Sulfate distribution in Populus tremula x P. alba: identification of phloem localized sulfate transporter genes and environmental influences on their transcript level
Thesis submitted in partial fulfilment of the requirements of the degree Doctor rer. nat. of the Faculty of Forest and Environmental Sciences Albert-Ludwigs-University Freiburg im Breisgau, Germany
by Jasmin Dürr
Freiburg im Breisgau, Germany 2009
Dean of Faculty: Prof. Dr. H. Rennenberg Supervisor: Prof. Dr. C. Herschbach Second Reviewer: Prof. Dr. S. Fink Date of thesis’ defence: July 1st 2009 Content
Abbreviations
1 Introduction...... 1
1.1 Sulfate transport from the soil solution into the plant shoot ...... 4 1.2 Sulfate transport in the phloem ...... 7 1.3 Sulfur storage and mobilization ...... 11 1.4 Molecular characterization of sulfate transporters ...... 13 1.5 Why research on poplar? ...... 20 1.6 Aims and research questions ...... 21
2 Materials and Methods...... 23
2.1 Plant material...... 23 2.1.1 Plant propagation...... 23 2.1.2 Planting on soil...... 24 2.1.3 Growth conditions in the greenhouse ...... 24 2.1.4 Field-grown poplar trees ...... 24
2.2 Plant treatments and sampling...... 25 2.2.1 Sulfate deprivation treatment on clay culture ...... 25 2.2.2 Sulfate deprivation treatment on sand culture ...... 25 2.2.3 Plant protection ...... 26 2.2.4 Sampling of plant material ...... 26
2.3 Molecular biological methods ...... 29 2.3.1 Preparation of RNA...... 29 2.3.2 Isolation of genomic DNA ...... 31 2.3.3 Preparation of DNA free RNA...... 31 2.3.4 Determination of nucleic acid concentration and quality ...... 32 2.3.5 Separation of DNA segments by agarose gel-electrophresis...... 32 2.3.6 Separation of RNA segments by agarose gel-electrophresis...... 33 2.3.7 Isolation of DNA segments from agarose gels and DNA purification.... 34 2.3.8 cDNA synthesis...... 34 2.3.9 Standart DNA amplification via PCR ...... 35 2.3.10 DNA ligation into plasmid...... 36 2.3.11 Preparation of competent Escherichia coli cells ...... 36 2.3.12 Transformation of Escherichia coli strain INVαF’ ...... 37 2.3.13 Isolation of sulfate transporter sequences...... 38 2.3.14 Northern blotting...... 41 2.3.15 Quantitative real-time RT PCR ...... 46
2.4 Histochemistry – in situ hybridization ...... 55 2.4.1 Tissue fixation and embedding in paraffin ...... 56 2.4.2 Preparation of tissue slices...... 57 2.4.3 Synthesis of DIG-labeled RNA probes ...... 57 2.4.4 In situ hybridization with DIG labeled probes ...... 58 2.4.5 Immunological detection of hybridized DIG probes...... 60
2.5 Determination of sulfate...... 61 2.5.1 Sample preparation...... 62 2.5.2 Ion chromatographic measurements ...... 62
2.6 Bioinformatic analysis...... 64
2.7 Statistical analyses and calculations...... 65
3 Results...... 67
3.1 Populus tremula x P. alba sulfate transporter gene family...... 67 3.1.1 Characterization of cDNAs encoding putative Populus tremula x P. alba sulfate transporters ...... 67 3.1.2 Tissue specific expression analyses of the putative Populus tremula x P. alba sulfate transporter gene family ...... 73 3.1.3 Structure analyses of predicted sulfate transporter amino acid sequences...... 77
3.2 Cellular transcript localization of SULTR1;1 and SULTR3;3a...... 81 3.2.1 Cellular transcript localization of SULTR1;1 in leaves...... 81 3.2.2 Cellular transcript localization of SULTR1;1 in stem and roots with secondary growth...... 83 3.2.3 Cellular transcript localization of SULTR3;3a in leaves...... 87 3.2.4 Cellular transcript localization of SULTR3;3a in stem tissues and roots with secondary growth ...... 88 3.2.5 Transcript localization of sulfate transporter SULTR1;1 and SULTR3;3a in sink tissues, i.e. fine roots and the shoot apex...... 91
3.3 Influence of sulfur deprivation on sulfate transporter mRNA expression...... 94 3.3.1 Influence of sulfate deprivation on sulfate transporter transcript accumulation in bark and leaf vein ...... 96 3.3.2 Influence of sulfate deprivation on sulfate transporter transcript accumulation in stem wood...... 99
3.4 Seasonal influences on sulfate transporter gene expression ...... 102 3.4.1 Seasonal influences on sulfate transporter gene expression in bark tissue...... 104 3.4.2 Seasonal influences on sulfate transporter gene expression in leaves107 3.4.3 Principle component, correlation and regression analysis ...... 110
4 Discussion ...... 120
