures, and provide the basis for a genetic approach to dissect the function of PRPs during growth and development. Plant cell walls are dynamic and complex structures that contribute to functional differences between cell types during plant growth and development. Pro-rich proteins (PRPs) represent one of five families of structural cell wall proteins that have been identified in higher plants (for review, see Carpita and Gibeaut, 1993; Showalter, 1993; Cassab, 1998). PRPs were first identified as proteins that accumulate in the cell wall in response to physical damage (Chen and Varner, 1985; Tierney et al., 1988) and have subsequently been shown to be temporally regulated during plant development. PRP gene expression is associated with early stages of legume root nodule formation (Franssen et al., 1987; van de Wiel et al., 1990; Wilson et al., 1994), soybean seedling, leaf, stem, and seed coat development (Hong et al., 1989; Kleis-San Francisco and Tierney, 1990; Lindstrom and Vodkin, 1991; Ye et al., 1991), bean seedling growth (Sheng et al., 1991), and with early stages of tomato fruit development (Santino et al., 1997). The spatial pattern of PRP expression is also tightly regulated, as shown by in situ hybridization and reporter gene expression analysis (Wyatt et al., 1992; Suzuki et al., 1993). For example, the soybean SbPRP1 and SbPRP2 transcripts have been localized to sclereids, the inner integument of the seed coat and the epidermal, cortical, and endodermoidal cells of young seedlings. Protein localization studies suggest that PRPs may function both in determining cell-type-specific wall structure during plant development and by contributing to defense reactions against physical damage and pathogen infection. Immunohistochemical analyses using antibodies raised against SbPRP2 localized PRP accumulation in soybean to protoxylem cells within the root and xylem and phloem fibers within the stem, indicating that these proteins are critical for maintaining structural integrity of mature tissues (Ye et al., 1991). PRPs may play a similar role during seed development, since seed coat integrity appears to be altered in soybean lines that fail to accumulated these proteins within their cell walls (Nicholas et al., 1993). PRPs are rapidly insolubilized within the cell wall in response to physical damage, treatment with fungal elicitors, and pathogen infection (Kleis-San Francisco and Tierney, 1990; Bradley et al., 1992; Brisson et al., 1994), indicating an active role in plant defense reactions. While the mechanism for PRP insolubilization is not known, there is evidence that this process involves the formation of intermolecular isodityrosine or di-isodityrosine residues through an oxidative coupling reaction (Cooper and Varner, 1984; Fry, 1982; Bradley et al., 1992; Waffenschmidt et al., 1993; Brady et al., 1996). DNA sequence analysis of PRP genomic and cDNA clones indicates that these proteins can be placed into more than one class based on their primary structure. The first of these classes is characterized by PRP genes isolated from carrot and soybean, which encode tandem copies of the pentapeptide PPVX(K/T), where X is often Y, H, or E (Chen and Varner, 1985; Hong et al., 1987, 1990). SbPRP1 and SbPRP2, two members of this class, have been purified from soybean (Averyhart-Fullard et al., 1988; Kleis-San Francisco and Tierney, 1990; Lindstrom and Vodkin, 1991). Neither of these proteins appears to be highly glycosylated (Datta et al., 1989), and N-terminal sequence analysis has shown that the repetitive unit for both mature proteins is ProHypVal(Tyr/Glu)Lys. In contrast, a second group of PRP cDNAs predicts two-domain proteins containing a Pro-rich N-terminal domain and a C-terminal domain that lacks Pro-rich or repetitive sequences. This group of PRP genes includes PvPRP1 in bean (Sheng et al., 1991) andTPRP-F1 in tomato (Salts et al., 1991; Santino et al., 1997). We present the molecular organization and expression patterns of four PRP genes from Arabidopsis. These genes encode two unique classes of PRPs based on DNA sequence identity, repetitive motifs, and domain organization. Northern hybridization and promoter/reporter gene analysis indicate that each of these AtPRP genes has a unique temporal and spatial pattern of expression, suggesting potential functions for these proteins in determining specific extracellular matrix structures throughout plant development. MATERIALS AND METHODS Plant Material and Growth Conditions For RNA isolation, Arabidopsis ecotype Columbia plants were grown in Promix:vermiculite:perlite (3:1:1) at 19°C using an 8-h light/16-h dark photoperiod, followed by a 12-h light/12-h dark regime to induce flowering. Leaf, stem, and floral tissues were harvested, frozen in liquid nitrogen, and stored at −80°C. Root tissue used for RNA isolation was obtained from plants grown in liquid culture (1× Murashige and Skoog salts, 1× Gamborg's B5 vitamins, 1% [w/v] Suc, and 10 mm 2-(N-morpholino)-ethanesulfonic acid [MES], pH 6.0) for 10 d under continuous light, frozen in liquid nitrogen, and stored at −80°C. Tissue from transgenic Arabidopsis lines expressing AtPRP/β-glucuronidase (GUS) constructs was obtained for histochemical analysis by growing plants in Magenta boxes on either Murashige and Skoog medium with 1% (w/v) Suc (for vegetative tissues) or in Promix:vermiculite:perlite (3:1:1) (for reproductive tissues) under a light/dark regime as described above. Isolation of AtPRP Genomic and cDNA Clones AtPRP genomic clones were isolated from a genomic library (Landsberg) constructed in λ-fix (Voytas and Ausubel, 1988) using carrot (pDC16; Chen and Varner, 1985) and soybean (SbPRP1; Hong et al., 1987 and SbPRP2; Datta and Marcus, 1990) PRP genes as probes. Nitrocellulose membrane filter lifts of bacteriophage λ plaques (Sambrook et al., 1989) were hybridized with the heterologous probes at 55°C in 6× SSC, 5× Denhardt's, 0.5% (w/v) SDS, and 100 μg mL−1 denatured salmon-sperm DNA, washed at 55°C in 2× SSC, 0.5% (w/v) SDS, and exposed to XAR-5 x-ray film (Kodak, Rochester, NJ) with intensifying screens at −80°C. Hybridizing plaques were purified and the AtPRP sequences were subcloned into plasmid vectors for DNA sequencing. AtPRP subclones were digested with Exonuclease III using an Erase-a-Base kit (Promega, Madison, WI) and the series of nested deletions were sequenced using either Sequenase (United States Biochemical, Cleveland) orTaq DNA polymerase and a dsDNA cycle sequencing kit (BRL, Gaithersburg, MD). AtPRP cDNA clones were isolated from a λPRL2 library (Newman et al., 1994) screened with AtPRP probes pH/R2 (Fig. 4, bp 717–1,455) and pH/Sau3A (Fig. 2, bp 190–865). Genomic Southern Hybridization Total DNA was isolated from leaves of 6-week-old plants using a miniprep method (Junghans and Metzlaff, 1990) scaled to accommodate 0.5-g samples. The DNA was digested with EcoRI, size-fractionated by electrophoresis through 0.7% (w/v) agarose gels, and transferred to nitrocellulose membranes using 20× SSC (Sambrook et al., 1989). The membranes were incubated at 80°C under vacuum for 1.5 h and hybridized at 55°C with both an AtPRP1 (pH/Sau3A; bp 190–865 in Fig. 2) and an AtPRP2 (pH/R2; bp 717–1,455 in Fig. 4) coding-region probe in 5.5× SCP, 0.925% (w/v) sodiumN-lauroylsarcosine, 10% (w/v) dextran sulfate (Pharmacia, Uppsala), 1 mg mL−1 heparin, and 100 μg mL−1 sheared, denatured fish-sperm DNA. The filters were washed for 15 min at room temperature in 2× SSC, 0.5% (w/v) SDS, followed by a wash in 2× SSC, 0.2% (w/v) SDS at 42°C for 15 min, and exposed to x-ray film at −80°C with two intensifying screens. AtPRP RNA Analysis RNA was extracted from various tissues using a Tris-HCl/SDS/phenol extraction method as described previously (DeVries et al., 1988). Poly(A+) RNA was isolated from total RNA preparations using the PolyATtract kit (Promega), according to the manufacturer's protocol. Poly(A+) RNA (1.5 μg/lane) was size-fractionated by electrophoresis in 1.4% (w/v) agarose gels containing 1× 3-(N-morpholino)-propanesulfonic acid (MOPS) buffer (Sambrook et al., 1989) and 0.44 mformaldehyde. The RNAs were capillary blotted to nitrocellulose membranes with 20× SSC, and the membranes were incubated at 80°C under vacuum for 2 h. RNA blots were hybridized individually at 42°C overnight with 32P-labeled gene-specific probes in 5× SSC, 5× Denhardt's, 0.5% (w/v) SDS, and 100 mg mL−1 single-stranded DNA. The individual probes used corresponded to: AtPRP1, bp 1,475–1,665 (Fig. 2); AtPRP2, bp 176–371 (Fig. 4); AtPRP3, bp 1,265–1,455 (Fig. 3); AtPRP4, bp 1,609–1,867 (Fig. 5). The formamide concentrations were adjusted for each probe to ensure gene-specific conditions: AtPRP1 and AtPRP3, 40% (w/v) formamide; AtPRP2, 50% (w/v) formamide; AtPRP4, 43% (w/v) formamide. Each of the filters was washed at high stringency using the following conditions: AtPRP1 and AtPRP3, 30 min at 65°C in 2× SSC, 0.5% SDS (w/v) followed by 30 min at 65°C in 1× SSC, 0.25% (w/v) SDS; AtPRP2, 30 min at 65°C in 1× SSC, 0.25% SDS (w/v) followed by 30 min at 65°C in 0.3× SSC, 0.2% (w/v) SDS; AtPRP4, 30 min at 65°C in 2× SSC, 0.5% SDS (w/v) followed by 30 min at 65°C in 0.7× SSC, 0. 5% (w/v) SDS. Filters were then exposed to x-ray film at −80°C with two intensifying screens. Intron Mapping Intron positions within the AtPRP2 and AtPRP4 genomic clones were determined by reverse transcriptase (RT)-PCR (Kawasaki et al., 1988). First-strand cDNA was synthesized using AMV RT (Boehringer Mannheim, Basel) and 50 μg of total RNA isolated from flower tissue. For the RT-PCR reactions, a common first-strand oligonucleotide primer (5′ GATA(A/G)AAACACGATCTTGG 3′) was used with both AtPRP2 and AtPRP4 transcripts. This primer has a single degeneracy that allows it to prime both transcripts at a conserved site 3′ of the splice junctions (AtPRP2 [Fig. 4], bp 785–804; AtPRP4 [Fig. 5], bp 640–659). Second-strand DNA synthesis was performed using oligonucleotide primers that allowed specific amplification of either AtPRP2 or AtPRP4 sequences. Reaction conditions used for the RT-PCR were 10 mm Tris-HCl, pH 9.0; 2.5 mmMgCl2; 50 mm KCl; 200 μm each of dATP, dCTP, dGTP, and dTTP; 0.1% (v/v) Triton-X; 6.6% of the flower cDNA (2 μL of 30 μL); 0.5 μm gene-specific oligonucleotide primer (AtPRP2; Fig. 4, bp 188–207); (AtPRP4: Fig. 5, bp 179–198); 1.0 μmdegenerate primer (see above); and 2.5 units of Taq DNA polymerase. Reactions were heated to 95°C for 5 min, followed by 50 cycles of 94°C for 30 s, 45°C for 60 s, 72°C for 60 s in a thermal cycler (Perkin-Elmer, Foster City, CA). The PCR fragments generated in this manner were gel-purified and ligated into pT7Blue(R) (Novagen, Madison, WI) for sequencing. Intron positions within the AtPRP1 and AtPRP3 genomic clones were determined by comparison of the genomic sequences to the sequences of corresponding partial or full-length cDNAs isolated from the λPRL2 library. Predicted Signal Peptide Cleavage Sites Cleavage sites for the signal peptide were predicted using the matrix method as described by von Heijne (1986). Construction of AtPRP Promoter/GUS Lines 5′-Flanking sequences for AtPRP2 (2.5 kb), AtPRP3 (1.5 kb), and AtPRP4 (1.4 kb) were fused to the bacterial uid gene encoding GUS (vector pBI101; Jefferson et al., 1987) and transformed into Arabidopsis ecotype Columbia (AtPRP2 and AtPRP3) or Landsbergerecta (AtPRP4), respectively, using an in planta transformation method (Bechtold et al., 1993). Kanamycin-resistant lines were identified, allowed to set seed, and T2transgenic plants were grown and analyzed for GUS expression. Histochemical GUS Staining Histochemical staining of plant tissue for GUS activity was performed as described by Jefferson et al. (1987). Samples were immediately placed in substrate solution (50 mm sodium phosphate, pH 7.5, 15% [v/v] methanol, 2 mm5-bromo-4-chloro-3-indolyl-glucuronide, and 0.05% [v/v] Tween 20), vacuum infiltrated for 2 min at 85 kPa, and incubated at 37°C for 8 to 18 h. Removal of pigments was achieved by several washes in 50% to 70% (v/v) ethanol. Samples were analyzed under a stereomicroscope (model 2000, Zeiss, Jena, Germany) and pictures were taken on Kodak 25 film. RESULTS Screening of an Arabidopsis (Landsberg erecta) genomic library with carrot and soybean PRP probes (Chen and Varner, 1985;Suzuki et al., 1993) resulted in the identification of four distinct genomic clones encoding Pro-rich proteins (AtPRPs). cDNA clones corresponding to each of these genomic clones were isolated from a λ-PRL2 library (Newman et al., 1994) obtained through the Arabidopsis Biological Resource Center (Ohio State University, Columbus). DNA sequencing of genomic and representative cDNA clones corresponding to these isolates indicated that the PRP genes in Arabidopsis can be separated into two classes (AtPRP1 and AtPRP3versus AtPRP2 and AtPRP4) based on DNA sequence homology, domain structure, and predicted amino acid sequence. Southern hybridization showed that AtPRP gene sequences within each class hybridized well with each other and poorly with clones encoding other PRPs. For example, at high stringency AtPRP1hybridizes with AtPRP3 but not with AtPRP2,AtPRP4, or PRP gene sequences from other plant species (data not shown). Figure 1 illustrates the pattern of restriction fragments that are detected when coding region probes for AtPRP1 and AtPRP2 were used in Southern hybridizations with EcoRI-digested genomic DNA. These fragments were analyzed by DNA sequencing and were shown to correspond to AtPRP3 (7 kb), AtPRP1 (3.8 kb),AtPRP4 (3.3 kb), and AtPRP2 (1.5 kb), indicating that each of these genes is represented as single copy within the Arabidopsis genome. Fig. 1. Open in new tabDownload slide Genomic Southern analysis of AtPRP sequences. One microgram of Arabidopsis genomic DNA was digested withEcoRI and analyzed by Southern hybridization using coding region probes for AtPRP1 and AtPRP2. The relative positions of the molecular mass markers are indicated. From top to bottom, the four restriction fragments correspond to AtPRP3, AtPRP1, AtPRP4, and AtPRP2. Fig. 1. Open in new tabDownload slide Genomic Southern analysis of AtPRP sequences. One microgram of Arabidopsis genomic DNA was digested withEcoRI and analyzed by Southern hybridization using coding region probes for AtPRP1 and AtPRP2. The relative positions of the molecular mass markers are indicated. From top to bottom, the four restriction fragments correspond to AtPRP3, AtPRP1, AtPRP4, and AtPRP2. Structure of AtPRP1 and AtPRP3 The DNA sequence of the AtPRP1 and AtPRP3 genomic and cDNA clones (Figs. 2 and 3) predicts Pro-rich proteins containing a signal peptide followed by two domains and having molecular masses of 36.5 and 34.4 kD, respectively. The N-terminal domain of AtPRP1 consists of 13 imperfect copies of the amino acid repeat KPTLSPPVYT. This decapeptide motif, which contains the pentapeptide motif PPVX(K/T) that is characteristic of other PRPs, is found five times within the N-terminal domain of AtPRP3 as part of a longer repeat unit, KPTIPPPVYTPPVYKPTLSPPVYT. The C-terminal domain of both of these proteins, while rich in P, Y, and K, is unique in sequence. While the amino acid sequence of AtPRP1 and AtPRP3 is highly conserved (76% amino acid identity overall), the C-terminal domain of these proteins was found to exhibit the greatest sequence identity (Table I). Fig. 2. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP1. The ORF for AtPRP1 and the predicted amino acid sequence are presented in uppercase, while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Fig. 2. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP1. The ORF for AtPRP1 and the predicted amino acid sequence are presented in uppercase, while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Fig. 3. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP3. The ORF for AtPRP3 and the predicted amino acid sequence is presented in uppercase, while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Fig. 3. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP3. The ORF for AtPRP3 and the predicted amino acid sequence is presented in uppercase, while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Table I. Comparison of the AtPRP protein sequences AtPRP1 MAITRASFAICILLSLATIATADYYAPSSPPVYTSPVNKPTLPPPVYTPPVHKPTLPPPVYTPPVHKPTLSPPVYT AtPRP3 .....S.L...LI...V..T.....S.......K..EH.....S.......Y....S.....––––.......... AtPRP1 ––––––––––––––KPTLPPPAYTPPVYNKPTLPAPVYT–––––PPVY–––––KPTLSPPVYTKPTLLPPVFKPTL AtPRP3 PPVYKHTPSPPVYT.......V......K–...SP....KPTIP....TPPVY...PD–––.....I––––––––– AtPRP1 SPPVYTKPTLSPTVYKPTLSPPVNNKPSLSPPVYKPTLSPPVYTKPTLPPPVYKKSPIY–SPPPPfapkptytppt AtPRP3 P.....–––––.P.....P....–––––––––––––––––––––––––––––.....S.S.....yv.......t. AtPRP1 kpyvpeiikavggiilckngyetypiqgakakivcsergsyeksknevviysdptdfkgyfhvvlthiknlsncrv AtPRP3 ..........................l...iq....dpa..g..nt......n...s......s..s....ay... AtPRP1 klytspvetcknptnvnkgltgvpfsmy–––––sdknlklfnvgpfyftagskaapatpry AtPRP3 ...l....................lal.gyrfyp....e..s.....y.–.p.......k AtPRP2 MRILPKSGGGALCLLFVFALCSVAHSLSRDVKVVGDVEVIGYSEISKIKIPNAFSGLRVTIECKAADSKGHFVT AtPRP4 .....EPR.SVP...LLV–––..LL.ATLSLAR.––...V..A.–....T...........D..V–N–...... ATPRP2 RGSGEVEETGKFHLNIPHDIVGDDGTLKEACYAHLQSAFGNPCPAHDGLEASKIVFLSKSGANHVLGLKQSLKF ATPRP4 K...NIDDK...G........S.N.A...E...Q.H..A.T.........ST.........DK.I.....N... ATPRP2 SPEVCISKF WHMPK––––––––––fplppplnlppltfpkikkpcppiyippvvip––vpiykpp–––––––– ATPRP4 ...I.V...F.....LPPFKGFDHP.......e...f––––.......k.s...ev.pp..v.e..pkkeippp AtPRP2 –––––––––––––––vpiykpp––vv––––ipkkpcppkih––––––––––––––––––––––––––––––––– AtPRP4 vpvydpppkkevppp..v....pk.elppp.........ppkiehpppvpvykpppkiehpppvpvykpppkie AtPRP2 –––––––––––kkpcppkvahkpiyk––––––––––––––––––ppk––––––––––––––––––––––––––– AtPRP4 hpppvpvhkpp.......–––––––kvdpppvpvhkpptkkpcppkkvdpppvpvhkpppkivipppkiehppp AtPRP2 –piykppvpiykppvvipkktfpplhkpiykh–––––––––pvpiykpifkppvvvipkkpc–––––––––––– AtPRP4 v.v....––––––––––..––––iehp...ippivkkpcpp.......––––.–........pppvpvykppvv AtPRP2 –––––––––––––pplpkfphfppkyiphpkfgkwppfpshp AtPRP4 vipkkpcpplpql.......pl.....h.........l.p.. AtPRP1 MAITRASFAICILLSLATIATADYYAPSSPPVYTSPVNKPTLPPPVYTPPVHKPTLPPPVYTPPVHKPTLSPPVYT AtPRP3 .....S.L...LI...V..T.....S.......K..EH.....S.......Y....S.....––––.......... AtPRP1 ––––––––––––––KPTLPPPAYTPPVYNKPTLPAPVYT–––––PPVY–––––KPTLSPPVYTKPTLLPPVFKPTL AtPRP3 PPVYKHTPSPPVYT.......V......K–...SP....KPTIP....TPPVY...PD–––.....I––––––––– AtPRP1 SPPVYTKPTLSPTVYKPTLSPPVNNKPSLSPPVYKPTLSPPVYTKPTLPPPVYKKSPIY–SPPPPfapkptytppt AtPRP3 P.....–––––.P.....P....–––––––––––––––––––––––––––––.....S.S.....yv.......t. AtPRP1 kpyvpeiikavggiilckngyetypiqgakakivcsergsyeksknevviysdptdfkgyfhvvlthiknlsncrv AtPRP3 ..........................l...iq....dpa..g..nt......n...s......s..s....ay... AtPRP1 klytspvetcknptnvnkgltgvpfsmy–––––sdknlklfnvgpfyftagskaapatpry AtPRP3 ...l....................lal.gyrfyp....e..s.....y.–.p.......k AtPRP2 MRILPKSGGGALCLLFVFALCSVAHSLSRDVKVVGDVEVIGYSEISKIKIPNAFSGLRVTIECKAADSKGHFVT AtPRP4 .....EPR.SVP...LLV–––..LL.ATLSLAR.––...V..A.–....T...........D..V–N–...... ATPRP2 RGSGEVEETGKFHLNIPHDIVGDDGTLKEACYAHLQSAFGNPCPAHDGLEASKIVFLSKSGANHVLGLKQSLKF ATPRP4 K...NIDDK...G........S.N.A...E...Q.H..A.T.........ST.........DK.I.....N... ATPRP2 SPEVCISKF WHMPK––––––––––fplppplnlppltfpkikkpcppiyippvvip––vpiykpp–––––––– ATPRP4 ...I.V...F.....LPPFKGFDHP.......e...f––––.......k.s...ev.pp..v.e..pkkeippp AtPRP2 –––––––––––––––vpiykpp––vv––––ipkkpcppkih––––––––––––––––––––––––––––––––– AtPRP4 vpvydpppkkevppp..v....pk.elppp.........ppkiehpppvpvykpppkiehpppvpvykpppkie AtPRP2 –––––––––––kkpcppkvahkpiyk––––––––––––––––––ppk––––––––––––––––––––––––––– AtPRP4 hpppvpvhkpp.......–––––––kvdpppvpvhkpptkkpcppkkvdpppvpvhkpppkivipppkiehppp AtPRP2 –piykppvpiykppvvipkktfpplhkpiykh–––––––––pvpiykpifkppvvvipkkpc–––––––––––– AtPRP4 v.v....––––––––––..––––iehp...ippivkkpcpp.......––––.–........pppvpvykppvv AtPRP2 –––––––––––––pplpkfphfppkyiphpkfgkwppfpshp AtPRP4 vipkkpcpplpql.......pl.....h.........l.p.. For each of the AtPRP proteins, the predicted signal sequences are underlined, the N-terminal domain is represented by uppercase letters, and the C-terminal domain is represented by lowercase letters. Dashes represent introduced sequence gaps, and dots indicate the identity of the amino acid sequence between the predicted PRPs. Open in new tab Table I. Comparison of the AtPRP protein sequences AtPRP1 MAITRASFAICILLSLATIATADYYAPSSPPVYTSPVNKPTLPPPVYTPPVHKPTLPPPVYTPPVHKPTLSPPVYT AtPRP3 .....S.L...LI...V..T.....S.......K..EH.....S.......Y....S.....––––.......... AtPRP1 ––––––––––––––KPTLPPPAYTPPVYNKPTLPAPVYT–––––PPVY–––––KPTLSPPVYTKPTLLPPVFKPTL AtPRP3 PPVYKHTPSPPVYT.......V......K–...SP....KPTIP....TPPVY...PD–––.....I––––––––– AtPRP1 SPPVYTKPTLSPTVYKPTLSPPVNNKPSLSPPVYKPTLSPPVYTKPTLPPPVYKKSPIY–SPPPPfapkptytppt AtPRP3 P.....–––––.P.....P....–––––––––––––––––––––––––––––.....S.S.....yv.......t. AtPRP1 kpyvpeiikavggiilckngyetypiqgakakivcsergsyeksknevviysdptdfkgyfhvvlthiknlsncrv AtPRP3 ..........................l...iq....dpa..g..nt......n...s......s..s....ay... AtPRP1 klytspvetcknptnvnkgltgvpfsmy–––––sdknlklfnvgpfyftagskaapatpry AtPRP3 ...l....................lal.gyrfyp....e..s.....y.–.p.......k AtPRP2 MRILPKSGGGALCLLFVFALCSVAHSLSRDVKVVGDVEVIGYSEISKIKIPNAFSGLRVTIECKAADSKGHFVT AtPRP4 .....EPR.SVP...LLV–––..LL.ATLSLAR.––...V..A.–....T...........D..V–N–...... ATPRP2 RGSGEVEETGKFHLNIPHDIVGDDGTLKEACYAHLQSAFGNPCPAHDGLEASKIVFLSKSGANHVLGLKQSLKF ATPRP4 K...NIDDK...G........S.N.A...E...Q.H..A.T.........ST.........DK.I.....N... ATPRP2 SPEVCISKF WHMPK––––––––––fplppplnlppltfpkikkpcppiyippvvip––vpiykpp–––––––– ATPRP4 ...I.V...F.....LPPFKGFDHP.......e...f––––.......k.s...ev.pp..v.e..pkkeippp AtPRP2 –––––––––––––––vpiykpp––vv––––ipkkpcppkih––––––––––––––––––––––––––––––––– AtPRP4 vpvydpppkkevppp..v....pk.elppp.........ppkiehpppvpvykpppkiehpppvpvykpppkie AtPRP2 –––––––––––kkpcppkvahkpiyk––––––––––––––––––ppk––––––––––––––––––––––––––– AtPRP4 hpppvpvhkpp.......–––––––kvdpppvpvhkpptkkpcppkkvdpppvpvhkpppkivipppkiehppp AtPRP2 –piykppvpiykppvvipkktfpplhkpiykh–––––––––pvpiykpifkppvvvipkkpc–––––––––––– AtPRP4 v.v....––––––––––..––––iehp...ippivkkpcpp.......––––.–........pppvpvykppvv AtPRP2 –––––––––––––pplpkfphfppkyiphpkfgkwppfpshp AtPRP4 vipkkpcpplpql.......pl.....h.........l.p.. AtPRP1 MAITRASFAICILLSLATIATADYYAPSSPPVYTSPVNKPTLPPPVYTPPVHKPTLPPPVYTPPVHKPTLSPPVYT AtPRP3 .....S.L...LI...V..T.....S.......K..EH.....S.......Y....S.....––––.......... AtPRP1 ––––––––––––––KPTLPPPAYTPPVYNKPTLPAPVYT–––––PPVY–––––KPTLSPPVYTKPTLLPPVFKPTL AtPRP3 PPVYKHTPSPPVYT.......V......K–...SP....KPTIP....TPPVY...PD–––.....I––––––––– AtPRP1 SPPVYTKPTLSPTVYKPTLSPPVNNKPSLSPPVYKPTLSPPVYTKPTLPPPVYKKSPIY–SPPPPfapkptytppt AtPRP3 P.....–––––.P.....P....–––––––––––––––––––––––––––––.....S.S.....yv.......t. AtPRP1 kpyvpeiikavggiilckngyetypiqgakakivcsergsyeksknevviysdptdfkgyfhvvlthiknlsncrv AtPRP3 ..........................l...iq....dpa..g..nt......n...s......s..s....ay... AtPRP1 klytspvetcknptnvnkgltgvpfsmy–––––sdknlklfnvgpfyftagskaapatpry AtPRP3 ...l....................lal.gyrfyp....e..s.....y.–.p.......k AtPRP2 MRILPKSGGGALCLLFVFALCSVAHSLSRDVKVVGDVEVIGYSEISKIKIPNAFSGLRVTIECKAADSKGHFVT AtPRP4 .....EPR.SVP...LLV–––..LL.ATLSLAR.––...V..A.–....T...........D..V–N–...... ATPRP2 RGSGEVEETGKFHLNIPHDIVGDDGTLKEACYAHLQSAFGNPCPAHDGLEASKIVFLSKSGANHVLGLKQSLKF ATPRP4 K...NIDDK...G........S.N.A...E...Q.H..A.T.........ST.........DK.I.....N... ATPRP2 SPEVCISKF WHMPK––––––––––fplppplnlppltfpkikkpcppiyippvvip––vpiykpp–––––––– ATPRP4 ...I.V...F.....LPPFKGFDHP.......e...f––––.......k.s...ev.pp..v.e..pkkeippp AtPRP2 –––––––––––––––vpiykpp––vv––––ipkkpcppkih––––––––––––––––––––––––––––––––– AtPRP4 vpvydpppkkevppp..v....pk.elppp.........ppkiehpppvpvykpppkiehpppvpvykpppkie AtPRP2 –––––––––––kkpcppkvahkpiyk––––––––––––––––––ppk––––––––––––––––––––––––––– AtPRP4 hpppvpvhkpp.......–––––––kvdpppvpvhkpptkkpcppkkvdpppvpvhkpppkivipppkiehppp AtPRP2 –piykppvpiykppvvipkktfpplhkpiykh–––––––––pvpiykpifkppvvvipkkpc–––––––––––– AtPRP4 v.v....––––––––––..––––iehp...ippivkkpcpp.......––––.–........pppvpvykppvv AtPRP2 –––––––––––––pplpkfphfppkyiphpkfgkwppfpshp AtPRP4 vipkkpcpplpql.......pl.....h.........l.p.. For each of the AtPRP proteins, the predicted signal sequences are underlined, the N-terminal domain is represented by uppercase letters, and the C-terminal domain is represented by lowercase letters. Dashes represent introduced sequence gaps, and dots indicate the identity of the amino acid sequence between the predicted PRPs. Open in new tab Comparison of cDNA and genomic clones showed that AtPRP1 andAtPRP3 each contain an intron within their second domain. In each case, the consensus GT/AC intron splice donor and acceptor sites are present at the intron/exon border. The relatedness of these two genes is emphasized by the conserved position of the intron that interrupts a Gly codon within the second domain of the open reading frame (ORF) (Figs. 2 and 3). Structure of AtPRP2 and AtPRP4 AtPRP2 and AtPRP4 constitute a second, novel class of PRPs in Arabidopsis. The genomic clones encoding these PRPs predict proteins with molecular masses of 32.6 and 46 kD, respectively. Their primary structure consists of a signal peptide followed by a unique, non-repetitive domain and ending with a basic domain containing Pro-rich repeats (Figs. 4 and5). Like AtPRP1 and AtPRP3, the non-PRP-like domain of these proteins shares the highest degree of amino acid identity (Table I). Within the C-terminal domains of AtPRP2 and AtPRP4, the PRP consensus motif PPVX(K/T) is present only degenerately as PPV and P(V/I)YK. Instead, AtPRP2 contains nine copies of the amino acid motif PIYKPPV (Fig. 4), while AtPRP4 contains eight imperfect copies of the sequence PPPKIEHPPPVPVYK (Fig. 5). In addition, AtPRP2 and AtPRP4 contain four and six copies, respectively, of the Cys-containing motif KKPCPP (Figs.4 and 5). Fig. 4. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP2. The ORF for AtPRP2 and the predicted amino acid sequence are presented in uppercase, while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Fig. 4. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP2. The ORF for AtPRP2 and the predicted amino acid sequence are presented in uppercase, while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Fig. 5. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP4. The ORF for AtPRP4 and the predicted amino acid sequence is presented in uppercase while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. Fig. 5. Open in new tabDownload slide DNA and predicted amino acid sequence of AtPRP4. The ORF for AtPRP4 and the predicted amino acid sequence is presented in uppercase while upstream, downstream, and intron genomic sequences are presented in lowercase. The predicted cleavage site for the signal peptide is indicated with an arrowhead. A potential TATA box and polyadenylation signal are underlined. RT-PCR was used to identify the position of a single intron withinAtPRP2 and AtPRP4. The intron was present at a conserved position within the non-Pro-rich domain of these proteins and was flanked by consensus GT/AC intron donor and acceptor sites (Figs. 4and 5). AtPRP Gene Expression Is Developmentally Regulated Previous analysis of structural cell wall proteins has shown that individual gene family members often exhibit distinct patterns of expression during plant development. As an initial step in characterizing AtPRP expression during plant growth, poly(A+) mRNA isolated from expanding and mature rosette leaves, cauline leaves, inflorescence stalks, flowers, and root tissue of Arabidopsis was analyzed using northern hybridization (Fig.6A). AtPRP1 and AtPRP3 transcripts were detected exclusively in root mRNA preparations. In contrast, both AtPRP2 and AtPRP4 transcripts accumulated in rosette and cauline leaves, stems, and floral tissue. A low amount of AtPRP4 transcript could also be detected in root mRNA preparations. Fig. 6. Open in new tabDownload slide AtPRP expression analyzed by northern hybridization. A, Poly(A+) RNA (1.5 μg) isolated from the following organs of soil-grown plants was loaded onto each lane: 1, expanding rosette leaves; 2, mature rosette leaves; 3, roots (obtained from plants grown in liquid culture); 4, stems; 5, cauline leaves; and 6, flowers. Equal loading was confirmed by ethidium bromide staining. B, The specificity of the probes was analyzed using northern hybridization. The left two panels illustrate the difference in transcript size that was observed using the AtPRP2 and AtPRP4 probes to analyze northern blots of mRNA from expanding rosette leaves. In the right two panels, parallel northern blots containing in vitro-synthesized sense AtPRP1 and AtPRP3 transcripts (which are of the same predicted size) were hybridized with gene-specific probes for either AtPRP1 or AtPRP3. Fig. 6. Open in new tabDownload slide AtPRP expression analyzed by northern hybridization. A, Poly(A+) RNA (1.5 μg) isolated from the following organs of soil-grown plants was loaded onto each lane: 1, expanding rosette leaves; 2, mature rosette leaves; 3, roots (obtained from plants grown in liquid culture); 4, stems; 5, cauline leaves; and 6, flowers. Equal loading was confirmed by ethidium bromide staining. B, The specificity of the probes was analyzed using northern hybridization. The left two panels illustrate the difference in transcript size that was observed using the AtPRP2 and AtPRP4 probes to analyze northern blots of mRNA from expanding rosette leaves. In the right two panels, parallel northern blots containing in vitro-synthesized sense AtPRP1 and AtPRP3 transcripts (which are of the same predicted size) were hybridized with gene-specific probes for either AtPRP1 or AtPRP3. Control hybridizations were used to demonstrate the gene-specific nature of the probes under these hybridization conditions (Fig. 6B). Since AtPRP2 and AtPRP4 encode transcripts of different sizes, we analyzed the same mRNA preparation used in Figure6A for cross-reactivity. In contrast, AtPRP1 and AtPRP3 encode transcripts that cannot be distinguished by size. Therefore, we compared the cross-reactivity of each probe using in vitro-transcribed sense RNAs for each gene. In all cases, the probes were found to hybridize specifically to a single transcript. Since developmentally regulated changes in cell wall structure may be critical for normal growth and differentiation processes, we characterized the temporal and spatial expression patterns of the AtPRP genes using promoter/reporter gene constructs in transgenic plants. 5′-Flanking sequences for AtPRP2, AtPRP3, and AtPRP4 were fused to the bacterial uid gene encoding GUS and these constructs were transformed into Arabidopsis using vacuum infiltration (Bechtold et al., 1993). A minimum of four independent T2 transgenic lines for each of the AtPRP promoter/GUS constructs were analyzed for their patterns of GUS expression. AtPRP3/GUS expression was exclusively detected in roots during plant development, which is consistent with data obtained using northern hybridization. Shortly after germination, expression was found in root epidermis and root hairs localized around the transition zone marking the root/shoot junction. With further growth of the root, GUS expression could be detected in root epidermis and root hairs along the length of the root and was the most intense in the root zone forming new hairs (Fig. 7a). No AtPRP3/GUS expression was observed in the root tip. In older seedlings, AtPRP3 expression continued to be restricted to the regions of the main root active in root hair development, and this pattern of expression was reiterated in lateral roots (Fig. 7b). Fig. 7. Open in new tabDownload slide Histochemical localization of AtPRP expression using AtPRP promoter/GUS analysis. a and b, AtPRP3/GUS. a, 2-d-old seedling; b, 8-d-old seedling. c to l, AtPRP4/GUS. c, 1-d-old seedling; d, 2-d-old seedling; e, 23-d-old seedling; f, detail stipules; g, detail roots; h, immature inflorescence; i, flower cluster; j, young silique; k, detail nectaries; l, maturing silique. m to o, AtPRP2/GUS. m, 23-d-old seedling; n, immature inflorescence; o, flower cluster. Fig. 7. Open in new tabDownload slide Histochemical localization of AtPRP expression using AtPRP promoter/GUS analysis. a and b, AtPRP3/GUS. a, 2-d-old seedling; b, 8-d-old seedling. c to l, AtPRP4/GUS. c, 1-d-old seedling; d, 2-d-old seedling; e, 23-d-old seedling; f, detail stipules; g, detail roots; h, immature inflorescence; i, flower cluster; j, young silique; k, detail nectaries; l, maturing silique. m to o, AtPRP2/GUS. m, 23-d-old seedling; n, immature inflorescence; o, flower cluster. Several aspects of AtPRP2 and AtPRP4 gene expression were found to be similar during plant development. In young seedlings, AtPRP4/GUS expression was detected in the hypocotyl, cotyledons (Fig. 7, c and d), and rosette leaves. Staining was most intense in expanding leaves and gradually disappeared with age (Fig. 7, e and m). After transition to the reproductive phase of growth, AtPRP4 was found to be expressed in stems, cauline leaves, and sepals (Fig. 7, h and n). Similar patterns of expression were observed for AtPRP2/GUS in these tissues (data not shown). The timing of AtPRP2 and AtPRP4 expression during anther development was temporally controlled, with AtPRP2/GUS transcription associated with anthers of closed flowers (Fig. 7o), while AtPRP4 expression was only detected in anthers of open flowers (Fig. 7i). Later in development, both genes were found to be expressed in pedicels of developing siliques, nectaries, and along the length of maturing siliques (Fig. 7, j–l). AtPRP4/GUS was found to be uniquely expressed in stipules of both rosette and cauline leaves (Fig. 7, e and h), the stigma surface of opening flowers (Fig. 7i), emerging lateral roots, and in spaced intervals along the root that may represent initials for lateral root development (Fig. 7g). DISCUSSION We have isolated and characterized genomic and cDNA clones encoding four Pro-rich cell wall proteins from Arabidopsis. The expression of each of these genes is temporally and spatially regulated during plant development and targets cell types and organs where they may function to determine cell wall structure. In addition, AtPRP2 and AtPRP4 represent novel members of this gene family of extracellular matrix proteins. Structure of the AtPRPs The overall structure of AtPRP1 and AtPRP3 consists of a signal sequence, an N-terminal PRP-like domain, and a highly charged, non-repetitive C terminus. This structural organization is similar to that predicted for a number of other cell wall proteins, including an extensin-like protein (ISG) from Volvox (Ertl et al., 1992), several AGP-like proteins (TTS) from tobacco (Cheung et al., 1995), and three PRP-like proteins from bean, tomato, and tobacco (Salts et al., 1991; Sheng et al., 1991; Chen et al., 1993; Santino et al., 1997). In both the Volvox and tobacco systems, the interaction of these proteins with other components within the extracellular matrix was found to be critical for proper development. Disruption of the interaction between ISG and other matrix components resulted in the inability of cells to complete gamete formation, while inhibition of TTS expression using antisense or sense co-suppression transgenic lines resulted in a reduced rate of pollen tube growth (Cheung et al., 1995). These studies support the potential importance of matrix interactions between AtPRP1/ AtPRP3 and other components within the cell wall, and indicate that such interactions may be critical for root or root hair development in Arabidopsis. AtPRP2 and AtPRP4 represent a second, newly described subset of PRPs in higher plants. These genes encode proteins containing a signal sequence, an N-terminal domain that is non-repetitive, and a PRP-like C-terminal region. The predicted amino acid sequence of these two proteins indicates that they are highly charged polypeptides. The PRP-like, repetitive motifs present within the C-terminal domain are more degenerate than those observed for AtPRP1 and AtPRP3 and are found to border a Cys-rich motif (KKPCPP). While Cys-rich motifs have been observed in other two-domain PRPs, they have previously been found within the non-repetitive domain of these proteins (Sheng et al., 1991;Chen et al., 1993; Wu et al., 1993). Comparison of the nucleotide sequence of the four AtPRP genes presented here indicates that these genes are likely to have evolved from two gene duplication events. This is supported by the conserved position of a single intron within the unique domain of each of the AtPRP genes and the high degree of amino acid and nucleotide identity observed in the non-Pro-rich domains. Sequence gaps between either AtPRP1 and AtPRP3 or AtPRP2 and AtPRP4 are flanked by the repetitive motifs PPVX(K/T) or PTL(P/S), suggesting a possible function for these sequences in recombination (Table I). In soybean, SbPRP1 and SbPRP2 variants differing in molecular mass and containing multiple deletions or additions of the pentapeptide PPVXK have been identified (Schmidt et al., 1994). This type of variation suggests that recombination within sequences encoding the repetitive, Pro-rich motifs characteristic of PRPs may provide a mechanism for generating new structural cell wall proteins. Possible Functions for AtPRPs in Determining Cell Wall Structure PRPs are thought to contribute to the cell wall structure of specific cell types based both on their patterns of gene expression during plant development and their ability to associate with and become cross-linked to components within the cell wall (for review, seeShowalter, 1993). The predicted pIs of the AtPRPs range between 9.6 and 10, suggesting that they may interact with the acidic pectin network within the cell wall. In addition, the localization of Cys-rich elements with the Pro-rich domain of AtPRP2 and AtPRP4 may facilitate disulfide bond formation between these PRPs themselves and/or other proteins within the plant extracellular matrix. Further analysis of these novel PRPs may provide clues about the relationship between structural matrix protein function and cellular aspects of growth and development. Tyr and Lys are an abundant amino acids in both PRPs and extensins (a second family of Hyp-rich structural cell wall proteins) and have been implicated as the substrate for the peroxidase-mediated insolubilization of PRPs in soybean (Kleis-San Francisco and Tierney, 1990; Bradley et al., 1992; Brisson et al., 1994) and in the cross-linking of extensins within the cell wall of suspension-cultured cells (Brady et al., 1996; Schnabelrauch et al., 1996). An extensin-specific peroxidase has been identified in tomato cell suspension cultures, and the substrate for this enzyme has tentatively been identified as Val-Tyr-Lys. Interestingly, two soybean PRPs containing this motif were not substrates for this enzyme in vitro (Schnabelrauch et al., 1996). Pectin/extensin cross-links have been identified in cotton cell walls (Qi et al., 1995) and are thought to occur through either a 3,6-linked galactan or a ferulated sugar/amino acid cross-link (Keegstra et al., 1973; Brownleader and Dey, 1993). Thus, the insolubilization of the AtPRPs may involve either protein/protein or protein/carbohydrate linkages within the cell wall, and further investigation will be needed to determine if and how the cross-linking of these proteins within the wall contributes to the structure of the extracellular matrix. As more structural cell wall proteins are characterized, it appears that extensins and PRPs may be considered members of a superfamily of Pro/Hyp-rich cell wall proteins, as has been suggested previously (Kieliszewski and Lamport, 1996). Several structural features of the AtPRP gene family support this suggestion. Database analysis indicated that AtPRP1 and AtPRP3 share 42% identity with a predicted extensin-like protein in Nicotiana alata. In addition, AtPRP1 and AtPRP3 contain multiple Ser-Pro-Pro repeats throughout their N-terminal domain and a single Ser-Pro4 sequence, both of which are reminiscent of the Ser-Hyp4repetitive motif characteristic of many extensin proteins. The potential relationship between PRP and extensin protein sequences is also apparent when a repetitive unit within the AtPRP4 gene product (PPPKIEHPPPVPVYK) is compared with a known peptide sequence found within a sugar beet extensin (SOOVHEYPOOTOVYK), where O represents Hyp. However, it will be necessary to gain a better understanding of the sequences critical for extensin and PRP function within the cell wall before we can interpret whether this level of sequence conservation represents the remnants of a common evolutionary history or simply reflects conserved functional motifs required for the interaction of HRGPs and PRPs with other extracellular matrix components. Developmental Regulation of AtPRP Gene Expression Each of the AtPRP genes was differentially expressed. AtPRP1 and AtPRP3 transcripts were only detected in root tissue. This was supported by histochemical promoter/GUS analysis, which localized expression of AtPRP3 to the regions of the root producing root hairs. GUS expression was not observed in older parts of the root or in the root tip, indicating that AtPRP3 may play an important role during root hair formation. Two extensin genes with root-hair-specific expression patterns have recently been identified in tomato and bean (Arsenijevic-Malisimovic et al., 1997; Bucher et al., 1997), suggesting that at least two families of structural proteins may dictate aspects of cell wall architecture necessary for the initiation and growth of root hairs in different plant species. Northern hybridization analysis of AtPRP2 and AtPRP4 gene expression indicated that transcripts for these cell wall protein genes are most abundant in leaf, stem, and reproductive tissue. Analysis of AtPRP2/GUS and AtPRP4/GUS expression patterns supported these observations and showed that both of these genes are highly expressed in the hypcotyl and cotyledons of young seedlings, immature rosette and cauline leaves, stems, sepals, anthers, siliques, and in nectaries at the silique-pedicel junction. AtPRP4 was also found to be expressed uniquely in the stipules and stigma of opening flowers. Furthermore, AtPRP4 may play an important role in establishing a cell wall matrix necessary for the initiation and early stages of lateral root development, as its expression was observed in spaced intervals along the root and at junctions between laterals and the main root. A similar pattern of expression has been observed for a tobacco extensin gene (Keller and Lamb, 1989). However, these two genes differ in their expression pattern, as AtPRP4 is strictly associated with the early steps of lateral root initiation, while the tobacco extensin gene is also associated with lateral tip growth. Analysis of the regulation of AtPRP4 expression in association with the hormonal regulation of lateral root development will provide additional insight into the possible relationship between AtPRP4 function and lateral root growth. In summary, we have characterized the structure and expression of four members of the PRP gene family in Arabidopsis. These genes predict cell wall proteins that fall into two classes based on domain structure, sequence identity, intron location, and patterns of gene expression during plant development. In addition, two of these proteins (AtPRP2 and AtPRP4) represent a newly described class of structural cell wall proteins whose function may involve novel interactions within the extracellular matrix and possibly with proteins within the cell membrane. Analysis of the protein products of these genes using genetic and biochemical approaches readily available in Arabidopsis will provide an opportunity to dissect the mechanism(s) by which PRPs interact with other cell wall polymers in distinct cell types during plant development and in response to environmental stimuli. ACKNOWLEDGMENTS We thank Keith Davis for providing the Arabidopsis genomic library, Doreen Ware for assistance with DNA sequence analysis, and Gary Ward and Eunice Froeliger for helpful discussions. 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IBN–9219712) and by the U.S. Department of Agriculture (grant no. NRICGP–95–02982). C.B. was supported by experiment station grant no. 0171655. 2 Present address: Microbiology and Molecular Genetics Department, University of Vermont, Burlington, VT 05405. * Corresponding author; e-mail [email protected]; fax 802–656–0440. 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)

