TY - JOUR
AU - Pawlowski, Katharina
AB - Abstract Two modes of phloem loading have been proposed, apoplastic and symplastic, depending on the structure of sieve element–companion cell complexes (SE–CCCs) in minor vein phloem. Species are usually classified as either apoplastic or symplastic loaders although the cytology of SE–CCCs in minor veins of the majority of plants indicates that both mechanisms can be simultaneously involved in phloem loading. The functions of structurally different SE–CCCs in minor veins of the stachyose-translocating plant Alonsoa meridionalis were examined. A stachyose synthase gene, AmSTS1, was expressed in intermediary cells but not in the ordinary companion cell of the same vein. In contrast, sucrose transporter AmSUT1 protein was present in ordinary companion cells but not in the neighbouring intermediary cells. These data reveal the principles of phloem sap formation in A. meridionalis and, probably, in many other dicots. The two types of SE–CCCs within one and the same minor vein load different carbohydrates, using contrasting mechanisms for their delivery into the phloem. Lateral sieve pores in the minor vein phloem lead to mixing of the carbohydrates soon after loading. While symplastic and apoplastic pathways can function simultaneously during phloem loading, they are separated at the level of different SE–CCCs combined in phloem endings. Apoplast, companion cells, phloem loading, stachyose synthase, sucrose transporter, symplast Introduction The main function of phloem companion cells in minor veins of leaves is the loading of sieve elements with assimilates, while companion cells from larger veins of leaves, petioles, stems, and other axial organs are engaged in the retrieval of leaked assimilates into the phloem. Two modes of phloem loading, symplastic and apoplastic, are distinguished on the basis of the cytology of companion cells in minor vein phloem (Gamalei, 1990). The companion cells of putative symplastic phloem loaders (termed ‘intermediary cells’) are connected to bundle sheath cells via numerous secondary plasmodesmata supposed to allow symplastic passage of photoassimilates from the mesophyll into the phloem. In species with scarce plasmodesmata, an apoplastic step is required for the phloem loading of assimilates. Companion cells in apoplastic phloem loaders are, for example, transfer cells characterized by the presence of cell wall ingrowths increasing the plasma membrane surface available for apoplastic transport processes (Pate and Gunning, 1969), and ordinary companion cells. The type of companion cells in minor vein phloem is conserved, with few exceptions, at the level of plant genera and even families (Gamalei, 1990, 1991; Turgeon et al., 2001; Gamalei et al., 2007). Based on this feature, plant species were formally classified as either ‘apoplastic’ or ‘symplastic’ loaders. This division seems well justified for species containing structurally identical sieve element–companion cell complexes (SE–CCCs) in minor veins, such as, for example, Cucumis melo, Zinnia elegans, and some other model plants used in studies on phloem loading. At the same time, more than one type of companion cells is present in the minor vein phloem in representative species from a number of families (Gamalei, 1990; van Bel and Gamalei, 1991). A typical example is Alonsoa meridionalis (Scrophulariaceae) with three SE–CCCs in minor veins; the two lateral SE–CCCs contain intermediary cells and the central SE–CCC contains an ordinary cell (Knop et al., 2001; Voitsekhovskaja et al., 2006). The assortment of cell types in the phloem of a single minor vein in dicotyledonous plants can be amazingly diverse (Batashev and Gamalei, 1996; Batashev, 1997), but no functional analysis revealing the roles of single SE–CCCs in the phloem loading has been done thus far. Phloem loading from the apoplast occurs via transporter proteins. In most plant species, the dominant transport sugar is sucrose. The localization of class SUT1/SUC2 sucrose transporters on the plasmalemma of phloem companion cells has been demonstrated for several species (reviewed by Sauer, 2007), and the primary role of these transporters in apoplastic phloem loading of sucrose is now recognized. In symplastic phloem loading, there is no obvious necessity for a specific assimilate transport form. Nevertheless, it has been established that plants with abundant symplastic connections between mesophyll and phloem always translocate raffinose family oligosaccharides (RFOs) in addition to sucrose, and the more plasmodesmata exist between the mesophyll and phloem, the higher is the proportion of RFOs in the phloem sap of the corresponding species (Gamalei, 1984; Turgeon et al., 1993; Flora and Madore, 1996). The ‘polymerization trap’ model postulates that the synthesis of RFOs occurs in intermediary cells, building up a sucrose concentration gradient which would enable symplastic movement of sucrose from the mesophyll into the phloem by diffusion (Turgeon, 1991, 1996). In accordance with the model, two enzymes of the RFO biosynthetic pathway, stachyose synthase from C. melo (Holthaus and Schmitz, 1991) and galactinol synthase from Cucurbita pepo and Ajuga reptans (Beebe and Turgeon, 1992; Sprenger and Keller, 2000), have been found to locate to intermediary cells. The model, however, still awaits confirmation by in vivo evaluation of concentration gradients and of size discrimination between sucrose and RFOs by plasmodesmata. The experimental data on the sucrose concentration gradient between the cytoplasm of mesophyll cells and in the phloem of A. meridionalis lent no support for the polymer trap mechanism (Voitsekhovskaja et al., 2006). This leaves open the question of the role of RFO synthesis in phloem loading at least for some putative symplastic loaders. Recently, a transformation system has been reported for Verbascum phoeniceum, a plant with