4.1 Populus tremula x P. alba sulfate transporter gene family...... 120 4.2 Tissue specific sulfate transporter gene expression ...... 123 4.3 Cellular transcript localization of two sulfate transporters in minor leaf veins and the lamina ...... 126 4.4 Sulfate transporter expression in the transport phloem and in bark ...... 128 4.5 Sulfate transporter expression in wood...... 133 4.6 Sulfate transporter expression in sink tissues...... 134 4.7 Influence of sulfate deprivation on the sulfate transporter transcript level.. 135 4.8 Seasonal variation in sulfate transporter expression ...... 138 4.9 Conclusion ...... 144
5 Summary ...... 146
6 Zusammenfassung...... 149
7 References ...... 153
Acknowledgement
Abbreviations
A adenine aa amino acids AP alkaline phosphatase APS adenosine 5’-phosphosulfate At Arabidopsis thaliana ATP adenosine triphosphate ATPase adenosine triphosphatase BCIP 5-bromo-4-chloro-3-indolyl phosphate bp base pairs BSA bovine serum albumine C cytosine Ca calcium
CaCl2 calcium chloride
Ca(NO3)2 calcium nitrate CC companion cell cDNA complementary DNA Ci curie
CoCl2 cobalt chloride CP crossing point CTAB cetyltrimethylammonium bromide CTP cytidine triphosphate
CuCl2 copper chloride
CuSO4 copper sulfate dATP deoxyadenosine triphosphate dCTP deoxycytidine triphosphate DEPC diethylpyrocarbonate dGTP deoxyguanosine triphosphate DIG digoxigenin DNA deoxyribonucleic acid DNase deoxyribonuclease dNTPs deoxynucleotide triphosphates dTTP deoxythymidine triphosphate EDTA ethylene-diamine-tetraacetic acid EF1beta elongation factor 1-beta EM electron microscopical et al. et alii (lat.), and others f forward FW fresh weight
FeCl2 iron chloride
FeSO4 iron sulfate G guanine g gram g acceleration of gravity γ gamma, wavelength GC guanidine-cytosine h hour H+ proton, hydrogen ion
H3BO3 boric acid HCl hydrogen chloride
H2O water
H2Odist distilled water
H2Odd deionised water i. e. id est (lat.), that is KCl potassium chloride
KNO3 potassium nitrate
KH2PO4 potassium dihydrogen phosphate KJ potassium iodide LB lysogeny broth LiCl lithium chloride LMM leaf margin meristem MAE (= MOPS) 3-(N-morpholino) propanesulfonic acid MDS membrane spanning domain MES 2-(N-morpholino) ethanesulfonic acid Mg magnesium
MgCl2 magnesium chloride
MgSO4 magnesium sulfate min minute
MnCl2 manganese chloride
MnSO4 manganese sulfate mRNA messenger ribonucleic acid N all four bases of DNA: ATGC NaCl sodium chloride
Na2CO3 sodium carbonate
NaHCO3 sodium hydrogen carbonate
NaH2PO4 sodium dihydrogen phosphate
Na2HPO4 disodium hydrogen phosphate
Na2MoO4 sodium molybdate
Na4P2O7 tetrasodium pyrophosphate
Na2SO4 sodium sulfate NBT nitroblue tetrazolium chloride
NH4NO3 ammonium nitrate OD optical density PCA principle component analysis PCR polymerase chain reaction PEG polyethylene glycol pH measure of the acidity or basicity of a solution PP polypropylene PVP polyvinylpyrrolidone PVPP polyvinylpolypyrrolidone Pta Populus tremula x P. alba r reverse RNA ribonucleic acid RNase ribonuclease rpm rotations per minute rRNA ribosomal RNA s second S sulfur SAM shoot apical meristem SD standard deviation SDS sodium dodecyl sulfate SE sieve element 2- SO4 sulfate SSC sodium chloride / sodium citrate STAS sulfate transporter anti sigma factor antagonist SULTR sulfate transporter gene or transcript SULTR sulfate transporter protein T thymine Tris tris-(hydroxymethyl) aminomethane tRNA transfer RNA U units UTR untranslated region UV ultraviolet % v/v percentage volume per volume % w/v percentage weight per volume x-gal 5-bromo-4-chloro-3-indolyl- beta-D-galactopyranoside
ZnCl2 zinc chloride
ZnSO4 zinc sulfate
Introduction
1 Introduction
Sulfur is one of six macronutrients (hydrogen, oxygen, carbon, nitrogen, phosphorus, sulfur), which plants require for growth and development. It is present in plants in oxidized form as sulfate that serves as the precursor for organic sulfur compounds but also as a storage form of sulfur (Martinoia et al., 2000). Sulfur in the oxidized form as a sulfonate group modifies a variety of metabolites like sulfolipids, proteins and polysaccharides (Kopriva, 2006). After assimilatory sulfate reduction the amino acid cysteine constitutes the first organic reduced sulfur compound (Figure 1.1). Cysteine is the metabolic precursor of other sulfur containing amino acids, such as cystine, homocysteine and methionine and also of many sulfur containing metabolites including phytoalexins and glycosinolates (Leustek and Saito, 1999). In addition, cysteine is a component in proteins as well as in peptides such as glutathione and its macromolecular derivatives the phytochelatins (Leustek et al., 2000).