Thomma, Bart P.H.J.; Eggermont, Kristel; Tierens, Koenraad F.M.-J.; Broekaert, Willem F.

doi: 10.1104/pp.121.4.1093pmid: 10594097

Abstract Inoculation of wild-type Arabidopsis plants with the fungus Alternaria brassicicola results in systemic induction of genes encoding a plant defensin (PDF1.2), a basic chitinase (PR-3), and an acidic hevein-like protein (PR-4). Pathogen-induced induction of these three genes is almost completely abolished in the ethylene-insensitive Arabidopsis mutant ein2-1. This indicates that a functional ethylene signal transduction component (EIN2) is required in this response. The ein2-1 mutants were found to be markedly more susceptible than wild-type plants to infection by two different strains of the gray mold fungusBotrytis cinerea. In contrast, no increased fungal colonization of ein2-1 mutants was observed after challenge with avirulent strains of either Peronospora parasitica or A. brassicicola. Our data support the conclusion that ethylene-controlled responses play a role in resistance of Arabidopsis to some but not all types of pathogens. Ethylene is a gaseous plant hormone that has been implicated in a range of physiological processes including seed germination, organ senescence, organ abscission, fruit ripening, and morphological responses of organs (Abeles et al., 1992). It has been proposed that ethylene also plays an important role in controlling defense responses of plants to microbial pathogens. Pathogen challenge often causes an increase in ethylene production (Ross and Williamson, 1951; Van Loon, 1977; Mauch et al., 1984; Boller, 1991; Penninckx et al., 1998). Moreover, exogenous application of ethylene to plants can result in the activation of genes encoding antimicrobial pathogenesis-related (PR) proteins (Boller et al., 1983; Mauch and Staehelin, 1989; Memelink et al., 1990; Eyal et al., 1992; Beffa et al., 1995; Penninckx et al., 1996; Knoester et al., 1998), cell wall-strengthening Hyp-rich glycoproteins (Esquerré-Tugayé et al., 1979; Ecker and Davis, 1987; Tagu et al., 1992), or enzymes involved in the synthesis of phenylpropanoids (Ecker and Davis, 1987). If ethylene plays a crucial role in plant defense mechanisms, one would predict that treatment of plants with exogenous ethylene would enhance resistance to subsequent challenge with microorganisms or, conversely, that treatment with ethylene inhibitors would adversely affect their resistance level. This has been demonstrated for a number of plant-pathogen interactions (Esquerré-Tugayé et al., 1979;El-Kazzaz et al., 1983a; Marte et al., 1993). However, for other plant-pathogen combinations, pretreatment with ethylene either had no effect on resistance or actually diminished the resistance level (El-Kazzaz et al., 1983b; Brown and Lee, 1993; Van Loon and Pennings, 1993). These contradictory results have made the role of ethylene in host defense a frequently debated matter of controversy. Recently, however, conclusive evidence has been presented that ethylene is indeed involved in host resistance, albeit only to particular classes of pathogens and not to others, thus reconciling previous conflicting data (Knoester et al., 1998; Hoffman et al., 1999). In their experiments, Knoester et al. (1998) made use of transgenic tobacco plants transformed with a dominant-negative mutant allele of the Arabidopsis ethylene receptor gene ETR1. The transgenic plants with a disrupted ethylene response were more susceptible than wild-type plants to normally nonpathogenic soil-bornePythium spp., whereas their level of resistance to tobacco mosaic virus was unaffected. Hoffman et al. (1999) found that some soybean mutants with reduced ethylene sensitivity had a tendency toward more severe symptoms compared with wild-type plants when challenged with virulent strains of the fungi Septoria glycines andRhizoctonia solani and some but not all avirulent strains ofPhytophthora sojae. On the other hand, some of the ethylene-insensitive soybean mutants showed less-severe chlorotic symptoms relative to their wild-type parents upon inoculation with virulent strains ofPseudomonas syringae pv glycinea. Less-severe chlorosis was also observed in the ethylene-insensitive Never ripe tomato strain compared with wild-type plants when inoculated with either Xanthomonas campestris pv vesicatoriaor Pseudomonas syringae pv tomato. In addition, the Never ripe tomato mutants also showed less-severe wilting symptoms upon challenge with the fungal vascular pathogenFusarium oxysporum f. sp. lycopersici (Lund et al., 1998). Ethylene is known to promote events such as chlorophyll degradation (Stall and Hall, 1984) and xylem occlusion (VanderMolen et al., 1983), which are positively correlated with severity of disease symptoms such as chlorosis and wilting, respectively. In conclusion, it appears that ethylene controls both disease resistance responses and symptom expression. Therefore, this hormone can influence particular plant-pathogen interactions in different ways, depending on the offensive strategies of the pathogen, the efficacy of the defense genes it controls, and the nature of the physiological reactions that are triggered by the pathogen. Although most of our current highly detailed knowledge on the process of ethylene perception and signal transduction comes from the study of Arabidopsis mutants (Kieber, 1997; McGrath and Ecker, 1998), the role of ethylene in the resistance of this plant to microbial pathogens has so far only been examined in a handful of cases. Bent et al. (1992)studied the interaction between Arabidopsis and the phytopathogenic bacteria Xanthomonas campestris pv campestris andPs. syringae pv tomato. They observed that mutant ein2-1, a mutant affected in a membrane-associated signal transduction component of the ethylene response (McGrath and Ecker, 1998), showed less macroscopically visible chlorosis and less chlorophyll degradation compared with wild-type plants. However, when the bacteria multiplying in ein2-1 and wild-type plants were counted, no significant difference was found. It therefore appears that ethylene does not play a role in actual resistance to these bacteria but, rather, in the development of pathogen-induced chlorosis symptoms. Suppression of chlorotic disease symptoms after challenge with these bacteria was not observed for the ethylene-insensitive mutant etr1-3 (Bent et al., 1992), which is affected in the ETR1 gene encoding an ethylene receptor (Chang et al., 1993). This result is apparently difficult to reconcile with the supposed role of ethylene in chlorotic symptom development. However, when testing alongside the allelic mutantsetr1-1 and etr1-3 for their ability to inducePDF1.2 in response to challenge with Alternaria brassicicola, Penninckx et al. (1998) observed thatetr1-3 is a very leaky allele in contrast toetr1-1, at least with respect to its impact on this pathogen-induced response in adult plants. Therefore, the observation that the etr1-3 mutation does not affect bacterially induced symptom development may well be due to leakiness of this allele. When the etr1-1 and the ein2-1 mutants were tested for susceptibility to the Oomycete Peronospora parasitica strain Noco, a strain that is virulent on the wild-type parental line Columbia (Col-0), no differences in susceptibility relative to wild-type plants were observed (Lawton et al., 1994). Inoculation of leaves of wild-type plants with an avirulent Ps. syringae pv tomato strain was found to trigger a systemic defense response that protected the leaves against subsequent inoculation with either virulent strains of P. parasitica orPs. syringae pv tomato (Lawton et al., 1995;Pieterse et al., 1998). This systemic response was equally effective in the ethylene-insensitive etr1-1 mutant (Lawton et al., 1995;Pieterse et al., 1998). On the other hand, Pieterse et al. (1998)observed that inoculating Arabidopsis roots with a nonpathogenic root-colonizing strain of Pseudomonas fluorescence conferred systemic resistance in wild-type plants but not etr1-1mutants to subsequent inoculation of the leaves with a virulentPs. syringae pv tomato strain. Therefore, a systemic resistance response triggered by leaf inoculation with an avirulent bacterium appears to be ethylene independent, while that induced by inoculating roots with a nonpathogenic bacterium is ethylene dependent. So far, however, no pathogens of Arabidopsis have been described for which ethylene plays a role in local resistance responses. One complication in the study of the role of ethylene in disease resistance is that there appears to be an interrelationship with another stress hormone, jasmonate. Our previous studies on the expression of Arabidopsis gene PDF1.2, encoding an antifungal plant defensin peptide, have shown that this gene can be activated systemically upon pathogen challenge and that this activation requires both functional components of the ethylene response pathway, including ETR1 and EIN2, and the jasmonate response pathway, including COI1 (Penninckx et al., 1996). Both hormone response pathways need to be triggered concomitantly in order for pathogen-induced activation ofPDF1.2 to occur (Penninckx et al., 1998). On the other hand, activation of PDF1.2 is independent of the salicylate response pathway (Penninckx et al., 1996), which controls pathogen-induced expression of other antimicrobial proteins such as PR-1, PR-2, and PR-5 (Uknes et al., 1992). When assessing the role of jasmonate in disease resistance, we observed that a jasmonate-insensitive mutant, coi1-1, showed enhanced disease susceptibility to the fungal pathogens A. brassicicola and Botrytis cinerea, but not to P. parasitica, whereas the opposite resistance responses were observed for the salicylate response mutant npr1-1 and the salicylate degrading transgenic line NahG (Thomma et al., 1998). The main objectives of the current study were to assess the effect of a mutation in the ethylene transduction gene EIN2on the resistance response to the above-mentioned pathogens and the induction of some PR genes. MATERIALS AND METHODS Biological Material and Plant Inoculations The mutant ein2-1 (Guzmán and Ecker, 1990) was obtained from the Arabidopsis Biological Resource Center (Columbus, OH). The Arabidopsis mutants coi1-1 (Feys et al., 1994),npr1-1 (Cao et al., 1994), and pad3-1 (Glazebrook and Ausubel, 1994) were obtained from Drs. J. Turner (University of East Anglia, Norwich, UK), X. Dong (Duke University, Durham, NC), and J. Glazebrook (University of Maryland, College Park), respectively. All of these mutants are derived from the Col-0 ecotype. Arabidopsis plants were essentially grown as described previously (Penninckx et al., 1996). Growth and spore harvesting of the fungi Alternaria brassicicola (strain MUCL20297; Mycothèque UniversitéCatholique de Louvain, Louvain-la-Neuve, Belgium), Botrytis cinerea (strains IMI169558, International Mycology Institute, Kew, UK; and MUCL30158, Mycothèque Université Catholique de Louvain) were done as described previously (Broekaert et al., 1990). The transgenic A. brassicicola strain (MUCL20297) containing a chimeric GUS-expressing transgene is described in Thomma et al. (1998). Peronospora parasitica strain Wela (Delaney et al., 1994) was maintained on living Arabidopsis plants of the Weiningen ecotype, and was kindly provided by Drs. R. Vogelsang and A. Slusarenko (Rheinisch-Westfälische Technischetlochschule Aachen, Germany). Inoculation of 4-week-old soil-grown Arabidopsis plants with A. brassicicola, B. cinerea, and P. parasiticawas performed as described previously (Thomma et al., 1998). For inoculation with A. brassicicola and B. cinerea, care was taken to place drops with inoculum on fixed positions left and right from the midvein. Detection of Fungi in Inoculated Plants A transgenic A. brassicicola strain containing a chimeric UidA (GUS) expressing transgene driven by a constitutive glyceraldehyde-3-P dehydrogenase promoter was used for quantifying fungal biomass in inoculated plants. Plants were inoculated with three 5-μL drops per leaf of a suspension in water of 5 × 105 conidial spores of this strain per milliliter. Inoculated plants were incubated at 100% RH. Quantification of fungal biomass was performed as described previously (Thomma et al., 1998), using a quantitative RNA dot-blot assay withUidA as a probe. The presence of P. parasitica in inoculated plants was detected by microscopic observation of leaves stained with lactophenol trypan blue as described by Mauch-Mani and Slusarenko (1996). RNA Gel-Blot Analysis RNA was extracted from tissues of Arabidopsis by the phenol-LiCl method according to the method of Eggermont et al. (1996). RNA gel-blot analysis was performed as described previously (Penninckx et al., 1996). Riboprobes for PDF1.2, PR-3,PR-4, and β-Tubulin 1 were synthesized as described previously (Penninckx et al., 1996; Thomma et al., 1998). Ethylene and Methyl Jasmonate Treatments For testing the protective effect on Arabidopsis plants of ethylene against A. brassicicola, 4-week-old soil-grownpad3-1 plants were placed in a gastight translucent chamber. Ethylene was applied by injecting the appropriate amount of ethylene gas with a syringe through a rubber septum in the chamber. Methyl jasmonate was applied by pipeting an appropriate amount of 1% (v/v) liquid methyl jasmonate in ethanol on a cotton plug inside the chamber. After 48 h of treatment, the chambers were opened and the plants were inoculated with either A. brassicicola orB. cinerea as described above, except that for B. cinerea inoculation only one inoculation spot per leaf was applied. Six days after inoculation, infections were analyzed macroscopically by measuring lesion diameters (for A. brassicicola-inoculated plants) or by counting the ratio of inoculated leaves showing spreading necrosis versus total amount of inoculated leaves (for B. cinerea-inoculated plants). RESULTS Requirement of EIN2 for Pathogen-Induced Expression ofPR-3 and PR-4 The Arabidopsis genes encoding the plant defensin PDF1.2, basic PR-3-type chitinase (also called ChitB), and the basic PR-4 protein (also called hevein-like protein or Hel) have all been shown previously to be inducible by exogenous application of ethylene (Samac et al., 1990; Potter et al., 1993; Chen and Bleecker, 1995; Penninckx et al., 1996), as well as by methyl jasmonate (Thomma et al., 1998). Pathogen-induced expression of all of these genes is known to require a functional jasmonate response pathway, as expression of these genes is abolished in the coi1-1 mutant (Thomma et al., 1998), whereas requirement of a functional ethylene response pathway for pathogen-induced expression has so far only been demonstrated forPDF1.2 (Penninckx et al., 1996, 1998). We now show that the expression of both PR-3 and PR-4 is, like that ofPDF1.2, severely reduced in A. brassicicola-inoculated leaves of the ethylene-insensitive mutantein2-1 compared with similarly treated leaves of wild-type (Col-0) plants (Fig. 1). In noninoculated leaves of A. brassicicola-inoculated wild-type plants, systemic induction was clearly observed for PDF1.2,PR-3, and PR-4 genes, but this response was completely abolished in the ein2-1 mutants (Fig. 1). These results indicate that functional EIN2 and COI1 (Thomma et al., 1998) are required for pathogen-induced expression of PDF1.2,PR-3, and PR-4, suggesting that these genes are controlled by a similar jasmonate/ethylene-dependent signal transduction pathway. Fig. 1. Open in new tabDownload slide Induction of the PR genes in Arabidopsis in response to infection with A. brassicicola. Four-week-old soil-grown wild-type (Col-0) and ein2-1plants were infected with A. brassicicola and harvested 48 h following treatment. RNA blots were hybridized with the various probes indicated on the left. Symbols on top of the lanes are as follows: −, Mock-inoculated with water; +, inoculated withA. brassicicola spore suspension; 1°, treated lower rosette leaves; 2°, untreated upper rosette leaves. Fig. 1. Open in new tabDownload slide Induction of the PR genes in Arabidopsis in response to infection with A. brassicicola. Four-week-old soil-grown wild-type (Col-0) and ein2-1plants were infected with A. brassicicola and harvested 48 h following treatment. RNA blots were hybridized with the various probes indicated on the left. Symbols on top of the lanes are as follows: −, Mock-inoculated with water; +, inoculated withA. brassicicola spore suspension; 1°, treated lower rosette leaves; 2°, untreated upper rosette leaves. Requirement of EIN2 for Resistance to Particular Fungi Thomma et al. (1998) have previously shown that the jasmonate-insensitive Arabidopsis mutant coi1-1 is more susceptible than wild-type plants to infection by the fungi B. cinerea strain IMI169558 and A. brassicicola strain MUCL20297, but not by the Oomycete P. parasitica strain Wela. To investigate whether the ethylene-insensitive ein2-1mutant shares the same defects in disease resistance as thecoi1-1 mutant, the ein2-1 mutants were challenged with these three different fungal pathogens under the same conditions described in Thomma et al. (1998). All of these tests were performed on 4-week-old plants. Strain IMI169558 of the gray mold fungus B. cinerea did not cause any single case of complete plant decay among 60 inoculated wild-type plants. In contrast, 42% of the inoculated ein2-1plants were completely macerated by this strain over a 16-d period following inoculation (Fig. 2). B. cinerea strain MUCL30158, which was apparently more aggressive than strain IMI169558, caused decay of 9% and 100% of the inoculated wild-type and ein2-1 plants, respectively, within 16 d (Fig. 2). Therefore, ein2-1 mutants are more susceptible than wild-type plants to infection by either of two different strains of B. cinerea, which is in line with the observations made for the jasmonate-insensitive mutant coi1-1. Fig. 2. Open in new tabDownload slide Disease development on Arabidopsis inoculated withB. cinerea. A, Four-week-old Arabidopsis plants were drop-inoculated with B. cinerea strain IMI169558, and photographs were taken 12 d later. Circles (heads of pipet tips) indicate positions of completely decayed plants. B, Decay of Arabidopsis plants drop-inoculated with B. cinereastrains IMI169558 and MUCL30158. The percentage of dead plants is expressed as a function of time after inoculation. Plants were considered dead when their hearts were completely rotten. Data represent averages ± se of three different experiments performed with 20 plants per genotype. Circles, Wild-type (Col-0) plants; squares, the mutant ein2-1; white symbols, plants inoculated with strain IMI169558; black symbols, plants inoculated with strain MUCL30158. Fig. 2. Open in new tabDownload slide Disease development on Arabidopsis inoculated withB. cinerea. A, Four-week-old Arabidopsis plants were drop-inoculated with B. cinerea strain IMI169558, and photographs were taken 12 d later. Circles (heads of pipet tips) indicate positions of completely decayed plants. B, Decay of Arabidopsis plants drop-inoculated with B. cinereastrains IMI169558 and MUCL30158. The percentage of dead plants is expressed as a function of time after inoculation. Plants were considered dead when their hearts were completely rotten. Data represent averages ± se of three different experiments performed with 20 plants per genotype. Circles, Wild-type (Col-0) plants; squares, the mutant ein2-1; white symbols, plants inoculated with strain IMI169558; black symbols, plants inoculated with strain MUCL30158. When challenged with A. brassicicola strain MUCL20297, theein2-1 mutant produced restricted necrosis symptoms indicative of an incompatible interaction (Fig.3A). The necrotic lesions formed onA. brassicicola-inoculated ein2-1 plants had an average diameter that was about 2-fold higher compared with the diameter of lesions on wild-type plants (Fig. 3B). However, measurements of fungal biomass in the infected zones by hybridization of RNA dot blots with a fungus-specific probe did not reveal increased colonization of ein2-1 plants by A. brassicicolacompared with wild-type plants (Fig. 3C). In contrast, inoculation of the jasmonate-insensitive coi1-1 mutant with A. brassicicola yielded spreading lesions with markedly enhanced fungal colonization (Fig. 3; Thomma et al., 1998). Therefore,ein2-1 does not respond in the same way as coi1-1to this particular fungus. Fig. 3. Open in new tabDownload slide Disease development on Arabidopsis inoculated withA. brassicicola. A, Necrotic lesions on leaves of 4-week-old Arabidopsis wild-type (Col-0), ein2-1, andcoi1-1 plants drop-inoculated with spores of A. brassicicola. B, Average diameter of lesions formed after 6 d on 4-week-old Arabidopsis plants inoculated with a spore suspension of A. brassicicola. Data points represent averages ± se of measurements from 60 lesions on 15 different plants. Bars with different letter labels indicate that the corresponding data are significantly different (P> 0.95) according to Tukey's studentized range test (Neter et al., 1996). C, Percentage fungal RNA of total RNA in infection sites at different times after inoculation of leaves with A. brassicicola. Data points represent measurements on RNA extracted from 30 leaf discs. ○, Col-0; ■, ein2-1; and ♦, coi1-1. The experiment was repeated twice with similar results. Fig. 3. Open in new tabDownload slide Disease development on Arabidopsis inoculated withA. brassicicola. A, Necrotic lesions on leaves of 4-week-old Arabidopsis wild-type (Col-0), ein2-1, andcoi1-1 plants drop-inoculated with spores of A. brassicicola. B, Average diameter of lesions formed after 6 d on 4-week-old Arabidopsis plants inoculated with a spore suspension of A. brassicicola. Data points represent averages ± se of measurements from 60 lesions on 15 different plants. Bars with different letter labels indicate that the corresponding data are significantly different (P> 0.95) according to Tukey's studentized range test (Neter et al., 1996). C, Percentage fungal RNA of total RNA in infection sites at different times after inoculation of leaves with A. brassicicola. Data points represent measurements on RNA extracted from 30 leaf discs. ○, Col-0; ■, ein2-1; and ♦, coi1-1. The experiment was repeated twice with similar results. P. parasitica strain Wela has previously been shown to be avirulent on Arabidopsis Col-0 wild-type plants and on the jasmonate-insensitive mutant coi1-1, whereas Arabidopsis lines showing a defect in the salicylate-dependent defense pathway (NahG and npr1-1) were found to be susceptible to infection by this pathogen (Delaney et al., 1994; Thomma et al., 1998). When ein2-1 plants were challenged withP. parasitica strain Wela, a fully incompatible interaction was observed (Fig. 4). No intercellularly growing hyphae or oospores could be detected in any of 20 ein2-1 leaf samples analyzed under the microscope. In contrast, npr1-1 plants subjected to the same treatment showed an abundance of intercellularly growing hyphae and oospores (Fig. 4). This indicates that the ethylene response pathway is, unlike the salicylate response pathway, not implicated in the resistance of wild-type plants to an avirulent P. parasitica strain. Previous work established that Arabidopsis mutants affected in the ethylene-response pathway (etr1-1, ein2-1) do not show enhanced disease susceptibility relative to wild-type Col-0 plants to the virulent P. parasitica strain Noco (Lawton et al., 1994). Fig. 4. Open in new tabDownload slide Disease development on Arabidopsis inoculated withP. parasitica. Microscopic view of leaves of 4-week-old Arabidopsis wild-type (Col-0), ein2-1, andnpr1-1 plants spray-inoculated with conidiospores ofP. parasitica strain Wela. Eleven days after inoculation, inoculated leaves were stained with lactophenol trypan blue prior to microscopic examination. Leaves of thenpr1-1 mutant reveal the presence of intracellular hyphae and oospores. Fig. 4. Open in new tabDownload slide Disease development on Arabidopsis inoculated withP. parasitica. Microscopic view of leaves of 4-week-old Arabidopsis wild-type (Col-0), ein2-1, andnpr1-1 plants spray-inoculated with conidiospores ofP. parasitica strain Wela. Eleven days after inoculation, inoculated leaves were stained with lactophenol trypan blue prior to microscopic examination. Leaves of thenpr1-1 mutant reveal the presence of intracellular hyphae and oospores. Protection against A. brassicicola and B. cinerea by Ethylene and Methyl Jasmonate Pretreatment The remarkable susceptibility to the fungus B. cinerea of the ethylene-insensitive ein2-1 mutant (Fig.2) and the jasmonate-insensitive coi1-1 mutant (Thomma et al., 1998) suggests that ethylene- and jasmonate- dependent pathogen-inducible effector molecules contribute to resistance against this pathogen. Based on these observations, one would expect that increased production of such effector molecules prior to infection attempts by B. cinerea would enhance the resistance level to this pathogen. To test this prediction, wild-type Col-0 plants were placed for 2 d in airtight chambers containing either air or air supplemented with 0.5, 5.0, or 50 μL L−1 ethylene or 150 nmmethyl jasmonate, whereafter plants were inoculated with B. cinerea. The number of leaves showing soft rot symptoms was reduced by 57% in plants pretreated with 50 μL L−1 ethylene, while pretreatment with 150 nm methyl jasmonate reduced the number of leaves showing soft rot symptoms by 80% (Fig.5A). Fig. 5. Open in new tabDownload slide Protective effect of exogenously applied ethylene and methyl jasmonate on infection by B. cinerea strain MUCL30158 and A. brassicicola. A, Percentage of inoculated leaves showing spreading necrosis symptoms 6 d after inoculation of Arabidopsis wild-type (Col-0) plants with a spore suspension of B. cinerea. Prior to inoculation, separate sets of plants were placed for 48 h in gastight translucent chambers with an atmosphere containing the gaseous compounds as indicated below the bars. Data points represent averages ±se of seven series of inoculations on 16 leaves from two plants. Bars with different letter labels indicate that the corresponding data are significantly different (P> 0.95) according to Tukey's studentized range test (Neter et al., 1996). B, Percentage of inoculated leaves showing spreading necrosis symptoms 6 d after inoculation of Arabidopsisein2-1 plants with a spore suspension of B. cinerea. Specifications are as in the legend to A. C, Average diameter of lesions formed after 6 d on 4-week-old Arabidopsispad3-1 mutants inoculated with a spore suspension ofA. brassicicola. Prior to inoculation, separate sets of plants were placed for 48 h in gastight translucent chambers with an atmosphere containing the gaseous compounds as indicated below the bars. Data points represent averages ± se of measurements from 40 lesions on 10 different plants. Bars with different letter labels indicate that the corresponding data are significantly different (P > 0.95) according to Tukey's studentized range test (Neter et al., 1996). MeJA, Methyl jasmonate. Fig. 5. Open in new tabDownload slide Protective effect of exogenously applied ethylene and methyl jasmonate on infection by B. cinerea strain MUCL30158 and A. brassicicola. A, Percentage of inoculated leaves showing spreading necrosis symptoms 6 d after inoculation of Arabidopsis wild-type (Col-0) plants with a spore suspension of B. cinerea. Prior to inoculation, separate sets of plants were placed for 48 h in gastight translucent chambers with an atmosphere containing the gaseous compounds as indicated below the bars. Data points represent averages ±se of seven series of inoculations on 16 leaves from two plants. Bars with different letter labels indicate that the corresponding data are significantly different (P> 0.95) according to Tukey's studentized range test (Neter et al., 1996). B, Percentage of inoculated leaves showing spreading necrosis symptoms 6 d after inoculation of Arabidopsisein2-1 plants with a spore suspension of B. cinerea. Specifications are as in the legend to A. C, Average diameter of lesions formed after 6 d on 4-week-old Arabidopsispad3-1 mutants inoculated with a spore suspension ofA. brassicicola. Prior to inoculation, separate sets of plants were placed for 48 h in gastight translucent chambers with an atmosphere containing the gaseous compounds as indicated below the bars. Data points represent averages ± se of measurements from 40 lesions on 10 different plants. Bars with different letter labels indicate that the corresponding data are significantly different (P > 0.95) according to Tukey's studentized range test (Neter et al., 1996). MeJA, Methyl jasmonate. In contrast, pretreatment of ein2-1 plants with 50 μL L−1 ethylene did not reduce the disease incidence (Fig. 5B), indicating that the events causing protection in ethylene-treated wild-type plants are indeed dependent on a functional ethylene-response pathway. Pretreatment of ein2-1 plants with methyl jasmonate, on the other hand, still caused a reduction of the disease incidence by 64% (Fig. 5B). Similar experiments were also performed using A. brassicicola as a pathogen. In this case, however, wild-type Col-0 plants could not be used because A. brassicicola causes highly restricted, non-spreading lesions on this genotype. Instead, the pad3-1 mutant was used in these experiments. The pad3-1 mutant is deficient in an enzyme involved in the biosynthesis of camalexin (Glazebrook and Ausubel, 1994; N. Zhou and J. Glazebrook, personal communication), an antimicrobial metabolite that is an important determinant for resistance to A. brassicicola (Thomma et al., 1999). Previous work certified that ethylene- and jasmonate-dependent defense responses are still fully operative in the pad3-1 mutant (Thomma et al., 1999). Pretreatment of this mutant with 0.5, 5.0, or 50 μL L−1 ethylene in the atmosphere for 2 d prior to inoculation failed to confer any protection against A. brassicicola (Fig. 5C). In contrast, pretreatment of the plants with 150 nm gaseous methyl jasmonate reduced the average lesion diameter by 80% (Fig. 5C). DISCUSSION The results presented here confirm that Arabidopsis possesses a jasmonate/ethylene-dependent pathway for the induction of a particular subset of PR genes, including a plant defensin gene (PDF1.2), a basic chitinase gene (PR-3), and a hevein-like gene (PR-4). The involvement of both ethylene and jasmonate in this pathway is based on the observations thatPDF1.2, PR-3, and PR-4 can be activated by exogenous treatment with either methyl jasmonate (Thomma et al., 1998) or ethylene (Samac et al., 1990; Potter et al., 1993;Penninckx et al., 1996; B.P.H.J. Thomma, unpublished results), while they are not or very weakly induced by exogenous application of salicylic acid (Thomma et al., 1998). Moreover, induction of this set of genes upon challenge of Arabidopsis plants with the fungus A. brassicicola is largely abolished in a mutant (coi1-1;Thomma et al., 1998) affected in the COI1 gene, a gene encoding a signal transduction component of the jasmonate response (Xie et al., 1998). We have now shown that A. brassicicola-induced expression of these genes is also dramatically reduced in an ethylene-insensitive mutant (ein2-1) with a dysfunctional EIN2 gene encoding a membrane-associated signal transduction component of the ethylene response (McGrath and Ecker, 1998). Therefore, we considerPDF1.2, PR-3, and PR-4 as a class of co-regulated jasmonate/ethylene-dependent PR-genes whose regulation is clearly distinct from that of the salicylate-dependent PR-genes such as PR-1, PR-2, and PR-5 (Uknes et al., 1992; Cao et al., 1994; Delaney et al., 1994). The occurrence of two subsets of differentially regulated PR-genes has also been demonstrated in tobacco. The genes encoding extracellular isoforms such as acidic PR-1, acidic β-1,3-glucanase, and acidic chitinase are efficiently induced by salicylic acid but less so by ethylene (Memelink et al., 1990; Ohshima et al., 1990; Ward et al., 1991). Pathogen-induced activation of these genes is abolished in a transgenic line expressing the salicylate-degrading NahGgene (Gaffney et al., 1993). Another subset of PR genes, those encoding vacuolar PR proteins such as basic PR-1, basic β-1,3-glucanase, and basic chitinase, are more efficiently induced by ethylene than by salicylate (Memelink et al., 1990; Eyal et al., 1992; Beffa et al., 1995) and their pathogen-induced expression is down-regulated in transgenic tobacco plants expressing a dominant-negative mutant form of the Arabidopsis ethylene receptor ETR1 (Knoester et al., 1998). The role of jasmonate in the induction of the latter subset of PR genes has not yet been intensively studied, but Niki et al. (1998)recently reported that these genes can be induced by floating tobacco leaf discs on a jasmonate-containing solution. Therefore, a jasmonate/ethylene-dependent pathway for induction of particular PR genes also appears to be operative in tobacco. Arabidopsis PDF1.2 and PR-3 have previously been purified and shown to possess antifungal activity in vitro (Verburg and Huynh, 1991; Penninckx et al., 1996). Arabidopsis PR-4, on the other hand, has not yet been isolated, but it is known to be highly homologous to CBP-20, a tobacco PR protein with proven antifungal properties (Ponstein et al., 1994). PDF1.2, PR-3, and PR-4 are therefore likely to contribute to the defensive capacity of Arabidopsis plants directed against fungal organisms. Our results clearly show that the ein2-1 mutation in Arabidopsis entails markedly enhanced susceptibility to at least two different strains of the pathogenic fungus B. cinerea (Fig. 2). On the other hand, the ein2-1mutation had no impact on either resistance to an avirulent strain ofA. brassicicola (Fig. 3) or to an avirulent or a virulent strain of P. parasitica (Fig. 4 and Lawton et al., 1994, respectively). This is in line with the data obtained by Knoester et al. (1998) on ethylene-insensitive tobacco plants that were more susceptible than control plants to soil-borne Pythium spp. but not to tobacco mosaic virus. The ein2-1 mutation in Arabidopsis results in a lack of pathogen-inducible expression of a subset of PR genes (Penninckx et al., 1996; Fig. 1), as does expression of a dominant-negative mutantETR1 gene in tobacco (Knoester et al., 1998). However, these observations by themselves do not prove that such PR proteins are responsible for the control of particular pathogens. Ethylene insensitivity is likely to have pleiotropic effects, which would therefore affect the expression of other effector molecules as well. It is conceivable that such ethylene-controlled effector events are effective at controlling particular pathogens but have no effect on others. The data in the present study indicate that necrotrophic pathogens (e.g. B. cinerea in the case of Arabidopsis orPythium spp. in the case of tobacco) are among those that are effectively contained by ethylene-controlled effector molecules, whereas biotrophic pathogens (e.g. P. parasitica in the case of Arabidopsis and tobacco mosaic virus in the case of tobacco) are more efficiently countered by other defense mechanisms, including salicylate-controlled effector events. However, this may be a matter of coincidence and at the present time, it is more cautious not to speculate beyond the observation that some pathogens are kept in check by ethylene-controlled effector events while others are not. Both the Arabidopsis ein2-1 and coi1-1 mutants are more susceptible than wild-type plants to B. cinerea(Fig. 2; Thomma et al., 1998), although coi1-1 is more susceptible than ein2-1 in comparative assays (B.P.H.J. Thomma, unpublished results). In addition, a jasmonate-deficient mutant (fad3/fad7/fad8), a jasmonate-insensitive mutant (jar1) of Arabidopsis, and ethylene-insensitive tobacco plants are more susceptible than their respective control lines to soil-borne Pythium spp. (Knoester et al., 1998; Staswick et al., 1998; Vijayan et al., 1998). This may be seen as an additional argument for the involvement of jasmonate/ethylene-dependent PR genes in resistance against these pathogens, as expression of jasmonate/ethylene-dependent PR genes depends on both ethylene and jasmonate signal response pathways. Consistent with this notion we found that treatment of Arabidopsis plants with either methyl jasmonate or ethylene, both of which increase the levels of jasmonate/ethylene-dependent PR proteins, resulted in enhanced protection to B. cinerea. On the other hand, neitherein2-1 nor coi1-1 Arabidopsis mutants were more susceptible to P. parasitica (Fig. 4; Lawton et al., 1994;Thomma et al., 1998), excluding a role for jasmonate/ethylene-dependent PR genes in resistance against this pathogen. One interesting observation was that the coi1-1 andein2-1 mutants differed in their response to challenge byA. brassicicola. The coi1-1 mutant showed enhanced tissue colonization by this fungus relative to wild-type plants (Thomma et al., 1998), while the ein2-1mutant did not (Fig. 3). The most likely explanation for these results is that the jasmonate/ethylene-dependent PR genes are not effective or are only very marginally effective against this fungus, while a presumed jasmonate-dependent/ethylene-independent effector molecule may contribute much more effectively. The fact that the camalexin-deficient pad3-1 mutant is also more susceptible to A. brassicicola compared with wild-type plants suggests that this hypothetical effector molecule might be camalexin, the major Arabidopsis phytoalexin. However, camalexin production is not induced by treatment with jasmonate (Thomma et al., 1999), so we believe the hypothetical effector molecule to be different from camalexin. Consistent with the presumed existence of a jasmonate-inducible yet ethylene-independent effector molecule, we observed that treatment of pad3-1 mutants with methyl jasmonate increased the level of resistance to A. brassicicola, whereas pretreatment with ethylene failed to do so (Fig. 5). The jasmonate-inducible yet ethylene-independent effectors may also be effective against B. cinerea, as inferred from the observation that the ein2-1 mutant can be protected against this fungus by pretreatment with methyl jasmonate but not by ethylene (Fig. 5). A full range of pathogens are now available for future research that either cause less-severe symptoms on ethylene-insensitive versus ethylene-sensitive Arabidopsis genotypes (Ps. syringae andX. campestris, Bent et al., 1992), no or weak differences in symptoms or multiplication (A. brassicicola and P. parasitica, this study; Lawton et al., 1994), or more severe symptoms and increased multiplication (B. cinerea, this study). These data provide strong support to the notion that ethylene can play a balanced role in mounting disease resistance responses as well as in aggravation of disease symptoms, the outcome of which is dependent on the nature of the pathogen. ACKNOWLEDGMENTS The authors thank Drs. J. Turner, X. Dong, and J. Glazebrook for providing the mutants coi1-1, npr1-1, andpad3-1, respectively. The authors also thank Drs. R. Vogelsang and A. Slusarenko for providing P. parasiticastrain Wela. 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K.T. is the recipient of a predoctoral fellowship of the Vlaams Instituut voor Bevordering van het Wetenschappelijk-Technologisch Onderzoek in de Industrie. * Corresponding author; [email protected]; fax 32–16–321966. 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)