two structural types of SE–CCCs in minor vein phloem, similar to A. meridionalis (McCaskill and Turgeon, 2007). Suppression of RFO synthesis in intermediary cells of V. phoeniceum resulted in severe inhibition of phloem transport in some transgenic lines. The interpretation of these data and of future experiments with this highly promising model system requires exact information on compartmentalization of symplastic loading and of the energized apoplastic loading of sucrose in V. phoeniceum and species with similar minor vein configuration. In this study, the functions of structurally different SE–CCCs present in minor veins of the stachyose-translocating plant A. meridionalis were examined, based on analyses of localization of expression of genes involved in the metabolism and transport of its major phloem-translocated carbohydrates. The following questions were addressed: (i) whether the two structurally contrasting types of SE-CCCs present in minor veins of A. meridionalis differ with regard to the carbon compounds they load into the phloem, and (ii) how symplastic and apoplastic phloem loading are compartmentalized in the minor veins of A. meridionalis. Materials and methods Plant material and vein anatomy Alonsoa meridionalis O. Kuntze (Scrophulariaceae) was grown in a greenhouse on pot soil at 600–700 μmol m−2s−1photosynthetic photon fluence rate (PPFR), 14/10 h light/dark period, and 22 °C/14 °C temperature. Mature fully expanded leaves were used for most studies, but RNA for expression analysis was also isolated from cotyledons of 2-week-old seedlings, roots, young leaves (up to 25% of the size of a mature fully expanded leaf), stems, flowers, and 1-week-old developing fruits of mature plants. Arabidopsis thaliana ecotype Columbia was grown on Mini-tray soil (Baster Einheitserdewerk GmbH, Froedenberg, Germany) at 22 °C with a PPFR of 130 μmol m−2s−1in a 16 h light/8 h dark cycle. The types of companion cells were determined on sections prepared for light microscopy based on their conserved position within the minor veins as established by transmission electron microscopic (TEM) studies (Gamalei, 1990; Turgeon et al., 1993; Haritatos et al., 2000b). In minor veins of A. meridionalis, two intermediary cells are laterally positioned and distinguished by their large diameter, whereas a single ordinary companion cell with a smaller diameter is situated between the two intermediary cells in the centre of the vein. SE–CCCs in minor veins of Arabidopsis contain only ordinary companion cells (two companion cells per sieve element) and are asymmetrically positioned within the minor veins. Cloning of AmSTS1 full-length cDNA cDNA was prepared from RNA isolated from mature leaves with the Invisorb Spin Plant-RNA Mini Kit (Invitek, Berlin, Germany) using RevertAid M-MuLV reverse transciptase (MBI Fermentas, Lithuania). Degenerate primers (5′-GGNTGGTGYCANTGGGAYGC-3′ and 5′-TGRAACATRTCCCARTCNGG-3′) were designed based on conserved motifs (FGWCTWD and PDWDMF) of amino acid sequences from Vigna angularis stachyose synthase (VaSTS; Peterbauer et al., 1999), C. sativus raffinose synthase (CsRS; accession no. AAD02832), and A. thaliana raffinose synthase (AtRS; accession no. BAB11595). Two PCR products, one of ∼0.8 kb and the other of ∼1.1 kb were amplified, cloned in pGEM®-T Easy (Promega, Madison, WI, USA), and sequenced. The ∼1.1 kb product showed >80% amino acid identity with VaSTS and was termed AmSTS1. Reverse transcription was performed with RevertAid M-MuLV-RT using the adaptor primer AP (5′-GGCCACGCGTCGACTAGTAGTTTTTTTTTTTTTTTTT-3′). The first 3′-RACE (rapid amplification of cDNA ends) PCR was performed with the gene-specific primer 5′-CTCTGGCAAATCAGATAACTTTGG-3′ and the UAP primer (5′-GGCCACGCGTCGACTAGTAG-3′). The second 3′-RACE PCR was performed with 2 μl of the first PCR mixture, the gene-specific primer 5′-ATGCATTGGCTGGTGCATGG-3′, and UAP. The 5′-RACE was performed twice with two independently designed sets of gene-specific primers (5′-GTCAGCCCTCCATCAGC-3′ and 5′-GCACACCATGGTAAATACCAGC-3′; 5′-GAGGAGCTCGAAAGAGTTGTCC-3′ and 5′-CAGGAAGTTCAAAATTGCGTTCATGG-3′) which produced similar results. The first primer of each pair was used for the reverse transcription with Thermoscript reverse transcriptase (Invitrogen, Eggenstein, Germany). The reaction products were treated with terminal deoxynucleotidyl transferase and dCTP, and used as templates in a PCR with the second primer and the 5′-RACE anchor primer (5′-GGCCACGCGTCGACTAGTACGGGIIGGGIIGGGIIG-3′). The PCR products were cloned and sequenced. After the 3′ end and 5′ end sequences of AmSTS1 cDNA had been obtained, the full-length AmSTS1 cDNA was amplified with Advantage Tth polymerase mix (Clontech, Palo Alto, CA, USA) using gene-specific primers (5′-CCAAACTAAGTAATAACTCAATAGTTCATGC-3′ and 5′-CAAAACACATTTTCATGGCCAATTCTGG-3′) and first-strand cDNA as template. The PCR product was ligated into pGEM®-T Easy and sequenced. Cloning of the AmSTS1 promoter A 1.85 kb portion of the AmSTS1 5′-upstream genomic sequence was obtained using the Genome Walker™ DNA Walking Kit (Clontech) with gene-specific primers (5′-GACGTTGCTCGGGATATCGGTCAGAATCGG-3′ and 5′-GGATGGGATCATATGGAGGTGCCATGGTTG-3′), cloned in pGEM®-T Easy, and sequenced (EMBL accession no. AJ487031). Sequence analysis Sequencing was performed by SEQLAB Sequence Laboratories Göttingen GmbH (Göttingen, Germany). Sequence data were analysed using the programs of the Wisconsin Genetics Computer Group (Devereux et al., 1984). Database searches were performed using the BLAST algorithm (Altschul et al., 1990) in the nucleotide sequence databases of the National Center of Biotechnology Information (NCBI), National Library of Medicine, NIH, in Bethesda, MD. RNA and DNA gel blot analysis RNA from A. meridionalis organs was separated on 1.2% agarose–formaldehyde gels and transferred onto a Hybond N nylon membrane (Amersham Pharmacia