The free sulfhydryl (H2S) group, called thiol group, of the cysteine molecule and its derivatives has a strong nucleophilic character and is capable of building a disulfide bridge with another thiol group. The reaction between the reduced thiol and the oxidized disulfide bridge built by two thiols is reversible. Therefore, nearly all aerobic forms of life, including plants, use this reaction as the predominant mechanism of redox control (Leustek and Saito, 1999). Beside biological redox processes, disulfide bridges have a central relevance in protein structures, functions and the regulation of enzyme activity (Leustek et al., 2000). For instance, sulfur serves as an active component in co-enzymes (vitamins) like biotin and thiamine, but also in the iron- sulfur-clusters of metalloenzymes (Leustek and Saito, 1999).
The most abundant low molecular thiol compound in higher plants is the tripeptide glutathione (γ-glutamylcysteinylglycin) (Bergmann and Rennenberg, 1993). A cytosolic glutathione concentration of about 0.3 mM has been determined in poplar leaves (Hartmann et al., 2003), whereas the highest relative glutathione content within Arabidopsis thaliana cells has been found in mitochondria (Zechmann et al., 2008). Mitochondria of both leaf and root cells contained 7-fold and 4-fold, respectively, higher glutathione levels than plastids while the density of glutathione 1 Introduction immunolabelling in the cytosol, nuclei, and peroxisomes was intermediate (Zechmann et al., 2008). Glutathione is relevant for many physiological processes in plants, for instance redox control, regulation of enzyme activities, protection against oxidative damage and pathogen defence (Rausch and Wachter, 2005). Glutathione is also involved in the detoxification of herbicides, xenobiotics and heavy metals (Lamoureux et al., 1993; May et al., 1998, Kopriva, 2006; Rausch et al., 2007). In many plants heavy-metal detoxification is mediated by glutathione derivatives called phytochelatins in which thiol groups serve as the metal ion ligand (Rauser, 1995; Marrs et al., 1996; Rausch et al., 2007). Additionally, glutathione serves as an important transport and storage form for reduced sulfur and as a signal for regulation of sulfur assimilation and cell division (Rennenberg et al., 1979; Herschbach et al., 1998; Noctor et al., 1998; Herschbach and Rennenberg, 2001; Potters et al., 2002; Kopriva and Rennenberg, 2004).
Many of the described metabolites contain sulfur in the reduced form. Therefore, sulfate has to be reduced in the assimilatory pathway to sulfide and subsequently bound to a carbon skeleton (Figure 1.1). This multistep pathway requires the transfer of eight electrons. Because sulfate is very stable, it has to be first activated with ATP by adenylation to adenosine 5’-phosphosulfate (APS) (Hawkesford and Wray, 2000) (Figure 1.1). This reaction is catalyzed by adenosine-5’-triphosphate (ATP) 2- sulfurylase. In a second step APS is reduced to sulfite (SO3 ) via APS reductase (Gutierrez-Marcos et al., 1996), with glutathione as the electron donor (Suter et al., 2000) (Figure 1.1). Subsequently, sulfite reductase transfers six electrons from reduced ferredoxin to sulfite for sulfide (S2-) formation. Finally cysteine is synthesized by binding sulfide to a carbon skeleton derived from serine (Kopriva and Rennenberg, 2004). Serine has to be activated with acetyl-CoenzymeA by serine acetyltransferase (SAT) before O-acetylserinethiol lyase (OASTL) transfers sulfide to O-acetylserine to synthesize cysteine plus acetate (Figure 1.1). SAT and OASTL form the multi-enzyme complex of cysteine synthase (Hell et al., 2002). APS reductase is the regulatory key enzyme of this pathway (Kopriva, 2006).