Walden, Adrian R.; Walter, Christian; Gardner, Richard C.

doi: 10.1104/pp.121.4.1103pmid: 10594098

Abstract We describe the isolation and characterization of 13 cDNA clones that are differentially expressed in male cones of Pinus radiata (D. Don). The transcripts of the 13 genes are expressed at different times between meiosis and microspore mitosis, timing that corresponds to a burst in tapetal activity in the developing anthers. In situ hybridization showed that four of the genes are expressed in the tapetum, while a fifth is expressed in tetrads during a brief developmental window. Six of the seven cDNAs identified in database searches have striking similarity to genes expressed in angiosperm anthers. Seven cDNAs are homologs of defense and pathogen response genes. The cDNAs identified are predicted to encode a chalcone-synthase-like protein, a thaumatin-like protein, a serine hydrolase thought to be a putative regulator of programmed cell death, two lipid-transfer proteins, and two homologs of the anther-specific A9 genes from Brassica napus and Arabidopsis. Overall, our results support the hypothesis that many of the reproductive processes in the angiosperms and gymnosperms were inherited from a common ancestor. The gymnosperm Pinus radiata is monoecious, with male cones on the lower branches of the tree and female cones on the upper branches (see Fig. 1). Male cones consist of a spiral arrangement of tightly packed pollen-bearing structures or microsporophylls formed around a central axis. The lower surfaces of the microsporophylls have two chambers called microsporangia in which the pollen develops. Nutrients for the developing pollen are supplied from an inner layer of microsporangial cells called the tapetum. Fig. 1. Open in new tabDownload slide P. radiata male (left) and female (right) cones photographed during male cone dehiscence. Fig. 1. Open in new tabDownload slide P. radiata male (left) and female (right) cones photographed during male cone dehiscence. Anther-expressed genes have been isolated from angiosperm flowers, nearly all of which are expressed in the microspores or tapetum during or after the burst in tapetal activity that occurs during meiosis (Williams and Heslop-Harrison, 1979; Pacini et al., 1985; Scott et al., 1991b). Most genes encode distinct proteins that share similar, tightly regulated temporal and spatial expression patterns (for reviews, seeScott et al., 1991b; Goldberg et al., 1993). Pollen development in gymnosperms shares several developmental and physiological processes with that of angiosperms (Stanley and Linskens, 1974; Pacini et al., 1985), and homologs to angiosperm floral transcription factors have been isolated from gymnosperm reproductive tissues (Tandre et al., 1995; Mellerowicz et al., 1998; Mouradov et al., 1998a, 1998b). However, there are no reports describing cDNAs that are differentially expressed in gymnosperm male reproductive structures. Anther development can be divided into a series of defined developmental stages (Koltunow et al., 1990; Scott et al., 1991b; Goldberg et al., 1993) that correlate with aspects of male cone development. In P. radiata, male cone primordia first become visible in early summer (Wang, 1995); microsporophylls develop after 3 months, and the archesporial cells differentiate into the sporogenous cells and the parietal layer after 5 months. The parietal cells form a microsporangium wall four to five cells thick (the innermost layer is the tapetum) and the sporogenous tissue divides. After 6 months, the sporogenous cells are mature meiocytes and meiosis begins. Meiosis is complete after 7 months, the microspores are released from the tetrad, and the tapetum begins to degenerate (Wang, 1995). During degeneration, the cells of the tapetum release accumulated quantities of starch, crystallized protein, lipid, and polysaccharide, which are used by the microspores (Pacini et al., 1985). The two sacci of each pollen grain begin to inflate prior to the first division of the microspore as the three- to four-cell layer surrounding the sporangium collapses (Wang, 1995). Some time after 8 months, the pollen mother cells undergo three mitotic divisions. When dehiscence occurs, after about 9 months, each pollen grain contains two nonfunctional prothallial cells, a central vegetative cell, and a generative cell (Stanley and Linskens, 1974). In New Zealand, Australia, and Chile, P. radiata is grown on large-scale plantations for timber and fiber. We were interested in engineering reproductive sterility as a means to manipulate biomass distribution and to control the spread and persistence of introduced genes (Kaul, 1988; Strauss et al., 1995). One strategy for engineering sterility involves directing the expression of cytotoxic genes to male cones using tissue-specific promoters. Such promoters are available from various angiosperm species, but their tissue specificity and long-term expression characteristics in conifers remain unknown. We describe here the isolation and characterization of 13 cDNAs that are differentially expressed in P. radiata male cones. Six of the seven cDNAs identified by database searches are homologs of angiosperm genes expressed in anthers, indicating that a significant proportion of differentially expressed anther genes evolved prior to the divergence of gymnosperms and angiosperms. All of the seven cDNAs identified are also homologs of genes involved in the plant response to stress and pathogens. MATERIALS AND METHODS Tissue Collection All tissues used in this study were collected from New Zealand Forest Research Institute nurseries in Rotorua (latitude 38° 24′, altitude 350 m). Vegetative shoot and cone tissue was harvested from mature trees. Root tissue was collected from 1-year-old cuttings, and needle tissue was collected from 4-year-old seedlings. Tissue was immediately stored in liquid nitrogen in transit to refrigeration at −80°C, or immersed in 10× volume of FAA fixative (ethanol 50% [v/v], glacial acetic acid 5% [v/v], and formalin 10% [v/v]). Assessment of Microsporocyte Development Male cones harvested during 1994 and 1995 were removed from FAA fixative, dissected, and the microsporangium contents were released into a solution of 3% (w/v) Suc. Samples were transferred onto microscope slides and viewed under a light microscope to assess microsporocyte development. The developmental stage of pollen in these cones (listed in Table I) was assessed according to the stages described by Scott et al. (1991b) andKoltunow et al. (1990). Images were captured with a video camera connected to a Power Macintosh 7500 computer using the Image program developed at the National Institutes of Health (available on the Internet at http://rsb.info.nih.gov/nih-image/). Table I. Developmental stage of pollen extracted from male cones Harvest Date (Months after Cone Primordia Appear) . Morphological Description . Stage .   3/23/94 (3.75) Archesporial tissue not differentiated* A2 (Archesporial)   4/12/94 (4.5) Sporogenous and parietal cells differentiated* SP (Sporogenous parietal)   4/25/94 (4.75) Meiocytes at late interphase Me-i (Meiocyte, interphase)   5/5/94 (5) Meiocytes at late interphase, some early prophase Me-iep (Meiocytes, interphase early prophase I)   5/26/94 (5.75) Meiocytes at prophase I Me-p (Meiocytes, prophase I)   6/6/94 (6.25) Meiocytes at late anaphase I, early telophase I Me-at (Meiocytes, anaphase I telophase I)   6/29/94 (7) Tetrads T1 (Tetrads)   7/15/94 (7.5) Microspore sacci partially inflated Mi (Microspores)   3/17/95 (3.5) Archesporial tissue not differentiated* A1 (Archesporial)   7/3/95 (7) Meiocytes at anaphase I Me-a (Meiocytes, anaphase I)   7/7/95 (7.25) Meiocytes at anaphase I Me-a′ (Meiocytes, anaphase I)   8/11/95 (8.25) Microspore sacci inflated Mi-si (Microspores, sacci inflated)   9/1/95 (9) Mature pollen (two prothallial cells, a generative cell and a tube nucleus) Mp (Mature pollen)   7/4/96 (7) Meiocytes at telophase I and prophase II Me-tIpII (Telophase I and prophase II)   7/16/96 (7.5) Tetrads T2 (Tetrads) Harvest Date (Months after Cone Primordia Appear) . Morphological Description . Stage .   3/23/94 (3.75) Archesporial tissue not differentiated* A2 (Archesporial)   4/12/94 (4.5) Sporogenous and parietal cells differentiated* SP (Sporogenous parietal)   4/25/94 (4.75) Meiocytes at late interphase Me-i (Meiocyte, interphase)   5/5/94 (5) Meiocytes at late interphase, some early prophase Me-iep (Meiocytes, interphase early prophase I)   5/26/94 (5.75) Meiocytes at prophase I Me-p (Meiocytes, prophase I)   6/6/94 (6.25) Meiocytes at late anaphase I, early telophase I Me-at (Meiocytes, anaphase I telophase I)   6/29/94 (7) Tetrads T1 (Tetrads)   7/15/94 (7.5) Microspore sacci partially inflated Mi (Microspores)   3/17/95 (3.5) Archesporial tissue not differentiated* A1 (Archesporial)   7/3/95 (7) Meiocytes at anaphase I Me-a (Meiocytes, anaphase I)   7/7/95 (7.25) Meiocytes at anaphase I Me-a′ (Meiocytes, anaphase I)   8/11/95 (8.25) Microspore sacci inflated Mi-si (Microspores, sacci inflated)   9/1/95 (9) Mature pollen (two prothallial cells, a generative cell and a tube nucleus) Mp (Mature pollen)   7/4/96 (7) Meiocytes at telophase I and prophase II Me-tIpII (Telophase I and prophase II)   7/16/96 (7.5) Tetrads T2 (Tetrads) The harvest dates (mm/dd/yy) of P. radiata cones used in this study are shown. The approximate time that male cone primordia become visible is based on data collected in previous years (Wang, 1995). Morphological descriptions were determined by microscopic analysis, except the earliest stages, which are marked by asterisks and are based on data collected in previous years by Wang (1995). The abbreviations are based on the description of B. napusanther development (Scott et al., 1991). Numbers are used to distinguish samples at the same stage from different years. The last two samples were used for in situ analysis only. Note that there were clear differences in the rate of development over the 2 years sampled, since the Me-a stage was reached almost 1 month later in 1995 (7/3/95) than in 1994 (6/6/94). Open in new tab Table I. Developmental stage of pollen extracted from male cones Harvest Date (Months after Cone Primordia Appear) . Morphological Description . Stage .   3/23/94 (3.75) Archesporial tissue not differentiated* A2 (Archesporial)   4/12/94 (4.5) Sporogenous and parietal cells differentiated* SP (Sporogenous parietal)   4/25/94 (4.75) Meiocytes at late interphase Me-i (Meiocyte, interphase)   5/5/94 (5) Meiocytes at late interphase, some early prophase Me-iep (Meiocytes, interphase early prophase I)   5/26/94 (5.75) Meiocytes at prophase I Me-p (Meiocytes, prophase I)   6/6/94 (6.25) Meiocytes at late anaphase I, early telophase I Me-at (Meiocytes, anaphase I telophase I)   6/29/94 (7) Tetrads T1 (Tetrads)   7/15/94 (7.5) Microspore sacci partially inflated Mi (Microspores)   3/17/95 (3.5) Archesporial tissue not differentiated* A1 (Archesporial)   7/3/95 (7) Meiocytes at anaphase I Me-a (Meiocytes, anaphase I)   7/7/95 (7.25) Meiocytes at anaphase I Me-a′ (Meiocytes, anaphase I)   8/11/95 (8.25) Microspore sacci inflated Mi-si (Microspores, sacci inflated)   9/1/95 (9) Mature pollen (two prothallial cells, a generative cell and a tube nucleus) Mp (Mature pollen)   7/4/96 (7) Meiocytes at telophase I and prophase II Me-tIpII (Telophase I and prophase II)   7/16/96 (7.5) Tetrads T2 (Tetrads) Harvest Date (Months after Cone Primordia Appear) . Morphological Description . Stage .   3/23/94 (3.75) Archesporial tissue not differentiated* A2 (Archesporial)   4/12/94 (4.5) Sporogenous and parietal cells differentiated* SP (Sporogenous parietal)   4/25/94 (4.75) Meiocytes at late interphase Me-i (Meiocyte, interphase)   5/5/94 (5) Meiocytes at late interphase, some early prophase Me-iep (Meiocytes, interphase early prophase I)   5/26/94 (5.75) Meiocytes at prophase I Me-p (Meiocytes, prophase I)   6/6/94 (6.25) Meiocytes at late anaphase I, early telophase I Me-at (Meiocytes, anaphase I telophase I)   6/29/94 (7) Tetrads T1 (Tetrads)   7/15/94 (7.5) Microspore sacci partially inflated Mi (Microspores)   3/17/95 (3.5) Archesporial tissue not differentiated* A1 (Archesporial)   7/3/95 (7) Meiocytes at anaphase I Me-a (Meiocytes, anaphase I)   7/7/95 (7.25) Meiocytes at anaphase I Me-a′ (Meiocytes, anaphase I)   8/11/95 (8.25) Microspore sacci inflated Mi-si (Microspores, sacci inflated)   9/1/95 (9) Mature pollen (two prothallial cells, a generative cell and a tube nucleus) Mp (Mature pollen)   7/4/96 (7) Meiocytes at telophase I and prophase II Me-tIpII (Telophase I and prophase II)   7/16/96 (7.5) Tetrads T2 (Tetrads) The harvest dates (mm/dd/yy) of P. radiata cones used in this study are shown. The approximate time that male cone primordia become visible is based on data collected in previous years (Wang, 1995). Morphological descriptions were determined by microscopic analysis, except the earliest stages, which are marked by asterisks and are based on data collected in previous years by Wang (1995). The abbreviations are based on the description of B. napusanther development (Scott et al., 1991). Numbers are used to distinguish samples at the same stage from different years. The last two samples were used for in situ analysis only. Note that there were clear differences in the rate of development over the 2 years sampled, since the Me-a stage was reached almost 1 month later in 1995 (7/3/95) than in 1994 (6/6/94). Open in new tab RNA Extraction RNA extraction of frozen tissue was by LiCl precipitation (Chang et al., 1993). Synthesis of cDNA for probing cDNA libraries was primed with oligo p(dTTP)15 primers (Boehringer Mannheim, Mannheim, Germany) and catalyzed by reverse transcriptase (Superscript II, Gibco-BRL, Gaithersburg, MD) according to the manufacturer's instructions. cDNA Library Construction Meiocyte (Me-i-stage) and tetrad (T1-stage) cDNA libraries were constructed using a plasmid cloning kit (Superscript, Gibco-BRL) according to the manufacturer's instructions. Escherichia coli DH10B cells competent for electro-poration were transformed with aliquots of the ligation reaction using an electroporation apparatus (Bio-Rad Laboratories, Hercules, CA) (2,500 V, 50 μFa, 200 Ω) according to the manufacturer's instructions with 2-mm gap cuvettes. Differential Screening of the cDNA Library E. coli transformants (500 per plate) from the meiocyte stage (16,000) and tetrad stage (8,000) libraries were grown on 132-mm colony/plaque screen nylon membranes (DuPont-NEN, Boston). Replica plating, colony lysis, and DNA fixation (using an alkali method) were carried out according to the manufacturer's instructions. Cellular debris were removed from the membranes according to the method ofVogeli and Kaytes (1987). A differential screening strategy was used to select clones that hybridized strongly to a [32P]dCTP-labeled cDNA mixture (Rediprime, Amersham, Buckinghamshire, UK) prepared from Me-i-stage- and T1-stage-male cone cDNA, but did not hybridize to a cDNA mixture prepared from needle, root, and shoot tissue. Hybridization, stringency washing, and autoradiography were carried out according to the manufacturer's instructions (DuPont-NEN). Duplicate filters were initially hybridized with the first probe, washed, and hybridized with the second probe. Selected clones that were highly expressed in male cones on both filters were picked and amplified for further analysis. For secondary screening, plasmid DNA was purified from selected colonies, digested with NotI/SalI, and fractionated on 1.2% (w/v) agarose gels. Nucleic acids were transferred to nylon membrane (Hybond N+, Amersham) using the procedure outlined for Southern blotting. The filters were screened with the radiolabeled needle, root, and shoot cDNA mixture and then with the radiolabeled meiocyte and microspore cone cDNA as described above. Selected clones were further analyzed by northern blots of Me-i- and T1-stage male cone, needle, root, and shoot RNA (data not shown). Dot-Blot Hybridization Analysis Plasmid DNA samples were diluted 10-fold, denatured for 5 min (95°C), made up to 5× SSC, and then 2-μL aliquots were placed on Hybond N+ membrane prewetted in 10× SSC. Filters were probed with purified cDNA fragments released from pSPORT1 byNotI/SalI digestion. Filters were hybridized, washed (final wash in 2× SSC, 75°C; 20× SSC is 0.3m sodium citrate and 3 mNaCl), and exposed to film for 3 to 12 h. Northern Hybridization Analysis Aliquots (5 μg) of total RNA were glyoxylated, fractionated on agarose gels, and transferred to Hybond N+ nylon membranes (Sambrook et al., 1989; Munch, 1994). Hybridization conditions were as described by Virca et al. (1990). Loading differences were assessed by probing blots with a probe to the 26S rRNA that was amplified using PCR with specific primers (kindly provided by M. Jacobs). Southern Hybridization Analysis Genomic DNA was isolated from cone or young needle tissue according to previously published methods (Doyle and Doyle, 1990; Fang et al., 1992). Southern hybridization was carried out using 10 μg of digested genomic DNA per lane according to standard methods (Sambrook et al., 1989). Sequence Analysis Plasmid DNA was prepared for automated sequencing or further analysis according to a previously published method (Feliciello and Chinali, 1993). Sequencing was carried out with automated sequencers (ABI Prism 373 or 377, Perkin-Elmer, Foster City, CA). Initial sequence data were obtained using M13/pUC forward and reverse sequencing primers; when required specific primers were designed to sites within each cDNA. Sequences were analyzed using the Genetics Computer Group package (versions 8.1 and 9.1, GCG, Madison, WI) and HOMED (Dr. Peter A. Stockwell, Department of Biochemistry, University of Otago, New Zealand) software packages. Unless stated otherwise, deduced amino acid sequences were translated from the first in-frame ATG in each cDNA and terminated at the first stop codon encountered. Homology inferences are based on the results of BLAST and FASTA database searches. Phylogenetic analysis was carried out using the GCG version of PAUP (version 4.0.0d55 for UNIX) with parsimony and heuristic search criteria and 100 boot strap replications to assess branching confidence. In Situ Hybridization In situ hybridization was based on existing protocols (Cox and Goldberg, 1988; Bochenek and Hirsch, 1990; Wilkinson, 1992) with the addition of proprietary reagents supplied in the RNA Color Kit (Amersham). Tissue was fixed overnight in freshly prepared ice-cold fixative (gluteraldehyde 0.1% [w/v] and formaldehyde 4% [w/v], in phosphate-buffered saline [PBS], pH 7.2), washed in PBS (room temperature), dehydrated in an ethanol dilution series and infiltrated with xylene. Xylene was gradually replaced with molten paraffin wax (Paraplast, Sigma, St. Louis) and infiltration was continued for up to 2 d. The tissue was embedded, sectioned (8–10 μm), and baked onto Lys-coated slides (1–2 d, 42°C). Sections were rehydrated, incubated in 0.2 m HCl (20 min, room temperature), equilibrated in 10 mm Tris and 1 mm EDTA, pH 8.0 (TE), and then digested in 1 μg mL−1 proteinase K in TE buffer (37°C for 30 min). Digestion was stopped with 2 mg mL−1 Gly in PBS (5 min). Sections were equilibrated in an aqueous solution of triethanolamine (0.1 m, pH 8.0), acetylated with freshly prepared acetic anhydride (0.5% [v/v] in triethanolamine solution, 10 min), washed in PBS, and dehydrated prior to hybridization. Probe preparation, hybridization, and signal detection were carried out using the Amersham RNA Color Kit (catalog no. RPN3300). Sections were probed with single-stranded fluorescein-labeled RNA probes corresponding to the sense and antisense strands of the cDNA. After hybridization, sections were rinsed in 2× SSC, treated with RNase A (10 μg mL−1, 2× SSC, room temperature, 20 min), and washed under stringent conditions (1× SSC, 0.1% [w/v] SDS, 5 min, room temperature, then twice in 0.2× SSC, 0.1% [w/v] SDS, 55°C, 10 min). Sections were washed in TBS (100 mm Tris HCl, pH 7.5, and 400 mm NaCl, 5 min), incubated in block solution (0.5% [w/v] Amersham proprietary blocking agent in TBS for 1 h), rinsed in TBS then drained. Bound probe was detected using an anti-fluorescein antibody conjugated to alkaline phosphatase. Sections were incubated with the antibody for 1 h (1/1,000 in 0.5% [w/v] BSA in TBS), rinsed three times in TBS, washed in detection buffer (100 mm Tris HCl, pH 9.5, 100 mm NaCl, and 50 mm MgCl2) for 5 min, and drained. Detection buffer with 5-bromo-4-chloro-3-indolyl phosphate (0.18 mg mL−1, Amersham) and nitroblue tetrazolium (0.34 mg mL−1, Amersham) was added to each section and left to develop in the dark for up to 24 h. After the required development, the slides were rinsed in distilled water, dried, and mounted. Images were recorded on 35 mm Kodak Ektachrome 25t film. RESULTS Assessment of Male Cone Development We use a system of abbreviations based on that of Scott et al. (1991b) to describe the stages of pollen development. A summary of cones collected is listed in Table I. Thirteen Differentially Expressed cDNAs Were Isolated from the Microspore Library Two cDNA libraries were prepared, one from Me-i-stage-male cones (the meiocyte library) and one from T1-stage-male cones (the microspore library). Sixteen thousand clones from the meiocyte library and 8,000 clones from the tetrad library were differentially screened by sequentially probing duplicate filters with each of two probes. One was prepared from mRNA isolated from Me-i- and T1-stage-male cones and one from a mix of mRNA isolated from roots, needles, and vegetative shoots. A total of 120 differential clones were selected from the microspore library and a strict secondary screen (see “Materials and Methods”) reduced this number to 37 clones. No differential clones were obtained from screening the meiocyte library. The 37 differential clones were placed into 13 groups of cross-hybridizing clones (summarized in TableII). Table II. Summary data for cDNAs isolated in this study Largest Clone (No. Cloned) . cDNA Size . mRNA Size . Temporal Expression . Spatial Expression . Homology . Genomic Copy No. . Accession No. .   bp PrMC6 2,020 2,300            T1 Mi ND Intermediate AA220862 PrThL1 (3) 1,040 1,480 Me-a Me-at T1 Mi ND Thaumatin/permatin Intermediate AA220863 PrLTP1 (10*) 630 950 Me-a Me-at T1 Mi Tapetum Non-specific lipid transfer protein High U90342 * (AI857146) PrMC75 620 750 Me-a Me-at T1 Mi Mi-si ND Low AA220866 PrMC1 (3) 611 650 Me-a Me-at T1 Mi Tapetum A9 tapetum-expressed gene Low U90350 * PrChS1 (4) 1,468 1,350 Me-a Me-at T1 Mi Mi-si Tapetum Chalcone synthase/ stilbene synthase Low U90341 * PrLTP2 (2) 582 800            T1 Tetrads Non-specific lipid transfer protein ND AF110332 * PrMC103 1,375 2,100      Me-at T1 Mi ND Low AA220871 PrMC104 703 620               Mi Mi-si ND Intermediate AA220872 PrMC136 450 2,800 Me-a Me-at T1 Mi ND Low AA220874 PrMC187 (5) 1,330 1,300      Me-at T1 Mi Mi-si ND Intermediate AI857147 PrMC2 750 820 Me-a Me-at T1 Mi Mi-si Tapetum A9 tapetum-expressed gene Low U90343 * PrMC3 (4) 1,107 1,700 Me-a Me-at T1 Mi ND Ser hydrolase Low AF110333 * Largest Clone (No. Cloned) . cDNA Size . mRNA Size . Temporal Expression . Spatial Expression . Homology . Genomic Copy No. . Accession No. .   bp PrMC6 2,020 2,300            T1 Mi ND Intermediate AA220862 PrThL1 (3) 1,040 1,480 Me-a Me-at T1 Mi ND Thaumatin/permatin Intermediate AA220863 PrLTP1 (10*) 630 950 Me-a Me-at T1 Mi Tapetum Non-specific lipid transfer protein High U90342 * (AI857146) PrMC75 620 750 Me-a Me-at T1 Mi Mi-si ND Low AA220866 PrMC1 (3) 611 650 Me-a Me-at T1 Mi Tapetum A9 tapetum-expressed gene Low U90350 * PrChS1 (4) 1,468 1,350 Me-a Me-at T1 Mi Mi-si Tapetum Chalcone synthase/ stilbene synthase Low U90341 * PrLTP2 (2) 582 800            T1 Tetrads Non-specific lipid transfer protein ND AF110332 * PrMC103 1,375 2,100      Me-at T1 Mi ND Low AA220871 PrMC104 703 620               Mi Mi-si ND Intermediate AA220872 PrMC136 450 2,800 Me-a Me-at T1 Mi ND Low AA220874 PrMC187 (5) 1,330 1,300      Me-at T1 Mi Mi-si ND Intermediate AI857147 PrMC2 750 820 Me-a Me-at T1 Mi Mi-si Tapetum A9 tapetum-expressed gene Low U90343 * PrMC3 (4) 1,107 1,700 Me-a Me-at T1 Mi ND Ser hydrolase Low AF110333 * The number of cross-hybridizing clones in each cDNA group is in brackets adjacent to the name of the largest clone. Transcript size was derived from northern analysis. Temporal and spatial expression data are summarized from northern-blot and in situ hybridization data; developmental stage abbreviations are from Table I. Copy number data are derived from Southern analysis. The accession numbers are for the sequence of the largest clone in each group. Asterisks indicate that the complete sequence of cDNA has been determined, the remaining sequences are lodged in the database as expressed sequence tags. Accession number of the largest clone in the second PrLTP1 subgroup is in parentheses. ND, Not determined. Open in new tab Table II. Summary data for cDNAs isolated in this study Largest Clone (No. Cloned) . cDNA Size . mRNA Size . Temporal Expression . Spatial Expression . Homology . Genomic Copy No. . Accession No. .   bp PrMC6 2,020 2,300            T1 Mi ND Intermediate AA220862 PrThL1 (3) 1,040 1,480 Me-a Me-at T1 Mi ND Thaumatin/permatin Intermediate AA220863 PrLTP1 (10*) 630 950 Me-a Me-at T1 Mi Tapetum Non-specific lipid transfer protein High U90342 * (AI857146) PrMC75 620 750 Me-a Me-at T1 Mi Mi-si ND Low AA220866 PrMC1 (3) 611 650 Me-a Me-at T1 Mi Tapetum A9 tapetum-expressed gene Low U90350 * PrChS1 (4) 1,468 1,350 Me-a Me-at T1 Mi Mi-si Tapetum Chalcone synthase/ stilbene synthase Low U90341 * PrLTP2 (2) 582 800            T1 Tetrads Non-specific lipid transfer protein ND AF110332 * PrMC103 1,375 2,100      Me-at T1 Mi ND Low AA220871 PrMC104 703 620               Mi Mi-si ND Intermediate AA220872 PrMC136 450 2,800 Me-a Me-at T1 Mi ND Low AA220874 PrMC187 (5) 1,330 1,300      Me-at T1 Mi Mi-si ND Intermediate AI857147 PrMC2 750 820 Me-a Me-at T1 Mi Mi-si Tapetum A9 tapetum-expressed gene Low U90343 * PrMC3 (4) 1,107 1,700 Me-a Me-at T1 Mi ND Ser hydrolase Low AF110333 * Largest Clone (No. Cloned) . cDNA Size . mRNA Size . Temporal Expression . Spatial Expression . Homology . Genomic Copy No. . Accession No. .   bp PrMC6 2,020 2,300            T1 Mi ND Intermediate AA220862 PrThL1 (3) 1,040 1,480 Me-a Me-at T1 Mi ND Thaumatin/permatin Intermediate AA220863 PrLTP1 (10*) 630 950 Me-a Me-at T1 Mi Tapetum Non-specific lipid transfer protein High U90342 * (AI857146) PrMC75 620 750 Me-a Me-at T1 Mi Mi-si ND Low AA220866 PrMC1 (3) 611 650 Me-a Me-at T1 Mi Tapetum A9 tapetum-expressed gene Low U90350 * PrChS1 (4) 1,468 1,350 Me-a Me-at T1 Mi Mi-si Tapetum Chalcone synthase/ stilbene synthase Low U90341 * PrLTP2 (2) 582 800            T1 Tetrads Non-specific lipid transfer protein ND AF110332 * PrMC103 1,375 2,100      Me-at T1 Mi ND Low AA220871 PrMC104 703 620               Mi Mi-si ND Intermediate AA220872 PrMC136 450 2,800 Me-a Me-at T1 Mi ND Low AA220874 PrMC187 (5) 1,330 1,300      Me-at T1 Mi Mi-si ND Intermediate AI857147 PrMC2 750 820 Me-a Me-at T1 Mi Mi-si Tapetum A9 tapetum-expressed gene Low U90343 * PrMC3 (4) 1,107 1,700 Me-a Me-at T1 Mi ND Ser hydrolase Low AF110333 * The number of cross-hybridizing clones in each cDNA group is in brackets adjacent to the name of the largest clone. Transcript size was derived from northern analysis. Temporal and spatial expression data are summarized from northern-blot and in situ hybridization data; developmental stage abbreviations are from Table I. Copy number data are derived from Southern analysis. The accession numbers are for the sequence of the largest clone in each group. Asterisks indicate that the complete sequence of cDNA has been determined, the remaining sequences are lodged in the database as expressed sequence tags. Accession number of the largest clone in the second PrLTP1 subgroup is in parentheses. ND, Not determined. Open in new tab Each of the 37 sequences was partially sequenced to confirm the placement into 13 hybridization groups. Of the seven groups with more than one member, six contain cDNAs that appeared identical in the overlapping regions that have been sequenced, and therefore are likely to represent independent clones from the same gene. One hybridization group (largest clone PrLTP1) had 10 cross-hybridizing clones that fell into two subgroups of five identical members. Identity between the two subgroups for the region corresponding to nucleotides 43 to 126 of PrLTP1 (predicted coding sequence is from 39–422) was 87%. Seven of the 13 cDNA clones shared homology to sequences in the databases; six of these genes were sequenced completely (Table II; see “Discussion”). Most cDNAs Are Expressed from Just Before to Just After Meiosis Northern blots using male cones collected over two flowering seasons were used initially to characterize the expression of the 13 genes. The results are shown in Figure2, and summarized in Table II and Figure3. No transcripts of any of the cDNAs were detected in roots, vegetative shoots, needles or female cones. In male cones, the timing and duration of expression varied widely for the 13 genes, but in each case occurred between meiosis and the first mitotic division of the microspore. This timing is similar to that of most angiosperm anther-specific genes (Scott et al., 1991b;Goldberg et al., 1993) and corresponds to the stage of a burst in tapetal activity in the developing anthers (Heslop-Harrison, 1968). The expression of five transcripts (PrThL1, PrLTP1, PrMC1, PrMC136, and PrMC3) was first detected in Me-a-stage cones and last detected in Mi-stage cones. Three other transcripts (PrMC75, PrCHS1, and PrMC2) were also first expressed in Me-a-stage cones, but their expression extended for a slightly longer period of development, lasting up to the formation of microspores with inflated sacci (Mi-si stage). Transcripts of PrMC103 and PrMC187 were first detected in Me-at-stage cones. The PrMC103 probe detected faint signals from Me-at- and T1-stage cones and was last detected in Mi-stage cones. PrMC187 was last detected in Mi-si–stage-cones. Transcripts of PrMC6 and PrLTP2 were expressed for short periods within the burst of tapetal activity, suggesting that their expression may be regulated differently. PrMC104 was notably later than all the other genes, with transcript undetected until the juvenile microspores were present and disappearing by the time mature pollen had formed. Fig. 2. Open in new tabDownload slide Temporal expression analysis. Northern blots of total RNA extracted from male cones during two flowering seasons. Abbreviations for male cones are given in Table I. Needle, root, and shoot data (where presented) are as indicated, and the abbreviations for female cones are as follows: Fc8, female cones 8 months before meiosis; Fc7, female cones 7 months before meiosis; Fc4, female cones 4 months before meiosis. Expression data and transcript size are summarized in Figure 3 and Table II. Loading differences were assessed by probing blots with a 26S rRNA probe as indicated. Note that there is wide variation in the loading of samples between lanes, with some samples underloaded (e.g. T1, Misi); this is the likely cause of the apparent transient down-regulation of genes such as PrLTP1 in T1-stage cones. Faint smears immediately adjacent to lanes containing strongly hybridizing bands (e.g. PrMC104, T1 sample) have been interpreted as noise in the summary in Figure 3. Fig. 2. Open in new tabDownload slide Temporal expression analysis. Northern blots of total RNA extracted from male cones during two flowering seasons. Abbreviations for male cones are given in Table I. Needle, root, and shoot data (where presented) are as indicated, and the abbreviations for female cones are as follows: Fc8, female cones 8 months before meiosis; Fc7, female cones 7 months before meiosis; Fc4, female cones 4 months before meiosis. Expression data and transcript size are summarized in Figure 3 and Table II. Loading differences were assessed by probing blots with a 26S rRNA probe as indicated. Note that there is wide variation in the loading of samples between lanes, with some samples underloaded (e.g. T1, Misi); this is the likely cause of the apparent transient down-regulation of genes such as PrLTP1 in T1-stage cones. Faint smears immediately adjacent to lanes containing strongly hybridizing bands (e.g. PrMC104, T1 sample) have been interpreted as noise in the summary in Figure 3. Fig. 3. Open in new tabDownload slide Most cDNAs are expressed from before to just after meiosis. This figure illustrates the expression of the 13 cDNAs at different stages of pollen development (top). Development progresses from left to right. The cDNAs are labeled on the left side of the figure. An oval spot represents expression. Abbreviations for male cones are given in Table I. Bars indicating the size of developing pollen are 9 μm. Me-iep- to Me-at-stage meiocytes are enlarged by a factor of two. Fig. 3. Open in new tabDownload slide Most cDNAs are expressed from before to just after meiosis. This figure illustrates the expression of the 13 cDNAs at different stages of pollen development (top). Development progresses from left to right. The cDNAs are labeled on the left side of the figure. An oval spot represents expression. Abbreviations for male cones are given in Table I. Bars indicating the size of developing pollen are 9 μm. Me-iep- to Me-at-stage meiocytes are enlarged by a factor of two. Most Genes Are Expressed in the Tapetal Layer Spatial expression patterns of five genes, PrChS1, PrLTP1, PrLTP2, PrMC1, and PrMC2 were analyzed in developing male cones by in situ hybridization analysis (Fig. 4). In Me-tIpII-stage cones, expression of PrChS1, PrLTP1, PrMC1, and PrMC2 was restricted to the tapetum. PrChS1, PrLTP1, and PrMC2 continued to be expressed in the tapetal tissues of T2-stage cones. PrLTP2, which was expressed only for a brief period in T1-stage cones, showed a different pattern, and appeared to be restricted to tetrads in a cluster of microsporangia in T2-stage cones. This pattern suggested that expression was restricted to a particular part of the cone or progressed acropetally as the tetrads reach a certain stage of development (Fig. 4). Fig. 4. Open in new tabDownload slide Spatial expression of five genes in P. radiata male cones. Me-tIpII-stage- and T2-stage male cones (top and bottom, respectively) were sectioned and probed with sense (control) and antisense transcripts of five cDNAs. The results for PrChS1, PrLTP1, PrMC1, and PrMC2 were similar and are represented above by the results of PrChS1 and PrLTP1. The result for PrLTP2 is illustrated separately above. The bar beneath each image represents 250 μm; rectangles indicate the origin of the enlarged images. Detection of transcript is indicated by the blue/black tetrazolium blue signal (arrow) and in the control sections arrows indicate corresponding tissue. Fig. 4. Open in new tabDownload slide Spatial expression of five genes in P. radiata male cones. Me-tIpII-stage- and T2-stage male cones (top and bottom, respectively) were sectioned and probed with sense (control) and antisense transcripts of five cDNAs. The results for PrChS1, PrLTP1, PrMC1, and PrMC2 were similar and are represented above by the results of PrChS1 and PrLTP1. The result for PrLTP2 is illustrated separately above. The bar beneath each image represents 250 μm; rectangles indicate the origin of the enlarged images. Detection of transcript is indicated by the blue/black tetrazolium blue signal (arrow) and in the control sections arrows indicate corresponding tissue. Most cDNAs Represent Genes of Low or Intermediate Copy Number Southern analysis was performed using the shortest isolate of each of the 12 hybridizing groups of cDNAs (no data were obtained for PrLTP2). Most cDNAs represented genes of low (1–2 bands per digest) or intermediate (3–6 bands) copy number (Fig.5; Table II) except PrLTP1, which showed an average of nine hybridizing bands in each digest. Fig. 5. Open in new tabDownload slide Southern-blot analysis. Blots of genomic DNA (10 μg per lane, digested as indicated) were probed with the cDNA clones indicated. Clones with an average of less than three signals per digest were scored as low (e.g. PrChS1 and PrMC3), three to seven bands were scored as intermediate (e.g. PrMC187), and eight or more bands were scored as high copy (e.g. PrLTP1, data summarized Table II). Fig. 5. Open in new tabDownload slide Southern-blot analysis. Blots of genomic DNA (10 μg per lane, digested as indicated) were probed with the cDNA clones indicated. Clones with an average of less than three signals per digest were scored as low (e.g. PrChS1 and PrMC3), three to seven bands were scored as intermediate (e.g. PrMC187), and eight or more bands were scored as high copy (e.g. PrLTP1, data summarized Table II). Nucleotide Sequence of the cDNAs PrChS1 Belongs to a Tapetal-Specific Subgroup of Chalcone Synthase-Like Proteins The longest clone of this group was sequenced completely in both directions and named PrChS1 because of its similarity to chalcone synthase (ChS) and stilbene synthase (StS) genes. The highest similarity was with five ChS sequences, which, together with PrChS1, form a distinct clade on phylogenetic trees derived from various ChS and StS sequences (Fig. 6). Transcripts of four of the five genes in the PrChS1 clade have been isolated from floral tissues (Shen and Hsu, 1992; Hihara et al., 1996; Turgut et al., 1996; Atanassov et al., 1998), and the fifth is from a genomic sequence (GenBank accession no. u89959). Expression of the rice gene was tapetal specific (Hihara et al., 1996), the Brassica rapa gene was expressed in the tapetum and vasculature of anthers as well as in young microspores (Shen and Hsu, 1992), the Brassica napus gene was isolated on the basis of its differential expression in anthers (Turgut et al., 1996), and expression of the tobacco gene was restricted to the tapetum and developing microspores (Atanassov et al., 1998). Both ChS and StS enzymes are induced in response to various pathogens and stresses (Koes et al., 1989a; Hain et al., 1993;Schubert et al., 1997) and ChS is developmentally regulated in various tissues including anthers (Koes et al., 1989b; van der Meer et al., 1990). Fig. 6. Open in new tabDownload slide Phylogenetic analysis of ChS and StS-like sequences. The figure illustrates the tree generated when ChS and StS amino acid sequences corresponding to bases 8 to 386 of PrChS1 were aligned and subjected to a phylogenetic analysis. Five of the six sequences (indicated by asterisks) in the PrChS1 clade are specific to male reproductive tissues. ChS sequences from plant species corresponding to those represented in the PrChS1 clade are shaded gray. The distribution of StS sequences (boxed) throughout the phylogram suggests that StS evolved from ChS several times independently during evolution (also see Tropf et al., 1994). ChS sequences from plant species corresponding to the StS sequences are underlined. Abbreviations are as follows: Ph, Petunia hybrida; Ah,Arachis hypogaea; Pl, Pueraria lobata;Gm, Glycine max; Ms, Medicago sativa; Ps,Pisum sativum; Ts, Trifolium subterraneum; Psy, Pinus sylvestris; Pst,Pinus strobus; Cs, Camellia sinensis; Vv,Vitis vinifera; Bn, Brassica napus; At, Arabidopsis; Br, Brassica rapa; Ns, Nicotiana sylvestris; Os, Oryza sativa; Pr, P. radiata; Hv, Hordum vulgare; Sc, S. cereale; Zm, Z. mays; Mm, Matthiola incana; Pc, Petroselinum crispum; Am,Antirrhinum majus; Le, Lycopersicon esculentum. Fig. 6. Open in new tabDownload slide Phylogenetic analysis of ChS and StS-like sequences. The figure illustrates the tree generated when ChS and StS amino acid sequences corresponding to bases 8 to 386 of PrChS1 were aligned and subjected to a phylogenetic analysis. Five of the six sequences (indicated by asterisks) in the PrChS1 clade are specific to male reproductive tissues. ChS sequences from plant species corresponding to those represented in the PrChS1 clade are shaded gray. The distribution of StS sequences (boxed) throughout the phylogram suggests that StS evolved from ChS several times independently during evolution (also see Tropf et al., 1994). ChS sequences from plant species corresponding to the StS sequences are underlined. Abbreviations are as follows: Ph, Petunia hybrida; Ah,Arachis hypogaea; Pl, Pueraria lobata;Gm, Glycine max; Ms, Medicago sativa; Ps,Pisum sativum; Ts, Trifolium subterraneum; Psy, Pinus sylvestris; Pst,Pinus strobus; Cs, Camellia sinensis; Vv,Vitis vinifera; Bn, Brassica napus; At, Arabidopsis; Br, Brassica rapa; Ns, Nicotiana sylvestris; Os, Oryza sativa; Pr, P. radiata; Hv, Hordum vulgare; Sc, S. cereale; Zm, Z. mays; Mm, Matthiola incana; Pc, Petroselinum crispum; Am,Antirrhinum majus; Le, Lycopersicon esculentum. Two Groups Encode Nonspecific Lipid-Transfer Proteins (nsLTPs) PrLTP1 has homology with various plant nsLTPs, but is most similar to two B. napus anther sequences (34% amino acid identity). The first, bif38 (L31938), has not been characterized, and the second, BNE2 (Foster et al., 1992) (X60318), is expressed in the tapetum and microspores from meiosis to the first round of mitosis in the microspore. The deduced protein of PrLTP1 has a hydrophobic N-terminal secretory sequence with a predicted mature protein of 95 amino acids. PrLTP2 also encodes a nsLTP showing 40% nucleotide and 25% amino acid identity with PrLTP1. Sequence alignments (Fig.7) indicate that, although the PrLTP2 reading frame is missing eight to 16 amino acids at the N terminus, it encodes a preprotein with a hydrophobic N terminus and a secreted protein of 99 amino acids. The most similar homolog is the rice NLT4 gene (accession no. q42976), which has 33% amino acid identity with PrLTP2. Expression of NLT4 has not been described. Fig. 7. Open in new tabDownload slide Comparison of deduced LTP and A9 anther-expressed proteins. Deduced sequences of nsLTPs (PrLTP1, PrLTP2, E2, OsLTP, and bLTP) and A9 homologs (PrMC1, PrMC2, Men-8, Lhm7, and A9) were aligned using the GCG program PileUp. The eight conserved Cys residues are shown beneath the alignment. The four helical domains of the barley LTP are labeled A, B, C, and D as indicated. The cleavage site for removal of the secretory sequence predicted by the GCG program SPSCAN is indicated by asterisk. The eight conserved Cys residues are shaded. Accession numbers are Men-8, y08780; LHm7, x80719; A9, q05772; barley LTP, p07597; OsLTP, u29176; and E2, x60318. Fig. 7. Open in new tabDownload slide Comparison of deduced LTP and A9 anther-expressed proteins. Deduced sequences of nsLTPs (PrLTP1, PrLTP2, E2, OsLTP, and bLTP) and A9 homologs (PrMC1, PrMC2, Men-8, Lhm7, and A9) were aligned using the GCG program PileUp. The eight conserved Cys residues are shown beneath the alignment. The four helical domains of the barley LTP are labeled A, B, C, and D as indicated. The cleavage site for removal of the secretory sequence predicted by the GCG program SPSCAN is indicated by asterisk. The eight conserved Cys residues are shaded. Accession numbers are Men-8, y08780; LHm7, x80719; A9, q05772; barley LTP, p07597; OsLTP, u29176; and E2, x60318. Two Groups Encode A9 Homologs PrMC1 and PrMC2 are 42% identical and encode proteins with 29% amino acid identity. Their reading frames are both predicted to have secretory sequences that are cleaved to give secreted 6.7-kD proteins. Both deduced proteins have a nsLTP-like Cys motif (Fig. 7). However, compared with the nsLTPs, the length of some of the intervening stretches is reduced. PrMC1 and PrMC2 share best similarity with several anther-expressed cDNAs of which the most well characterized is A9 from B. napus (Scott et al., 1991a) and Arabidopsis (Paul et al., 1992). Within this anther-expressed group, the nearest homolog to PrMC2 is the Silene latifolia Men-8 mRNA that was isolated from anthers and is maximally expressed just prior to tapetum degradation; it was not detected in sepals, petals, filaments leaf, or root tissue (Scutt et al., 1997). PrMC1 is most similar to the tapetum-expressed M7 gene, which was isolated from aLilium henryi meiocyte library and is expressed in the tapetum from early prophase until microspore mitosis. It was not expressed in gynoecia, leaf, or root tissue (Crossley et al., 1995). PrThL1 Encodes a Thaumatin Based on sequence alignments with various plant thaumatins, PrThL1 encodes a full-length protein with a hydrophobic N-terminal signal sequence. The secreted protein is a member of the 24- to 25-kD family of thaumatin proteins with 16 conserved Cys residues, rather than the deleted 17-kD version with 10 conserved Cys residues found in some monocotyledons (Hu and Reddy, 1997). The closest homolog to PrThL1 is Tomf216, a floral-specific transcript from tomato with 45% amino acid identity. Tomf216 is expressed in immature inflorescences, in flowers prior to meiosis, in stamens from tetrad dissolution through to anthesis, and in petals at anthesis. The closest non-floral homolog is an Arabidopsis sequence, ATLF (40% amino acid identity; Arro et al., 1997), which is induced by parasites and by compounds inducing parasite resistance (Uknes et al., 1992). PrMC3 Is Similar to a Hypersensitive Response Protein PrMC3 is most similar to a tulip arylacylamidase (42% amino acid identity; GenBank accession no. e03271), a P. radiataexpressed sequence tag from embryo tissue culture (accession no.AA220894) and the tobacco protein hsr203J (Pontier et al., 1994) (36% amino acid identity). The tulip and P. radiata homologs are not described in the literature. However, hsr203J is rapidly and specifically expressed in the hypersensitive response (HR) to various pathogens (Pontier et al., 1994). Other homologs in the database include a peptide encoded by an expressed sequence tag from elongating root hairs and root tips of Medicago truncatula (amino acid identity 40%; GenBank accession no. AA660803) and the peptide encoded by an unknown Mycobacterium tuberculosis gene (26% amino acid identity; GenBank accession no. Z80108). All of these sequences, including PrMC3, include the Ser hydrolase motif GXSXG (Fig.8). Fig. 8. Open in new tabDownload slide Alignment of PrMC3 with four homologs. The figure shows an alignment of deduced amino acid sequences for four homologs of PrMC3. The Ser hydrolase GXSXG motif is enclosed in a box, and bases identical in three or more sequences are shaded. Accession numbers for the sequences are as follows: Medicago expressed sequence tag, aa660803; tobacco hsr203J, s42807; tuliparylacylamidase, e03271; andMycobacterium seq., z80108. Fig. 8. Open in new tabDownload slide Alignment of PrMC3 with four homologs. The figure shows an alignment of deduced amino acid sequences for four homologs of PrMC3. The Ser hydrolase GXSXG motif is enclosed in a box, and bases identical in three or more sequences are shaded. Accession numbers for the sequences are as follows: Medicago expressed sequence tag, aa660803; tobacco hsr203J, s42807; tuliparylacylamidase, e03271; andMycobacterium seq., z80108. DISCUSSION A differential screening strategy comparing gene expression in P. radiata male cones with expression in needles, roots, and vegetative shoots resulted in the isolation of 13 cDNAs, all of which were isolated from a tetrad-stage cone library. No differential cDNAs were identified among 16,000 clones from the early meiocyte library, indicating that there were few transcripts that were both abundant and differential in the Me-i-stage cones. All of the cDNAs were expressed within the period in which the tapetum is known to undergo a burst in RNA synthetic activity, beginning with meiosis and ending before the first mitotic division in the microspore (Williams and Heslop-Harrison, 1979; Pacini et al., 1985). The temporal expression of 10 of the 13 genes (from meiosis until after the tetrads have dissolved) is consistent with their being expressed in the tapetum. In situ hybridization confirmed that transcripts of four genes (PrLTP1, PrMC1, PrChS1, and PrMC2) are expressed in the tapetum of Me-tIpII- and T2-stage cones. However, three cDNAs, PrMC6, PrLTP2, and PrMC104, were expressed only for short periods within this developmental window, suggesting that their expression is regulated differently and may not be part of the burst in tapetum expression. PrLTP2 transcript, with the shortest duration of expression (Figs. 2and 3), was restricted to the tetrads (Fig. 4). Of the 13 groups of cross-hybridizing cDNAs isolated in this study, six represented novel genes with no homologs in the databases. The remaining seven showed homology to angiosperm sequences, six of which (corresponding to PrChS1, PrLTP1, PrLTP2, PrMC1, PrMC2, and PrThL1) are specifically expressed in anthers or flowers (Table II). The pine sequences generally exhibit similar temporal and spatial expression patterns to their angiosperm homologs (for reviews, see Scott et al., 1991b; Goldberg et al., 1993). Considering the evolutionary distance between angiosperms and gymnosperms and the morphological differences between flowers and cones, the conservation of gene expression is striking and supports previous hypotheses that angiosperm and gymnosperm reproductive structures share common ancestry (discussed in Hickey and Taylor, 1996). Microspores from both angiosperms and gymnosperms develop in a microsporangium surrounded by a tapetum, and the genes expressed in these organs have probably been retained through evolution because they play important roles in pollen development. Selective pressure appears to have maintained this intricate microspore-tapetum relationship, as disruptions to it are known to be a frequent cause of male sterility (Kaul, 1988). The temporal and spatial expression patterns of the genes expressed in cones is also consistent with the idea that the conifer homologs (Tandre et al., 1995; Mellerowicz et al., 1998; Mouradov et al., 1998a, 1998b) of angiosperm floral meristem and floral organ genes play similar roles in regulating gymnosperm cone development. A P. radiata homolog of ChS that grouped on a phylogenetic tree with a subset of five previously isolated ChS clones was isolated. The PrChS1 clade includes members from pine, rice, tobacco, andBrassica sp., so evolution of this clade must have occurred before the divergence of gymnosperms and angiosperms. In contrast, stilbene synthase genes evolved from ChS genes on several occasions in the course of evolution within both the angiosperm and gymnosperm divisions of the plant kingdom (Fig. 6; Tropf et al., 1994). Comparison of alignments of this clade with secondary structure predictions made by Tropf et al. (1994) supports the idea that genes in this clade may possess stilbene synthase activity. The role of the PrChS1 clade in male reproductive tissues remains to be determined. ChS enzymes have been demonstrated to be essential for pollen development. Disruptions to ChS activity in the anthers of petunia (Taylor and Jorgensen, 1992; van der Meer et al., 1992), maize (Coe et al., 1981), and tobacco (Fischer et al., 1997) resulted in the production of sterile pollen that could be rescued by the application of exogenous flavanoids (Taylor, 1995; Fischer et al., 1997). We identified two homologs of nsLTPs and two A9 homologs that were expressed in male cones. nsLTPs have been isolated from various species based on their expression in anthers and in response to infection and stress (Nacken et al., 1991; Foster et al., 1992; Aguirre and Smith, 1993; Kader, 1996, 1997). Plant nsLTPs bind lipids with low affinity and are able to transfer them between membranes (for review, see Kader, 1996, 1997). Some inhibit proteases and limit the growth of microbial pathogens (Cammue et al., 1995; Kader, 1996, 1997). The A9 proteins might have a function related to the nsLTPs, based on their eight conserved Cys residues (see Fig. 7) and their overall structural similarity (Paul et al., 1992; Crossley et al., 1995). Alignments indicate that helix C of the nsLTPs is probably modified or missing in the A9 proteins; see Figure 7 and Paul et al. (1992). This helix is important in accommodating binding to large lipids (Shin et al., 1995; Lerche et al., 1997). We suggest that deletion of this helix might confer different substrate specificity on the smaller A9 proteins. The high copy number of some of these nsLTP genes in the genome suggests that they may be in the process of acquiring new functions (Cammue et al., 1995; Kader, 1996). In vivo roles suggested for nsLTPs and A9 homologs in anthers include: shuttling lipids from the tapetum to the meiocytes and developing microspores (Foster et al., 1992;Kader, 1996, 1997), acting as a sulfur store (Aguirre and Smith, 1993), protecting the microspores from hydrolytic enzymes released during degradation of the tapetum (Crossley et al., 1995), and protecting the pollen from pathogens either before or after dehiscence (Paul et al., 1992; Crossley et al., 1995). To date, no anther-expressed nsLTP or A9 protein has been tested for antimicrobial, proteinase inhibition, or lipid transfer activity. The expression of PrLTP2 in the tetrads during a brief developmental window suggests that it is unlikely to play a role in protection from pathogens, but rather plays a specialized role associated with early microspore development. The screen for male-cone-specific genes also identified a P. radiata homolog of thaumatin. Many thaumatin proteins are induced by the plant in response to infection by pathogens and exposure to environmental stresses (Singh et al., 1989; Vigers et al., 1992;Stintzi et al., 1993; Abad et al., 1995; Griffith et al., 1997; Hu and Reddy, 1997). Some thaumatins have anti-microbial activity and are thought to permeabilize fungal hyphae by forming a pore or channel, allowing the release of the cytoplasmic contents (Abad et al., 1996;Cheong et al., 1997). Some thaumatin genes are expressed in floral tissues (Richard et al., 1992; Chen et al., 1996), but none of this group has been tested for antimicrobial activity. A possible role for PrThL1 in P. radiata may be to permeabilize the plasma membranes of the tapetum, facilitating transport of compounds to the developing microspores. PrMC3 is a member of a family of proteins that all contain a Ser hydrolase motif (GxSxG) and have similarity to lipases and esterases of prokaryotic origin. PrMC3 is the first member of the family differentially expressed in male reproductive structures of a plant. The timing of PrMC3 expression, which occurs right through the burst of tapetal layer activity, would be consistent with a role hydrolyzing stored lipids for transfer from the tapetum to the microspores by nsLTPs and A9 homologs. However, the spatial expression of PrMC3 has not been determined, and the fact that the protein encoded by the tobacco homolog hsr203J is unable to hydrolyze lipids (Baudouin et al., 1997) suggests that this scenario is unlikely. Recombinant hsr203J protein degrades p-nitrophenylbutyrate, a general substrate for carboxylesterases, which suggests it is an esterase (Baudouin et al., 1997). The tobacco gene hsr203J is induced specifically and early in the hypersensitive response, after challenge by pathogens, and probably plays a role in regulating or limiting programmed cell death (Pontier et al., 1994, 1998). We suggest that PrMC3 might play a similar role in regulating the developmentally regulated programmed cell death of the tapetal layer during male cone development. The association between genes expressed in P. radiata male cones and angiosperm genes involved in the pathogenesis response was striking. If the A9 proteins have activities related to the nsLTPs, then all seven of the genes that were identified in this study could be considered pathogenesis related. Vigers et al. (1992) proposed that tissues with vital reproductive capacities but limited resources for counterattack, such as seeds and tubers and, by inference, pollen, store thaumatin as protection from future microbial infections. Similar statements have been made in relation to the nsLTP and A9 proteins (Paul et al., 1992; Turgut et al., 1994; Crossley et al., 1995), and the same could be said of PrChS1. However, we consider it unlikely that the expression of these genes in male cones occurs solely in response to, or as protection against, pathogens or stress. First, the timing and pattern of expression of some of the genes (e.g. PrLTP2) is highly specific. Second, most of these genes are expressed in the tapetal tissue, which is protected from the external environment prior to anthesis and seems unlikely to be subject to large numbers of pathogen attacks. Third, the sheer number of genes involved in defense seems very high, given that pollen development involves an intense burst of metabolic activity over the period in which the genes are expressed (Pacini et al., 1985). Fourth, none of the nsLTP, A9, or thaumatin proteins expressed in male reproductive structures has been shown to have antimicrobial activity. Finally, most of the transcripts identified in this study have plausible functions in pollen development, such as shuttling lipids for the nsLTPs. In some cases, these functions are common to pathogen or stress responses. For example, both reproductive development and the plant response to stress or pathogens can involve the programmed reorganization and degradation of tissue (Greenberg, 1998). In male reproductive tissues the genes involved in tapetum degradation and mobilization are expected to be developmentally regulated, whereas in the pathogen response, the genes are induced as a result of an external stimuli. In summary, we have shown that homologous genes are expressed differentially in male reproductive development in the angiosperm and gymnosperm divisions of the plant kingdom. The genes isolated here will prove useful for the isolation of tapetal-specific promoters and the genetic manipulation of male sterility in conifers. ACKNOWLEDGMENTS We thank Dr. Marc Jacobs for 26S rRNA PCR primers, Keith Richards for help with cDNA library construction, and Dr. Dale Smith for advice on tissue collection and storage. 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Persans, Michael W.; Yan, Xiange; Patnoe, Jean-Marc M.L.; Krämer, Ute; Salt, David E.