Biotech, Freiburg, Germany) using 10× SSC. The blot was probed with the 1.1 kb AmSTS1 cDNA fragment labelled with the HexaLabel Kit from MBI Fermentas (Vilnius, Lithuania). The membrane was washed twice with 2× SSC and 0.1% SDS (w/v) for 20 min each, and once with 0.5× SSC and 0.1% SDS (w/v). Signal was detected by phosphoimaging (Fuji BAS-1000, Raytest, Sprockhoevel, Germany) for at least 3 h and evaluated with the computer program Tina 2.0 (Raytest, Sprockhoevel, Germany). Genomic DNA was isolated from leaves of A. meridionalis using the DNeasy™ Plant Maxi Kit (Qiagen, Hilden, Germany). A 20 μg aliquot of total DNA was restricted by EcoRI, HindIII, PstI, and BamHI, respectively, separated on a 0.8% agarose gel at 20 V, and transferred to a Hybond N+nylon membrane (Amersham-Pharmacia) using alkaline transfer. The blot was probed with the full size AmSTS1 cDNA labelled with the HexaLabel Kit from MBI Fermentas. Hybridization and washing conditions were as for RNA blots. Two independent blots were prepared and hybridized with similar results. To check whether additional restriction sites were present within AmSTS1 introns, PCRs were performed with genomic DNA. Primers were designed from the AmSTS1 cDNA sequence to obtain three DNA fragments covering the whole length of the AmSTS transcribed region (5′-GTAATAACTCAATAGTTCATGCATAAGC-3′ and 5′-AAATCCCACCACACTCTTCATACC-3′; 5′-GTGTGCAAGAGTTTGCTGATGG-3′ and 5′-CCTCCATCAGCAAACTCTTGC-3′; 5′-TCCAGCATGCAGCAATGCAACG-3′ and 5′-TCGTTGCATTGCTGCATGCTGG-3′). The PCR products were cloned in pGEM®-T Easy; they included the AmSTS1 gene introns whose size was estimated as 50, 100, and 180 bp. Restriction analysis showed that the PCR products contained no restriction sites for EcoRI, HindIII, BamHI, and PstI in addition to those already known for the AmSTS1 cDNA sequence (data not shown). In situ RNA hybridization RNA antisense probes for in situ hybridization were produced using the AmSTS1 1.1 kb cDNA fragment as a template. The fragment cloned in pGEM®-T Easy was excised with EcoRI and ligated into pBluescript®IIKS+(Stratagene, La Jolla, CA, USA). The plasmid was linearized with XbaI for production of antisense RNA. For production of sense RNA, the 0.3 kb fragment of AmSUT1 cDNA (Knop et al., 2001) was cloned into pBluescript®IIKS+and the plasmid was linearized with HindIII. In vitro transcription was performed with T3 RNA polymerase for antisense RNA and T7 RNA polymerase for sense RNA in the presense of digoxigenin-UTP (Roche Biochemicals, Mannheim, Germany). Leaves and stems were cut into pieces in fixative [4% (w/v) paraformaldehyde, 0.25% (w/v) glutaraldehyde in 10 mM sodium phosphate buffer with 100 mM NaCl, pH 6.8] and dehydrated in a graded ethanol series. Plant material was embedded in a methacrylate resin mixture (Sigma-Aldrich, Taufkirchen, Germany) and 2 μm thick sections were cut on an ultratome (Ultracut, Reichert-Jung, Vienna, Austria). Methacrylate was removed by acetone, and the sections were hydrated in a graded ethanol series. The hybridization of the probes (100 ng of digoxigenin-RNA probe per slide) proceeded overnight at 45 °C under conditions described previously (van der Wiel et al., 1990), and washing and detection were done as described by Jackson (1991). Transformation of Arabidopsis pBIstsA and pBIstsB plasmids carrying either 1.85 kb of the AmSTS1 promoter region plus the 108 bp 5′-untranslated region (UTR) (‘PmrA’), or the 1.85 kb without the 5′-UTR (‘PmrB’), respectively, were prepared as follows. PCRs with A. meridionalis genomic DNA were performed using sequence-specific primers (i) 5′-GGTTGAAAAAACAGAGCAAAAACAGG-3′ and 5′-ATCCTCTTGGCCTTGACACCTAGG-3′ for amplification of the PmrA region; and (ii) 5′-AGAAAAATGTGAAGGGGAGGCAGAC-3′ and 5′-ATCCTCTTGGCCTTGACACCTAGG-3′ for amplification of the PmrB region. The PCR products were cloned in pGEM®-T Easy, excised with EcoRI, and the 5′ overhangs were filled up with Klenow polymerase. The fragments were ligated into the SmaI-restricted plant transformation vector pBI101.3 (Jefferson et al., 1987). Agrobacterium tumefaciens GV3101 (pMP90) cells were transformed with pBIstsA and pBIstsB, respectively, via electroporation. Positive transformants were used for the infiltration of A. thaliana (Clough and Bent, 1998). Selection of AmSTS1–GUS transformants Kanamycin-resistant transformants were grown on Mini-Tray soil with a 13/11 h light/dark period, 24 °C, 63 μmol m−2s−1PPFR. Leaves of these plants were used for β-glucuronidase (GUS) activity staining (see below). Thirty-six independent lines of F1 transformants and 20 lines of F2 and further generations (10 lines for pBIstsA and pBIstsB, respectively) were analysed. All experiments presented in this study were performed on homozygous plants of F3 and further generations. Homozygosity was analysed via the segregation of kanamycin resistance. Histochemical localization of GUS expression Whole seedlings or organs from mature plants were infiltrated with buffer [50 mM Na phosphate pH 7.5, 1 mM EDTA, 0.1% (v/v) Tween-20] containing 1 mM X-Gluc and 6 mM K-ferri/ferrocyanide. After 16–20 h incubation at 37 °C in darkness, plants were fixed in 4% (w/v) paraformaldehyde, 0.25% (w/v) glutaraldehyde in 100 mM sodium phosphate buffer, pH 7.2 and cleared with ethanol. For localization of the GUS reaction product at the cellular level, staining was performed with 10 mM K-ferri/ferrocyanide in the buffer. Stained leaves were fixed (see above) and embedded in methacrylate. Sections of 2 μm were cut on an ultratome and counterstained in 0.1% ruthenium red for 8 min. Growth of pAmSTS–GUS plants on agar plates with different sucrose concentrations Six independent lines of the pAmSTS–GUS plants (three lines for each pBIstsA or pBIstsB construct) were grown on MS-agar at 24 °C either with a 8/16 h light/dark period and 250 μmol m−2s−1PPFR, or with a 16/8 h light/dark period and 75 μmol m−2s−1PPFR, respectively. The agar plates were supplemented with 1% and 4% sucrose, respectively. After 3 weeks, GUS activity staining was performed. Immunolocalization of AmSUT1 in minor vein phloem of A. meridionalis Fixation, embedding, and sectioning of mature leaves were performed as described in Stumpe et al.