2 Introduction
Figure 1.1. Sulfate assimilation pathway in plants. Enzymes which are involved in the sulfate reduction process are shown. All enzymes are indicated in italic letters, whereas intermediates and products are shown in bold letters. Abbreviations: adenosine-5’-triphosphate (ATP), adenosine-5’- phosphosulfate (APS), reduced glutathione (GSH), ferredoxin (Fd) (modified image from Kopriva et al., 2007).
Whereas sulfate reduction is localized in plastids only, cysteine synthesis can take place in plastids, mitochondria and the cytosol, because the involved enzymes have been found in all three compartments (Renosto, 1993; Kopriva, 2006). The assimilatory sulfate reduction is supposed to be predominantly localized in the chloroplasts of leaf cells (Brunold, 1990) because of the availability of electrons from photosynthetic electron transport. But the enzymes involved are also present and active in roots (Kopriva et al., 2001) and even the apex of poplar trees is capable to reduce and assimilate sulfate (Herschbach, 2003). Sulfate can be converted into numerous metabolites in different plant parts and accordingly it has to be available in the respective tissues.
3 Introduction
1.1 Sulfate transport from the soil solution into the plant shoot
Sulfur is available to plants primarily in the form of anionic sulfate in the soil (Rennenberg, 1984). Therefore, sulfate is supposed to be the main sulfur form taken up into the roots. Transport of sulfate across the plasma membrane into root cells is an active process driven by a proton gradient, the so-called proton motive force, which is build up by a proton ATPase (Lass and Ulrich-Eberius, 1984). At the plasma membrane, the extracellular H+ concentration is generally higher than in the cytosol. These conditions enable the symport of H+ with other inorganic solutes into the cell. Sulfate uptake is supposed to be a symport of three protons together with one sulfate molecule (Lass and Ulrich-Eberius, 1984; Hawkesford et al., 1993; Smith et al., 1995). For many herbaceous plant species high affinity sulfate transporters have been identified, which are mainly responsible for the sulfate uptake from the soil into the roots. AtSULTR1;1 and AtSULTR1;2 from Arabidopsis thaliana (A. thaliana) are mainly expressed in the epidermis and cortex of roots (Yoshimoto et al., 2002). High affinity sulfate transporters with specific localization in root cells and sulfate uptake functions have also been identified from Hordeum vulgare HvST1, Lycopersicon esculentum LeST1;1 / LeST1;2, Zea mays ZmST1;1 and from the tropical forage legume Stylosanthes hamata ShST1 / ShST2 (Smith et al., 1995, 1997; Howarth et al., 2003; Nocito et al., 2006). Hydropathy analyses of the amino acid sequences revealed that plant sulfate transporters span the membrane twelve times (Smith et al., 1997, 2000; Takahashi et al., 1997), which is a wide spread structure for sulfate transporters (Hästbacka et al., 1994). Once inside the root symplast, sulfate can move radially to the central stele through the cortical and endodermal cells via plasmodesmata connections, building a cytoplasmic continuum, without traversing cell membranes. Furthermore, in higher plants nutrients are distributed within the whole plant by long distance transport via the vascular system. Allocation of sulfate from root cells to shoot cells occurs in the xylem (Figure 1.2). This upward transport is driven by the water evaporation / transpiration from leaves through the stomata. Because trees are connected with the very low water potential of the air and a moderate low water potential of the soil, the water loss in plant leaves diminishes the water potential of leaf cells which continues up to the root cells. Hence, water is sucked through the stable lignified xylem cells 4 Introduction which are connected to the xylem parenchyma cells of the root stele. Soluble compounds have to be loaded into the xylem before they are transported with the transpiration stream to the upper part of the plant. Thus, sulfate has to be transported out of the symplast of living root cells for loading into the xylem because the xylem is part of the apoplastic space. In Arabidopsis thaliana one member of low affinity sulfate transporters, AtSULTR2;1, is localized specifically in xylem parenchyma cells of roots and leaves as well as in the phloem of leaves (Takahashi et al., 2000). As sulfate transport depends on the proton motive force this sulfate transporter is supposed to be involved in sulfate uptake from the apoplast of the xylem and not vice versa. Consistently, its heterologous expression in a yeast mutant lacking own sulfate transporter protein revealed sulfate uptake into cells (Takahashi et al., 2000). Still, loading of the xylem by AtSULTR2;1 has been discussed due to its expression in parenchyma cells of roots surrounding xylem vessels. AtSULTR3;5 is co-localized together with AtSULTR2;1 and amplifies the sulfate uptake capacity into xylem parenchyma cells of roots (Kataoka et al., 2004b). For this purpose, these two well investigated transporters mainly manage retrieval of apoplastic sulfate into xylem parenchyma cells. However, sulfate transport out of xylem parenchyma cells and uptake into the xylem is largely unknown. Nevertheless, for many tree species it has been shown that sulfate is loaded into the xylem (beech: Kreuzwieser et al., 1996; oak: Seegmüller et al., 1996; poplar: Herschbach et al., 2000) where it is the main sulfur component (Herschbach and Rennenberg, 2001).