doi: 10.1104/pp.121.4.1117pmid: 10594099

Abstract To understand the role of free histidine (His) in Ni hyperaccumulation in Thlaspi goesingense, we investigated the regulation of His biosynthesis at both the molecular and biochemical levels. Three T. goesingense cDNAs encoding the following His biosynthetic enzymes, ATP phosphoribosyltransferase (THG1, GenBank accession no. AF003347), imidazoleglycerol phosphate dehydratase (THB1, GenBank accession no. AF023140), and histidinol dehydrogenase (THD1, GenBank accession no. AF023141) were isolated by functional complementation of Escherichia coli His auxotrophs. Northern analysis of THG1,THD1, and THB1 gene expression revealed that each gene is expressed in both roots and shoots, but at the concentrations and dosage times of Ni treatment used in this study, these genes failed to show any regulation by Ni. We were also unable to observe any increases in the concentration of free His in root, shoot, or xylem sap of T. goesingense in response to Ni exposure. X-ray absorption spectroscopy of root and shoot tissue fromT. goesingense and the non-accumulator speciesThlaspi arvense revealed no major differences in the coordination of Ni by His in these tissues. We therefore conclude that the Ni hyperaccumulation phenotype in T. goesingense is not determined by the overproduction of His in response to Ni. There are certain plants, such as Thlaspi goesingense, that have the ability to accumulate concentrations of Ni in their shoots far exceeding those observed in the soil, without suffering the detrimental effects of Ni toxicity (Reeves and Brooks, 1983). Brooks et al. (1977) first used the term “hyperaccumulator” to describe plants that contain >1,000 μg g−1 (0.1%) Ni in their dried leaves, a concentration at least an order of magnitude higher than Ni levels in nonaccumulator species. Progress is being made in understanding the mechanisms involved in metal hyperaccumulation, but the molecular basis of this intriguing phenomenon remains elusive (Salt and Krämer, 1999). Recent work on the mechanism of Ni hyperaccumulation in T. goesingense has established that Ni tolerance is a primary determinant of the hyperaccumulation phenotype in hydroponically cultured plants (Krämer et al., 1997). An important component of this Ni tolerance mechanism appears to be based on the efficient intracellular compartmentalization of the Ni into the vacuole (U. Krämer and D.E. Salt, unpublished data). Recently, Krämer et al. (1996) observed a 36-fold increase in the concentration of free His in the xylem exudate of the Ni hyperaccumulator Alyssum lesbiacum after exposure to Ni. However, no significant change was observed in the nonaccumulatorAlyssum montanum. The authors also observed a significant linear correlation in the xylem exudate concentrations of free His and Ni in several Ni hyperaccumulators in the genus Alyssum(Krämer et al., 1996). Because His is an effective chelator of Ni at cytoplasmic pH (Dawson et al., 1986), the authors suggested that His may be involved in chelating Ni during the transport and or storage of Ni in the Alyssum Ni hyperaccumulators. This was supported by the use of x-ray absorption spectroscopy, which identified putative Ni-His complexes in the xylem sap, root, and shoot tissue of A. lesbiacum (Krämer et al., 1996). However, the mechanism by which Ni hyperaccumulation is achieved through the action of His has not been established. Both Alyssum and Thlaspi Ni hyperaccumulator species are members of the Brassicaceae family, suggesting that free His may also play a role in the mechanism of Ni hyperaccumulation inT. goesingense. This is supported by the recent identification of putative Zn-His complexes in the roots of the closely related Zn hyperaccumulator Thlaspi caerulescens (Salt at al., 1999b). Therefore, to determine if free His is involved in Ni hyperaccumulation in T. goesingense, we investigated the regulation of His biosynthesis at both the molecular and biochemical levels in T. goesingense. To determine if Ni regulates the expression of genes involved in His biosynthesis in T. goesingense, we cloned genes encoding ATP phosphoribosyltransferase (THG1), imidazoleglycerol phosphate dehydratase (THB1), and histidinol dehydrogenase (THD1), enzymes that catalyze potentially rate-limiting steps in His biosynthesis. Previously, several authors have published the sequences of various His biosynthetic genes, including HisD (encoding histidinol dehydrogenase [HDH]) from Arabidopsis (Bevan et al., 1998) andBrassica oleracea (Nagai et al., 1991);HisB (encoding imidazoleglycerol phosphate dehydratase [IGPD]) from Pisum sativum (Kim and Theologis, 1996),Triticum aestivum, and Arabidopsis (Tada et al., 1994); HisC (encoding l-histidinol phosphate aminotransferase [HPA]) from Nicotiana tabacum(El Malki et al., 1998); and HisIE (encoding phosphoribosyl-ATP pyrophosphohydrolase/phosphoribosyl-AMP cyclohydrolase [PR-ATP/PR-AMP]) from Arabidopsis (Fujimori and Ohta, 1998). However, to our knowledge, this is the first study to isolate ATP phosphoribosyltransferase (ATP-PRT), the enzyme that catalyzes the first committed step in His biosynthesis, from plants. To examine the effects of Ni exposure on His biosynthesis, we also measured the concentration of free His in the roots, shoots, and xylem sap in both the Ni hyperaccumulator T. goesingense and the nonaccumulator Thlaspi arvense. Additionally, we quantified putative Ni-His complexes in both T. goesingense andT. arvense using x-ray absorption spectroscopy. MATERIALS AND METHODS Plant Growth Conditions and Ni Treatment For cDNA library construction, Thlaspi goesingenseseeds were germinated and grown hydroponically according to the method of Krämer et al. (1997). The plants were exposed to 25 μmNi(NO3)2 for 5 weeks with two exchanges of hydroponic solution per week. After 5 weeks, whole plants were harvested, immediately frozen in liquid nitrogen, and stored at −80°C. Plants for RNA and genomic DNA studies were grown as follows. Seeds were germinated on filter papers moistened with double-distilled water for 7 d and subsequently transferred to hydroponic culture solution according to the method of Krämer et al. (1997). Plants were maintained on a 12-h d period and day/night temperature of 25°C/20°C, with weekly changes of hydroponic culture solution. Both fluorescent and incandescent lights were used to provide 170 μmol m−2 s−1 of photosynthetic photon flux (PPF) at the level of the plants. After 6 weeks of growth from the date of transfer into the hydroponic culture solution, plants were exposed to 50 μm Ni for 24 or 48 h by the addition of Ni(NO3)2 to the culture solution. Both roots and shoots were harvested separately, immediately frozen in liquid nitrogen, and stored at −80°C. For the efficient isolation of RNA from shoot material, it was necessary to deplete the plants of starch. This was achieved by harvesting treated plants just before the onset of the light period. His Levels in T. goesingense For analysis of the free His concentration, root and shoot material was extracted as follows. Approximately 2 g of tissue was frozen in liquid nitrogen and ground to a fine powder in a pestle and mortar. To the frozen powder 6 mL of 3% (w/v) sulfosalicylic acid was added and the slurry ground until the plant tissue had completely thawed. The slurry was then centrifuged at 1,550g for 15 min at room temperature. The supernatant was filtered through a 0.45-μm filter to remove suspended particulate material. Phenylthiocarbamyl derivatization was carried out following the method of Fierabracci et al. (1991). The non-biological amino acid Met sulfoxide was added to 10 μL of tissue extract or xylem exudate to a final concentration of 40 μm and used during analysis as an internal standard. Samples were vacuum-dried (model SVC200, Savant Instruments, Holbrook, NY), redissolved in 20 μL of ethanol:water:TEA (2:2:1, v/v), and vacuum-dried again. To the dried samples 20 μL of ethanol:triethylamine (TEA):water:phenyl isothiocyanate (7:1:1:1, v/v) was added, and the samples incubated at room temperature for 20 min. Samples were vacuum-dried to remove excess reagent and reconstituted in 250 μL of phosphate buffer (pH 7.4). Separation of the phenylthiocarbamyl amino acid derivatives was performed on a Nucleosil C18 5-μm HPLC column (Sigma, St Louis) (250 × 4.6 mm) at 38°C. The elution solvent consisted of 0.14m sodium acetate in water plus 0.5 mL/L TEA, titrated to pH 6.4 with glacial acetic acid, with the addition of 60 mL/L acetonitrile (solvent A); and 60% (v/v) acetonitrile in water (solvent B). Phenylthiocarbamyl amino acid derivatives were detected at 254 nm using an UV spectrophotometer (Spectroflow 783, Kratos Analytical, Ramsey, NJ). Determination of Ni Speciation in Thlaspi Plant samples were shipped to the Stanford Synchrotron Radiation Laboratory (Stanford University, Stanford, CA) on dry ice. To minimize breakdown and mixing of cellular components within the plant material, care was taken to keep the tissue frozen at all times prior to measurement. To this end, frozen plant tissues were carefully ground under liquid nitrogen and compacted into liquid-nitrogen-cooled 1-mm pathlength lucite sample holders with mylar windows. Aqueous model compounds were diluted by 30% to 50% (v/v) with glycerol (to avoid ice crystal formation) before being pipetted into holders and rapidly frozen in liquid nitrogen. During data collection, samples were held at approximately 15 K using a flowing liquid helium cryostat. X-ray absorption spectroscopy was carried out on beamline 7–3 of the Stanford Synchrotron Radiation Laboratory using a Si(220) double crystal monochromator, 1-mm upstream vertical aperture, and no focusing optics. Incident intensity was measured using a nitrogen-filled ion chamber, and the absorption spectrum was collected in fluorescence using a 13-element germanium detector (Cramer et al., 1988) by monitoring the Ni Kα afluorescence line at 7,472 eV. Spectra were energy calibrated with respect to a spectrum of Ni foil, and collected simultaneously with the spectrum of each sample, the first energy inflection of which was assumed to be 8,333 eV. X-ray absorption spectroscopy data reduction was carried out using the EXAFSPAK suite of programs (George, 1998) according to standard methods (Koningsberger and Prins, 1988). Quantitative edge-fitting analysis was performed using the program DATFIT (George et al., 1991). Here, the near-edge spectrum of the plant material is fit using a least-squares algorithm to a linear combination of edge spectra from a library of Ni model compounds. The fractional contribution of each model spectrum to the fit is then directly proportional to the percentage of Ni present in that form in the plant material. By analyzing the total Ni content of the tissue samples, the percentage fractional contribution can then be simply converted into the absolute amount of complex present. Total RNA Isolation and λTriplEx cDNA Library Construction Total RNA was isolated according to the method of Murphy and Taiz (1995) as follows. Frozen Ni-treated plant material (3–4 g) was ground in liquid nitrogen to a fine powder using a pre-chilled mortar and pestle. The ground tissue was added to 3.5 mL of extraction buffer (1% [w/v] triisopropylnapthylenesulfonic acid, 6% [w/v]p-aminosalicylic acid, 1% [w/v] NaCl, 3% [w/v] polyvinylpyrrolidine, 5% [v/v] β-mercaptoethanol, 4m guanidine thiocyanate, and 25 mm sodium citrate), shaken gently to mix, and incubated for 5 min at room temperature after the tissue had thawed. To the samples, 3.5 mL of dilution buffer (6× SSC, 10 mm Tris, 1 mm EDTA, and 0.25% [w/v] SDS) was added and vortexed for 1 min, then incubated at 65°C for 5 min. Phenol (6 mL) was added and the mixture was vortexed for 1 min and incubated at room temperature for 5 min. Chloroform (6 mL) was added and the mixture was vortexed for 1 min. Samples were then centrifuged at 5,000g for 10 min at 4°C to separate the phases. The upper aqueous phase was removed, 7 mL of phenol and 10 mL of chloroform were added, and the mixture was vortexed for 1 min, incubated at room temperature for 5 min, and vortexed again for 1 min. The phases were then separated by centrifugation at 3,000g for 10 min at 4°C. The upper aqueous layer was removed and 7 mL of isopropanol was added and the sample incubated for 4 h at −20°C to precipitate the RNA. RNA was collected by centrifugation at 13,500g for 15 min, and the pellet was resuspended in 400 μL of diethyl pyrocarbonate (DEPC)-treated water. The resuspended pellet was then re-extracted in 1 mL of 5 m LiCl by vortexing for 1 min, and RNA was reprecipitated by incubation at −20°C for 45 min. The RNA was pelleted by centrifugation at 27,000g for 25 min at 4°C. The RNA pellet was washed by the addition of 1 mL of 70% (v/v) ethanol and collected by centrifugation at 27,000g for 25 min at 4°C. The final purified RNA was resuspended in 100 μL of DEPC-treated water and stored at −80°C. The total T. goesingense RNA was sent to CLONTECH Laboratories (Palo Alto, CA), where mRNA isolation, cDNA synthesis, and construction of recombinant λTriplex were performed. The cDNA synthesis resulted in 1.6 × 106 independent clones with an average insert size of 2.0 kb (range 0.7–4.0 kb). AllT. goesingense cDNAs were cloned into the EcoRI and XhoI sites within the λTriplEx multiple cloning site and converted to the phagmid pTriplEx using a cre-recombinaseEscherichia coli expressing strain (BM25.8). The choice of the λTriplEx vector was made because of several of its unique features. Each vector has separate initiation codons in two differing frames, both followed by a single poly-dT tract (slip site) that allows either the RNA polymerase or the ribosomes to skip nucleotides, thereby allowing reading in all three frames. Furthermore, the vector incorporates the 5′-untranslated region (UTR) of theompA gene from E. coli to increase mRNA stability. These features allow every recombinant vector to express some protein, regardless of the initial frame of the insert. Cell Transformation and Functional Complementation in E. coli T. goesingense cDNAs encoding functional homologs of the E. coli His biosynthetic enzymes ATP-PRT, IGPD, and HDH were isolated by screening for cDNAs that could complement the His requirement of various His auxotrophic E. coli mutants.E. coli lacking a functional copy of the gene encoding ATP-PRT (strain KL738), IGPD (strain SB3930), or HDH (strain UTH4758) were transformed with the T. goesingense pTriplEx cDNA library by electroporation (model EC100 Electroporator, EC Apparatus, St. Petersburg, FL), per the manufacturer's protocol. Transformed cells were selected on Luria-Bertani medium and ampicillin (100 μg mL−1) agar plates, resulting in 106 to 107transformants/μg pTriplEx plasmid DNA. The resulting colonies were replica plated onto M-9 minimal media supplemented with 19 amino acids (excluding His), each at a concentration of 25 μg mL−1, 0.1 mmisopropylthiogalactoside, and ampicillin (100 μg mL−1). Colonies that were able to grow were retested for growth on M-9 minimal medium lacking His. Plasmid DNA was isolated from colonies that grew after replating, and the ability of these plasmids to complement the His requirement of the E. coli auxotrophic mutants was confirmed by retransformation of the appropriate His auxotrophic E. coli mutant. Sequencing and Analysis Double-stranded DNA from all three genes was completely sequenced using pTriplEx plasmid primers and primers based on previous sequence data using a DNA sequencer (model 373, Applied Biosystems, Foster City, CA) and a dye terminator cycle sequencing ready reaction kit (catalog no. P/N 402078, ABI PRISM, Perkin-Elmer, Foster City, CA). Predicted translations of the THG1, THD1, andTHB1 genes were generated and a search was performed to identify any homologous sequences in GenBank. The THG1predicted amino acid sequence was aligned with existing ATP-PRT sequences acquired from GenBank using the CLUSTAL_W algorithm (Thompson et al., 1994). The predicted amino acid sequence of THB1 andTHD1 was aligned with existing plant sequences using ALIGN (Myers and Miller, 1989). A phylogenetic construction of the resulting aligned sequences for THG1 (ATP-PRT) was also performed using the PHYLIP algorithm (Phylogeny Inference Package, version 3.57c, Department of Genetics, University of Washington, Seattle) (Felsenstein, 1993). Both the alignment and phylogenetic analysis was performed at the University of Illinois Biology Workbench (http://biology.ncsa.uiuc.edu /). Analysis for the presence and type of protein targeting sequence was performed using the PSORT algorithm (Nakai and Kanehisa, 1992) found athttp://psort.nibb.ac.jp:8800 /. The targeting sequence cleavage site was predicted using the SignalP (version 1.1) algorithm (Nielsen et al., 1997) found athttp://www.cbs.dtu.dk/services/SignalP/. Nucleotide Probe Preparation One-hour restriction digests of pTriplEx-(THG1) and pTriplEx-(THD1) with XhoI, pTriplEx-(THB1) with EcoRI and XhoI, and an Arabidopsis actin gene (GenBank accession no. U37281) withBamHI and EcoRI were performed at 37°C. The resulting fragments were run on a 1.5% (w/v) agarose gel and the appropriate size fragment was excised for the gel and recovered by electroelution. For THG1 and THD1, the resulting fragments were approximately 400 and 500 bp in size, respectively, and both contained the 3′-UTR of the gene. For THB1, the entire cDNA was recovered. The resulting actin probe was approximately 950 bp and did not contain either 5′- or 3′-UTRs, only the protein coding sequence. In advance of blot hybridization, 75 to 150 ng of DNA (approximately 5 μL) was denatured at 100°C for 10 min in 35 μL of double-distilled water, and the sample was snap-cooled on ice for 30 s. Then, 2 μL of bovine serum albumin (BSA) (10 mg/mL), 10 μL of 5× OLB buffer (250 mm Tris-HCl [pH 8.0], 25 mm MgCl2, 0.35% [v/v] β-mercaptoethanol, 100 μm each dGTP, dCTP, and dTTP, 1 m HEPES [pH 6.0], and 0.54 μg/μL pdN6 random hexamers), 2 to 3 μL of [α-32P]dATP (10 μCi/μL), and 5 units of DNA polymerase I Klenow fragment was added to the denatured DNA. The labeling reaction was incubated for at least 5 h at room temperature. The labeled DNA probe was boiled for 10 min, snap-cooled on ice for 30 s, centrifuged for 10 s at 14,000g, and added directly to the hybridization buffer. Southern Analysis To obtain genomic DNA, 0.5 to 1.0 g of frozen T. goesingense shoot tissue was placed in a 15-mL centrifuge tube (Falcon 2059, Becton Dickinson, Lincoln Park, NJ), frozen in liquid nitrogen, and ground to a fine powder with a glass rod. Urea extraction buffer (700 μL; 7 m urea, 312 mm NaCl, 20 mm EDTA, 1% [w/v] N-lauroyl sarkosine, and 50 mm Tris-HCl [pH 8.0) was added, and the sample was thawed to room temperature with frequent gentle mixing. Phenol/chloroform (1:1, 500 μL) was added and the sample was incubated for 15 min at 37°C in a rotary shaker. The sample was then transferred to a 1.5-mL microfuge tube and the aqueous phase was separated by centrifugation at 14,000g for 10 min. The upper aqueous phase (approximately 500 μL) was removed and placed in a fresh 1.5-mL microfuge tube. To the aqueous phase 50 μL of 4.4m ammonium acetate and 700 μL of isopropanol were added, the sample was mixed well, and the genomic DNA pelleted by centrifugation at 14,000g for 1 min. The genomic DNA pellet was resuspended in 500 μL of sterile water and reprecipitated as above. The final DNA pellet was washed once with 70% (v/v) ethanol, spun at 14,000g for 3 min, air-dried for 10 to 15 min inverted on a paper towel, and resuspended in 50 to 100 μL of sterile water. For genomic Southern analysis, genomic DNA was digested with 60 units of each of the following restriction enzymes: BamHI,EcoRI, HindIII, XbaI, XhoI, or PstI, for 6 to 8 h at 37°C. The resulting fragments were electrophoresed in a 0.8% (w/v) agarose Tris-boric acid-EDTA gel, and capillary blotted onto nylon membranes (catalog no. 80–6221–93, Pharmacia Biotech, Piscataway, NJ) overnight using 10× SSC. The genomic DNA was UV-crosslinked (model FB–UVXL–1000, Fisher Scientific, Loughborough, Leicestershire, UK) to the membrane. The blots were pre-hybridized at 65°C for at least 2 h in 10 mL of a pre-hybridization solution containing 50 mm Tris-HCl (pH 8.0), 10 mmEDTA (pH 8.0), 5× SSC, 5× Denhardt's solution, 0.2% (w/v) SDS, 7.5% (w/v) dextran sulfate, and 100 μg mL−1 sheared salmon-sperm DNA. The blots were then probed with a denatured 32P-labeled probe added directly to the hybridization solution and the blots incubated at 65°C for 12 to 16 h. After hybridization the blots were rinsed with 50 mL of 2× SSC and 0.1% (w/v) SDS for 5 min at room temperature and then washed two to four times for 15 min at 65°C with 50 mL of 0.1× SSC and 0.1% (w/v) SDS. After washing, the membranes were placed on x-ray film and exposed at −80°C for 10 to 16 h. Northern Analysis Total RNA was isolated according to the method of Puissant and Houdebine (1990). Approximately 5 to 10 g of 6-week-old T. goesingense shoots or roots were frozen in liquid nitrogen and ground to a fine powder in a chilled mortar and pestle. The ground tissue was placed in four to six centrifuge tubes each containing 5 mL of GuISCN extraction buffer (4 m guanidinium isothycyanate, 25 mm sodium citrate [pH 7.0], 0.5% [w/v] N-lauroyl sarkosine, and 0.1m β-mercaptoethanol), and the tubes were mixed by inversion. The samples were mixed with 0.1 volume of 2 m sodium acetate (pH 4.0) and 5 mL of phenol:chloroform (5:1) was added. The samples were mixed and centrifuged at 5,000g for 15 min at 4°C. The aqueous phase (approximately 7 mL) was removed and placed in a fresh 15-mL centrifuge tube, and RNA was precipitated by adding an equal volume of isopropanol at 4°C. The RNA was collected by centrifugation at 4,000g for 10 min at 4°C, and each pellet was resuspended in 2 mL of 4 m LiCl, mixed well, and re-centrifuged at 4,000g for 10 min at 4°C. Each pellet was resuspended in 2 mL of Tris-EDTA buffer containing 0.5% (w/v) SDS, and an equal volume of chloroform was added. After mixing and centrifugation at 4,000g for 10 min at 4°C, the upper aqueous phase was removed and the total RNA precipitated after adding 0.1 volume of 2 m sodium acetate (pH 5.0) and an equal volume of isopropanol. The total RNA was collected by centrifugation at 4,000g for 15 min at 4°C and washed with 70% (v/v) ethanol and 100% ethanol. The samples were air-dried for 15 min, resuspended in 300 μL of sterile, DEPC-treated water, and stored at −80°C. For northern analysis, 30 μg of total RNA was electrophoresed on 1.2% (w/v) agarose-formaldehyde gels and capillary blotted overnight onto nylon membranes (catalog no. 80–6221–93, Pharmacia Biotech) using 10× SSC. The RNA was UV-crosslinked to the membrane and the blot prehybridized in 10 mL of a pre-hybridization solution containing 200 mm Na2PO4 (pH 7.2), 5% (w/v) SDS, 1 mm EDTA, 10 mg/mL BSA, and 0.1 mg/mL sheared salmon-sperm DNA for at least 2 h at 65°C. The blots were probed with denatured α-32P-labeled probes added directly to the hybridization solution and incubated at 65°C for 12 to 16 h. The blots were washed twice for 15 to 20 min at 65°C with 50 mL of a solution containing 40 mmNa2PO4 (pH 7.2), 5% (w/v) SDS, 1 mm EDTA, and 5 mg/mL BSA. The blots were then washed for a second time in 50 mL of a solution containing 40 mm Na2PO4 (pH 7.2), 1% (w/v) SDS, and 1 mm EDTA for 30 to 35 min at 60°C to 65°C. After washing, blots were placed on x-ray film for 1 to 4 d at −80°C. RESULTS Functional Complementation of E. coli His Mutants T. goesingense cDNAs that encode ATP-PRT, IGPD, and HDH were isolated. These genes were designated THG1,THB1 (Persans et al., 1998), and THD1 (Persans et al., 1998), and their sequences were submitted to GenBank. When expressed in E. coli HisG, HisB, and HisD mutant strains, these cDNAs were able to complement the E. coli mutants' inability to grow in the absence of His (Fig.1). This suggests that the T. goesingense THG1, THB1, and THD1 cDNAs encode ATP-PRT, IGPD, and HDH, respectively. THG1 is the first cDNA sequence that encodes ATP-PRT to be isolated from plants. Fig. 1. Open in new tabDownload slide Functional complementation ofHisG−, HisB−,andHisD− His auxotropic E. colimutants with T. goesingense cDNAs. Three T. goesingense cDNAs were isolated by screening a pTriplEx cDNA library for complementation of HisG− (A),HisB− (B), and HisD−(C) mutations in E. coli. The three clones represent ATP phosphoribosyltransferase (THG1, accession no.AF003347), imidazolglycerol phosphate dehydratase (THB1, accession no. AF023140), and histidinol dehydrogenase (THD1, accession no. AF023141). Each plate contained the following: a mutant unable to grow in the absence of His (1); a mutant transformed with complementing T. goesingense cDNA (2); and a wild-type TOPP10F′ E. coli strain (3). Fig. 1. Open in new tabDownload slide Functional complementation ofHisG−, HisB−,andHisD− His auxotropic E. colimutants with T. goesingense cDNAs. Three T. goesingense cDNAs were isolated by screening a pTriplEx cDNA library for complementation of HisG− (A),HisB− (B), and HisD−(C) mutations in E. coli. The three clones represent ATP phosphoribosyltransferase (THG1, accession no.AF003347), imidazolglycerol phosphate dehydratase (THB1, accession no. AF023140), and histidinol dehydrogenase (THD1, accession no. AF023141). Each plate contained the following: a mutant unable to grow in the absence of His (1); a mutant transformed with complementing T. goesingense cDNA (2); and a wild-type TOPP10F′ E. coli strain (3). The predicted amino acid sequences derived from the cDNA all had high identity to several protein sequences in GenBank. The THG1 amino acid sequence had 83% identity with a putative Arabidopsis ATP-PRT expressed sequence tag (EST) (GenBank accession no. Z31670) amino acid sequence. Also, the THG1 amino acid sequence had 29% and 26% identity to the ATP-PRT from E. coli (HisG) (GenBank accession no. X13462) and Saccharomyces cerevisiae (His1) (GenBank accession no. V01306) amino acid sequences, respectively (Fig.2A). The THB1 amino acid sequence had 86%, 87%, and 84% identity to an Arabidopsis IGPD (GenBank accession no. 2244848), a Triticum aestivum IGPD (GenBank accession no. 551331), and a Pisum sativum IGPD (GenBank accession no. 2495230) amino acid sequence, respectively. (Fig. 2B). The THD1 amino acid sequence had 89% and 86% identity with Brassica oleracea HDH (GenBank accession no. 60466) (Fig. 2C) and a putative Arabidopsis EST HDH (GenBank accession no.T42850) sequence. Fig. 2. Open in new tabDownload slide Open in new tabDownload slide Alignment of the predicted amino acid sequences of THG1, THB1, andTHD1. Sequences used are as follows. A, T. goesingense ATP-PRT (THG1; accession no. AF003347), Arabidopsis EST sequence (At-HisG; accession no. Z31670), E. coliATP-PRT (Ec-HisG; accession no. X13462), S. cerevisiaeATP-PRT (Sc-His1; accession no. V01306). Green, Amino acids conserved in all aligned sequences; yellow, amino acids conserved in at least two of the aligned sequences; blue, conservative amino acid substitutions. B, T. goesingense (THB1; accession no. AF023140), and Arabidopsis IGPD (At-HisB; accession no. U02689). C, T. goesingense (THD1; accession no. AF023141), and B. oleracea HDH (Bo-HisD; accession no. M60466). Colons, Identity; periods, conservative replacements. Fig. 2. Open in new tabDownload slide Open in new tabDownload slide Alignment of the predicted amino acid sequences of THG1, THB1, andTHD1. Sequences used are as follows. A, T. goesingense ATP-PRT (THG1; accession no. AF003347), Arabidopsis EST sequence (At-HisG; accession no. Z31670), E. coliATP-PRT (Ec-HisG; accession no. X13462), S. cerevisiaeATP-PRT (Sc-His1; accession no. V01306). Green, Amino acids conserved in all aligned sequences; yellow, amino acids conserved in at least two of the aligned sequences; blue, conservative amino acid substitutions. B, T. goesingense (THB1; accession no. AF023140), and Arabidopsis IGPD (At-HisB; accession no. U02689). C, T. goesingense (THD1; accession no. AF023141), and B. oleracea HDH (Bo-HisD; accession no. M60466). Colons, Identity; periods, conservative replacements. An analysis of the protein leader sequences for all three T. goesingense His biosynthetic genes predicts that both THG1 and THD1 proteins are targeted to the chloroplast, and the THB1 protein appears to be targeted to the mitochondria (TableI). Table I. Predicted target sequence of the cloned T. goesingense His biosynthetic genes Gene Identification (Enzyme ID) . Predicted Target Sequence Type . PSORT Score (Most Likely Candidate) . GenBank Accession No. . THG1(ATP-PRT) Chloroplast 5.23 AF003347 THB1 (IGPD) Mitochondria 2.41 AF023140 THD1 (HDH) Chloroplast 3.04 AF023141 Gene Identification (Enzyme ID) . Predicted Target Sequence Type . PSORT Score (Most Likely Candidate) . GenBank Accession No. . THG1(ATP-PRT) Chloroplast 5.23 AF003347 THB1 (IGPD) Mitochondria 2.41 AF023140 THD1 (HDH) Chloroplast 3.04 AF023141 Open in new tab Table I. Predicted target sequence of the cloned T. goesingense His biosynthetic genes Gene Identification (Enzyme ID) . Predicted Target Sequence Type . PSORT Score (Most Likely Candidate) . GenBank Accession No. . THG1(ATP-PRT) Chloroplast 5.23 AF003347 THB1 (IGPD) Mitochondria 2.41 AF023140 THD1 (HDH) Chloroplast 3.04 AF023141 Gene Identification (Enzyme ID) . Predicted Target Sequence Type . PSORT Score (Most Likely Candidate) . GenBank Accession No. . THG1(ATP-PRT) Chloroplast 5.23 AF003347 THB1 (IGPD) Mitochondria 2.41 AF023140 THD1 (HDH) Chloroplast 3.04 AF023141 Open in new tab A phylogenetic tree was constructed to determine the evolutionary placement of the THG1 protein sequence in relation to existing protein sequences (Fig. 3). The THG1 amino acid sequence was closely grouped with an Arabidopsis EST encoding a putative ATP-PRT. The THG1 amino acid sequence is more distantly related to nine archaebacteria, eubacteria, and unicellular eukaryotic ATP-PRT sequences, and falls within its own unique group. Fig. 3. Open in new tabDownload slide Unrooted phylogenetic tree of T. goesingense ATP-PRT and nine other ATP-PRT sequences. T. goesingense ATP-PRT (THG1; accession no. AF003347), Arabidopsis EST sequence (At-HisG; accession no. Z31670), S. cerevisiae ATP-PRT (Sc-His1; accession no. V01306),Yarrowia lipoytica (Yl-His1; accession no. U40563), Schizosaccharomyces pombe (Sp-His1; accession no. Z70691), Hemophilus influenzae (Hi-HisG; accession no. U32729),Salmonella typhimurium (St-HisG; accession no. X13464),E. coli ATP-PRT (Ec-HisG; accession no. X13462),Archaeoglobus fulgidus (His1-Arcfu; accession no. AE001064), Methanococcus jannaschii(His1-Metja; accession no. U67562), and Methanobacterium thermoautotrophicum (His1-Metth; accession no. AE000911). Fig. 3. Open in new tabDownload slide Unrooted phylogenetic tree of T. goesingense ATP-PRT and nine other ATP-PRT sequences. T. goesingense ATP-PRT (THG1; accession no. AF003347), Arabidopsis EST sequence (At-HisG; accession no. Z31670), S. cerevisiae ATP-PRT (Sc-His1; accession no. V01306),Yarrowia lipoytica (Yl-His1; accession no. U40563), Schizosaccharomyces pombe (Sp-His1; accession no. Z70691), Hemophilus influenzae (Hi-HisG; accession no. U32729),Salmonella typhimurium (St-HisG; accession no. X13464),E. coli ATP-PRT (Ec-HisG; accession no. X13462),Archaeoglobus fulgidus (His1-Arcfu; accession no. AE001064), Methanococcus jannaschii(His1-Metja; accession no. U67562), and Methanobacterium thermoautotrophicum (His1-Metth; accession no. AE000911). Southern Blot of THG1 To examine the number of THG1 genes present in theT. goesingense genome, genomic DNA was digested with various restriction enzymes, run on an agarose gel, and Southern blotted. The genomic DNA was probed with an internal 400-bp XhoI fragment containing the 3′-UTR of the THG1 cDNA (Fig.4.) Of the restriction enzymes used,BamHI, EcoRI, HindIII,PstI, and XbaI do not cut within theTHG1 cDNA sequence and XhoI cuts once. As expected, BamHI, PstI, and HindIII digestion resulted in a single band, while digestion withXhoI resulted in two bands. Interestingly, EcoRI and XbaI digestion produced two bands. This is inconsistent with the restriction map of the cloned THG1 cDNA. This result implies that there may be more than one THG1 gene present in the genome. However, these results are consistent with the assumption that two or perhaps a small family of THG1 genes is present in the genome. Fig. 4. Open in new tabDownload slide Southern-blot analysis of T. goesingense THG1. Genomic DNA was cut with BamHI (lane A),EcoRI (lane B), HindIII (lane C),XbaI (lane D), XhoI (lane E), andPstI (lane F), and probed with a 32P-labeledTHG1 cDNA fragment. Fig. 4. Open in new tabDownload slide Southern-blot analysis of T. goesingense THG1. Genomic DNA was cut with BamHI (lane A),EcoRI (lane B), HindIII (lane C),XbaI (lane D), XhoI (lane E), andPstI (lane F), and probed with a 32P-labeledTHG1 cDNA fragment. Ni Regulation of THG1, THD1, and THB1 Gene Expression Northern analysis of total RNA isolated from T. goesingense exposed to 50 μm Ni for 0, 24, and 48 h showed that mRNA levels of THG1,THB1, or THD1 in both the roots and shoots are not affected by exposure to Ni in the hydroponic culture solution (Fig.5). Fig. 5. Open in new tabDownload slide Northern analysis of total RNA isolated fromT. goesingense exposed to 50 μm Ni for 0, 24, and 48 h showing the expression of THG1,THD1, and THB1. Total RNA was isolated from T. goesingense shoot (A) and root (B) tissue and probed with 32P-labeled THG1 (a),THB1 (b), and THD1 (c) cDNAs probes. Bottom rows in both A and B show blots from total RNA being probed with an Arabidopsis 32P-labeled actin cDNA fragment as a loading control. Fig. 5. Open in new tabDownload slide Northern analysis of total RNA isolated fromT. goesingense exposed to 50 μm Ni for 0, 24, and 48 h showing the expression of THG1,THD1, and THB1. Total RNA was isolated from T. goesingense shoot (A) and root (B) tissue and probed with 32P-labeled THG1 (a),THB1 (b), and THD1 (c) cDNAs probes. Bottom rows in both A and B show blots from total RNA being probed with an Arabidopsis 32P-labeled actin cDNA fragment as a loading control. Free His Concentrations and Ni Speciation Free His concentrations in both shoots and xylem sap of the Ni hyperaccumulator T. goesingense did not differ significantly from those observed in the nonaccumulator Thlaspi arvense(Table II). However, His concentrations in the roots of T. goesingense were significantly higher than those observed in T. arvense. Interestingly, after exposure to 50 μm Ni for 7 d, His concentrations in both the shoot and xylem exudate of T. goesingense remained unchanged. However, His concentrations in the roots dropped to levels observed in unexposed T. arvense(Table II). Acid hydrolysis of selected samples showed that there were no increases in the amount of His associated with peptides and proteins (data not shown). Table II. Free His content of T. goesingense tissues exposed to 50 μm Ni for 7 d . Shoot . Root . Xylem Sap . nmol g−1fresh biomass μmol L−1 T. goesingense  Control 136 ± 37 (4) 742 ± 188 (4) 7.4 ± 3 (4)  Nickel-treated 107 ± 62 (5) 68 ± 30 (7) 18.2 ± 9 (6) T. arvense  Control 73 ± 16 (4) 43 ± 9 (3) 57 ± 31 (8)  Nickel-treated n.a. n.a. n.a. . Shoot . Root . Xylem Sap . nmol g−1fresh biomass μmol L−1 T. goesingense  Control 136 ± 37 (4) 742 ± 188 (4) 7.4 ± 3 (4)  Nickel-treated 107 ± 62 (5) 68 ± 30 (7) 18.2 ± 9 (6) T. arvense  Control 73 ± 16 (4) 43 ± 9 (3) 57 ± 31 (8)  Nickel-treated n.a. n.a. n.a. His was measured by HPLC as the phenylthiocarbamyl amino acid derivative with methionine sulfoxide as the internal standard. n.a., Not available. Data are the means ± sd of between three and eight independent plant samples (nos. in parentheses represent n). Open in new tab Table II. Free His content of T. goesingense tissues exposed to 50 μm Ni for 7 d . Shoot . Root . Xylem Sap . nmol g−1fresh biomass μmol L−1 T. goesingense  Control 136 ± 37 (4) 742 ± 188 (4) 7.4 ± 3 (4)  Nickel-treated 107 ± 62 (5) 68 ± 30 (7) 18.2 ± 9 (6) T. arvense  Control 73 ± 16 (4) 43 ± 9 (3) 57 ± 31 (8)  Nickel-treated n.a. n.a. n.a. . Shoot . Root . Xylem Sap . nmol g−1fresh biomass μmol L−1 T. goesingense  Control 136 ± 37 (4) 742 ± 188 (4) 7.4 ± 3 (4)  Nickel-treated 107 ± 62 (5) 68 ± 30 (7) 18.2 ± 9 (6) T. arvense  Control 73 ± 16 (4) 43 ± 9 (3) 57 ± 31 (8)  Nickel-treated n.a. n.a. n.a. His was measured by HPLC as the phenylthiocarbamyl amino acid derivative with methionine sulfoxide as the internal standard. n.a., Not available. Data are the means ± sd of between three and eight independent plant samples (nos. in parentheses represent n). Open in new tab X-ray absorption spectroscopy clearly demonstrated that the amount of Ni coordinated by His in both roots and shoots of T. arvensealways exceeded that found in T. goesingense during 1 to 7 d of exposure to 10 μm Ni (TableIII). The x-ray absorption edge spectra for Ni-His were significantly different from Ni-imidazole (data not shown); therefore, the x-ray absorption spectroscopy data presented suggest that the Ni-His complex observed in Thlaspi tissues represents Ni coordinated with free His and not His residues in proteins. Table III. Nickel coordination by His ligands in T. goesingense and T. arvense measured by x-ray absorption spectroscopy Time (d) . T. goesingense . T. arvense . Shoot . Root . Shoot . Root . mmol kg−1dry biomass 1 0.2  (3.0)3-a 2.5  (8.7) 0.4  (0.5) 2.9  (6.2) 3 0.1  (4.2) 2.5  (6.7) 1.0  (2.3) 3.9  (10.6) 5 0.4  (6.5) 2.6  (8.1) 1.2  (2.4) 4.3  (13.0) 7 0.2  (5.3) 4.1  (9.4) 1.3  (3.3) 6.1  (14.8) Time (d) . T. goesingense . T. arvense . Shoot . Root . Shoot . Root . mmol kg−1dry biomass 1 0.2  (3.0)3-a 2.5  (8.7) 0.4  (0.5) 2.9  (6.2) 3 0.1  (4.2) 2.5  (6.7) 1.0  (2.3) 3.9  (10.6) 5 0.4  (6.5) 2.6  (8.1) 1.2  (2.4) 4.3  (13.0) 7 0.2  (5.3) 4.1  (9.4) 1.3  (3.3) 6.1  (14.8) Values in parentheses represent the total Ni content of the tissue mmol kg−1 dry biomass. F3-a Plants were exposed to 10 μmNi(NO3)2 in the hydroponic solution. By fitting x-ray absorption spectra of aqueous Ni2+ and Ni2+ coordinated with His: 6.66 mmNi(NO3)2, 80 mm His, 30% (w/v) glycerol, pH 7.0; citrate: 6.66 mmNi(NO3)2, 70 mm citrate, 30% glycerol, pH 8.0; Gln: 1 mmNi(NO3)2, 4 mm Gln, 30% glycerol, pH 7.3; and isolated T. goesingense shoot cell wall material (Lasat et al., 1996), we were able to determine the percentage contribution of His as a ligand of Ni2+. Total Ni content of the tissues was analyzed by inductively coupled plasma emission spectroscopy and this was used to calculate the absolute amount of Ni coordinated by His in the tissues. The values represent data collected from a single plant sample at each time point, and each x-ray spectrum used for the fits represents the mean of three independent scans, each being composed of data acquired from 13 independent detectors. Open in new tab Table III. Nickel coordination by His ligands in T. goesingense and T. arvense measured by x-ray absorption spectroscopy Time (d) . T. goesingense . T. arvense . Shoot . Root . Shoot . Root . mmol kg−1dry biomass 1 0.2  (3.0)3-a 2.5  (8.7) 0.4  (0.5) 2.9  (6.2) 3 0.1  (4.2) 2.5  (6.7) 1.0  (2.3) 3.9  (10.6) 5 0.4  (6.5) 2.6  (8.1) 1.2  (2.4) 4.3  (13.0) 7 0.2  (5.3) 4.1  (9.4) 1.3  (3.3) 6.1  (14.8) Time (d) . T. goesingense . T. arvense . Shoot . Root . Shoot . Root . mmol kg−1dry biomass 1 0.2  (3.0)3-a 2.5  (8.7) 0.4  (0.5) 2.9  (6.2) 3 0.1  (4.2) 2.5  (6.7) 1.0  (2.3) 3.9  (10.6) 5 0.4  (6.5) 2.6  (8.1) 1.2  (2.4) 4.3  (13.0) 7 0.2  (5.3) 4.1  (9.4) 1.3  (3.3) 6.1  (14.8) Values in parentheses represent the total Ni content of the tissue mmol kg−1 dry biomass. F3-a Plants were exposed to 10 μmNi(NO3)2 in the hydroponic solution. By fitting x-ray absorption spectra of aqueous Ni2+ and Ni2+ coordinated with His: 6.66 mmNi(NO3)2, 80 mm His, 30% (w/v) glycerol, pH 7.0; citrate: 6.66 mmNi(NO3)2, 70 mm citrate, 30% glycerol, pH 8.0; Gln: 1 mmNi(NO3)2, 4 mm Gln, 30% glycerol, pH 7.3; and isolated T. goesingense shoot cell wall material (Lasat et al., 1996), we were able to determine the percentage contribution of His as a ligand of Ni2+. Total Ni content of the tissues was analyzed by inductively coupled plasma emission spectroscopy and this was used to calculate the absolute amount of Ni coordinated by His in the tissues. The values represent data collected from a single plant sample at each time point, and each x-ray spectrum used for the fits represents the mean of three independent scans, each being composed of data acquired from 13 independent detectors. Open in new tab DISCUSSION To test the hypothesis that free His plays a role in the mechanism of Ni hyperaccumulation in T. goesingense, we investigated the regulation of His biosynthesis at the molecular and biochemical levels, and studied the role of His in Ni coordination in planta. By functionally complementing His auxotrophic E. coli mutants with T. goesingense cDNAs, we isolated genes (THG1, THB1, and THD1) encoding enzymes that catalyze three steps in the His biosynthetic pathway. The first of these, THG1, encodes ATP-PRT, a functional homolog of an enzyme that catalyzes the production of N-(5′-phosphoribosyl)-ATP (PR-ATP) from ATP and phosphoribosyl pyrophosphate, the first committed step in His biosynthesis in E. coli. This is the first conclusive evidence that ATP-PRT exists in plants and confirms the earlier observation of ATP-PRT-like enzymatic activity in plant tissue extracts (Waiter et al., 1971). The presence of ATP-PRT in T. goesingense also supports the growing body of evidence suggesting that His biosynthesis in plants follows a very similar pathway to that observed in E. coli(Nagai et al., 1991; Tada et al., 1993, 1994; Kim and Theologis, 1996;Bevan et al., 1998; El Malki et al., 1998; Fujimori and Ohta, 1998). Because THG1 is the first example of a gene encoding ATP-PRT in plants, we performed a genomic Southern analysis to determine how many copies of this gene occur in the T. goesingense genome. This analysis suggested that there are only a small number ofTHG1 genes present in T. goesingense, similar to the low copy number of genes encoding HDH and IGPD in B. oleracea and Arabidopsis (Nagai et al., 1991; Tada et al., 1994). An investigation of the evolutionary relationships between the T. goesingense ATP-PRT and other archaebacteria, eubacteria, and unicellular eukaryotic ATP-PRT sequences placed the plant ATP-PRT on a separate evolutionary branch from the three other groups (Fig. 3). TheT. goesingense ATP-PRT did cluster with an Arabidopsis EST clone (Z31670) that showed 81% amino acid identity to the T. goesingense ATP-PRT amino acid sequence. Expression of the T. goesingense THG1 in E. coligenerated a protein with an apparent molecular mass of 49,000 D, based on SDS-PAGE (data not shown). This corresponds closely to a predicted molecular mass of 49,335 D based on the protein translation product of the fusion of the THG1 cDNA sequence and the expression vector sequence. The predicted molecular mass of the T. goesingense ATP-PRT after cleavage of the putative chloroplast target sequence, at the predicted cleavage site between amino acids residues 45 and 46, was calculated to be 39,056 D. This is similar to the molecular masses of other ATP-PRTs of approximately 32,500 D (Alifano et al., 1996). In E. coli ATP-PRT is an important control point for His biosynthesis, being regulated at the level of transcription, translation, and allosteric activation/inhibition (Alifano et al., 1996). The highly regulated nature of ATP-PRT in E. coli and its role in the catalysis of the first committed step in His biosynthesis suggests that it might also play a key regulatory role in plants, making it a likely target for the regulation of His biosynthesis by Ni. However, northern analysis of the THG1RNA message clearly demonstrated that Ni does not induce or suppress transcription of the THG1 mRNA in either the roots or shoots of T. goesingense (Fig. 5). Because it is possible that ATP-PRT is not the key regulated step in His biosynthesis in plants, we also analyzed expression of two other genes in the His biosynthetic pathway, THB1 and THD1, which encode the enzymes IGPD and HDH, respectively. The predicted amino acid sequence of bothTHB1 and THD1 show very high homology to other known plant homologs, suggesting that the His biosynthetic pathway is highly conserved in plants. The IGPD enzyme catalyzes the conversion of imidazolglycerol phosphate to imidazoleacetol phosphate, the first step after the branch point that feeds into the purine recycling pathway. This enzyme also catalyzes the conversion ofl-histidinol phosphate tol-histidinol. Because of its key position at a branch point in the His biosynthetic pathway this enzyme may also be regulated. HDH catalyzes the oxidation ofl-histidinol tol-hisitidine, the final step in His biosynthesis. Because this catalytic step uses NAD+ as an oxidant, it is possible that it is also regulated in plants. Northern analysis of the mRNA levels for both THB1 andTHD1 clearly showed that expression of these mRNAs is not induced or repressed by Ni treatment in either the roots or the shoots of T. goesingense (Fig. 5). Because THG1,THB1, and THD1 mRNA expression levels were not changed by Ni treatment, it is unlikely that control of the His biosynthetic pathway at the transcriptional level by Ni is involved in Ni hyperaccumulation in T. goesingense. To determine if Ni modifies His biosynthesis at the post-transcriptional level in T. goesingense, we also analyzed the concentration of free His in root, xylem sap, and shoot tissue. It is clear from this data (Table II) that His concentrations remain basically unchanged after Ni exposure in both the xylem sap and the shoots. These biochemical data strongly support the molecular evidence that free His concentrations in T. goesingense are not increased by Ni exposure. It is possible, however, that the constitutive concentration of free His observed in T. goesingense is sufficient to fulfill its theorized role in Ni hyperaccumulation. To test this hypothesis, we compared the His concentration in T. goesingense and the nonaccumulatorT. arvense. This comparison revealed that the nonaccumulatorT. arvense contained equal concentrations of His in roots, shoots, and xylem sap as that found in T. goesingense during Ni exposure. The His concentration in the xylem sap of T. goesingenseafter Ni exposure was also similar to that measured in the nonaccumulators Vitis rotundifolia and Lagerstroemia indica (Anderson and Brodbeck, 1989; Anderson et al., 1993). However, we did observe that roots of T. goesingense before Ni exposure had a significantly higher concentration of free His compared with T. arvense. An interesting possibility is that His is overproduced in the roots of T. goesingense as a Ni-scavenging mechanism to enhance Ni acquisition under low external Ni conditions. Once plants were exposed to a higher Ni concentration, the free His concentration in the roots of the hyperaccumulator were observed to significantly decrease (Table II), and this may reflect the fact that Ni scavenging is no longer required. However, this reduction in His concentration in the roots of the hyperaccumulator was not reflected in reduced expression of THG1,THB1, or THD1. Also, this His loss was not accounted for by increased His concentrations in xylem sap or shoots. It is possible that reduced His concentrations may reflect increased catabolism or efflux of His into roots. However, recent analysis ofT. goesingense root exudate showed no increases in the rates of His exudation from roots after exposure to Ni (Salt et al., 1999a). If free His is involved in the hyperaccumulation of Ni, as has been suggested to occur in Alyssum species (Krämer et al., 1996), we would predict that His binds Ni within the plant. To directly address this hypothesis we used x-ray absorption spectroscopy to determine the in planta coordination environment of the Ni in both the hyperaccumulator and nonaccumulator Thlaspi species (TableIII). From these data it was clear that free His or a free-His-like molecule is involved in coordinating Ni in both the roots and shoots ofT. goesingense and T. arvense. However, the concentration of the Ni-His complex in the shoots of the nonaccumulatorT. arvense appears to be approximately 5- to 10-fold higher than in T. goesingense, and equal in the roots, suggesting that increased Ni coordination by His is not a primary determinant of the Ni hyperaccumulation mechanism in Thlaspi. Furthermore, the addition of l-His to the culture solution of hydroponically grown T. arvense had no effect on the accumulation of Ni in either the root or shoot tissue (data not shown). Our data suggest that Ni hyperaccumulation in T. goesingenseis not simply related to an enhanced ability of the hyperaccumulator to accumulate more free His in response to Ni. We would also caution that the role of free His in Ni hyperaccumulation in Alyssumremains speculative and will remain so until more detailed mechanistic data are available. For example, using Arabidopsis probes for northern and western analysis in the Ni hyperaccumulator Alyssum pintodasilvae (Baker and Brooks, 1989), expression levels of the His biosynthetic enzymes ATP-PRT, IGDH, and HDH were found not to be regulated by Ni (data not shown). However, there is certain limited evidence suggesting that His may be involved in Ni transport in Thlaspi species in general. For example, in this study a significant proportion of root Ni was found to be coordinated by free His ligands in both hyperaccumulator and nonaccumulator Thlaspi species. Also, exposure of the nonaccumulator T. arvense to d-His was observed to reduce the accumulation of Ni in shoots, but was found to have no effect on root Ni concentrations. This suggests that thed-His-Ni complex may compete with an endogenousl-His-Ni complex for transport to the shoot. However, we would again like to stress that the primary determinant of the Ni hyperaccumulation phenotype in T. goesingense is not governed by the overproduction of free His, as has been suggested forAlyssum Ni hyperaccumulators. ACKNOWLEDGMENTS The authors wish to extend their appreciation to Ingrid Pickering and Roger Prince for their help with x-ray absorption spectroscopy data collection and analysis. The Stanford Synchrotron Radiation Laboratory is funded by the Department of Energy, Office of Basic Energy Sciences, Divisions of Chemical and Materials Science. The Biotechnology Program is supported by the National Institutes of Health, National Center for Research Resources, Biomedical Technology Program. Further support is provided by the Department of Energy, Office of Biological and Environmental Research. We would also like to thank Pamella Motely and Isaac Shaffer for technical assistance, Ilya Raskin for his support of this project, and Richard Meager for providing the Arabidopsis actin cDNA. We would also like to thank theE. coli Genetic Stock Center (http://cgsc.biology.yale.edu) for providing the E. coli mutant strains. LITERATURE CITED 1 Alifano P Fani R Lio Pietro Lazcano A Bazzicalupo M Carlomagno MS Bruni CB Histidine biosynthetic pathway and genes: structure, regulation and evolution. 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Nebenführ, Andreas; Gallagher, Larry A.; Dunahay, Terri G.; Frohlick, Jennifer A.; Mazurkiewicz, Anna M.; Meehl, Janet B.; Staehelin, L. Andrew