(2006). For detection of AmSUT1, the AmSUT1 antibody (Knop et al., 2004) was diluted 1:500 in Tris-buffered saline (TBS) containing 1% (w/v) bovine serum albumin (BSA). In negative controls, the primary antibody was omitted. The secondary antibody, goat anti-rabbit Alexa Fluor 488 (Molecular Probes, Eugene, OR, USA), was diluted 1:250 in TBS with 1% BSA. Sections were mounted using the ProLong Gold Antifade kit (Molecular Probes). Light microscopy Seedlings and organs of mature plants were observed with a SZX-12 stereomicroscope (Olympus Deutschland GmbH, Hamburg, Germany). For conventional light microscopy a BX51 microscope (Olympus Deutschland GmbH) was used. Images were captured using a ColorView II digital camera and DP-Soft image-analytical software (Olympus Soft Imaging Solutions, Münster, Germany). Confocal laser scanning microscopy (CSLM) was performed using the LSM 510 META NLO system based on the AxioImager.Z1 microscope (Carl Zeiss, Göttingen, Germany). The argon laser band 488 nm was used to excite Alexa Fluor 488 fluorescence. The emission was observed by using a 505 nm long pass filter and the Plan-Apochromat 100×/1.4 oil objective. Sections were viewed and captured by using the differential interference contrast channel merged with the corresponding confocal images with the help of Carl Zeiss LSM v. 4.2 software. Z-stacks of 20 optical sections throughout the minor veins were obtained from 10 μm cross-sections. 3D reconstructions were performed using LSM Software Module ImageVisArt and Inside4D (Carl Zeiss). Transmission electron microscopy Pieces of mature leaves (4×4 mm) were fixed in 2.5% glutaraldehyde in 50 mM potassium phosphate buffer (pH 7.2) for 3 h, and post-fixed overnight in 2% osmium tetroxide in the same buffer at 4 °C. The tissue was dehydrated in an ethanol–acetone series, contrasted with 1% uranyl acetate in 70% ethanol, and embedded in Epon812-AralditeM epoxy resin. Ultrathin sections (40–60 nm) were cut with glass knives on an ultratome, contrasted on grids using 2% lead citrate, and viewed and photographed at 75 kV with a high resolution electron microscope H-600 (Hitachi High Technologies, Tokyo, Japan). Statistics Cytological data were obtained from at least three experiments performed independently, all of which gave consistent results. Data from molecular biological experiments and analyses of A. thaliana transgenic plants were from at least five experiments with 6–20 independent lines. Results Cloning and analysis of cDNA encoding stachyose synthase from leaves of Alonsoa meridionalis A full-length cDNA clone of AmSTS1(EMBL accession no. CAD31704) was 2889 bp including a 5′-UTR of 108 bp and a 3′-UTR of 172 bp. The open reading frame (ORF) of AmSTS1 encoded a protein of 868 amino acids with a calculated molecular weight of 95.6 kDa and an isoelectric point of 5.37. The 5′-UTR contained two ATG codons upstream of the putative initiation codon, representing the beginnings of two small upstream ORFs (uORFs) encoding peptides of four and 11 amino acids, both in-frame with the AmSTS1 ORF (Supplementary Fig. S1A available at JXB online). The AmSTS1 sequence showed 82% amino acid identity with the sequence from the other member of the Lamiales, Stachys affinis (SaSTS, GenBank accession no. CAC86963), and 63% and 66% identity with the sequences from Fabales, V. angularis (VaSTS; Peterbauer et al., 1999) and Pisum sativum (PsSTS; GenBank accession no. CAC38094), respectively (Supplementary Fig. S1B). AmSTS1 is encoded by more than one gene in the A. meridionalis genome A Southern blot with total DNA of A. meridionalis was hybridized with the full size cDNA of AmSTS1 (Fig. 1). The total DNA was digested with either of four different restriction endonucleases. EcoRI recognizes one restriction site within the AmSTS1 cDNA sequence, HindIII recognizes two sites, BamHI one site, and PstI none. The sites for EcoRI, HindIII, and BamHI were mapped on the AmSTS1 gene sequence including its known cDNA sequence, PCR-amplified introns, and 1850 bp promoter region. This allowed prediction of the size of the DNA fragments hybridizing on the Southern blot (marked with asterisks on Fig. 1). The appearance of two additional DNA fragments in the HindIII and BamHI digests suggested the presence of at least one additional member of the gene family. The appearance of three weak bands (arrowheads on Fig. 1) might be explained by cross-hybridization between AmSTS1 and raffinose synthase sequences. It is less likely that such a cross-hybridization could produce the strongly labelled additional fragments found in the HindIII and BamHI digests, as the blot was washed under stringent conditions. The presence of only one hybridizing fragment in the PstI restriction suggests that the number of gene family members is small; probably there are no more than two AmSTS genes. Similarly, genomic DNA gel blot analysis of the gene family for stachyose synthase from adzuki bean suggested the presence of two highly homologous genes and probably one or two more distantly related genes (Peterbauer et al., 1999). Fig. 1. View largeDownload slide Southern blot hybridization of genomic DNA of A. meridionalis with the AmSTS1 full size cDNA. Arrowheads point at weak bands. Asterisks mark internal fragments of the AmSTS1 gene. The numbers of hybridizing fragments as expected from the restriction sites within the AmSTS1 cDNA and its introns are compared with the numbers of hybridizing fragments observed. PstI-digested λ DNA was used as a size marker. Fig. 1. View largeDownload slide Southern blot hybridization of genomic DNA of A. meridionalis with the AmSTS1 