After reaching the shoot via the transpiration stream of the xylem, sulfate has to be transported into leaf mesophyll cells and further into plastids for assimilatory reduction. Therefore, several membranes have to be passed. Until now no sulfate transporter has been identified in leaf cell membranes. Probably sulfate that has been taken up into xylem parenchyma cells (Takahashi et al., 2000) reaches the leaf mesophyll cells symplastically via plasmodesmata. Leaf specific SULTR expression without precise cellular localization has been shown for BSULTR3;3, in Brassica oleracea (Buchner et al., 2004a) and for three investigated sulfate transporter sequences from Arabidopsis thaliana (AtSULTR3;1, AtSULTR3;2, AtSULTR3;3; Takahashi et al., 2000). Once inside the leaf cell, sulfate has to be translocated into the chloroplast for assimilatory reduction. But until now no sulfate transporter has been localized in the chloroplast membrane. One suggestion is that sulfate transport
5 Introduction into plastids is connected to phosphate influxe via a triose phosphate / phosphate translocator. Also a proton / sulfate symport mechanism or a phosphate / dicarboxylate exchanger is possible (Leustek and Saito et al., 1999; Hawkesford, 2003). The triose phosphate / phosphate translocator across the chloroplast membrane is an antiport mechanism which mediates the export of fixed carbon in form of triose phosphates in strict counter-exchange with inorganic phosphate or 3- phosphoglycerate (Flügge, 1992; Flügge et al., 2003). It has been shown that in spinach (Spinacia oleracea), sulfate is transported across the inner envelope membrane of chloroplasts in exchange with inorganic phosphate and phosphoglycerate. This observation gives evidence that a phosphate translocator or a similar mechanism is involved in sulfate influx into chloroplasts (Hampp and Ziegler, 1977).
Excess sulfate transported into leaf cells accumulates mainly in the vacuoles and constitutes a large internal sulfur reserve (Kaiser et al., 1989; Bell et al., 1994). Sulfate is the major anionic component of vacuolar saps (Leustek and Saito, 1999). Therefore, sulfate has to be transported across the tonoplast membrane. Two vacuolar sulfate transporters have been identified in Arabidopsis thaliana, AtSULTR4;1 and AtSULTR4;2. Both are localized in the tonoplast membrane of xylem parenchyma cells in roots and hypocotyls where they facilitate the efflux of sulfate from the vacuoles into the cell cytoplasm (Kataoka et al., 2004a). The importance of sulfate efflux from the vacuole for xylem loading under conditions of low sulfur supply has been demonstrated (Kataoka et al., 2004a). This sulfate transport across the tonoplast membrane is supposed to be driven by the electrical gradient between the vacuolar sap and the cytoplasm. The proton concentration of the cytosol is typically two orders of magnitude lower compared to the concentration measured in the vacuole (Kurkdjian and Guern, 1989). Furthermore, the H+-ATPase at the tonoplast membrane generates an inside positive membrane potential that serves as potential driving force for anion influx across the tonoplast into the vacuole (Martinoia et al., 2000). Physiological studies with isolated barley mesophyll vacuoles showed that sulfate uptake into the vacuoles is stimulated by addition of MgATP (Kaiser et al., 1989). This requirement allows the assumption that sulfate can be transported by a proton co-transport out of the vacuole and via the inside positive potential into the vacuole. Nevertheless, import of sulfate was saturable but,
6 Introduction depended on the sulfate concentration. This observation suggests that a specific transporter for the import of sulfate exists which requires the inside positive potential generated by H+-ATPase (Kaiser et al., 1989). The exact transport mechanism into vacuoles however is still unknown and needs to be investigated in further studies.