doi: 10.1104/pp.121.4.1127pmid: 10594100

Abstract The Golgi apparatus in plant cells consists of a large number of independent Golgi stack/trans-Golgi network/Golgi matrix units that appear to be randomly distributed throughout the cytoplasm. To study the dynamic behavior of these Golgi units in living plant cells, we have cloned a cDNA from soybean (Glycine max), GmMan1, encoding the resident Golgi protein α-1,2 mannosidase I. The predicted protein of approximately 65 kD shows similarity of general structure and sequence (45% identity) to class I animal and fungal α-1,2 mannosidases. Expression of a GmMan1::green fluorescent protein fusion construct in tobacco (Nicotiana tabacum) Bright Yellow 2 suspension-cultured cells revealed the presence of several hundred to thousands of fluorescent spots. Immuno-electron microscopy demonstrates that these spots correspond to individual Golgi stacks and that the fusion protein is largely confined to the cis-side of the stacks. In living cells, the stacks carry out stop-and-go movements, oscillating rapidly between directed movement and random “wiggling.” Directed movement (maximal velocity 4.2 μm/s) is related to cytoplasmic streaming, occurs along straight trajectories, and is dependent upon intact actin microfilaments and myosin motors, since treatment with cytochalasin D or butanedione monoxime blocks the streaming motion. In contrast, microtubule-disrupting drugs appear to have a small but reproducible stimulatory effect on streaming behavior. We present a model that postulates that the stop-and-go motion of Golgi-trans-Golgi network units is regulated by “stop signals” produced by endoplasmic reticulum export sites and locally expanding cell wall domains to optimize endoplasmic reticulum to Golgi and Golgi to cell wall trafficking. The Golgi apparatus of plant cells consists of a large number of small, independent stack-trans-Golgi network (TGN) units that are distributed throughout the cytoplasm (Driouich and Staehelin, 1997;Andreeva et al., 1998b; Dupree and Sherrier, 1998). This seemingly random organization within plant cells is in striking contrast to the highly ordered Golgi complex in animal cells (Rambourg and Clermont, 1997). Presumably, these different forms of spatial organization reflect different mechanisms for controlling the localization of Golgi membranes. Similarly, the distinct distributions found in the two organisms likely impose different requirements on the transport of vesicles to and from the Golgi. However, while many of the molecules underlying Golgi positioning and functioning have been identified in animal systems (Barlow, 1998; Lowe and Kreis, 1998), very little is known about these processes for the plant Golgi (Andreeva et al., 1998a). The juxtanuclear position of the Golgi apparatus in animal cells depends on the presence of an intact microtubule (MT) cytoskeleton (Burkhardt, 1998). Golgi membranes assemble around the centrosome in close proximity to the nucleus, where the cis-cisternae are anchored to the minus ends of MTs (Infante et al., 1999). Vesicular transport from the peripheral endoplasmic reticulum (ER) to this central Golgi complex occurs in a targeted fashion along MTs (Presley et al., 1997; Scales et al., 1997), and post-Golgi transport also often follows MT tracks into the periphery of the cell (Hirschberg et al., 1998; Toomre et al., 1999). In contrast, the Golgi stack-TGN units of angiosperms are dispersed throughout the cytoplasm (Robinson and Kristen, 1982). This distribution conceivably reduces the distance that transport vesicles have to travel, both from ER export sites to the Golgi and from the Golgi to the plasma membrane or vacuole. However, it is unclear whether this traffic is directed, as is the case in animal cells, or whether it occurs by passive diffusion or cytoplasmic streaming (Williamson, 1993). Cytoplasmic streaming is a process found in many plant cells that moves large quantities of cytoplasm (including organelles) around the cell. This phenomenon is most pronounced in larger, highly vacuolated cells and is generally assumed to allow for efficient mixing and distribution of solutes. Cytoplasmic streaming has been studied extensively in internodal cells of characean algae (Kuroda, 1990; Shimmen and Yokota, 1994). It usually is driven by the acto-myosin system, although instances of microtubule-based organelle movement have been described (e.g. Mizukami and Wada, 1981; Mineyuki and Furuya, 1986). Studies of streaming events in the past have relied on visualization of moving organelles with phase-contrast or Nomarski microscopy. The nature of the organelles was therefore mostly unknown. The participation of Golgi vesicles in cytoplasmic streaming was inferred based only on the effects of inhibitor studies (e.g. Mollenhauer and Morré, 1976). It also has been proposed that entire Golgi stacks might participate in the streaming motion (Staehelin and Moore, 1995). This postulated movement of Golgi stacks has recently been demonstrated in tobacco (Nicotiana clevelandii) leaf epidermal cells by means of green fluorescent protein (GFP) transiently expressed in the Golgi as a fusion with the targeting domain of mammalian sialyltransferase (Boevink et al., 1998). Golgi stack movement may also provide the mechanism responsible for their dispersed distribution throughout the cytoplasm. At the same time, this movement imposes additional constraints on the possible mechanisms for transport to and from the Golgi. To investigate these Golgi dynamics in greater detail, we set out to develop a plant-gene-based Golgi marker in a stably transformed cell line. Such a system would allow quantitative analysis of this Golgi movement in great detail, and also enable us to study Golgi dynamics in dividing cells and to carry out novel biochemical fractionation studies. In this report we describe the isolation of a cDNA from soybean (Glycine max) encoding an α-1,2 mannosidase I,GmMan1, the first enzyme of the N-linked oligosaccharide pathway cloned from plants. Fusion of this protein to GFP when expressed in stably transformed tobacco Bright Yellow 2 (BY-2) cells is localized to the cis-Golgi. In living cells, Golgi stacks show characteristic saltatory movements throughout the cytoplasm. Inhibitor treatments demonstrate that this movement is dependent on the acto-myosin system, and that MTs can limit Golgi mobility in a subset of cells. MATERIALS AND METHODS Cloning of GmMan1 A partial soybean (Glycine max) α-1,2 mannosidase I clone was generated using degenerate oligonucleotides corresponding to conserved regions 1 and 3 as described previously (Herscovics et al., 1994). The clone (kindly provided by A. Herscovics, McGill University) was sequenced, and gene-specific primers to this partial soybean sequence (MannoN: 5′-TGGTTTTATGARTAYTTGYTGAAA, MannoC: 5′-ATACTTCAGCGTCTCCGCAAG) were designed (marked by thin underlines in Fig. 1). These primers were used in a PCR-based screen of a soybean cDNA library (kindly provided by J. Mullet, Texas A&M University, College Station). Plates of plaques were top-laid with SM medium (Sambrook et al., 1989), which was then tested by PCR for the presence of inserts that could be amplified with MannoN and MannoC. Positive pools were then repeatedly plated at lower plaque densities until a PCR product could be detected on a plate of about 100 plaques. A clone of approximately 2.6 kb was then isolated by plaque-lift screening. Fig. 1. Open in new tabDownload slide Predicted amino acid sequence of GmMan1 and its alignment with selected other α-1,2 mannosidase I protein sequences. The sequences and their accession nos. are: soybean, AF126550 (this report); mouse1b, U03458 (Herscovics et al., 1994);Drosophila, X82641 (Kerscher et al., 1995); and yeast,M63598 (Camirand et al., 1991). Homologous residues are highlighted by shading; residues identical in soybean and other sequences are boxed. The solid bar highlights the predicted membrane-spanning domain of GmMan1. The predicted catalytic domain is bracketed by arrows. Dots indicate conserved cysteyl residues that form a disulfide bridge (Lipari and Herscovics, 1996). Conserved acidic residues required for enzymatic activity are highlighted with asterisks (Lipari and Herscovics, 1999). The primers used for cDNA library screening were designed against the underlined sequences. Fig. 1. Open in new tabDownload slide Predicted amino acid sequence of GmMan1 and its alignment with selected other α-1,2 mannosidase I protein sequences. The sequences and their accession nos. are: soybean, AF126550 (this report); mouse1b, U03458 (Herscovics et al., 1994);Drosophila, X82641 (Kerscher et al., 1995); and yeast,M63598 (Camirand et al., 1991). Homologous residues are highlighted by shading; residues identical in soybean and other sequences are boxed. The solid bar highlights the predicted membrane-spanning domain of GmMan1. The predicted catalytic domain is bracketed by arrows. Dots indicate conserved cysteyl residues that form a disulfide bridge (Lipari and Herscovics, 1996). Conserved acidic residues required for enzymatic activity are highlighted with asterisks (Lipari and Herscovics, 1999). The primers used for cDNA library screening were designed against the underlined sequences. Sequence Analysis Multiple sequence alignments were initially performed with the PileUp program from the Genetics Computer Group (Madison, WI) and subsequently optimized manually. Prediction of transmembrane domains was done with TMpred (http://www.ch.embnet.org/software/TMPRED_form.html; Hofmann and Stoffel, 1993). Coiled-coil prediction was carried out with the COILS program (http://www.ch.embnet.org/software/COILS_form.html) using the algorithm of Lupas et al. (1991). Construction of GmMan1::GFP Fusion Protein The gene encoding a modified GFP (HBT-SGFP-TYG-nos in pUC18, obtained from J. Sheen, Massachusetts General Hospital) was subcloned with BamHI and EcoRI into pBluescript (Stratagene, La Jolla, CA) to obtain an in-frame XbaI site at its 5′ end. This was then spliced to an internal XbaI site in GmMan1, thus removing the C-terminal 11 amino acids of the mannosidase-coding region. To drive expression in plant cells a modified cauliflower mosaic virus 35S promoter (with a dual-enhancer element) generated by PCR from a plasmid (pZEV Ω, J. Oliver and K. Danna, University of Colorado, Boulder) was used. This expression cassette was then inserted into the SacI and KpnI sites of pBIN20 (Hennegan and Danna, 1998) and transformed intoAgrobacterium tumefaciens strain LBA4404 to yield strain BP37. Transformation of tobacco (Nicotiana tabacum) Bright Yellow 2 (BY-2) suspension-cultured cells (3 d after subculture) was achieved by co-cultivation with strain BP37 for 2 d at 27°C. Cells were then transferred onto selective medium (BY-2 medium plus 500 μg/mL carbenicillin and 100 μg/mL kanamycin). Growth Conditions Transformed cells were grown in a modified Linsmaier and Skoog medium (Nagata et al., 1982) with constant shaking (120 rpm) at 27°C in the dark. Cells were subcultured weekly into fresh medium at a dilution of 1:50. Cells were harvested by low-speed centrifugation at 500g for 2 min 6 to 8 d after subculturing and immediately used for experiments. Electron Microscopy Transformed BY-2 cells were high-pressure-frozen/freeze-substituted and embedded for transmission electron microscopy, as described in Samuels et al. (1995). For immunogold detection, the following modifications of the standard protocol were applied. After staining with osmium, samples were embedded in LR White. Sections (90 nm) of the samples were cut and placed on formvar-coated 300 mesh nickel grids. After a 20-min exposure to saturated sodium metaperiodate, the grids were washed briefly and blocked with 5% (w/v) nonfat milk in phosphate-buffered saline containing 0.1% (w/v) Tween (PBST). The sections were then exposed to the primary anti-GFP antibody (kindly provided by J. Kahana, Harvard University) for 2 h. Following a thorough rinse with PBST, the sections were labeled with 15 nm of goat anti-rabbit IgG secondary antibody (British BioCell International, Cardiff, UK) for 1 h. The final rinse was with PBST followed by water. The grids were then post-stained in 2% aqueous uranyl acetate and Reynold's lead citrate. Sections were observed on an electron microscope (model CM10, Philips, Eindhoven, The Netherlands). Fluorescence Microscopy Cells were observed using the standard fluorescein isothiocyanate filter set. Confocal images were obtained on a Sarastro MultiProbe system (Molecular Dynamics, Sunnyvale, CA) using a ×100 objective (Nikon, Tokyo). Conventional fluorescence microscopy was on an Axioscope microscope (Zeiss, Jena, Germany) with a ×100 objective and a Nikon Eclipse microscope with a ×60 objective (for video capture). Videos of streaming Golgi stacks were captured with a color CCD camera (Optronics, Goleta, CA) at an exposure setting of 1/8 s. Video frames were recorded on a PowerMacintosh computer equipped with a graphics digitizer board (RasterOps) at a rate of 10 frames per s. Individual frames in 1-s intervals were imported into the public domain NIH Image program (developed at the National Institutes of Health and available on the Internet athttp://rsb.info.nih.gov/nih-image), optically enhanced, and used for determination of x/y coordinates of individual stacks. To quantify the streaming behavior of individual stacks, the “streaming coefficient” was calculated according to the formula: sc=vnet×dirEquation 1 where sc is the streaming coefficient,v net is the net velocity, anddir is a directionality factor. The net velocity is calculated as: vnet=nd/tEquation 2 where nd is the net displacement during the observation period (i.e. the distance between the first and last position of the tracing), and t is time (i.e. the duration of the observation period). The directionality factor dir is defined as: dir=nd/tdtEquation 3 where tdt is the total distance traveled during the observation period. The streaming coefficient for straight trajectories therefore equals the average velocity of the stack. For curved trajectories, the streaming coefficient roughly equals the average velocity in the preferred direction of movement, corrected by the directionality factor. Inclusion of this factor ensures that stacks displaying a high degree of random motion will have a low streaming coefficient, even in the presence of substantial amounts of drift. Drug Treatments Drugs (from Sigma, St. Louis) were prepared as a 1,000-fold concentrated stock solution in DMSO and stored at −20°C, except for 2,3-butanedione monoxime (BDM), which was freshly dissolved in BY-2 medium just prior to the experiment. Drug treatments were performed for 15 to 45 min. Controls containing an equivalent concentration of DMSO (0.1%, w/v) did not show any response to the solvent. RESULTS Cloning of Soybean α-1,2 Mannosidase I The enzyme α-1,2 mannosidase I belongs to the class I α-mannosidases, a family of enzymes which remove α-1,2-mannosyl residues from the high Man N-linked oligosaccharides that are synthesized in the ER (Moremen et al., 1994). These are the first modification reactions to occur in the Golgi. A number of class I α-1,2 mannosidases have been cloned from animal and fungal organisms (Herscovics, 1999b), but no enzymes of the N-linked oligosaccharide pathway have been cloned from plants. Degenerate primers based on α-1,2-mannosidase sequences from mouse and yeast (Herscovics et al., 1994) have previously been used to amplify cDNA fragments from other species, among them soybean (A. Herscovics, personal communication). Specific primers were designed based on the termini of this partial soybean clone (target sequences are underlined in Fig. 1) and used in a PCR-based screen of cDNA libraries from soybean, Arabidopsis, and tobacco. Using this approach, a clone was isolated from the soybean library with an insert of approximately 2.5 kb, which could serve as a template for PCR with our gene-specific primers and also hybridized with the original gene fragment from degenerate PCR on a DNA gel blot (data not shown). Sequencing of the cDNA clone revealed an open reading frame of 1,734 bp, encoding a hypothetical protein of 578 amino acids and a calculated molecular mass of 65,345 D (Fig. 1, GenBank accession no. AF126550). Sequence analysis predicts a single, very short transmembrane domain (amino acids 30… 45 [black line in Fig. 1]) with the N terminus of the protein on the cytoplasmic face of the membrane. This type II orientation is typical of Golgi proteins and is also found in all other known α-1,2-mannosidases (Herscovics, 1999a, 1999b). The lumenal domain consists of a putative stalk region (46… 101), the catalytic domain (102… 548), and a C-terminal tail (549… 578). The stalk region has a high probability of forming a coiled-coil structure. The stalk and tail show no sequence similarity to other known mannosidases. In contrast, the putative catalytic domain is approximately 55% similar and 45% identical to corresponding parts of α-1,2-mannosidases from either mouse or yeast and also contains several features predicted to be important for enzymatic activity of α-1,2-mannosidases. In particular, two conserved Cys residues that form a required disulfide bond in yeast (Lipari and Herscovics, 1996) are present in the predicted soybean protein (C387 and C420; bullets in Fig. 1). Several carboxyl residues shown by site-directed mutagenesis to be crucial for enzymatic activity (Lipari and Herscovics, 1999) are also conserved (asterisks in Fig. 1). Based on these extensive similarities, we predict that the isolated soybean cDNA encodes a class I α-1,2-mannosidase (EC 3.2.1.113; glycosyl hydrolase family 47,Henrissat and Bairoch, 1996) and propose to call it GmMan1. Localization of a GmMan1::GFP Fusion to the cis-Golgi in Tobacco Cells Some α-1,2 mannosidases are localized to the Golgi (Herscovics et al., 1994; Lal et al., 1994), although other members of the family were found in the ER (e.g. Roth et al., 1990; Burke et al., 1996). The localization of GmMan1 was tested by creating an in-frame fusion to a modified GFP (GmMan1::GFP) and expressing it in stably transformed tobacco suspension-cultured cells (BY-2, Nagata et al., 1982). Most of the recovered cell lines displayed a punctate pattern of GFP fluorescence, as would be expected from the dispersed organization of Golgi stacks in plant cells (Fig. 2). Some cell lines showed an additional reticulate fluorescence that resembled the fluorescence seen in cells expressing ER-targeted GFP (data not shown). Fig. 2. Open in new tabDownload slide Distribution of GmMan1::GFP in living tobacco BY-2 suspension-cultured cells. A, Single optical section from a confocal microscope through the middle of a group of cells. B, Optical section through the cortical cytoplasm of a single cell. Brightly fluorescing spots can be found throughout the cytoplasm in both cortical regions and transvacuolar strands but not the vacuole (V) or the nucleus (N). The insets (×3 enlargement) demonstrate that some spots appear as short lines (arrowheads), whereas others resemble discs (arrows). C, Conventional epifluorescence picture of cortical cytoplasm. Several Golgi stacks appear as ring-like structures (arrow). Some of the other stacks changed in appearance from lines to rings over time (arrowhead; compare video sequence athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1). Fig. 2. Open in new tabDownload slide Distribution of GmMan1::GFP in living tobacco BY-2 suspension-cultured cells. A, Single optical section from a confocal microscope through the middle of a group of cells. B, Optical section through the cortical cytoplasm of a single cell. Brightly fluorescing spots can be found throughout the cytoplasm in both cortical regions and transvacuolar strands but not the vacuole (V) or the nucleus (N). The insets (×3 enlargement) demonstrate that some spots appear as short lines (arrowheads), whereas others resemble discs (arrows). C, Conventional epifluorescence picture of cortical cytoplasm. Several Golgi stacks appear as ring-like structures (arrow). Some of the other stacks changed in appearance from lines to rings over time (arrowhead; compare video sequence athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1). Immuno-electron microscopy on high-pressure frozen/freeze-substituted samples with antibodies against the GFP part of the fusion protein was performed to determine whether the punctate fluorescence does indeed highlight individual Golgi stacks. As shown in Figure3C, gold label accumulated predominantly over the cis and medial cisternae of stacks, with very little staining in trans and TGN cisternae. In some cells, weak labeling of ER (Fig.3D) and multivesicular bodies could also be observed. These localizations could mark fusion proteins in transit to the Golgi or targeted for degradation, respectively. Therefore, the GmMan1::GFP fusion construct is predominantly targeted to the cis-Golgi in BY-2 cells. The fluorescent spots shown in Figure 2 thus represent Golgi stacks and can be used to investigate Golgi dynamics in living cells. The overall morphology of Golgi stacks is not altered by overexpression of the fusion protein (compare Fig. 3, A and B). However, we frequently noticed an increase in staining intensity of the cisternal membranes on the cis side of the stack (Fig. 3B). In addition, Golgi stacks in the transgenic lines had a slight reduction of diameter (0.636 versus 0.803 μm, n = 33), which was accompanied by an marginal decrease in average number of cisternae per stack (5.0 versus 5.4, n = 33). Fig. 3. Open in new tabDownload slide Ultrastructure of Golgi stacks in untransformed (A) and transformed (B) BY-2 cells and immunogold-localization of the GmMan1::GFP fusion protein (C and D). A, Thin section image of single Golgi stack in high-pressure-frozen/freeze-substituted control cell showing normal appearance of Golgi stacks in BY-2 cells. B, In transformed cells, the cisternae generally have a normal appearance, except for stronger staining of the cis-cisternae. As in control cells, intercisternal elements are present between the trans-cisternae. C, Localization of GmMan1::GFP, as detected by antibodies against the GFP protein, is mostly restricted to the cis-side of the Golgi stack. D, In some cells, weak, non-Golgi labeling was observed over the ER (arrows). M, Mitochondrion; G, Golgi stack. Bar, 0.2 μm. Fig. 3. Open in new tabDownload slide Ultrastructure of Golgi stacks in untransformed (A) and transformed (B) BY-2 cells and immunogold-localization of the GmMan1::GFP fusion protein (C and D). A, Thin section image of single Golgi stack in high-pressure-frozen/freeze-substituted control cell showing normal appearance of Golgi stacks in BY-2 cells. B, In transformed cells, the cisternae generally have a normal appearance, except for stronger staining of the cis-cisternae. As in control cells, intercisternal elements are present between the trans-cisternae. C, Localization of GmMan1::GFP, as detected by antibodies against the GFP protein, is mostly restricted to the cis-side of the Golgi stack. D, In some cells, weak, non-Golgi labeling was observed over the ER (arrows). M, Mitochondrion; G, Golgi stack. Bar, 0.2 μm. As illustrated in Figure 2, BY-2 cells contain several hundred individual Golgi stacks. The stacks were more or less evenly distributed throughout the cytoplasm in both the cortical region underlying the plasma membrane and in cytoplasmic strands that traverse the large central vacuole. Careful examination of individual fluorescent spots revealed that they either appeared as discs (arrows in Fig. 2) or as short lines (arrowheads in Fig. 2). The length of the lines was identical to the diameter of the discs and uniform throughout the cell. In living cells, we could observe individual spots change from one shape to the other. These data suggest that the fluorescently labeled Golgi cisternae can be seen in both face-on views, when they appear as discs, and from the side, when they appear as lines. In a few cells we could also observe ring-like fluorescent structures (arrow in Fig. 2C). The diameter of these rings was approximately 1 μm, i.e. similar to Golgi stacks (Fig. 2, A and B). Occasionally these rings could change into short lines of the same length (arrowhead in Fig.2C), suggesting that the rings represent a face-on view of Golgi stacks in which the fusion protein is restricted to the rim region of the cisternae. Analysis of Golgi Stack Movement in Living Cells Observation of living BY-2 cells expressing the GmMan1::GFP fusion protein demonstrated that Golgi stacks can participate in cytoplasmic streaming and move throughout the cell (a video of Golgi stack movement can be viewed athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1). This movement was most pronounced in transvacuolar strands and in certain regions of the cortical cytoplasm. Golgi stacks that were not participating in directed movement displayed random “wiggling” reminiscent of Brownian motion. Those stacks that appeared to stream showed saltatory movement in which episodes of rapid movement alternated with periods of relative inactivity and wiggling. Golgi stacks that followed the same track occasionally paused at the same position along the track, suggesting that directed movement is inhibited at these sites. Within a given culture, only about two-thirds of the cells displayed “active streaming” of Golgi stacks. In the remaining cells, the stacks exhibited only wiggling motions (Table II). The percentage of active cells depended on the status of the culture, with younger cultures, which typically contained smaller cells, being less active in terms of their Golgi streaming. Most of the observations of streaming cells were therefore conducted on 7- to 8-d-old cultures. Within a given population, larger cells tended to have more active streaming, whereas small, round cells were mostly inactive. Movement of individual Golgi stacks was quantitated by video microscopy. Cells with bright green spots that showed active streaming were selected for recording over a period of 10 to 75 s with 1/8-s exposure time. The x-y coordinates of individual stacks were extracted in 1-s intervals to obtain tracings reflecting their movements (Fig. 4). This approach allowed for visualization of the relative activity of different regions of the cortical cytoplasm. In Figure 4, the tracings of individual stacks are color-coded according to the level of streaming they displayed over the entire observation period. Streaming stacks are marked with warm colors (yellow–red), whereas tracings of “wiggling” stacks are blue. The streaming level is defined by the “streaming coefficient,” which is calculated as the net velocity of a stack during a certain time interval multiplied by a directionality factor (see “Materials and Methods”). The resulting pattern shows that streaming Golgi stacks follow preferred paths (Fig. 4). These tracks are separated by regions of limited or random movement. It is also evident that large differences in mobility can exist between stacks that are spatially close together. Fig. 4. Open in new tabDownload slide Tracing of selected Golgi stacks over 30 s. A, Cortical region in untreated cell. B, Transvacuolar strands in untreated cell. C, Cortical region in cell treated with 40 μm cytochalasin D for 30 min. D, Cortical region in cell treated with 3.3 μm nocodazole for 30 min. The positions of individual Golgi stacks are marked in 1-s intervals. The tracings of the stacks are color coded according to the streaming coefficient of movement over the entire observation period (see text). Tracings in warm colors (red and yellow) represent stacks with a high degree of streaming, tracing in cold colors (blue) represent stacks that are mostly wiggling (compare color scale in C). Note that streaming stacks follow straight, preferred paths. Arrows denote the direction of movement. Cytochalasin D treatment eliminated streaming (C), while nocodazole did not affect directed movement (D). Video sequences can be viewed athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1. Fig. 4. Open in new tabDownload slide Tracing of selected Golgi stacks over 30 s. A, Cortical region in untreated cell. B, Transvacuolar strands in untreated cell. C, Cortical region in cell treated with 40 μm cytochalasin D for 30 min. D, Cortical region in cell treated with 3.3 μm nocodazole for 30 min. The positions of individual Golgi stacks are marked in 1-s intervals. The tracings of the stacks are color coded according to the streaming coefficient of movement over the entire observation period (see text). Tracings in warm colors (red and yellow) represent stacks with a high degree of streaming, tracing in cold colors (blue) represent stacks that are mostly wiggling (compare color scale in C). Note that streaming stacks follow straight, preferred paths. Arrows denote the direction of movement. Cytochalasin D treatment eliminated streaming (C), while nocodazole did not affect directed movement (D). Video sequences can be viewed athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1. The percentage of stacks displaying streaming was higher in cytoplasmic strands than in cortical regions. When stacks were selected at random for tracing analysis (n = 50 per cell, two cells each for cortical regions and transvacuolar strands, respectively), it was found that 15% to 50% of the stacks in cytoplasmic threads had a streaming coefficient greater than 0.2 over the entire observation period, while this fraction was less than 2% for cortical regions (Table I). This difference confirms the observation that large areas of the cortical cytoplasm do not show directed movement (Fig. 4A). Table I. Statistics of Golgi stack movement in cortical regions and cytoplasmic strands of two cells per experiment . Control . 3.3 μmNocodazole . Cortical region . Transvacuolar strand . Cortical region . Transvacuolar strand . Average velocity (μm/s) 0.03–0.46 0.11–1.55 0.09–1.33 0.12–2.37 Maximal velocity (μm/s) 0.34–2.41 0.49–3.85 0.34–2.78 0.49–4.19 % Fast stacks 0–6 4–42 0–7 2–73 % Streaming stacks (30 s) 0, 0 14, 47 0, 2 44, 94 . Control . 3.3 μmNocodazole . Cortical region . Transvacuolar strand . Cortical region . Transvacuolar strand . Average velocity (μm/s) 0.03–0.46 0.11–1.55 0.09–1.33 0.12–2.37 Maximal velocity (μm/s) 0.34–2.41 0.49–3.85 0.34–2.78 0.49–4.19 % Fast stacks 0–6 4–42 0–7 2–73 % Streaming stacks (30 s) 0, 0 14, 47 0, 2 44, 94 Fifty randomly selected stacks in two different cells for either region per treatment (eight cells total) were traced for 30 s. Instantaneous velocities were calculated for every stack at all time intervals. Ranges of maximal and average velocities for all stacks from both cells per region are given. “% Fast stacks” indicates the range of percentages of stacks at any moment that display velocities ≥1 μm/s. “% Streaming stacks” gives the fraction of stacks in the two cells whose movement shows a streaming coefficient ≥ 0.2 over the entire observation period. Open in new tab Table I. Statistics of Golgi stack movement in cortical regions and cytoplasmic strands of two cells per experiment . Control . 3.3 μmNocodazole . Cortical region . Transvacuolar strand . Cortical region . Transvacuolar strand . Average velocity (μm/s) 0.03–0.46 0.11–1.55 0.09–1.33 0.12–2.37 Maximal velocity (μm/s) 0.34–2.41 0.49–3.85 0.34–2.78 0.49–4.19 % Fast stacks 0–6 4–42 0–7 2–73 % Streaming stacks (30 s) 0, 0 14, 47 0, 2 44, 94 . Control . 