full size cDNA. Arrowheads point at weak bands. Asterisks mark internal fragments of the AmSTS1 gene. The numbers of hybridizing fragments as expected from the restriction sites within the AmSTS1 cDNA and its introns are compared with the numbers of hybridizing fragments observed. PstI-digested λ DNA was used as a size marker. AmSTS1 is expressed in intermediary companion cells of mature leaves RNA gel blot hybridization analysis (Fig. 2) showed that the expression levels of AmSTS1 were highest in mature leaves and in cotyledons, followed by young leaves, while no hybridization was detected for RNA from stems, roots, flowers, and fruits. Expression of AmSTS1 was studied at the cellular level in stems and in leaves of A. meridionalis by in situ RNA hybridization. It should be noted that the in situ hybridization with RNA probes directed against highly conserved sequences cannot distinguish between closely related members of gene families (Morris, 1995). Therefore, the expression of all putative AmSTS genes was expected to be detected with antisense RNA corresponding to the complete AmSTS1 cDNA. The AmSTS1 transcripts were present in intermediary cells of minor veins in A. meridionalis leaves, while no expression signal was detected in ordinary companion cells of minor veins, nor in mesophyll cells (Fig. 3A, B). No expression was detected in major veins in leaves and in stems (data not shown). Hence, in leaves of A. meridionalis, stachyose synthesis seems only to take place in intermediary cells of minor vein phloem. Fig. 2. View largeDownload slide Northern blot hybridization of RNA from various organs of A. meridionalis with an AmSTS1 cDNA fragment. R, roots; S, stems; C, cotyledons; YL, young leaves; L, mature leaves; Fl, flowers; Fr, fruits. Fig. 2. View largeDownload slide Northern blot hybridization of RNA from various organs of A. meridionalis with an AmSTS1 cDNA fragment. R, roots; S, stems; C, cotyledons; YL, young leaves; L, mature leaves; Fl, flowers; Fr, fruits. Fig. 3. View largeDownload slide Localization of AmSTS1 expression and AmSUT1 protein in minor veins of A. meridionalis leaves (A–D), and pattern of pAmSTS1–GUS expression in Arabidopsis (E–J). (A and B) In situ hybridization on cross-sections of metacrylate-embedded leaves of A. meridionalis with a digoxigenin-labelled AmSTS1 antisense RNA probe (A) and a sense control (B). Arrows point at intermediary cells and arrowheads point at ordinary cells. (C) Immunolocalization of AmSUT1 protein in a leaf minor vein. The protein is localized on the plasma membrane of the ordinary cell (arrow). (D) TEM of part of the A. meridionalis minor vein. Arrowheads point at lateral sieve pores between the two sieve elements (SE) one of which is connected to an ordinary cell (OC) and the second one to an intermediary cell (IC). Arrows point at plasmodesmata connecting an OC and IC to the bundle sheath cell (BS). (E and F) GUS activity staining of pAmSTS–GUS A. thaliana seedlings grown on MS agar supplemented with 1% sucrose (E) and 4% sucrose (F), respectively. (G– I) GUS activity in soil-grown pAmSTS–GUS A. thaliana plants: G, mature leaves; H, flowers; I, siliques. (J) Localization of GUS activity in minor vein phloem of pAmSTS–GUS A. thaliana leaves in two SE–CCCs (arrows), each consisting of two companion cells and one sieve element between them. PP, phloem parenchyma; X, xylem; XP, xylem parenchyma Scale bars denote 10 μm (A, B, J), 5 μm (C), 1 μm (D); 3 mm (E, F); 1 mm (G, I), and 500 μm (H). Fig. 3. View largeDownload slide Localization of AmSTS1 expression and AmSUT1 protein in minor veins of A. meridionalis leaves (A–D), and pattern of pAmSTS1–GUS expression in Arabidopsis (E–J). (A and B) In situ hybridization on cross-sections of metacrylate-embedded leaves of A. meridionalis with a digoxigenin-labelled AmSTS1 antisense RNA probe (A) and a sense control (B). Arrows point at intermediary cells and arrowheads point at ordinary cells. (C) Immunolocalization of AmSUT1 protein in a leaf minor vein. The protein is localized on the plasma membrane of the ordinary cell (arrow). (D) TEM of part of the A. meridionalis minor vein. Arrowheads point at lateral sieve pores between the two sieve elements (SE) one of which is connected to an ordinary cell (OC) and the second one to an intermediary cell (IC). Arrows point at plasmodesmata connecting an OC and IC to the bundle sheath cell (BS). (E and F) GUS activity staining of pAmSTS–GUS A. thaliana seedlings grown on MS agar supplemented with 1% sucrose (E) and 4% sucrose (F), respectively. (G– I) GUS activity in soil-grown pAmSTS–GUS A. thaliana plants: G, mature leaves; H, flowers; I, siliques. (J) Localization of GUS activity in minor vein phloem of pAmSTS–GUS A. thaliana leaves in two SE–CCCs (arrows), each consisting of two companion cells and one sieve element between them. PP, phloem parenchyma; X, xylem; XP, xylem parenchyma Scale bars denote 10 μm (A, B, J), 5 μm (C), 1 μm (D); 3 mm (E, F); 1 mm (G, I), and 500 μm (H). AmSUT1 in minor veins of A. meridionalis localized to ordinary companion cells The sucrose transporter AmSUT1 had been localized in minor veins of leaves of A. meridionalis at the cellular level by using an anti-AmSUT1 antiserum but, importantly, it was not possible to distinguish between companion cell types (Knop et al., 2004). The present detailed analysis showed that the fringing of the ordinary companion cell was labelled (Fig. 3C), indicating the presence of AmSUT1 on the plasmalemma (Knop et al., 2004) of this cell, while no label was detected in the intermediary cells. Reconstruction of a Z-stack of optical sections of another minor vein of A. meridionalis obtained by CLSM showed that the ordinary cell, but not intermediary cells, was labelled throughout the length of the cell (Supplementary Fig. S2 at JXB online). Hence, sucrose loading via AmSUT1 in minor veins of A. meridionalis leaves seems to