1.2 Sulfate transport in the phloem
In higher plants the major flow out of the leaves down to the roots and up to the apex takes place in the phloem (Figure 1.2). Phloem and xylem build together the vascular bundle and are located next to each other (Figure 1.2 C). The vascular cambium is situated between phloem and xylem in the stem of trees (Figure 1.2 B). The phloem consists of living nuclei free sieve elements (SEs) forming a conduit for mass flow called the sieve tube system. The angiosperm sieve elements lacking any apparent capacity for transcription or translation and are dependent on the neighbouring companion cells (CCs). Companion cells provide the cellular components that are required for maintenance and function of the sieve elements (van Bel et al., 2003; Lough and Lucas, 2006). This obligatory association is known as the sieve element- companion cell complex (Figure 1.2 D).
Sieve element and companion cell originate from one mother cell (Esau, 1969) and are connected by many plasmodesmata. The plasmodesmata gateway between sieve elements and companion cells differ from the general plasmodesmata structure. They are branched on the companion cell side of the common cell wall (van Bel et al., 1996). Beside small molecules like anions, organic acids and small peptides like glutathione, RNA molecules and proteins up to 20 kDa can be transported from companion cells to sieve elements via these connections (Imlau et al., 1999; Lee et al., 2003; Lough and Lucas, 2006). In the shoot apex the phloem exists at an early developmental stage not divided in sieve elements and companion cells and is part of the provascular strands (Figure 1.2 A). Equivalent undifferentiated phloem is found in the central cylinder of the root tip.
7 Introduction
Figure 1.2. The vascular plant architecture. Direction of the xylem (blue) and phloem (red) vascular flow is indicated by arrows (modified from Lough and Lucas, 2006). Histochemical transversal (B, C) and longitudinal (A) sections of the vascular system in stem (B, magnification x 238), roots (C, magnification x 183) and the shoot apex (A, magnification x 53) of plants are shown (modified from Beck, 2005). Electron micrograph of the sieve element (SE) - companion cell (CC) complex of Vicia faba at a sieve plate (D) (modified from van Bel et al., 2003). Abbreviations: shoot apical meristem (SAM), cambium (C).
Phloem loading in general occurs at the side of so-called ‘source tissues’, which can be different parts of a plant and which can differ due to environmental influences and developmental requirements. Typical 'source tissues' are related to the carbon / sugar household of plants. Leaves produce a surplus of sugar, which is loaded into the phloem and transported to the so-called 'sink tissues'. 'Sink tissues' for sugar are heterotrophic tissues like roots and meristems as well as storage tissues in the stem and roots. Plant parts which have a high energy demand due to high biosynthetic activities like the shoot apex including local developing leaves or seeds are also 'sink tissues'. The mass flow in the phloem is based on the hydrostatic pressure gradient that is build between source and sink tissues due to sugar accumulation in the phloem of source and sugar depletion in sink tissues (Münch, 1930; van Bel, 1995).
8 Introduction
According to these functions the phloem can be separated into the loading phloem of source tissues, the transporting phloem of major leaf veins, petioles, stem and roots with secondary growth (van Bel et al., 2003) and the releasing phloem of sink tissues. Within the mass flow generated by sugar transport, the transport of other nutrients including sulfate into upper and lower plant parts is supposed to take place. Beside carbohydrates, amino acids and many other compounds, also sulfate has been detected in the phloem sap of herbaceous plants (Rennenberg et al., 1979; Bonas et al., 1982) and deciduous trees like beech and poplar (Herschbach et al., 2000; Herschbach and Rennenberg, 2001). Sulfate is the predominant sulfur form in phloem exudates (Herschbach and Rennenberg, 2001) which requires loading into this tissue. Loading of sulfate into the phloem of leaves has been demonstrated for beech (Herschbach and Rennenberg, 1996) and poplar (Hartmann et al., 2000). In herbaceous plants, sulfur translocation via the phloem takes place mainly to the roots and the apex (Bonas et al., 1982). In perennial plants like deciduous trees, sulfur transported in the phloem is also unloaded along the transport path and is stored in tissues of the trunk (Herschbach and Rennenberg, 1995, 1996; Hartmann et al., 2000). Sulfur from developing poplar leaves is exported mainly into apical tree parts and only to a minor extent to basipetal plant parts (Hartmann et al., 2000). Thereby, sulfate is the only sulfur containing compound transported in apical direction. It seems that developing leaves and the apex of poplar trees are not completely dependent on the supply with reduced sulfur from mature leaves. This assumption is supported by the finding that all enzymes of the assimilatory sulfate reduction pathway are highly active in young developing poplar leaves (Hartmann et al., 2000). Even the apex tissue of poplar trees is capable to reduce and assimilate sulfate (Herschbach, 2003). In contrast to expanding leaves, mature and old mature leaves export 35S taken up via flap-feeding mainly in basipetal direction with the roots constituting the main sink. These results indicate that leaves of different developmental stages are source for different sink organs in trees which previously has been shown for carbohydrates in poplar (Dickson, 1991). Accordingly, sulfate distribution seems to be complex in trees and dependent on the developmental status of a particular organ.