3.3 μmNocodazole . Cortical region . Transvacuolar strand . Cortical region . Transvacuolar strand . Average velocity (μm/s) 0.03–0.46 0.11–1.55 0.09–1.33 0.12–2.37 Maximal velocity (μm/s) 0.34–2.41 0.49–3.85 0.34–2.78 0.49–4.19 % Fast stacks 0–6 4–42 0–7 2–73 % Streaming stacks (30 s) 0, 0 14, 47 0, 2 44, 94 Fifty randomly selected stacks in two different cells for either region per treatment (eight cells total) were traced for 30 s. Instantaneous velocities were calculated for every stack at all time intervals. Ranges of maximal and average velocities for all stacks from both cells per region are given. “% Fast stacks” indicates the range of percentages of stacks at any moment that display velocities ≥1 μm/s. “% Streaming stacks” gives the fraction of stacks in the two cells whose movement shows a streaming coefficient ≥ 0.2 over the entire observation period. Open in new tab The tracing data were used to calculate the average speed of individual stacks as well as instantaneous velocities. As shown in Figure5, the saltatory movement observed under the microscope is immediately evident in the variations of speed derived from the tracings. Every stack alternated between episodes of rapid movement and periods of relative inactivity. These alterations did not follow any pattern, and Fourier analysis revealed no periodicity (data not shown). The maximal velocity observed for any stack was 4.2 μm/s. Golgi stacks in cortical regions tended to stream more slowly than those in transvacuolar strands (Table I). The variations in instantaneous velocity are also reflected in changes between streaming and wiggling events. Figure 5 exemplifies some of the patterns that could be observed when the streaming coefficient was calculated for 4-s intervals. In particular, some stacks initially displayed a high degree of directional movement (i.e. streaming) that later changed to reduced directionality (i.e. wiggling) (Fig. 5A). Other stacks showed the opposite behavior, with a sudden start of streaming late in the observation period (Fig. 5B). In addition, many stacks showed several reversals of streaming behavior over the 75-s observation period (Fig. 5C). Periods of predominantly translational movement (Fig. 5B, between arrows 2 and 4) could be interrupted by short wiggling events (arrow 3 in Fig. 5B). Fig. 5. Open in new tabDownload slide Variation of instantaneous velocity of individual Golgi stacks in control cells over 75 s. Tracing data of three different stacks (A–C) was used to calculate the instantaneous velocity between the marked positions (broken lines). The streaming coefficient was calculated for a 4-s interval centered on the respective time points (solid lines). Note the rapid changes in velocity that do not follow any predictable pattern. Also note that the streaming coefficient of the movement can change over time. Low values for the streaming coefficient indicate wiggling motion; high values indicate streaming motion. Corresponding points on the tracing and the graph are marked with numbered arrows. Fig. 5. Open in new tabDownload slide Variation of instantaneous velocity of individual Golgi stacks in control cells over 75 s. Tracing data of three different stacks (A–C) was used to calculate the instantaneous velocity between the marked positions (broken lines). The streaming coefficient was calculated for a 4-s interval centered on the respective time points (solid lines). Note the rapid changes in velocity that do not follow any predictable pattern. Also note that the streaming coefficient of the movement can change over time. Low values for the streaming coefficient indicate wiggling motion; high values indicate streaming motion. Corresponding points on the tracing and the graph are marked with numbered arrows. The movement of Golgi stacks showed a high degree of specificity. Rapidly moving stacks were often seen to pass slower stacks in close proximity (e.g. Fig. 6). The differences in instantaneous velocities could be as high as 2.2 μm/s for stacks that were less than 1 μm apart (Fig.7). When stacks were seen in an edge-on view as they were traveling through the cytoplasm, the edge was usually aligned with the direction of movement and maintained this orientation over the entire streaming event (double lines in Fig. 6). Fig. 6. Open in new tabDownload slide Stacks maintain their orientation during streaming events. Sequence of 12 video frames from cortical cytoplasm taken in 1-s intervals. General direction of movement is from lower right to upper left. The stack marked with a double line maintains its orientation from 2 to 8 s and from 10 to 12 s. At 9 s it appears to dive under a slowly moving stack and its orientation cannot be resolved unambiguously. The Golgi stack marked with the arrow first shows a rotational movement (2–4 s), followed by translational movement to the end of the sequence. Asterisks denote a stack that did not move during the entire observation period. Fig. 6. Open in new tabDownload slide Stacks maintain their orientation during streaming events. Sequence of 12 video frames from cortical cytoplasm taken in 1-s intervals. General direction of movement is from lower right to upper left. The stack marked with a double line maintains its orientation from 2 to 8 s and from 10 to 12 s. At 9 s it appears to dive under a slowly moving stack and its orientation cannot be resolved unambiguously. The Golgi stack marked with the arrow first shows a rotational movement (2–4 s), followed by translational movement to the end of the sequence. Asterisks denote a stack that did not move during the entire observation period. Fig. 7. Open in new tabDownload slide Movement of several Golgi stacks along one track in cortical cytoplasm of a control cell. A, Positions of individual stacks were marked in 1-s intervals for up to 30 s. Tracing coordinates were transformed so that the abscissa runs along the track, and the ordinate lies perpendicular to it. B, Temporal relationship between stacks that are at the same position along the strand. Note that stacks that are in close proximity (less than 1-μm distance, arrows in A and circles in B) can have drastically different velocities. Fig. 7. Open in new tabDownload slide Movement of several Golgi stacks along one track in cortical cytoplasm of a control cell. A, Positions of individual stacks were marked in 1-s intervals for up to 30 s. Tracing coordinates were transformed so that the abscissa runs along the track, and the ordinate lies perpendicular to it. B, Temporal relationship between stacks that are at the same position along the strand. Note that stacks that are in close proximity (less than 1-μm distance, arrows in A and circles in B) can have drastically different velocities. When individual cells were observed for longer periods of time, it was found that the level of activity varied (compare E and F in Fig.8) and that the regions of the cell with streaming tracks also shifted laterally (compare Fig. 8, D and E). The phenomenon of variable streaming activity was quantified by counting the number of tracks with active streaming in cells that were kept in a perfusion chamber with constant supply of fresh, aerated growth medium (flow rate 0.5 mL/min). Figure 9A shows typical examples for this variability in streaming activity. Cells usually maintained a fairly constant level of streaming activity, which was interrupted by short episodes of reduced activity. Interestingly, sister cells that were connected to each other sometimes displayed parallel fluctuations in activity level (cells A1 and A4 in Fig. 9A), suggesting that a global signal influencing cytoplasmic streaming can travel through plasmodesmata. Most cells also showed a decrease in activity over the course of an experiment (approximately 2 h), with some of them losing streaming activity during the observation period (e.g. cells A1 and A2 in Fig. 9B). Fig. 8. Open in new tabDownload slide Shift of regions with active streaming in cortical cytoplasm. Ten-second video sequences of the same cortical region taken in 10-min intervals were analyzed for movement of Golgi stacks. A to C, Single enhanced video images of cortical region in 10-min intervals. D to F, Movement analysis of the video sequences corresponding to A to C, respectively. The positions of Golgi stacks were automatically detected using a peak-finding algorithm. Positions where stacks were detected most of the time (slowly moving and wiggling stacks) are coded in light gray and as hollow spots; positions where stacks were detected only rarely (fast-moving stacks) are coded in black. Video sequences can be viewed athttp://www.plantphysiol.org/cgi/content/full/ 121/4/1127/DC1. Fig. 8. Open in new tabDownload slide Shift of regions with active streaming in cortical cytoplasm. Ten-second video sequences of the same cortical region taken in 10-min intervals were analyzed for movement of Golgi stacks. A to C, Single enhanced video images of cortical region in 10-min intervals. D to F, Movement analysis of the video sequences corresponding to A to C, respectively. The positions of Golgi stacks were automatically detected using a peak-finding algorithm. Positions where stacks were detected most of the time (slowly moving and wiggling stacks) are coded in light gray and as hollow spots; positions where stacks were detected only rarely (fast-moving stacks) are coded in black. Video sequences can be viewed athttp://www.plantphysiol.org/cgi/content/full/ 121/4/1127/DC1. Fig. 9. Open in new tabDownload slide Level of streaming activity of individual cells over time. Individual cells were observed every 5 min, and the number of strands with active streaming in the entire cell were counted. Times are given in minutes from the start of perfusion. In some cases, strands could not be classified unambiguously, since only single stacks showed directed movement or the velocity was slow. A, Activity levels in control cells. ▪, A1; ■, A4; ▴, B1; ▵, B2. B, Activity levels prior, during, and after treatment with 30 mmbutanedione monoxime. ▾, A1; ▿, A2; ▵, B1; ●, C1; ▪, D1. C, Activity levels prior, during, and after treatment with 10 μm propyzamide. ○, A1; ●, A2; ▪, B1; ▴, C1; ▵, C3. Shaded areas indicate the duration of drug treatments. Cells coded with the same letter are sister cells that share a cell wall. Differences in basal activity are usually related to cell size, with smaller cells showing less Golgi streaming. Fig. 9. Open in new tabDownload slide Level of streaming activity of individual cells over time. Individual cells were observed every 5 min, and the number of strands with active streaming in the entire cell were counted. Times are given in minutes from the start of perfusion. In some cases, strands could not be classified unambiguously, since only single stacks showed directed movement or the velocity was slow. A, Activity levels in control cells. ▪, A1; ■, A4; ▴, B1; ▵, B2. B, Activity levels prior, during, and after treatment with 30 mmbutanedione monoxime. ▾, A1; ▿, A2; ▵, B1; ●, C1; ▪, D1. C, Activity levels prior, during, and after treatment with 10 μm propyzamide. ○, A1; ●, A2; ▪, B1; ▴, C1; ▵, C3. Shaded areas indicate the duration of drug treatments. Cells coded with the same letter are sister cells that share a cell wall. Differences in basal activity are usually related to cell size, with smaller cells showing less Golgi streaming. Molecular Basis of Golgi Movement To determine the molecular basis for the observed movement, the streaming behavior of Golgi stacks was investigated after treatment with different drugs that disrupt specific components of the cytoskeleton. Golgi stacks in cells that had been treated with 40 μm cytochalasin D for 30 min to disrupt actin filaments were still wiggling, but showed essentially no translational movement (see tracings in Fig. 4C). Occasionally, single stacks were observed that followed a clear trajectory (not shown). We attribute these few streaming stacks to residual actin filaments that were not affected by the drug. Latrunculin A, another actin-filament-disrupting drug, also inhibited movement of Golgi stacks. At the concentration used (0.1 μm), latrunculin A appeared more potent in that no moving stacks were observed (data not shown). Movement was also reversibly inhibited by 30 mm BDM, an inhibitor of the myosin ATPase (Herrmann et al., 1992; Fig. 9B). All drug effects could be reversed by washing with growth medium; however, most cells were not able to recover fully. It is unclear whether this reduced level of streaming activity reflects normal decrease in cell viability during prolonged observation (see above) or some irreversible component of the drug effect. While the inhibition of directed movement by actin-disrupting drugs was usually accompanied by a breakdown of transvacuolar strands, BDM did not show such an effect (not shown). A number of microtubule-disrupting drugs was tested, namely nocodazole (at 3.3 μm), colchicine (at 250 μm), and propyzamide (at 6 μm). All had similar effects on the streaming behavior. Here we report data mostly from the nocodazole experiments. Casual observation of nocodazole-treated cells under the microscope revealed that microtubule-disrupting drugs did not inhibit streaming. Instead we noticed an apparent slight increase in streaming activity over control cells. The percentage of cells with actively streaming stacks was increased to a small extent (Table II). This difference, albeit small, is statistically significant when the results from matched drug and control treatments are compared (pairedt test, P < 0.004). Table II. Fraction of cells with active streaming in a population (%) Experiment . 3.3 μm Nocodazole . Control . Ratio . 1 81 74 1.09 2 83 82 1.01 3 84 84 1.00 4 77 64 1.20 5 73 64 1.14 6 78 57 1.37 7 68 60 1.13 8 81 78 1.04 9 53 46 1.15 10 61 56 1.09 Mean ±sd 74 ± 11 67 ± 13 1.12 ± 0.11 Experiment . 3.3 μm Nocodazole . Control . Ratio . 1 81 74 1.09 2 83 82 1.01 3 84 84 1.00 4 77 64 1.20 5 73 64 1.14 6 78 57 1.37 7 68 60 1.13 8 81 78 1.04 9 53 46 1.15 10 61 56 1.09 Mean ±sd 74 ± 11 67 ± 13 1.12 ± 0.11 Aliquots of a suspension cultures were treated with 3.3 μm nocodazole or 0.1 % (w/v) DMSO (control). After about a 45-min incubation, 100 cells per treatment were classified as displaying active streaming or not. Ten independent experiments were performed. While the average fraction of cells displaying active streaming was not significantly changed by the drug treatment, a consistent small increase was observed in most experiments (P = 0.004, paired t test). Open in new tab Table II. Fraction of cells with active streaming in a population (%) Experiment . 3.3 μm Nocodazole . Control . Ratio . 1 81 74 1.09 2 83 82 1.01 3 84 84 1.00 4 77 64 1.20 5 73 64 1.14 6 78 57 1.37 7 68 60 1.13 8 81 78 1.04 9 53 46 1.15 10 61 56 1.09 Mean ±sd 74 ± 11 67 ± 13 1.12 ± 0.11 Experiment . 3.3 μm Nocodazole . Control . Ratio . 1 81 74 1.09 2 83 82 1.01 3 84 84 1.00 4 77 64 1.20 5 73 64 1.14 6 78 57 1.37 7 68 60 1.13 8 81 78 1.04 9 53 46 1.15 10 61 56 1.09 Mean ±sd 74 ± 11 67 ± 13 1.12 ± 0.11 Aliquots of a suspension cultures were treated with 3.3 μm nocodazole or 0.1 % (w/v) DMSO (control). After about a 45-min incubation, 100 cells per treatment were classified as displaying active streaming or not. Ten independent experiments were performed. While the average fraction of cells displaying active streaming was not significantly changed by the drug treatment, a consistent small increase was observed in most experiments (P = 0.004, paired t test). Open in new tab While the number of cells with active streaming was increased by microtubule-disrupting drugs, no consistent effect on the activity levels of individual cells could be observed (Fig. 9C), although some cells seemed to show an increase in streaming activity during the drug treatment (e.g. cell A2 in Fig. 9C). Analysis of tracing data for four nocodazole-treated cells (two cortical regions, two thread regions, 50 stacks per cell) showed no significant differences from control cells (Table I). Some of the parameters analyzed appeared elevated in drug-treated cells (e.g. percent streaming stacks in the transvacuolar strands of one of the cells; Table I), but the high degree of variability for the control cells precludes any firm conclusions. From these experiments it can be concluded that the directed movement of Golgi stacks in plant cells requires intact actin filaments and is probably propelled by myosin motors. Microtubules do not appear to have an effect on movement, except for a subset of cells in which they seem to limit streaming. DISCUSSION The Plant α-1,2 Mannosidase Is Homologous to Corresponding Animal and Fungal Enzymes We have isolated a cDNA from soybean, GmMan1, encoding a protein with a high degree of similarity to animal and fungal class I α-1,2 mannosidases. While the final confirmation of this assignment has to await demonstration of the appropriate enzymatic activity, it appears that the strong conservation of key features found in other mannosidases supports the conclusion that the isolated message encodes this enzymatic function in soybean (Fig. 1). To our knowledge, this is the first report of an enzyme of the N-linked oligosaccharide processing pathway cloned from plants. However, an Arabidopsis expressed sequence tag with 80% similarity toGmMan1 has been reported previously (GenBank accession no.W43154). GmMan1 shows the typical type II orientation of other Golgi membrane proteins, with a short cytoplasmic tail at its amino terminus followed by a single transmembrane domain. This region, as well as the following putative stalk domain, show little similarity to other α-1,2 mannosidases at the sequence level. In contrast, the catalytic domain is 45% identical in amino acid sequence to either mouse or yeast α-1,2 mannosidase and contains all residues that were shown in other species to be essential for enzymatic function (Fig. 1). This striking distribution of sequence conservation is also found between the α-1,2-mannosidase homologs of other species (Herscovics, 1999a). The soybean cDNA described in this report therefore displays all the hallmarks of known α-1,2 mannosidases. The high degree of conservation in the catalytic domain is contrasted by the near complete absence of sequence similarity at the N terminus of the protein. This part of the protein is most likely responsible for correct localization of the mannosidase (for review, see Colley, 1997). The lack of sequence conservation led to the formulation of two models that explain the targeting/retention of Golgi proteins in terms of structural features. According to one model, Golgi proteins with their typically short transmembrane helices preferentially partition into the thinner membranes of the early secretory pathway (Pelham and Munro, 1993). The other model proposes a “kin-recognition” mechanism, where resident proteins of the same compartment can physically interact and thereby maintain their specific localization (Nilsson et al., 1993). The sequence of GmMan1 is compatible with both models. The predicted membrane-spanning domain is unusually short, only 16 amino acids. At the same time, the stalk region has a high probability of forming a coiled-coil structure, which is often indicative of protein-protein interactions. Interestingly, two recent reports describe proper targeting of mammalian sialyltransferase to the trans-Golgi in plant cells (Boevink et al., 1998; Wee et al., 1998), suggesting that the targeting mechanism of Golgi oligosaccharide processing enzymes could be a common structural motif conserved between animals and plants. GmMan1::GFP Fusion Can Be Used as an in Vivo Marker for cis/Medial Golgi Cisternae The fusion protein of GmMan1 to GFP is localized to the cis side of plant Golgi stacks of suspension-cultured tobacco BY-2 cells (Fig.3) and can therefore act as a marker of Golgi stack localization in living plant cells. The Golgi stacks of BY-2 cells appear to be randomly distributed throughout the cytoplasm. This is consistent with predictions based on electron micrographs (Robinson and Kristen, 1982) and observations made in a number of other systems. For example, similar patterns of fluorescent dots have been observed in chemically fixed maize root cells stained with the monoclonal antibody JIM84 (Satiat-Jeunemaitre and Hawes, 1992), which recognizes Golgi- and plasma membrane-specific Lewis a type epitopes (Fitchette-Lainéet al., 1997), as well as in transgenic Arabidopsis cells expressing an epitope-tagged sialyltransferase (Wee et al., 1998) and in tobacco leaf epidermal cells transiently expressing GFP fusions of sialyltransferase and the KDEL receptor protein AtERD2 (Boevink et al., 1998). Occasionally, we could observe cells that had a green fluorescent ER in addition to Golgi stacks (data not shown). It is not clear whether this reflected a bottleneck in the export from the ER or an increased rate of retrograde transport from the “saturated” Golgi. A few cells also contained Golgi-sized fluorescent structures that appeared as small rings (Fig. 2C). Similar structures have been observed when a GFP-tagged sialyltransferase was transiently expressed in tobacco leaf epidermis cells; the images were interpreted as edge-on views of highly curved stacks (Boevink et al., 1998). The observation that the rings can change into lines with a length similar to that of the ring diameter (compare video sequence athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1), however, suggests that they represent face-on views of Golgi stacks in which the fusion protein is concentrated in the rims of the cisternae. This distribution could be caused by conditions that lead to an osmotic collapse of the cisternal lumen and the exclusion of the bulky fusion proteins from the central region of the cisternae where the membranes became appressed. This interpretation could also explain the distribution of the sialyltransferase-GFP fusion construct, since the trans-cisternae in cryo-fixed and freeze-substituted cells typically exhibit a collapsed central domain with minimal intracisternal volume (Staehelin et al., 1990). The Saltatory Movement of Plant Golgi Stacks Is Acto-Myosin Based The Golgi stacks of BY-2 cells usually alternate between episodes of random “wiggling” motion reminiscent of Brownian motion and directed movement along linear tracks. The wiggling periods can last from seconds to minutes and during such periods individual stacks can “drift” several micrometers without participating in any directed translocation events. In contrast, the stacks displaying directed movement appear to follow the same tracks as cytoplasmic streaming (Williamson, 1993). This was confirmed by staining mitochondria with a rhodamine-labeled dye (MitoTracker, Molecular Probes) and observing the movement of the two organelles in the same cell. Streaming Golgi stacks and mitochondria typically traveled along the same paths and also showed qualitatively similar stop-and-go movement (data not shown). Individual Golgi stacks followed more or less straight trajectories, often with several stacks following the same track (Fig. 7). Progress along these tracks was intermittent, so individual stacks showed stop-and-go movement (Figs. 5 and 7). Interestingly, stacks moving along the same track often paused at the same position, suggesting that these sites can inhibit movement (see below). To determine the molecular basis for this saltatory motion, BY-2 cells were treated with a number of inhibitors that interfere with normal functioning of cytoskeletal elements. Drugs that disrupt the actin cytoskeleton (cytochalasin D, latrunculin A) also stopped the streaming of Golgi stacks (Fig. 4C). “Wiggling” motion usually was not affected. Cytochalasin D used at 40 μm often did not lead to a complete block of directed movement, and in some instances produced localized circular movements. We assume that this drug concentration did not disrupt all actin filaments and that the few remaining filaments allowed Golgi stack movement to occur. Streaming of Golgi stacks therefore requires intact actin microfilaments. A similar conclusion was reached by Boevink et al. (1998). Interestingly, these authors found a tight co-localization of actin filaments and tubular ER, suggesting that the movement of Golgi stacks occurs in close association with the ER. This is consistent with the finding ofLichtscheidl et al. (1990) that actin filament bundles in the epidermal cells of Drosera tentacles often co-localize with tubular ER cisternae. In our transformed BY-2 cells this ER-Golgi streaming relationship was less evident. The involvement of myosin motors in this movement was tested by applying the myosin inhibitor BDM, which has been shown to inhibit the ATPase function on isolated skeletal muscle myosin II (Herrmann et al., 1992). Movement of Golgi stacks could be stopped by adding 30 mm BDM to the perfusion medium of the cells (Fig. 9B), suggesting that myosin provides the motive force for the observed motion. This conclusion is consistent both with observations that have implicated myosin as the motor for cytoplasmic streaming (Shimmen and Yokota, 1994) and with reports in which the translocation of organelles in pollen tubes has been linked to myosin (Kohno and Shimmen, 1988). We assume that the movement of plant Golgi stacks occurs by active translocation along actin filaments as opposed to passive drift in a general cytoplasmic stream. This interpretation is based on, among others, on the high degree of specificity observed in the movement of individual stacks (see e.g. Fig. 6). This implies that Golgi movement is mediated by a specific myosin, presumably associated with the Golgi-matrix that surrounds the individual Golgi stack-TGN units. Microtubule Disruption Does Not Inhibit Golgi Saltatory Movements In animal cells, the localization of the Golgi complex and the movement of transport carriers to and from the Golgi depend on the presence of intact microtubules (Presley et al., 1997; Scales et al., 1997; Burkhardt, 1998; Hirschberg et al., 1998; Toomre et al., 1999). Therefore, we tested the role of microtubules in directed movement of plant Golgi stacks. Treatment of BY-2 tobacco cells with 3.3 μm nocodazole, 250 μm colchicine, or 6 μm propyzamide did not inhibit streaming (Figs. 4D and9C; data not shown), indicating that microtubules are not required for the movement of Golgi stacks. Quite unexpectedly, we noticed an apparent increase in streaming activity during the drug treatments. Such a stimulatory effect of microtubule-disrupting drugs on cytoplasmic streaming has to our knowledge not been described before. Careful analysis of a variety of streaming parameters revealed that the enhanced streaming effect of microtubule disruptors is brought about by increasing the percentage of cells displaying saltatory Golgi movement (Table II). Movement of individual stacks was unaffected with respect to average and maximal velocities, maximal accelerations, and changes between streaming and wiggling events. A few cells appeared to have a higher percentage of streaming stacks after drug treatment (Fig. 9C, cell A2; Table I, cell 2 of transvacuolar strands), while the other cells were not affected. It is not known whether the cells with higher activity are examples for the increase in number of cells with streaming Golgi stacks (see Table II), or whether they represent extremes within the normal variability that was also seen in control cells. It is unclear how disruption of microtubules can trigger cytoplasmic streaming in a subset of cells. One possibility is that cells in a certain phase of the cell cycle have reduced streaming, and that this inhibition could be mediated in part by microtubules. Disruption of the microtubule scaffold would then release the constraints on the Golgi translocation machinery. For example, it is known that cytoplasmic streaming ceases during mitosis (Mineyuki et al., 1984). However, the percentage of cells in mitosis is too small to account for the observed increase in numbers of cells with streaming. Therefore, we speculate that Golgi movement may also be constrained during another stage of the cell cycle. We are currently investigating this possibility in synchronized cells. Is the Stop-and-Go Movement of Plant Golgi Stacks a Regulated Process? A Hypothesis The rapid stop-and-go movement of plant Golgi stacks raises the question of how the secretory pathway in plant cells can function efficiently. One of the consequences of the streaming behavior of Golgi stacks is that there is no fixed spatial relationship between the ER and Golgi. The variable spatial orientation of Golgi stacks to nearby ER cisternae has been known to plant cell biologists for over 20 years (for review, see Robinson and Kristen, 1982), but the mechanistic basis for this variability has remained an enigma. This report and that byBoevink et al. (1998) help to explain this apparent random distribution as a steady-state intermediate of continuously moving stacks. At the same time, these findings raise the question of whether vesicle trafficking between the ER and Golgi is based strictly on random encounters, or if it is regulated by mechanisms yet to be discovered. The observation that in leaf epidermal cells Golgi stacks usually track along the well-defined ER elements underlying the plasma membrane has led to the suggestion that the Golgi stacks act as “vacuum cleaners” that move around to pick up products from the ER (Boevink et al., 1998). However, this movement was predominantly observed along tubular ER strands (Boevink et al., 1998), which may not be very active in protein synthesis. It is also difficult to envision the targeting of ER-derived transport vesicles to cis-Golgi cisternae when the stacks travel at speeds most likely greater than vesicle diffusion rates. In the model depicted in Figure 10, we offer an alternative hypothesis, in which the stop-and-go motion of Golgi stacks is postulated to be regulated to increase the efficiency of ER-to-Golgi transport, as well as the delivery of secretory products to specific cell wall domains. Fig. 10. Open in new tabDownload slide Model of regulated stop-and-go movement of Golgi stacks and its relationship to the transport of products through the secretory pathway. A, Golgi stacks with myosin motors attached to the Golgi matrix move along actin filaments. B, Activation of ER exit sites may release a local signal that inhibits movement of Golgi stacks and allows for uptake of ER-to-Golgi transport vesicles. C, Local cell wall expansion or sites of secondary wall thickenings may lead to a signal resulting in stopping of Golgi stack movement and to the release of Golgi-derived secretory vesicles. Fig. 10. Open in new tabDownload slide Model of regulated stop-and-go movement of Golgi stacks and its relationship to the transport of products through the secretory pathway. A, Golgi stacks with myosin motors attached to the Golgi matrix move along actin filaments. B, Activation of ER exit sites may release a local signal that inhibits movement of Golgi stacks and allows for uptake of ER-to-Golgi transport vesicles. C, Local cell wall expansion or sites of secondary wall thickenings may lead to a signal resulting in stopping of Golgi stack movement and to the release of Golgi-derived secretory vesicles. Our model postulates that active ER export sites produce a localized signal that leads to the uncoupling of nearby Golgi stacks from the actin tracks and to their pausing in the vicinity of the activated ER export site (Fig. 10B). This would increase the efficiency of ER-to-Golgi (and Golgi-to-ER) trafficking. Upon completion of the transfer, the ER stop signal would be turned off, allowing the stacks to resume their movement. A similar stop signal may be produced by regions where Golgi products are required, such as areas of wall expansion or sites of secondary cell wall thickenings (Fig. 10C). It is well-known that secretion can be directed to specific cell wall sites (Fowler and Quatrano, 1997), and a patch-like distribution of secretory vesicle profiles in freeze-fracture images of plasma membranes of root tip and cultured cells has been reported (Staehelin and Chapman, 1987;Craig and Staehelin, 1988). The gradual shifting of Golgi streaming domains over larger time intervals (Fig. 8) may help to ensure an even deposition of cell wall products over time. A potential candidate for the postulated stop signal is calcium, since it is known that elevated calcium concentrations can block cytoplasmic streaming (Shimmen and Yokota, 1994) by a calmodulin-mediated inhibition of myosin (Yokota et al., 1999). Such sites of localized inhibition of movement may be recognized by studying the behavior of several stacks within one streaming strand. Indeed, we have found positions along a strand where two or three sequentially arriving stacks stop briefly before moving on (compare videos athttp://www.plantphysiol.org/cgi/content/full/121/4/1127/DC1). Of course, this circumstantial evidence is not conclusive, as discontinuities in actin filaments could cause similar effects. We are currently initiating experiments to address this question more directly. ACKNOWLEDGMENTS We are indebted to Dr. Annette Herscovics (McGill University, Montreal) for providing the partial soybean clone and insightful comments concerning the manuscript. 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Plant Physiol 119 1999 231 239 Google Scholar Crossref Search ADS PubMed WorldCat Author notes 1 This work was supported by the National Institutes of Health (grant no. GM18639) to L.A.S. * Corresponding author; e-mail [email protected]; fax 303–492–7744. 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)