occur only via ordinary companion cells. Lateral symplastic connections are present between SE–CCCs in minor vein phloem of A. meridionalis The AmSTS1 expression pattern and the localization of Am SUT1 in minor veins of A. meridionalis suggested that the two major transport carbohydrates, stachyose or sucrose, are loaded into the phloem via different SE–CCCs. To find out where in the phloem these carbohydrates mix to form the ‘export phloem sap’, the presence of lateral sieve pores between sieve elements of different SE–CCCs within minor veins was investigated. Figure 3D shows such pores found between a sieve element associated with an ordinary companion cell and a sieve element associated with an intermediary cell. The presence of lateral sieve pores suggests that RFOs and sucrose can mix already in the minor veins, forming the export composition of the phloem sap. Expression pattern of pAmSTS1–GUS fusions in Arabidopsis confirmed the AmSTS1 specificity for phloem companion cells in source organs Arabidopsis plants were transformed with either of the two transcriptional GUS fusion constructs, 1.85 kb of the AmSTS1 5′-upstream sequence with and without the 5′-UTR of the cDNA, respectively (PmrA and PmrB constructs; see Materials and methods). As no influence of the 5′-UTR on pAmSTS1–GUS expression levels or pattern was found (data not shown), the data for both constructs will be discussed together. In seedlings grown on agar plates, AmSTS1–GUS expression was confined to the vascular system of cotyledons, as indicated by GUS activity staining (Fig. 3E). In small developing leaves, GUS activity was absent from veins but sometimes detected in leaf trichomes. When the leaves became larger, GUS activity was still absent from the veins in the proximal part of the leaf lamina but appeared in the veins of the distal part of the lamina and proceeded basipetally, indicating that the onset of AmSTS1–GUS expression in veins coincided with the sink–source transition of the leaf (Truernit and Sauer, 1995). In mature flowering plants grown on pot soil, GUS activity was detected in the vascular tissue of leaves (Fig. 3G), green sepals (Fig. 3H), and siliques (Fig. 3I). In siliques, GUS activity was not observed in the seeds (sink organs) but in the veins of the pods which are known to represent local source organs for the seeds in Brassicaceae (Allen et al., 1971). In flowers, GUS activity was only found in veins of green sepals which photosynthesize and can contribute to growth and development of the flowers (see, for example, Ganelevin and Zieslin, 2002). Thus, in both flowers and siliques, AmSTS1–GUS expression was confined to source tissues. The staining of veins in sepals and siliques could not be due to the transport of the GUS reaction products from source leaves to sink tissues, because it was also observed in inflorescences cut off from the plants before staining. No GUS activity was detected in inflorescence shoots and in roots (data not shown). In minor veins of mature leaves, GUS activity was confined to both SE–CCCs (Fig. 3J). It should be mentioned that Arabidopsis has ordinary companion cells in its minor vein phloem (Haritatos et al., 2000b). Some precipitation of reaction product was also seen in adjacent parenchyma cells; however, the amount of the dye was much lower than in the two SE–CCCs. The level of pAmSTS1–GUS expression in Arabidopsis was strongly enhanced by sucrose under high light growth conditions The activation of the AmSTS1 promoter during sink–source transition of leaves of A. meridionalis and Arabidopsis suggested that the increase of sucrose levels in the phloem companion cells of minor veins due to the onset of phloem loading might be involved in the regulation of the AmSTS1 expression level. To test whether sucrose can specifically influence the level of AmSTS1 promoter activity in Arabidopsis, pAmSTS1–GUS plants (PmrA and PmrB lines) were grown on agar plates supplemented with either 1% sucrose (control) or 4% sucrose. Plantlets on both types of plates possessing cotyledons and one to two pairs of fully developed true leaves were used for GUS activity staining. Two series of plates with 1% and 4% sucrose were grown under either high light or low light conditions. In seedlings grown under high light conditions, GUS activity levels increased dramatically on plates with 4% sucrose (Fig. 3F) compared with control plates (Fig. 3E), indicating that elevated sucrose levels strongly enhanced the activity of the AmSTS1 promoter. In seedlings grown under low light, no differences in GUS activity staining were detectable between plantlets grown on 1% and 4% sucrose (data not shown). Darkening of soil-grown pAmSTS1–GUS plants for 48 h led to a visible decrease of GUS activity levels in leaf veins (data not shown). Discussion A phloem ending represents an entity which is highly specialized for efficient loading of assimilates. Whereas structural heterogeneity of the SE–CCCs assembled in the phloem endings of a number of dicotyledonous plants was discovered a long time ago, no evidence was obtained for functional heterogeneity of the SE–CCCs within a phloem ending (van Bel, 2003). As this study shows, in A. meridionalis, phloem loading of sucrose from the apoplast occurs via the sucrose transporter AmSUT1 present on the plasmalemma of ordinary companion cells (Fig. 3C), whereas RFOs are synthesized in the intermediary cells, i.e. directly in the phloem (Fig. 3A, B). Hence the two types of transport carbohydrates in A. meridionalis enter the phloem by different mechanisms separated at the level of specific SE–CCCs in phloem endings. Several lines of evidence indicate that the expression of the stachyose synthase gene AmSTS1 in A. meridionalis is specific for intermediary cells in minor vein phloem and that it is activated during sink–source transition. (i) In situ hybridization revealed that AmSTS1 RNA was present in