9 Introduction
Phloem loading occurs either from the apoplast with specific transport proteins or symplastically via plasmodesmata. It is very likely that the phylogenetic development of apoplastic or symplastic phloem loading is species specific (van Bel, 1993) and even changes during tissue growth can be observed. Based on plasmodesmata frequency, many deciduous tree species are supposed to load the phloem symplastically (Gamalei, 1989). However, if symplastic diffusion would be the uptake mechanism for several solutes the accumulation of these compounds in the sieve element-companion cell complex would not be possible because solutes can not passively flow against a concentration gradient. An apoplastic loading would feature the advantage that sulfate and other solutes which are transported into the cytosol can not diffuse back and accumulate for further translocation by phloem mass flow. Phloem unloading can also precede symplastically via plasmodesmata or apoplastically with specific transport proteins (Lalonde et al., 2003). Symplastic phloem unloading driven by the large concentration differences of assimilates between sieve elements and importing sink cells seems to be more common (Fisher and Oparka, 1996; Patrick et al., 2001). For example in the case of developing seeds, the uptake of photoassimilates into seed cells from the maternal tissue precedes symplastically (Oparak and Gates, 1981; Patrick and Offler, 1995; Patrick et al., 2001). In some cases, symplastic transport is interrupted by an apoplastic step. In typical sink tissues, companion cells are very small or missing. Under these circumstances nutrients and other metabolites can leak directly from sieve elements into the apoplastic space where they can be taken up by the surrounding cells (van Bel, 2003). In developing seeds, sucrose release from maternal tissues occurs by facilitated diffusion through carriers in cereals (Porter et al., 1987; Wang and Fisher, 1995) and non-selective pores in pea (De Jong et al., 1996). A non-selective channel that is permeable to a wide range of electrolytes including large organic ions such as glutamate has been detected in release cells of Phaseolus seed coats (De Jong et al., 1997; Zhang et al., 2002). During the further transport course, sucrose influx into filial seed tissues of grain legumes (McDonald et al., 1996; Tegeder et al., 1999), temperate cereals (Bagnall et al., 2000; Weschke et al., 2000) and developing tomato fruits (Gear et al., 2000) is mediated by a sucrose / proton symport. In the seeds, sucrose transporter and H+-ATPase are co-localized to the plasma membranes of filial cells of the maternal-filial interface (Bagnall et al., 2000; Weschke et al., 2000; Patrick and Offler, 2001). Also a seed-specific amino acid / H+-symporter
10 Introduction
(AtAAP1) is expressed strongly in the endosperm and the cotyledons of developing Arabidopsis seeds (Hirner et al., 1998). In summery, there are many examples that transport proteins are necessary in the case of assimilate and nutrient accumulation in cells of sink tissues after phloem unloading in many plant species.
Also various sucrose and amino acid transporters, which are involved in phloem loading have been identified in many plant species (Hellmann et al., 2000; Patrick et al., 2001; Ramsperger-Gleixner et al., 2004; Carpaneto et al., 2005; Schmitt et al., 2008). Until now only one phloem specific sulfate transporter AtSULTR1;3 has been identified in Arabidopsis thaliana which seems involved in sulfate redistribution from source to sink tissues (Yoshimoto et al., 2003). This transporter is located in the sieve element-companion cell complexes of the phloem in cotyledons and roots. AtSULTR1;3 belongs to the group 1 of high affinity sulfate transporters and is essential for sulfate loading into the phloem of cotyledons to supply young sink organs with sulfate during early plant development (Yoshimoto et al., 2003). As these investigations were performed with ten days old seedlings of an herbaceous plant it remains questionable whether sulfate loading into the phloem of mature source leaves is mediated equally and whether these results are transferable to perennial plants such as trees. Because phloem development is at an early stage in the Arabidopsis thaliana seedlings, a clear differentiation between companion cells and sieve elements was largely not possible. Thus, the documented localization of AtSULTR1;3 in the hypocotyl and cotyledons of Arabidopsis thaliana phloem missed cell specificity. In summery, our knowledge about the mechanism of phloem loading and unloading with sulfate is still far from being complete and implies many open questions.