Hueros, Gregorio; Gomez, Elisa; Cheikh, Nordine; Edwards, Janice; Weldon, Marcia; Salamini, Francesco; Thompson, Richard D.

doi: 10.1104/pp.121.4.1143pmid: 10594101

Abstract The maize (Zea maysL.) betl1 locus, encoding a basal endosperm transfer layer-specific protein, has been mapped and molecularly cloned in its entirety. The locus is shown to consist of three gene copies in the maize inbred line A69Y. To distinguish the three transcription units from the locus name, we have termed them BETL1a, BETL1b,and BETL1c. Two of the copies are expressed, whereas one is inactive and contains retrotransposon-like insertions in both promoter and intron regions. Based on this information, and a restriction site map covering 17 kb around the BETL1locus, a DNA fragment putatively containing an active promoter sequence was identified. This fragment was tested for its ability to confer transfer-cell-specific expression in transient and stably transformed maize tissues. The transgenic maize plants obtained showed the predicted cell-type specificity of expression restricted to the basal endosperm transfer cells, although there were minor deviations in promoter strength and timing and accumulation of the transgene product from the corresponding BETL-1 endogene expression pattern. The endosperm is the main storage organ in maize seeds, nourishing the embryo while the seed develops and providing nutrients to the seedling on germination. Thus, the uptake of assimilates by the growing endosperm is a critical process in seed development. There are no symplastic connections between maternal and embryonic tissues (Thorne, 1985); instead, phloem unloading releases nutrients into the apoplastic compartment of the pedicel. Uptake of nutrients by the endosperm from the pedicel is facilitated by the conversion of the cells at the base of the endosperm to transfer cells (Davis et al., 1990). Basal endosperm transfer cells possess anatomical modifications such as extensive cell wall ingrowths, which increase the membrane surface area and therefore the solute transport capacity (Pate and Gunning, 1972). The absence of this layer is correlated with reduced rates of grain filling and eventual abortion of the seeds (Brink and Cooper, 1947). The endosperm is a triploid tissue, mainly consisting of two cell types, the central endosperm cells, which accumulate starch and proteins, and outer aleurone cells at the periphery of the endosperm. Aleurone cell differentiation takes place between 6 and 10 d after pollination (DAP), producing a single cell layer of small cuboidal cells that accumulate spherosomes and protein bodies. Endosperm transfer cells develop instead of the aleurone cell phenotype in the basal endosperm cell layer bordering the pedicel. The most basal layer (also termed “modified aleurone”) consists of a sheet of elongated cells densely covered on the basal surface by cell wall ingrowths. Two or three adjacent endosperm cell layers also possess cell wall ingrowths, successively decreasing in extent toward the center of the kernel. (Schel et al., 1984; Shannon et al., 1986; Davis et al., 1990). To date, physiological and cytological studies have been carried out on the endosperm transfer cell layer, but little is known about the regulation of development of this cell type. The lack of development of endosperm transfer cells in 4n endosperm has been attributed to a possible regulatory effect of genomic imprinting (Charlton et al., 1995). A number of genes expressed specifically in different tissues of the seed have been isolated. The corresponding promoter sequences have been functionally analyzed, and sequences conferring tissue specificity in starchy endosperm (Thomas and Flavell, 1990; Thompson et al., 1990; Quayle and Feix, 1992; Muller and Knudsen, 1993) and aleurone (Leah et al., 1994; Kalla et al., 1996) cells have been identified. In contrast, only two transfer-cell-specific cDNAs have been reported, BET1 in maize (Hueros et al., 1995, synonymous with BETL1 [basal endosperm transfer layer-specific protein] in this article) and END1 in barley (Doan et al., 1996). The basis for transfer-cell-specific expression is currently unknown. We report here the identification of functional promoter sequences for the BETL1 gene. A genomic fragment of 9 kb isolated from the maize line A69Y contained three tandem copies of the gene present at the betl1 locus. On sequencing, the genes were found to be embedded in a complex array of transposon relic and satellite DNA repeats. A search for transcripts derived from each of the three BETL1 genes demonstrated that only two of the three copies were functionally active. Based on these data, we defined a promoter region having a maximum length of 1.8 kb. Particle bombardment experiments showed that a 930-bp promoter fragment proximal to the coding sequence was sufficient to drive β-glucuronidase (GUS) expression in the endosperm transfer cell layer. Finally, maize transgenic plants containing the 930-bp promoter region fused to GUS directed reporter gene expression in a manner resembling that of theBETL1 transcript, both in timing and cellular localization. MATERIALS AND METHODS Plant Material Maize (Zea mays L. cv A69Y) shoots, leaves, immature seeds, tassels, and roots were extracted either from greenhouse-grown plants or from seedlings germinated on moist filter paper in Petri dishes. Plant Transformation Transgenic maize plants transformed with vector pMON17570 containing the BETL1 promoter sequence fused to the GUS gene were produced using microprojectile bombardment (Klein et al., 1988; Gordon-Kamm et al., 1990; Walters et al., 1992). Embryogenic callus initiated from immature maize embryos was used as a target tissue. Plasmid DNA at 1 mg mL−1 in Tris-EDTA (TE) buffer was precipitated onto M10 tungsten particles using a calcium chloride plus spermidine procedure, essentially as described byKlein et al. (1988). The plasmids also contained the neomycin phosphotransferase II gene (nptII) driven by the 35S promoter from cauliflower mosaic virus and fused to the HSP70 intron. The gene of interest and the selectable marker were transcribed in the same direction. Embryogenic callus target tissue was pretreated on culture medium osmotically buffered with 0.2 mmannitol plus 0.2 m sorbitol for approximately 4 h prior to bombardment (Vain et al., 1993). Tissue was bombarded two times with DNA-coated tungsten particles using the gunpowder version of the particle delivery system (PDS 1000 device, Bio-Rad, Hercules, CA). Approximately 16 h after bombardment, tissue was subcultured onto a medium minus mannitol and sorbitol, but containing an appropriate aminoglycoside antibiotic, e.g. G418 to select for cells expressing the 35S/nptII gene. Actively growing tissue sectors were transferred to fresh selection medium approximately every 3 weeks. About 3 months after bombardment, plants were regenerated from surviving embryogenic callus essentially as described by Duncan and Widholm (1988). Isolation of RNA and DNA and Analysis by Filter Hybridization Standard methods of DNA and RNA manipulation were carried out as described previously (Hueros et al., 1995). For the filter hybridizations presented in Figure 5, digoxigenin PCR-generated probes were labeled with digoxigenin, as recommended by the supplier (Boehringer Mannheim, Mannheim, Germany), CPD-Star was used as the substrate for the alkaline phosphatase (Boehringer Mannheim), and the filters were exposed for 30 min unless otherwise stated. Inverse-PCR Reactions To clone the genomic sequences immediately upstream of the coding region contained in the cDNA clone, a protocol based on that designed for genomic sequencing (Pfeifer et al., 1989) was used. Genomic DNA was digested with XbaI, denatured, and annealed to an 18-mer oligonucleotide derived from the cDNA sequence. The oligonucleotide was used in a primer extension reaction using Sequenase (Amersham-Pharmacia, Buckinghamshire, UK). The resulting blunt ends were ligated to an unphosphorylated synthetic double-stranded linker. The genomic sequences located between the restriction sites and the 5′ end of the cDNA were subsequently amplified by PCR using primers from the cDNA and synthetic linkers. Transient Expression Analysis Maize kernels harvested 10 DAP were surface-sterilized and hand-dissected to isolate the endosperm. Endosperms were maintained in the solid medium described below until being bombarded with DNA-coated gold particles. The coating and bombardment was performed according to the method of Knudsen and Müller (1991). After bombardment, tissue samples were incubated at 25°C for 24 h in the dark, in solid (0.5% [w/v] agarose) Murashige and Skoog medium containing 100 mg L−1 myo-inositol, 2 g L−1 Asn, 2 g L−1Gln, 30 g L−1 Suc, and Murashige and Skoog vitamins (Sigma, St. Louis). GUS Assay Expression of the GUS gene was detected by histochemical staining according to the method of Jefferson et al. (1987). In vitro-cultured endosperms and fresh tissues were stained for GUS in a medium containing: 0.5 mg mL−1 X-glucuronide (CLONTECH, Palo Alto, CA), 0.5 mmK+-ferrocyanide, 0.5 mmK+-ferricyanide, 10 mmNa2EDTA, 50 mm phosphate buffer (pH 7.0), and 0.1% (w/v) Triton X-100. A blue background, caused by endogenous glucuronidase activity observed in the pedicel, was eliminated by including 20% (v/v) methanol in the staining solution. Immunological Quantification of Proteins in Transgenic Kernel Extracts Immature kernels were removed, cut longitudinally, and half-kernels were stained for GUS activity as above. The remaining half-kernels from GUS-positive samples were ground in 100 μL of 3× SDS-PAGE loading buffer (Rotiload, Roth, Karlsruhe, Germany) and centrifuged in an Eppendorf minifuge at 13,000 rpm for 5 min. The supernatant (30 μL) was fractionated by 15% (w/v) SDS-PAGE, electroblotted onto polyvinylidenefluoride (PVDF, Millipore, Bedford, MA) membrane, and proteins detected by enhanced chemiluminescence (ECL, Amersham, Little Chalfont, UK) using antibodies raised to BETL1 (Hueros et al., 1995), immunophilin mzFKBP66 (Hueros et al., 1998), and GUS (A-5790, Molecular Probes, Eugene, OR). Images were quantitated using a CCD recording camera (Lumimager, Boehringer Mannheim) RESULTS BETL1 Is Present in Three Copies in the Maize Genome Analysis of the organization of BETL1 coding sequences by Southern hybridizations indicated that the locus probably consisted of a short array of three copies of the gene (Hueros et al., 1995). To confirm this hypothesis, a gene copy number reconstruction was carried out by serially diluting BETL1 plasmid DNA with maize genomic DNA (Fig. 1). Aliquots (8 μg) of HindIII-digested genomic DNA (corresponding to approximately 1 million maize nuclei) were mixed with increasing amounts of linearized plasmid containing the BETL1 cDNA. After Southern blotting, the filter was hybridized to aBETL1 probe. The genomic band intensity seen was equal to the intensity of 8.9-pg plasmid DNA (Fig. 1, arrowhead), which is equivalent to 2.4 million copies. Therefore, each HindIII genomic band contains at least one copy of BETL1 per haploid genome. The existence of more than one copy was further confirmed by physical mapping of the restriction sites upstream of the coding sequence. For this purpose, double digests including NsiI (which cuts at the 3′ end of the cDNA) and various other restriction enzymes were blotted and hybridized to a 5′-specific cDNA probe (data not shown). Fig. 1. Open in new tabDownload slide Three copies of the BETL1 gene are present in maize variety A69Y. HindII-digested genomic DNA (5 μg [lane 1] or 8 μg [lanes 2–5]) was run along with increasing amounts of plasmid DNA containing the BETL1cDNA. Lane 1, 0 pg of plasmid DNA; lane 2, 2.22 pg; lane 3, 4.44 pg; lane 4, 8.88 pg; lane 5, 17.76 pg. The resulting Southern blot was hybridized with a BETL1 cDNA probe. Genomic fragments are located just below the 8-, 6-, and 2-kb markers, with theBETL1 plasmid DNA control located at 3.5 kb (arrowhead). Fig. 1. Open in new tabDownload slide Three copies of the BETL1 gene are present in maize variety A69Y. HindII-digested genomic DNA (5 μg [lane 1] or 8 μg [lanes 2–5]) was run along with increasing amounts of plasmid DNA containing the BETL1cDNA. Lane 1, 0 pg of plasmid DNA; lane 2, 2.22 pg; lane 3, 4.44 pg; lane 4, 8.88 pg; lane 5, 17.76 pg. The resulting Southern blot was hybridized with a BETL1 cDNA probe. Genomic fragments are located just below the 8-, 6-, and 2-kb markers, with theBETL1 plasmid DNA control located at 3.5 kb (arrowhead). Inverse-PCR Cloning of BETL-1 Genes To clone the sequences upstream of the coding regions, inverse-PCR (I-PCR) was used. Genomic DNA was digested with XbaI, denatured, and allowed to anneal to a reverse primer derived from the cDNA sequence; after primer extension, a blunt-ended adaptor was ligated and PCR was performed using a nested reverse primer derived from the cDNA and a forward primer derived from the adaptor sequence. The XbaI-digested I-PCR reaction produced two distinct bands of 1,475 and 942 bp. After cloning and sequencing, it was shown that the 942-bp band contained a single species (BETL1b), while the 1,475-bp band was a mixture of two DNA fragments, BETL1aand BETL1c, having slightly different sequences. The nested primers derived from the cDNA were designed to amplify a large portion of the coding sequence along with the promoter. In this way, we were able to identify three distinct copies of the gene, all of which contained an intron sequence inserted after T-80, taking the A of the translation start codon as nucleotide no. 1. Two features distinguished BETL1b: first, the acceptor splice site was TG instead of AG; second, the intron sequence was larger than that present in BETL1a or BETL1c at 463 bp instead of 123 bp. As a result, the promoter sequence contained in the I-PCR fragment ofBETL1b was only 177 bp long. Isolation of Intergenic Regions and Structure of theBETL1 Cluster From the information obtained from the physical mapping of the upstream sequences and the restriction sites identified after sequencing of the I-PCR products, a physical map was constructed that was confirmed by further Southern-blot analysis (data not shown). The three copies of BETL1 were found to be located on a singleEcoRV restriction fragment (Fig.2A). All of the copies of the gene are orientated in the same direction. The map was confirmed by amplifying and cloning the intergenic regions using the primers indicated in Figure 2A. A contig of 9,167 bp containing the three genes was assembled, which is summarized in Figure 2B. The cluster showed a rather complex organization with the three copies of the gene interspersed with a number of distinct repetitive sequences. Fig. 2. Open in new tabDownload slide Structure of the BETL1 cluster. A, Physical map of the 17-kb genomic fragment containing BETL-1a, BETL-1b, and BETL-1c. V, EcoR5; N, NcoI; E,EcoRI; X, XbaI; H, HindII; Ns, Nsi1; B, Bgl2; BI, BamHI. Lines shown under the map represent the PCR fragments used to sequence the 9-kb region outlined in B. Elements identified are shown as numbered boxes. 1,Spm-like transposon sequence; 2,Tourist-like transposable element; 3, 27 × (TTA) microsatellite repeat; 4, sleepy-like transposable element; black boxes, BETL1 and proton ATPase coding regions (transcription direction is indicated below the map); gray boxes and attached empty boxes, putative retrotransposon LTRs. C, Sequences possibly mobilized through a retrotransposition-like event (direct 5-bp repeats generated after insertion are shown in B). Fig. 2. Open in new tabDownload slide Structure of the BETL1 cluster. A, Physical map of the 17-kb genomic fragment containing BETL-1a, BETL-1b, and BETL-1c. V, EcoR5; N, NcoI; E,EcoRI; X, XbaI; H, HindII; Ns, Nsi1; B, Bgl2; BI, BamHI. Lines shown under the map represent the PCR fragments used to sequence the 9-kb region outlined in B. Elements identified are shown as numbered boxes. 1,Spm-like transposon sequence; 2,Tourist-like transposable element; 3, 27 × (TTA) microsatellite repeat; 4, sleepy-like transposable element; black boxes, BETL1 and proton ATPase coding regions (transcription direction is indicated below the map); gray boxes and attached empty boxes, putative retrotransposon LTRs. C, Sequences possibly mobilized through a retrotransposition-like event (direct 5-bp repeats generated after insertion are shown in B). The 135 bp of the BETL1b promoter obtained from the I-PCR product did not possess any homology with the promoter sequences ofBETL1a or BETL1c. However, it did match (95% identity) a repeated sequence found in the promoter of theZEMa gene and shown to be related to the Spm/En element (Montag et al., 1996). The sequence contained in theBETL1 contains two repeats of 36 bp, each formed by a palindromic sequence. Southern-blot analysis confirmed that the promoter region of BETL1b contains highly repeated DNA (not shown). The intron sequence present in BETL1b was also modified by the presence of a second transposable element insertion. The transposon was 346 bp long and possessed 15-bp inverted repeats at both ends, flanked by 3-bp direct repeats as follows: TAAgggcatgtacagtgg… … .ccactatacatgccc TAA (capital letters = direct repeat; lowercase letters = inverted repeat). The sequences identify this element as a miniature inverted repeat transposable element (MITEs, Wessler et al., 1995), in the same subfamily as Tourist (Bureau and Wessler, 1992). Hybridization of the transposon sequence to maize genomic DNA showed that related sequences are dispersed throughout the maize genome (not shown). TheBETL-1b gene is also modified by the exchange of the splicing acceptor site of the intron to TG instead of the standard AG found in BETL1a and BETL1c. The promoter sequences of BETL1a andBETL1c are nearly identical up to position −1,609, numbering from the A of the start codon. At −1,609,BETL1c contains two tandem repeats of 365 bp, whileBETL1a contains a microsatellite repeat of (ATT)27 (labeled “3” in Fig. 2B). A 230-bp fragment homologous to the 365-bp repeats is found downstream of the microsatellite in BETL1a. These findings suggest that a retrotransposition event may have been responsible for the integration of BETL1a between BETL1b andBETL1c. A fragment of about 5.5 kb, delimited by the 365-bp elements described above, functioning as the long terminal repeats (LTR) of a retrotransposon element (Fig. 2C), would have moved into the tandem duplication previously formed by BETL1b andBETL1c to give the present structure (Fig. 2B). In support of this hypothesis, it is known that retrotransposons possess a primer binding site at the 5′ end, whose sequence resembles that of a tRNA. A 15-bp sequence, partially homologous to a primer binding site, is present in the 365-bp repeated units at 6,494-TGGTCCTCGCCGAAGG-6,479. Further evidence for a transposon relic is the presence of fragments of an unrelated gene between the two LTRs. A sequence downstream of BETL1a shows homology to plasma membrane proton ATPases. The most similar sequence was that of a maize H+ ATPase, with a stretch displaying 75% identity extending over 238 residues. Intriguingly, this sequence was also found to be part of the maize retrotransposon-like sequence Bs1 (Young-Kwan and Bennetzen, 1994). However, the sequence in the BETL1 cluster is fragmented into six segments and contains two stop codons and seven frameshifts, indicating an accumulation of mutations. An additional feature found in the BETL1a andBETL1c promoters is the presence of a 242-bp element showing 62% identity with the transposable element “sleepy” (labeled “4” in Fig. 2B, Winkler and Helentjaris, 1995). A BETL1 Gene Cluster Is Present in Different Maize Varieties To investigate the degree of conservation of the BETL1gene cluster, we compared the organization of the locus in different maize lines by filter hybridizations. DNA was prepared from maize lines of diverse origin and from teosinte (Zea diploperennis). The DNA was digested with HindIII, which cuts once at the 5′ end of the BETL1 transcribed sequence, and thus gives an estimate of the number of copies of BETL1 present in each line (Fig. 3). Several of the samples, including the teosinte DNA, contain more than one BETL1 gene copy. Interestingly, the highest number of copies (three) was found in lines that had been produced after intensive breeding programs (lanes 2, 4, 5, and 12) as compared with those considered as primary lines or first derivatives (lanes 1, 3, 7–11). The process of gene amplification is illustrated by the line F2 (Fig. 3, lane 11, one copy) and its derivatives, F252 (lane 12, one copy), F1110 (lane 13, two copies), and F1444 (lane 14, three copies). Fig. 3. Open in new tabDownload slide BETL1 gene copy number in different maize varieties. Genomic DNAs (8 μg per line) were digested withHindII and the resulting Southern blot was hybridized with a BETL1 cDNA probe. Lane M, 1-kb DNA ladder; lane 1, teosinte (Zea diploperennis); lane 2, maize variety A69Y; lane 3, A239; lane 4, A632; lane 5, B73; lane 6, H99; lane 7, Pa91; lane 8, W64A; lane 9, FR16; lane 10, F2; lane 11, F252; lane 12, F1110; lane 13, F1444; lane 14, DBTS; lane 15, Y204. Fig. 3. Open in new tabDownload slide BETL1 gene copy number in different maize varieties. Genomic DNAs (8 μg per line) were digested withHindII and the resulting Southern blot was hybridized with a BETL1 cDNA probe. Lane M, 1-kb DNA ladder; lane 1, teosinte (Zea diploperennis); lane 2, maize variety A69Y; lane 3, A239; lane 4, A632; lane 5, B73; lane 6, H99; lane 7, Pa91; lane 8, W64A; lane 9, FR16; lane 10, F2; lane 11, F252; lane 12, F1110; lane 13, F1444; lane 14, DBTS; lane 15, Y204. Expression Analysis of BETL1 Genes The determination of copy number and mapping of each copy at thebetl1 locus enabled us to approach the issue of which copies of the gene were transcribed. The alignment of the threeBETL1 coding sequences, which was deduced from the genomic sequence after removing the introns, showed up to 23 base substitutions between genes. However, they are evenly distributed, which precluded the design of gene-specific probes for analyzing the expression of BETL1a, BETL1b, andBETL1c separately by RNA filter hybridizations. Therefore, two alternative approaches were used. First, RT-PCR analysis was performed using either a BETL1b or aBETL1a/BETLc specific primer (Fig.4). Two primers annealing to all BETL1 copies amplified a single band from endosperm cDNA (lane 1), two bands from genomic DNA (due to the different sizes of theBETL1b and BETL1a/BETLcintrons, lane 2) and one band from each genomic clone containing eitherBETL1a or BETL1b (lanes 3 and 4, respectively). When a BETL1b gene-specific primer was used, only a faint band was amplified from endosperm cDNA (lane 5), while the BETL1a gene specific primer produced an intense band from the same cDNA preparation (lane 8). Fig. 4. Open in new tabDownload slide RT-PCR analysis of the expression of theBETL1 genes. Endosperm cDNA samples (lanes 1, 5, and 8), genomic DNA (lane 2), and genomic clones containing eitherBETL1a (lanes 3, 6, and 9) or BETL1b(lanes 4, 7, and 10) were amplified using unspecific forward and reverse primers (lanes 1–4), BETL1b specific primers (lanes 5–7), or BETL1a/BETL1c specific primers (lanes 8–10). Lane M, 1-kb DNA ladder. Fig. 4. Open in new tabDownload slide RT-PCR analysis of the expression of theBETL1 genes. Endosperm cDNA samples (lanes 1, 5, and 8), genomic DNA (lane 2), and genomic clones containing eitherBETL1a (lanes 3, 6, and 9) or BETL1b(lanes 4, 7, and 10) were amplified using unspecific forward and reverse primers (lanes 1–4), BETL1b specific primers (lanes 5–7), or BETL1a/BETL1c specific primers (lanes 8–10). Lane M, 1-kb DNA ladder. Gene specificity of the primers was confirmed by the fact that PCR from genomic clones containing either BETL1b (Fig. 4, lanes 7 and 10) or BETL1a (lanes 6 and 9) rendered a PCR product only when the corresponding specific primers were used in the reaction. This experiment suggests that BETL1b is only weakly expressed, which is consistent with numerous modifications found in this gene, including transposon insertions in promoter and intron sequences. RT-PCR cannot be easily used to distinguish betweenBETL1a and BETL1c. To know if they are expressed at the same level, 16 independent BETL1 cDNA clones were sequenced and classified into three groups, a, b, or c. No clone was found that contained the sequence derived fromBETL1b, 10 clones contained the BETL1asequence, and six clones, in addition to the original BETL1clone, contained the BETL1c sequence. We conclude that both BETL1a and BETL1c promoters are functional. Comparison of the 5′ end sequences of the cDNA clones indicates a probable transcription start site at −53 or −51 upstream of the translation start codon. A 985-bp Promoter Sequence from BETL1a Directs the Expression of the GUS Reporter Gene in the Endosperm Transfer Cell Layer The promoter sequences of BETL1a are delineated at −1.8 kb by the insertion of BETL1b sequences. Because we have evidence that BETL1a is expressed, it is likely that a 1.8-kb promoter fragment will be sufficient to confer this expression pattern. Furthermore, the presence of a transposon-like sequence inserted at position −761 suggests that only the sequences downstream of this element might be needed. A −985 (KpnI–HindIII) BETL1apromoter fragment was fused to GUS, and the cassette was used in transient expression experiments by particle bombardment of immature dissected endosperms (not shown). Immature (12 DAP) endosperms expressed GUS at the sites of bombardment on the outer endosperm layer when a positive control, pAHC25 (Christensen et al., 1992), containing a maize polyubiquitin promoter fused to GUS was used. No signal was observed after bombardment with a promoterless GUS construct. Finally, blue spots were restricted to the basal area of the endosperm, when a −985-bp-BETL1apromoter-GUS construct was used. The GUS expression obtained in basal endosperm cells was weak and subject to large variation between individual experiments, thus preventing any quantitative conclusions. Generation and Analysis of Transgenic Maize Plants Containing theBETL1 Promoter-GUS Construct The 985-bp BETL1 promoter fragment was active specifically in the transfer cell layer of the kernel. The ability of this promoter fragment to confer transfer-cell-specific expression in transgenic maize was tested by maize transformation with the GUS reporter gene. Three independent transgenic lines, numbered 1, 11, and 17, were randomly chosen to be analyzed in detail. Plants containing the GUS gene were identified by Southern filter analysis (Fig. 5, top). Since the plants analyzed derived from the crosses between the primary transformants and non-transgenic plants, the segregation observed (1:1, transformant:non-transformant) was consistent with a single-copy integration. Nevertheless, the additional hybridizing bands present in line 17 (lanes 15 and 16) and the complex hybridizing patterns observed in all the lines when other restriction enzymes were used (Fig. 5, bottom) indicate the presence of multiple copy insertions, presumably linked or catenated. Fig. 5. Open in new tabDownload slide Top, Southern-blot analysis of putative transgenic plants. Genomic DNA from a negative control plant (10 μg, lane 1, A69Y) and five plants segregating the transgene locus from each transgenic line (line 1, lanes 2–6; line 11, lanes 7–11; and line 17, lanes 12–16) were digested with EcoR5, blotted onto a positively charged nylon filter, and hybridized with a probe derived from the GUS gene coding sequence. Size markers are shown on the left. Bottom, Southern-blot analysis of the transgenic loci present in three transgenic lines. Genomic DNA from a negative control plant (10 μg; lane 1, A69Y) and one plant from each transgenic line (line 1, lane 2; line 11, lane 3; and line 17, lane 4) were digested withDraI, blotted, and hybridized with either a 300-bp fragment from the proximal promoter of BETL1 (A) or a fragment derived from the GUS gene coding sequence (B). Lane M, DNA size marker (1-kb ladder). Fig. 5. Open in new tabDownload slide Top, Southern-blot analysis of putative transgenic plants. Genomic DNA from a negative control plant (10 μg, lane 1, A69Y) and five plants segregating the transgene locus from each transgenic line (line 1, lanes 2–6; line 11, lanes 7–11; and line 17, lanes 12–16) were digested with EcoR5, blotted onto a positively charged nylon filter, and hybridized with a probe derived from the GUS gene coding sequence. Size markers are shown on the left. Bottom, Southern-blot analysis of the transgenic loci present in three transgenic lines. Genomic DNA from a negative control plant (10 μg; lane 1, A69Y) and one plant from each transgenic line (line 1, lane 2; line 11, lane 3; and line 17, lane 4) were digested withDraI, blotted, and hybridized with either a 300-bp fragment from the proximal promoter of BETL1 (A) or a fragment derived from the GUS gene coding sequence (B). Lane M, DNA size marker (1-kb ladder). The transgenic lines were further analyzed using DraI, an enzyme that does not cleave within the reporter gene. ProbingDraI-digested DNA with the GUS coding sequence gave different patterns of hybridization for all three transgenic lines, confirming their independent origin (Fig. 5, bottom, lanes 2–4). Additionally, the comparison between blots A and B of Figure 5, which were probed with a fragment of the BETL1 promoter and the GUS coding sequence, respectively, demonstrates that the integration sites detected contain both GUS gene and BETL1 promoter sequences. Transgenic plants from each line were grown to maturity under greenhouse conditions. Histochemical staining of leaves, roots, adventitious roots, anthers, silks, and female flowers for GUS did not give signals for any transgenic line analyzed (not shown). The transgenic plants were either self-pollinated or crossed in both directions with a non-transgenic maize line (A69Y), and the developing kernels were stained for GUS enzyme activity at various stages during development. The position of the immature embryo, endosperm with transfer cells, and phloem terminals in the pedicel are indicated schematically in Figure 6F. Fig. 6. Open in new tabDownload slide GUS staining of immature seeds from transgenic plants. Immature kernels at various stages of development were hand-dissected and stained for 24 h as described in “Materials and Methods.” A, Negative control at 16 DAP; B, 11-DAP kernel; C, 27-DAP kernel; D, 16-DAP kernel; E, phase-contrast image at higher magnification of transfer cell region shown in D; F, a schematic representation of a longitudinal section of the kernel. GUS activity is seen as a blue precipitate of dichloro-dibromoindigo. Magnification: B, ×16; A, C, and D, ×6.4; and E, ×200. Bars: D, 1 mm; E, 50 μm. F, Schematic representation of the component tissues of the developing kernel. En, Endosperm; Em, embryo; TC, transfer cell layer of endosperm; Ph, placentochalaza. Fig. 6. Open in new tabDownload slide GUS staining of immature seeds from transgenic plants. Immature kernels at various stages of development were hand-dissected and stained for 24 h as described in “Materials and Methods.” A, Negative control at 16 DAP; B, 11-DAP kernel; C, 27-DAP kernel; D, 16-DAP kernel; E, phase-contrast image at higher magnification of transfer cell region shown in D; F, a schematic representation of a longitudinal section of the kernel. GUS activity is seen as a blue precipitate of dichloro-dibromoindigo. Magnification: B, ×16; A, C, and D, ×6.4; and E, ×200. Bars: D, 1 mm; E, 50 μm. F, Schematic representation of the component tissues of the developing kernel. En, Endosperm; Em, embryo; TC, transfer cell layer of endosperm; Ph, placentochalaza. Some of the seeds stained were GUS negative (Fig. 6A), as would be expected for plants segregating for the transgene. In other seeds, however, a pale blue staining corresponding to weak GUS activity appeared after 11 DAP, which was highly specific for the transfer cell layer (Fig. 6B). At later stages of seed development, the GUS signal remained confined to the basal transfer cells and the intensity peaked by 16 DAP (Fig. 6D), with no decrease in intensity observed until 27 DAP (Fig. 6C). Figure 6E shows a higher magnification of the 16 DAP staining pattern photographed under phase contrast microscopy; the presence of cell wall ingrowths in the transfer cell layer is evident, and the concentration of GUS activity in these cells can be seen. The comparison between the staining intensity obtained from seeds resulting from reciprocal crosses indicated staining proportional to gene endosperm dosage (not shown). A very weak endogenous GUS activity in the placentochalazal region of the pedicel appeared in both transgenic and non-transgenic kernels at approximately 16 DAP, and could be largely eliminated by the inclusion of 20% (v/v) methanol in the staining solution, a technique that did not affect staining in the transgenic basal layer. Comparison of the Accumulation of BETL1 and GUS Gene Products in Transgenic Maize GUS protein has been reported to be very stable in the plant cell, and consequently might not accurately reflect either the rate of transcription or the steady-state mRNA concentration derived from the transgene. Therefore, the steady-state GUS mRNA concentration was estimated by northern filter hybridization of poly(A+) RNA (Fig.7, top). Seeds from the cross between the transgenic line 11 and a wild-type parent (A69Y) were collected at five different developmental stages. Half of each kernel was stained for GUS activity, and the remainder stored at −80°C for RNA extraction if GUS activity was seen. A comparison between GUS mRNA (Fig. 7A) and that of the endogenous BETL1 (Fig. 7B) shows that the kinetics of accumulation of GUS mRNA differs from that of the endogenousBETL1 transcript. GUS mRNA is first detected at 13 DAP, later than that for BETL1, while the dramatic decrease in BETL1 mRNA after 17 DAP contrasts with a much more modest decline in GUS mRNA concentration. Fig. 7. Open in new tabDownload slide Top, Northern-blot analysis of the GUS gene expression. mRNA from non-transgenic (A69Y) seeds at 10 DAP (lane 1, 1 μg) or transgenic seeds (line 11 crossed by A69Y) at various stages of development (0.7 μg per lane) were electrophoresed in a formaldehyde gel, blotted, and sequentially hybridized with a GUS probe (A), a BETL1 probe (B), and a ubiquitin probe (C). Lane 2, 9 DAP; lane 3, 13 DAP; lane 4, 17 DAP; lane 5, 21 DAP; lane 6, 27 DAP. Filters were exposed for 2 h (A), 1 min (B), or 15 min (C). Bottom, Accumulation of BETL1 (hatched bars), GUS (white bars), and immunophilin (shaded bars) in BETL1/GUS transgenic maize kernels. Total protein extracts (15% of one kernel per track) were prepared from kernels harvested at different days after fertilization as indicated, loaded onto 15% SDS-polyacrylamide gels, and electroblotted onto PVDF membranes. Immunoblots were probed successively with antisera against BETL1 (Hueros et al., 1995), GUS (A-5790, Molecular Probes), and mzFKBP-66 (Hueros et al., 1998), detected by enhanced chemiluminescence, and changes in protein concentration (expressed in arbitrary units) derived by image quantitation. Fig. 7. Open in new tabDownload slide Top, Northern-blot analysis of the GUS gene expression. mRNA from non-transgenic (A69Y) seeds at 10 DAP (lane 1, 1 μg) or transgenic seeds (line 11 crossed by A69Y) at various stages of development (0.7 μg per lane) were electrophoresed in a formaldehyde gel, blotted, and sequentially hybridized with a GUS probe (A), a BETL1 probe (B), and a ubiquitin probe (C). Lane 2, 9 DAP; lane 3, 13 DAP; lane 4, 17 DAP; lane 5, 21 DAP; lane 6, 27 DAP. Filters were exposed for 2 h (A), 1 min (B), or 15 min (C). Bottom, Accumulation of BETL1 (hatched bars), GUS (white bars), and immunophilin (shaded bars) in BETL1/GUS transgenic maize kernels. Total protein extracts (15% of one kernel per track) were prepared from kernels harvested at different days after fertilization as indicated, loaded onto 15% SDS-polyacrylamide gels, and electroblotted onto PVDF membranes. Immunoblots were probed successively with antisera against BETL1 (Hueros et al., 1995), GUS (A-5790, Molecular Probes), and mzFKBP-66 (Hueros et al., 1998), detected by enhanced chemiluminescence, and changes in protein concentration (expressed in arbitrary units) derived by image quantitation. A comparison of the accumulation of BETL1 and GUS protein during endosperm development was made by immunoblotting with specific antisera. As control for a constitutively expressed intracellular protein, an immunophilin antibody was used (Fig. 7). It is evident that in contrast to the quantitative turnover of BETL1 after 20 DAP, GUS continues to accumulate and is present in mature kernels. DISCUSSION The betl1 locus, encoding an abundant, endosperm transfer-cell-specific transcript (Hueros et al., 1995), has been found to consist of a cluster of three tandemly arranged gene copies. In the maize line A69Y, the three gene copies BETL1a, BETL1b, andBETL1c, (Figs. 1 and 2) are located on a 9-kb DNA fragment. The distribution of these sequences allowed us to define a region of maximum 1.8 kb in which an active promoter sequence resides; 985 bp of this promoter is sufficient to confer expression specifically in basal endosperm transfer cells. The structure of the gene cluster (Fig. 2) and the sequence comparisons suggest a model in which BETL1band BETL1c have arisen by spontaneous duplication (Ohno, 1979), followed by the insertion of the BETL1a copy by a transposition event (see below). Interestingly, the different maize inbred lines examined all show evidence of gene duplication, having two to three copies (Fig. 3). The BETL1a gene is located on a DNA fragment (from position 1,259–6,745, Fig. 2B) with similarities to TY3/gypsy retrotransposons, such as “reina” (SanMiguel et al., 1996). Both reina and the element described here are flanked by LTRs of around 0.3 kb. The total length of both elements is 5.5 kb and both apparently are flanked by 5-bp repeats of GGTTG (only detected at the 3′ end in the case of reina) at the integration site. The potential transcription direction of the retro-element, as deduced from the position of the putative primer-binding site, is opposite to that of BETL1a. This has two consequences. First, this orientation would explain why the 3′ flanking LTR adjoins a microsatellite repeat (27× AAT, if read on the noncoding strand). The A-rich microsatellites found at the 3′ end of LTR-like repeats are thought to be derived from the poly-dA tail of retrotransposons, as suggested for Alu repeats (Nadir et al., 1996) or Artiodactyl retroposons (Kaukinen and Varvio, 1992). This would also explain why the 3′ LTR found in the BETL1 cluster is shorter than the elements found at the 5′ end. Retrotransposon LTRs have a tripartite structure, i.e. are formed by three different elements, when integrated into the host genome. After transcription, however, the 3′ LTR lacks the 3′ element and the 5′ LTR lacks the 5′ element; complete LTRs are regenerated only after successful integration. A second consequence of the transcription polarity is the presence of introns in the genes contained in the retrotransposon. Had BETL1a been inserted in the sense orientation within the retrotransposon, theBETL1 intron would have been spliced out after transcription, as reported for the H-ATPase fragment found in Bs1 (Young-Kwan and Bennetzen, 1994). These considerations strongly suggest that the 365 bp repeats, and the truncated repeat associated with the (AAT)27microsatellite, are the LTRs of a maize retrotransposon, although this sequence configuration could also derive from homologous recombination between Solo-LTR elements. In this case, LTRs would have provided recombination sites used in generating multiple copies ofBETL1. In addition to the putative retrotransposon described above, the 9-kbBETL1 cluster harbors a tourist-like transposon element (Bureau and Wessler, 1992; Rio et al., 1996), a sleepy-like transposon fragment (Winkler and Helentjaris, 1995), and a sequence highly homologous to a member of the suppressor/mutator family (Montag et al., 1996). Such a high level of interspersion between repeated mobile elements and genes seems to be a characteristic of the maize genome.SanMiguel et al. (1996) found that retrotransposons accounted for more than 60% of a 280-kb region containing the ADH1-F maize gene. Furthermore, the comparison of the region containing the genesSh2 and a1 from maize, sorghum, and rice showed that, despite the conserved order of genes, the intergenic regions had accumulated extensive differences due to the integration of unrelated, repeated sequences (Chen et al., 1997). Once those regions of the cluster belonging to repeated/mobile DNA families had been identified, a promoter sequence was selected for testing specificity of expression in transgenic plants. AsBETL1b was not expressed (Fig. 4 and cDNA sequencing results), a −983-bp fragment of BETL1a was selected for functional analysis. Preliminary experiments using particle bombardment with promoter-GUS constructs (data not shown) showed specific expression in the transfer cells of immature maize kernels. Nevertheless, the GUS expression seen was too weak to be quantitated. This may be due to the technical difficulty of exposing the basal cells of immature endosperms to particle bombardment without extensive cell damage. Another possibility is that when the transfer cell layer is removed from the influence of solute flux through the pedicel, it may be altered in its expression characteristics. Interestingly, a similar finding was made for expression of an aleurone-specific promoter in transient assays (Kalla et al., 1996). The functionality of the BETL1 promoter was demonstrated by generation of transgenic maize plants containing the BETL1promoter-GUS gene construct. Inspection of transgene organization indicates that multiple copies are clustered in a few discrete regions of the maize genome (Fig. 5, top). The transgenes were integrated in different restriction fragments in the three lines analyzed, indicating that they are of independent origin (Fig. 5, bottom). Finally, histochemical GUS staining demonstrated that the −983 BETL1promoter-GUS construct introduced in transgenic plants can direct reporter gene activity (Fig. 6) in a way that resembles the spatial and temporal pattern of BETL1 expression (Hueros et al., 1995). Comparison of transgene-derived GUS mRNA with the endogene-derivedBETL1 transcript (Fig. 7, top) shows that the behavior of the BETL1 promoter fragment in the GUS fusion differs in two respects from that of the native promoter. First, the reporter gene mRNA was present at a much lower concentration than BETL1 mRNA. Second, the pattern of accumulation of GUS mRNA showed a delay compared with that from the BETL1 gene. These effects could be due to the lack of enhancer sequences, for example, located upstream of −983 or in the BETL1 intron, which were not in the region used for transformation, but alternative explanations such as the influence of position effects or differential mRNA stability are possible. Despite these minor differences in expression profiles, GUS protein accumulates in seeds approaching seed maturity, in contrast to the BETL1 protein (Fig. 7, bottom). It may be that BETL1 and other secreted proteins of the transfer cell layer are selectively degraded by extracellular proteases. An alternative might be their quantitative incorporation in insoluble cell wall material, which would render them non-extractable. The presence of transfer cells in the basal endosperm region suggests that this layer may promote solute transfer into the kernel, but the relative inaccessibility of the tissue makes this contribution difficult to assess by physiological techniques. Furthermore, to date, no mutant has been unequivocally identified whose primary site of action is in the transfer cells, although this may very well be the case for mn1, which affects one enzyme located in the transfer layer, cell wall-bound invertase (Cheng et al., 1996). We have shown that a 983-bp BETL1 promoter fragment directs expression exclusively in basal endosperm cells of maize. TheBETL1 promoter may become a valuable tool for the identification of components influencing solute transfer into the endosperm, and could potentially be used to manipulate grain filling. ACKNOWLEDGMENTS We thank Ursula Seul and Brigitte Piegeler for dedicated technical assistance and Dr. Bernd Weisshaar and the Max-Planck-Institut fuer Zuechtungsforschung sequencing facility for sequencing. LITERATURE CITED 1 Bureau TH Wessler SR Tourist, a large family of small inverted repeat elements frequently associated with maize genes. 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Plant Mol Biol 18 1992 189 200 Google Scholar Crossref Search ADS PubMed WorldCat 36 Wessler SR Bureau TE White SE LTR-retrotransposons and MITEs: important players in the evolution of plant genes. Curr Opin Genet Dev 5 1995 814 821 Google Scholar Crossref Search ADS PubMed WorldCat 37 Winkler RG Helentjaris T The maize Dwarf3 gene encodes a cytochrome P450-mediated early step in gibberellin byosynthesis. Plant Cell 7 1995 1307 1317 Google Scholar PubMed OpenURL Placeholder Text WorldCat 38 Young-Kwan J Bennetzen JL Integration and nonrandom mutation of a plasma membrane proton ATPase gene fragment within the Bsl retroelement of maize. Plant Cell 6 1994 1177 1186 Google Scholar PubMed OpenURL Placeholder Text WorldCat Author notes 1 This work was supported by the Deutsche Forschungsgemeinschaft (grant nos. SFB274 and SPP322 1005) and by European Community contract no. BIO4 CT–972158. * Corresponding author; e-mail [email protected]; fax 49–221–5062–413. 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)