leaves in the intermediary cells of higher order veins (Fig. 3A, B) and not in companion cells of lower order veins (data not shown). (ii) RNA gel blot hybridization showed that AmSTS1 expression was much stronger in mature than in young leaves of A. meridionalis (Fig. 2); the presence of AmSTS1 transcripts not only in mature but also in young leaves reflected the fact that in some parts of the younger leaves collected for the RNA isolation, sink–source transition had already started. (iii) Expression of pAmSTS1–GUS fusions in Arabidopsis leaves was confined to phloem companion cells and appeared in veins in concert with the sink–source transition of leaves. In Arabidopsis, the AmSTS1 promoter was active in the vascular tissue of not only leaves but also other source organs such as cotyledons and green sepals, and inactive in the heterotrophic organs (Fig. 3E–J). In siliques, expression of pAmSTS1–GUS was found in veins of green pods, which were shown to be source organs in Brassicaceae (Allen et al., 1971). This strict specificity to the phloem of source organs distinguishes the AmSTS1 promoter from the CmGAS (melon galactinol synthase) promoter that was active in both source and sink (filaments and petals) organs in Arabidopsis (Haritatos et al., 2000a), although in tobacco pCmGAS–GUS expression was confined to the vascular tissue of source organs only (Haritatos et al., 2000a). In addition, the fact that sucrose can tune up the activity of the AmSTS1 promoter is in line with its proposed regulation by the source activity of leaves because sucrose starts entering companion cells in leaf minor veins en masse only with the onset of assimilate export. It is noteworthily that differences in the pattern of AmSTS1 promoter activity were found between Arabidopsis and A. meridionalis regarding its expression in flowers and siliques. This might be due to the fact that the AmSTS1 promoter is active in ordinary companion cells of Arabidopsis, but not of A. meridionalis. In veins of sepals and green pods, only ordinary companion cells are expected to be present in both Arabidopsis and A. meridionalis, as the vein network shows fewer orders than in leaves, although no ultrastructural studies were performed for sepals. Consequently, the activity of the AmSTS1 promoter in those veins in Arabidopsis but not in A. meridionalis, might reflect the fact that stachyose synthesis takes place only in intermediary companion cells. Similarly, the observation that in Arabidopsis but not in A. meridionalis, AmSTS1 was active in lower order veins and, in some transgenic lines, even in midribs, probably reflected the fact that ordinary companion cells are the only companion cells present in those veins in both Arabidopsis and A. meridionalis. These data seem to suggest that the two structurally close companion cell types, i.e. ordinary cells in Arabidopsis and in A. meridionalis, might not be functionally identical: ordinary cells in Arabidopsis seem to contain transcription factors involved in the expression of genes encoding RFO biosynthetic enzymes (Haritatos et al., 2000a, b; this study) whereas those in A. meridionalis do not. Plant sucrose transporters are encoded by gene families whose members show specificity for organs, tissues, and cell types (Sauer, 2007). In Arabidopsis, out of seven functional sucrose transporters, only one, AtSUC2, is responsible for phloem loading and is located in the plasmalemma of companion cells (Stadler and Sauer, 1996; Sauer et al., 2004; Sauer, 2007), suggesting that there is no redundancy in companion cell-specific isoforms in this species. If, in a similar way, AmSUT1 is the sucrose transporter responsible for apoplastic phloem loading of sucrose in A. meridionalis, then the SE–CCCs with intermediary companion cells (SE–ICCs) would load the phloem predominantly with RFOs, and SE–CCCs with ordinary cells (SE–OCCs) with sucrose. This could lead to the origin of two distinct phloem transport fluxes, the RFOs flux starting from the SE–ICCs in minor veins, and the sucrose flux starting from SE–OCCs. However, the presence of lateral sieve pores in minor veins (Fig. 3D) indicates that in A. meridionalis these fluxes mix directly in minor veins. Knop et al.(2004) reported labelling of sieve elements with the anti-AmSUT1 antibody which was not present in our study. This can be explained by the fact that different protocols for the preparation of sections and for immunofluorescence were used in both studies. In the present experiments, much less background was found and label in sieve elements was never detected. Knop et al. (2004) suggested that a cross-reactivity of the anti-AmSUT1 antibody might be responsible for the labelling of sieve tubes in their experiments. Previous studies have shown that the sucrose concentration in the phloem sap of A. meridionalis was significantly higher than the concentration in the cytoplasm of mesophyll cells (Voitsekhovskaja et al., 2006), rendering the diffusion of sucrose from the mesophyll into the phloem postulated by the ‘polymer trap’ mechanism impossible. Rather, in A. meridionalis, the concentration gradient favours sucrose diffusion from the phloem to bundle sheath cells. It is easy to see that under stationary conditions, i.e. when sucrose moves from cell to cell mainly by diffusion, the leakage of sucrose from sieve elements via intermediary cells into the bundle sheath could be prevented by the activity of stachyose synthase in intermediary cells converting sucrose into stachyose, provided that the assumption of the polymer trap model about plasmodesmatal discrimination between stachyose and sucrose is correct. On the other hand, the expenditure of sucrose in intermediary cells due to stachyose synthesis should contribute to the creation of a steep sucrose concentration gradient at the sieve element/intermediary cell boundary, promoting sucrose diffusion from sieve elements into intermediary cells. This would interfere with the long-distance transport of sucrose, and also increase the probability of losing sucrose from the intermediary cells via plasmodesmata, unless stachyose synthesis in intermediary cells would always perfectly match sucrose arrival from the sieve elements, which seems improbable. The cytological data show that even sucrose loaded into the phloem via ordinary cells could be lost in this way because SE–CCCs within minor veins are interconnected at the SE level. However, during phloem translocation, i.e. under non-stationary conditions, sucrose movement is defined by pressure gradients rather than by concentration gradients, and therefore the considerations described above are not applicable. The proposed turgor pressure regulation of plasmodesmata at the intermediary cells/bundle sheath boundary would ‘close’ the plasmodesmata even for small molecules such as sucrose (Voitsekhovskaja et al., 2006). In spite of the fact that AmSUT1 is present in the phloem of both minor and major veins of leaves, and of stems (Knop et al., 2004), and AmSTS1 is only present in minor veins, RFOs dominate over sucrose in the phloem sap of A. meridionalis (Knop et al., 2001, Voitsekhovskaja et al., 2006), reflecting the fact that only a negligible part of phloem loading occurs outside of minor veins. The phloem sap concentrations measured in A. meridionalis were 174 mM for sucrose, 397 mM for stachyose, and 249 mM for raffinose (Voitsekhovskaja et al., 2006). Assuming that RFOs in the intermediary cells are synthesized from sucrose, 397 mM of the tetrasaccharide stachyose and 249 mM of the trisaccharide raffinose in the phloem sap of A. meridionalis would correspond to 1167 mM of the disaccharide sucrose without its metabolic conversion into RFOs. If this estimate is correct, then, roughly 85% of sucrose destined for export passes through intermediary cells and is converted into RFOs. Thus, intermediary cells seem to be more efficient in phloem loading in A. meridionalis than ordinary cells. This can explain the finding that phloem loading was severely inhibited in transgenic V. phoeniceum by the inhibition of the RFO synthesis in the phloem through antisense repression of galactinol synthase expression (McCaskill and Turgeon, 2007). The present study on A. meridionalis indicates that the residual driving force for transport in the transgenic V. phoeniceum may be the apoplastic loading of sucrose via ordinary companion cells. Obviously, in A. meridionalis, this loading component is responsible for the higher concentration of sucrose in the phloem compared with the cytoplasm of mesophyll cells (Voitsekhovskaja et al., 2006). It is noteworthily that Coleus blumei and Ajuga reptans, also often used as model symplastic phloem loaders in studies on phloem loading (Turgeon and Gowan, 1990; Sprenger and Keller, 2000), have minor vein configurations similar to that of A. meridionalis. It seems reasonable to assume that phloem loading in these species, as well as in many other representatives of various dicot families with similar configuration of the minor vein phloem, occurs in the same way as in A. meridionalis, i.e. that the intermediary cells in their minor veins are responsible for symplastic phloem loading of RFOs (and maybe some passive loading of sucrose with the bulk flow via plasmodesmata), and the ordinary cells for apoplastic phloem loading of sucrose, eventually causing phloem sucrose concentrations to be higher than those in the bundle sheath cells. In species with exclusively SE–ICCs in the minor veins such as C. melo (Holthaus and Schmitz, 1991) and some representatives of the Lamiaceae, Oleaceae, and other families (Gamalei, 1990; Batashev, 1997), the question about the presence and localization of the apoplastic phloem loading appears especially important as there seem to be no structures in their minor veins specialized for the energized uptake of assimilates from the apoplast. In conclusion, the data presented here demonstrate that differences in the structure of SE–CCCs are related to functional differences. Evidence is provided for the persistent idea that both symplastic and apoplastic pathways can function simultaneously during phloem loading (Fisher, 1986; Gamalei, 1990; van Bel et al., 1992; Turgeon et al., 1993), showing that in a large group of putative symplastic phloem loaders, these pathways are separated at the level of different SE–CCCs combined in their phloem endings. The question of how the combination of two loading strategies in a phloem ending may be beneficial for the process of phloem loading in these species awaits further investigation. Abbreviations Abbreviations CC companion cell CLSM confocal laser scanning microscopy IC intermediary cell OC ordinary cell GUS β-glucuronidase ORF open reading frame RACE rapid amplification of cDNA ends RFO raffinose family oligosaccharide SE sieve element SE–CCC sieve element–companion cell complex TEM transmission electron microscopy UTR untranslated region We are grateful to Professor H-W Heldt and Professor R Turgeon for stimulating discussions, and to Dr C Knop for providing AmSUT1 cDNA and Genome Walking libraries of A. meridionalis. This work was supported by a grant from the German Research Council (DFG) to GL, and grants from the Russian Foundation for Basic Research (#04-04-48388 and #07-04-01707) and from the Russian Ministry for Science and Education (#02.442.11.7101) to OVV. 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TI - Evidence for functional heterogeneity of sieve element–companion cell complexes in minor vein phloem of Alonsoa meridionalis
JO - Journal of Experimental Botany
DO - 10.1093/jxb/erp074
DA - 2009-03-25
UR - https://www.deepdyve.com/lp/oxford-university-press/evidence-for-functional-heterogeneity-of-sieve-element-companion-cell-0gxw1SzgEs
SP - 1873
EP - 1883
VL - 60
IS - 6
DP - DeepDyve
ER -