van der Krol, Alexander R.; van Poecke, Remco M.P.; Vorst, Oscar F.J.; Voogd, Charlotte; van Leeuwen, Wessel; Borst-Vrensen, Tanja W.M.; Takatsuji, Hiroshi; van der Plas, Linus H.W.

doi: 10.1104/pp.121.4.1153pmid: 10594102

Tsiantis, Miltos; Brown, Matthew I.N.; Skibinski, Gaia; Langdale, Jane A.

doi: 10.1104/pp.121.4.1163pmid: 10594103

Abstract Despite recent progress, the mechanisms governing shoot morphogenesis in higher plants are only partially understood. Classical physiological studies have suggested that gradients of the plant growth regulator auxin may play a role in controlling tissue differentiation in shoots. More recent molecular genetic studies have also identified knotted1 like homeobox (knox) genes as important regulators of shoot development. The maize (Zea mays L.) mutant rough sheath2 (rs2) displays ectopic expression of at least three knox genes and consequently conditions a range of shoot and leaf phenotypes, including aberrant vascular development, ligular displacements, and dwarfism (R. Schneeberger, M. Tsiantis, M. Freeling, J.A. Langdale [1998] Development 125: 2857–2865). In this report, we show that rs2 mutants also display decreased polar auxin transport in the shoot. We also demonstrate that germination of wild-type maize seedlings on agents known to inhibit polar auxin transport mimics aspects of thers2 mutant phenotype. The phenotype elaborated in inhibitor-treated plants is not correlated with ectopic KNOX protein accumulation. The majority of the aerial part of higher plants is derived from the shoot apical meristem (SAM). Despite recent progress, the exact process by which cells derived from the SAM give rise to the different parts of the vegetative plant body are still unclear. Molecular genetic analysis has suggested that the regulation of knox genes is instrumental both to maintenance of the SAM and to the initiation of lateral shoot organs (Kerstetter and Hake, 1997). Ectopic expression ofknox genes in dicotyledonous plants results in a range of plant phenotypes, including lobed leaves, shoot vivipary, and decreased apical dominance. Intriguingly, these phenotypes are also observed in transgenic plants that either overexpress a cytokinin biosynthetic gene or underproduce auxin, and therefore have elevated cytokinin to auxin ratios (Estruch et al., 1991; Li et al., 1992; Klee and Lanahan, 1995). Such findings have led to the suggestion that the developmental pathways defined by plant growth regulators and knox genes are somehow interrelated (Kerstetter et al., 1997; Brutnell and Langdale, 1998; Tsiantis and Langdale, 1998). Another tentative area of convergence between hormone- and homeobox-specified pathways is vascular development. It is known that exogenous auxin can induce vascular differentiation and affect the path of vascular strand differentiation in different plant systems (Aloni, 1995). In addition, correlations have been found in Arabidopsis between aberrations in vascular tissue development (twisting, midvein bifurcation) and decreased polar auxin transport (PAT) (Bennett et al., 1995; Carland and McHale, 1996). Notably, maize (Zea maysL.) mutants that ectopically express the homeobox genes kn1and rough sheath1 (rs1) also display abnormalities in vascular differentiation patterns (Volbrecht et al., 1991; Becraft and Freeling, 1994). Moreover, in the stem of wild-type maize plants, both the rs1 and kn1 homeobox genes are expressed in close association with provascular strands (Smith et al., 1992; Jackson et al., 1994; Schneeberger et al., 1995). These observations suggest that auxin may be involved in mediating certain aspects of the phenotype that result from inappropriate knoxgene expression. The rough sheath 2 (rs2) mutant of maize displays ectopic expression of three knox genes due to loss of function of the rs2 gene that encodes a myb-like transcription factor (Schneeberger et al., 1998; Timmermans et al., 1999; Tsiantis et al., 1999). The resulting phenotype includes midrib duplication, leaf twisting, dwarfism, and vascular tissue aberrations. In this report, we assess whether perturbations in auxin homeostasis are a component of the maize rs2 mutant phenotype. Furthermore, we investigate the effects of PAT inhibitors on the growth of wild-type maize seedlings. MATERIALS AND METHODS Plant Material Seeds of the maize (Zea mays L.) inbred line B73 were a gift from Pioneer Hi-Bred International (Des Moines, IA). Thers2-twd allele was isolated as in Schneeberger et al. (1998). The mutation was induced by transposon insertion into the region of the rs2 gene that encodes the mybdomain (Tsiantis et al., 1999). Measurement of PAT Auxin transport measurements were conducted according to the method of Okada et al. (1991). Elongated mesocotyls were harvested from seedlings grown in the dark at 25°C for a week. Mesocotyl segments used were 2.2 to 2.4 cm. Tissue samples were incubated in a microfuge tube containing 40 μL of C-14 indole acetic acid for 16 h. After this time, the upper 2 mm of tissue was removed, placed in scintillant, and counted in a multipurpose scintillation counter (model LS6500, Beckman Instruments, Fullerton, CA). Treatment of Plants with Inhibitors of PAT Seeds of the inbred line B73 were sterilized and germinated on Murashige-Skoog medium in the presence or absence of 2,3,5-triiodobenzoic acid (TIBA) (28 μm) or naphthylphthamic acid (15 μm). Plants were grown in sterile pots at 25°C under a 16-h light/8-h dark photoperiod (100 μmol m−2 s−1), and after 2 weeks seedling morphology was examined. Histology Leaf samples were fixed in formalin acetic acid for 30 min, dehydrated through an ethanol series, paraffin embedded, and sectioned as in Langdale (1994). Sections (10 μm) were stained with Safranin/Fast Green as described in Schneeberger et al. (1998). Mutant and wild-type shoot apices were fixed in formalin acetic acid for 2 h, dehydrated, and embedded as above. Apices were sectioned completely and the number of axillary buds was noted per plant. Ten wild-type and seven mutant plants were examined. Immunolocalization Assays Tissue was fixed as described above and sections were reacted with anti-KNOX antibody as described in Schneeberger et al. (1998). RESULTS AND DISCUSSION Auxin Transport Aberrations in rs2 Mutant Plants To assess the auxin transport capacity of rs2 mutant plants, PAT measurements were conducted on etiolated mesocotyls of wild-type and rs2 maize seedlings. These measurements revealed that there was a clear difference between basipetal and acropetal transport in wild-type plants, whereas in mutant seedlings such a difference was not apparent (Fig.1). This indicates that auxin gradients may be perturbed in the shoots of rs2-twd seedlings. Auxin is generally thought to be produced in young emerging leaves and transported basipetally through the shoot (Sachs, 1991). A block in basipetal transport would be expected to result in the disruption of auxin gradients both within the leaves (where entrapment of excess auxin could occur) and across the vegetative axis (where less auxin could flow). The latter event would result in reduced internode elongation and could therefore explain the reduced stature ofrs2 plants. Fig. 1. Open in new tabDownload slide Perturbed auxin transport in mesocotyls ofrs2 mutant seedlings. Acropetal transport (black bars) and basipetal (white bars) transport of exogenously supplied C-14 IAA in wild-type and mutant (rs2-twd) seedlings. In wild-type plants, a significant difference is seen between the acropetal and basipetal measurements, demonstrating the presence of active polar basipetal transport mechanisms. In mutant plants, no significant difference is seen between basipetal and acropetal measurements, suggesting that the ability to basipetally transport auxin is significantly reduced in mutant tissue. Fig. 1. Open in new tabDownload slide Perturbed auxin transport in mesocotyls ofrs2 mutant seedlings. Acropetal transport (black bars) and basipetal (white bars) transport of exogenously supplied C-14 IAA in wild-type and mutant (rs2-twd) seedlings. In wild-type plants, a significant difference is seen between the acropetal and basipetal measurements, demonstrating the presence of active polar basipetal transport mechanisms. In mutant plants, no significant difference is seen between basipetal and acropetal measurements, suggesting that the ability to basipetally transport auxin is significantly reduced in mutant tissue. Auxin gradients are also believed to influence both vascular strand patterning and cytoskeletal organization. Thus, disruption of auxin gradients within the leaves could explain both the twisting growth pattern of rs2 leaves and the bifurcation of midribs. Indeed, it has recently been shown that two Arabidopsis mutants with deficiencies in PAT show similar characteristics. The lop1mutant is both dwarfed and twisted (Carland and McHale, 1996), and thepin1 mutant often shows midrib bifurcation and leaf twisting (Bennett et al., 1995). Disrupted auxin gradients may also account for the changes of vascular patterning and overt vascularization of leaves that have been observed in both rs2 and Rs1mutants (Becraft and Freeling, 1994; Schneeberger et al., 1995, 1998). Indeed, it has already been suggested that the Rs1 mutation, which conditions increased vascular size, could interfere with auxin-regulated developmental pathways (Becraft and Freeling, 1994). Thus, our findings suggest that perturbations to auxin physiology could mediate certain facets of the rs2 mutant phenotype. Growth of Wild-Type Maize on Auxin Transport Inhibitors Mimics Aspects of the rs2 Phenotype To ascertain whether reductions in PAT in wild-type maize could cause phenotypic perturbations similar to those seen in rs2mutants, we germinated wild-type maize seedlings in the presence of compounds known to inhibit PAT. Treatment with TIBA (28 μm) resulted in pronounced effects on seedling development (Table I). Roots were agravitropic and showed inhibition of lateral root growth (data not shown). Treated seedlings showed similar phenotypes to rs2mutant plants in that they were dwarfed, with compressed internodes and twisted leaves (Fig. 2, A–C). Occasionally, the second leaf to emerge exhibited a non-discrete blade/sheath boundary (Fig. 2E) as opposed to the discrete boundary defined by the ligule of untreated plants (Fig. 2D). Notably,rs2 mutant leaves show similar perturbations at the blade/sheath boundary (Fig. 2F). Histological examination of leaf sections revealed the presence of hypertrophic vascular bundles in both TIBA-treated wild-type plants and in rs2 mutant plants (for example, compare phloem tissue in Fig. 2, G–I). Qualitatively similar results were obtained after treatment of plants with naphthylphthamic acid (15 μm). Thus, aspects of thers2 phenotype are phenocopied by treating wild-type maize seedlings with PAT inhibitors. Table I. Phenotypes exhibited by wild-type plants treated with the auxin transport inhibitor TIBA . TIBA . Control . no. Agravitropic roots 28 0 Dwarfism 25 4 Twisted leaves/stem 15 0 Aberrant ligule 3 0 Feiled to germinate 5 2 Severe growth arrest 3 1 . TIBA . Control . no. Agravitropic roots 28 0 Dwarfism 25 4 Twisted leaves/stem 15 0 Aberrant ligule 3 0 Feiled to germinate 5 2 Severe growth arrest 3 1 Seventy-six plants were grown, half on control medium and half on medium containing TIBA. The number of plants showing specific phenotypes is indicated. Open in new tab Table I. Phenotypes exhibited by wild-type plants treated with the auxin transport inhibitor TIBA . TIBA . Control . no. Agravitropic roots 28 0 Dwarfism 25 4 Twisted leaves/stem 15 0 Aberrant ligule 3 0 Feiled to germinate 5 2 Severe growth arrest 3 1 . TIBA . Control . no. Agravitropic roots 28 0 Dwarfism 25 4 Twisted leaves/stem 15 0 Aberrant ligule 3 0 Feiled to germinate 5 2 Severe growth arrest 3 1 Seventy-six plants were grown, half on control medium and half on medium containing TIBA. The number of plants showing specific phenotypes is indicated. Open in new tab Fig. 2. Open in new tabDownload slide Effects of PAT inhibitors on maize seedling growth. Dwarfism and twisting: A and C, Plants on the left have been germinated on control medium and allowed to grow in sterile pots for 2 weeks; plants on the right have been germinated in the presence of 28 μm TIBA and grown for the same time. B, The plant on the left is a wild-type sibling of the rs2 mutant plant shown on the right. Displaced ligule formation: D, Seedling leaf of an untreated wild-type plant. White arrow points to the ligule. Leaf twisting and aberrant ligular formation in a TIBA-treated wild-type plant (E) and in a rs2 mutant plant (F). White arrows point to the non-discrete ligular boundary. Hypertrophic vascularization: G, Vascular morphology of a lateral vein in an untreated wild-type seedling. Vascular hypertrophy seen in a TIBA-treated plant (H) and in a rs2 mutant (I) plant. Xylem and phloem are labeled X and P, respectively. Gray lines indicate the edge of the phloem in each case. Bar = 30 μm. Fig. 2. Open in new tabDownload slide Effects of PAT inhibitors on maize seedling growth. Dwarfism and twisting: A and C, Plants on the left have been germinated on control medium and allowed to grow in sterile pots for 2 weeks; plants on the right have been germinated in the presence of 28 μm TIBA and grown for the same time. B, The plant on the left is a wild-type sibling of the rs2 mutant plant shown on the right. Displaced ligule formation: D, Seedling leaf of an untreated wild-type plant. White arrow points to the ligule. Leaf twisting and aberrant ligular formation in a TIBA-treated wild-type plant (E) and in a rs2 mutant plant (F). White arrows point to the non-discrete ligular boundary. Hypertrophic vascularization: G, Vascular morphology of a lateral vein in an untreated wild-type seedling. Vascular hypertrophy seen in a TIBA-treated plant (H) and in a rs2 mutant (I) plant. Xylem and phloem are labeled X and P, respectively. Gray lines indicate the edge of the phloem in each case. Bar = 30 μm. As discussed above, most of the phenotypic perturbations observed in treated plants can be rationalized on the basis of disrupted auxin gradients. However, little is known about the early signals involved in ligular formation so it is more difficult to establish why disrupted auxin gradients in emerging leaves led to the observed perturbations in the ligular area. Signals involved in ligule differentiation originate near the midrib at plastochron (P) 1–2 (Sylvester et al., 1990). Interestingly, the leaves in which we observed an abnormal blade/sheath boundary are established during embryogenesis and thus would be predicted to have already formed the blade/sheath boundary at the time of inhibitor treatment. Our data therefore suggest that there is a degree of plasticity in the formation of the ligule and that the boundary can be influenced somewhat later than P2. It is conceivable that auxin gradients may play a role in this process. For example, it was recently suggested that the steep radial gradient of auxin that exists in pine leaves acts as a morphogenetic field to direct the development of different cell types (Uggla et al., 1996). It is possible that similar gradients exist in leaves of other higher plants and that cellular differentiation within the leaf depends on such gradients. Despite apparent similarities between wild-type maize seedlings treated with PAT inhibitors and rs2 mutant plants, there are also notable differences. Most obviously, the root phenotypes observed in TIBA-treated plants are not seen in rs2 mutants. This finding implies that at least some component of the PAT system is functional in rs2 mutants. Decreased PAT in the rs2 Mutant Is Accompanied by Precocious Axillary Meristem Development Basipetal auxin transport is believed to be at least partly responsible for axillary meristem arrest (Cline, 1994). Thus, we would predict that a reduction in PAT may lead to overdevelopment of axillary buds. Consistent with this idea, we observed overdevelopment of lateral buds in rs2-twd mutant apices (Fig.3B). Detailed examination showed that 10 d after germination more axillary meristems were developed inrs2 mutants than in wild-type plants (Fig. 3C). This phenotype is consistent with the measured reduction in PAT since apical dominance is thought to involve basipetal flow of auxin across the vegetative axis. Notably, however, the number of lateral buds in wild-type and mutant plants was not significantly different 21 d after germination. Thus, the rs2-twd allele shows precocious rather than ectopic development of lateral buds. Although extra axillary meristems are initiated early in development, rs2mutants do not produce increased numbers of tillars (side shoots) or ear shoots. This would suggest that the reduction in PAT is either transient or is not sufficient to fully derepress axillary bud development. Fig. 3. Open in new tabDownload slide Precocious axillary bud development inrs2 mutant seedlings. A, Median section through the apical region of a 10-d-old wild-type seedling. Bar = 1 mm. B, Median section through the apical region of a 10-d-oldrs2 mutant seedling. Arrowheads point toward the axillary buds. Size bar = 1 mm. C, Number of axillary buds developed in 10-d-old wild-type (wt) and rs2 mutant seedlings. Fig. 3. Open in new tabDownload slide Precocious axillary bud development inrs2 mutant seedlings. A, Median section through the apical region of a 10-d-old wild-type seedling. Bar = 1 mm. B, Median section through the apical region of a 10-d-oldrs2 mutant seedling. Arrowheads point toward the axillary buds. Size bar = 1 mm. C, Number of axillary buds developed in 10-d-old wild-type (wt) and rs2 mutant seedlings. Ectopic Expression of knox Genes and Disruptions of PAT How does the observed reduction in PAT relate to thers2 mutation and to ectopic knox gene expression? It is known that ectopic accumulation of KNOX proteins inrs2 mutants disrupts cell fate acquisition in the leaf and leads in particular to vascular-tissue-related aberrations. However, very little is known regarding the exact nature of the developmental pathways in which KNOX proteins operate. It is possible that genes involved in plant growth regulator function (including auxin) could be among the knox gene targets. Changes in expression patterns of such genes could impair auxin function, resulting in vascular tissue abnormalities. Alternatively, changes in auxin homeostasis could alterknox gene expression patterns. The latter possibility is suggested by a recent report showing that perturbations in growth regulator levels affect knox gene expression levels in Arabidopsis (Rupp et al., 1999). To distinguish these possibilities in our experimental system, KNOX protein accumulation patterns were examined in TIBA-treated wild-type plants. In both TIBA-treated and untreated wild-type plants, KNOX proteins accumulated in shoot meristems (both apical and axillary) (Fig.4). No ectopic KNOX accumulation was observed in leaves. Thus, aberrant PAT in rs2 mutants is likely to result from rather than cause ectopic knox gene expression. Fig. 4. Open in new tabDownload slide KNOX protein accumulation patterns in TIBA-treated wild-type plants. A, Median section through the apical region of a 10-d-old untreated wild-type seedling. B, Median section through the apical region of a 10-d-old TIBA-treated wild-type seedling. Arrows denote the position of axillary meristems. C, Transverse section through the apical region of a 10-d-old TIBA-treated wild-type seedling. Bars = 100 μm. Fig. 4. Open in new tabDownload slide KNOX protein accumulation patterns in TIBA-treated wild-type plants. A, Median section through the apical region of a 10-d-old untreated wild-type seedling. B, Median section through the apical region of a 10-d-old TIBA-treated wild-type seedling. Arrows denote the position of axillary meristems. C, Transverse section through the apical region of a 10-d-old TIBA-treated wild-type seedling. Bars = 100 μm. Support for the idea that ectopic KNOX gene expression could alter hormonal function comes from studies in tobacco, rice, and lettuce (Tamaoki et al., 1997; Kusaba et al., 1998; Tanaka-Ueguchi et al., 1998; Frugis et al., 1999). In all cases, ectopic expression of KNOX protein was reported to drastically alter hormonal levels and in particular to lead to elevated cytokinin levels (Ori et al., 1999). The idea that KNOX proteins may directly affect hormonal production has been reinforced by a recent study demonstrating that targeted expression of kn1 in a novel developmental context increases cytokinin levels. Interestingy, disruptions in PAT would be predicted to condition phenotypic effects similar to those resulting from elevated cytokinin, since certain cells in PAT-inhibited plants would have reduced auxin and therefore an increased cytokinin to auxin ratio. Despite the fact that a reasonable amount of evidence suggests tight connections between KNOX genes and hormonal function, an indirect link between ectopic KNOX protein accumulation and hormonal regulation cannot be ruled out. For example, it is possible that KNOX proteins alter cellular identities such that the normal transport canals of auxin are disrupted and therefore a reduction in PAT occurs as a secondary effect. To further this work, it will be essential to identify maize mutants that perturb auxin function. Analysis of double mutants obtained by crossing such lines and the already existing leaf development mutants (such as Rs1 and rs2) should help define the role of auxin in maize leaf development more accurately. 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M.T. is the recipient of a University of Oxford Glasstone Postdoctoral Fellowship. M.I.N.B. and G.S. were recipients of Nuffield Foundation Undergraduate Bursaries. * Corresponding author; e-mail [email protected]; fax 44–1865–275147. 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)