1.3 Sulfur storage and mobilization
Storage of starch and proteins (Zimmermann, 1974; Dickson, 1991; Cooke and Weih, 2005) as well as mobilization of sugar and amino acids is well established during the seasonal growth of deciduous trees (Bonicel et al., 1987, Sauter and van Cleve, 1992, 1994, Cooke and Weih, 2005). Starch is stored during late summer and mobilized during spring when current year flushes are developed (Sauter and van Cleve, 1994). Studies with several tree species showed that storage proteins 11 Introduction synthesized during autumn are localized in the vacuoles of rays and phloem parenchyma cells (Sauter and Wellenkamp, 1988; Sauter et al., 1994). These storage proteins disappeared during spring, when remobilization of nitrogen compounds in form of amino acids has been observed. As a consequence, the content of amino acids in the xylem sap increased, when buds became sink organs, and decreased after expansion in Salix (Sauter, 1981) as well as in polar trees (Sauter and van Cleve, 1992, Schneider et al., 1994).
Several observations support the idea, that also sulfur compounds are stored in trees during winter and are mobilized during bud break in spring (Herschbach and Rennenberg, 1996). During leaf senescence the sulfur fraction in beech leaves moved from living cells into the apoplastic space (Eschrich et al., 1988). In green leaves sulfur was mainly located in the cytoplasm of palisade parenchyma cells, bundle sheath cells and in sieve elements. During senescence, when beech leaves become brown, sulfur was shifted into apoplastic compartments, in particular into the sieve element wall, bundle sheath cell wall and the cell wall of palisade parenchyma cells. Simultaneously, the sulfur content in ray parenchyma and pith cells of beech stems increased (Eschrich et al., 1988). In principle, 35S-sulfate which was fed to beech leaves was mainly translocated into rays, medullary sheath and pith cells (Ziegler, 1965). It seems that sulfate is removed from senescent leaves for storage in stem tissues during dormancy. Storage of sulfur in living cells of the wood and bark can also be assumed from studies in which 35S-cysteine was fed to a mature beech leaf in summer. Under these conditions cysteine was partially metabolized to sulfate and both sulfate and cysteine were allocated through the phloem to basipetal sections of the trunk to storage tissues in the bark and wood (Herschbach and Rennenberg, 1995). Comparable results were found when 35S-sulfate was fed to a mature beech leaf in late summer (Herschbach and Rennenberg, 1996). The finding that 35S-sulfate or 35S-glutathione was not transported to apical tree parts in appreciable amounts led to the assumption that sulfate and glutathione was stored in living cells of wood and bark and did not exchange between phloem and xylem in beech trees. However, in poplar trees sulfate exchanged between phloem and xylem because interrupted phloem transport by girdling the stem apical to the fed leaf did not enhance 35S-sulfate transport to apical tree parts (Hartmann et al., 2000). Export of sulfate and of reduced sulfur from mature leaves and further storage in bark and
12 Introduction wood tissues is also evident for oak (Schulte et al., 1998) and poplar (Hartmann et al., 2000).
The 35S-sulfur fed during late summer to a mature beech leaf was retained in stem tissues throughout dormancy (Herschbach and Rennenberg, 1996). Until bud break 35S was mainly located in the wood of trunks and main roots while after leaf expansion 35S was found in the newly developed leaves. In spring cysteine and sulfate concentrations in the xylem sap of beech trees (Fagus sylvatica) increased and during bud break peak values of cysteine, glutathione and sulfate were observed (Schupp et al., 1991; Rennenberg et al., 1994). Whereas, cysteine was found to be the main thiol in xylem sap of beech, sulfate was the predominant sulfur compound (Rennenberg et al., 1994). All these investigations clearly show that sulfur stored along the tree axis in wood and bark during winter is remobilized during spring to fulfil the sulfur need of the developing leaves (Herschbach and Rennenberg, 1996). For this purpose, sulfur and especially sulfate from living cells in the wood and / or bark must be loaded from ray parenchyma cells, pith cells and from bark parenchyma cells into the xylem and / or phloem in order to reach the new developing leaves. Furthermore, sulfate has to be transported out of the xylem and/or phloem into the cells of sink tissues. It is totally unknown in which way these processes proceed in trees and whether transport proteins are involved. But obviously, this demand driven sulfate allocation is highly regulated during the seasons.
1.4 Molecular characterization of sulfate transporters
During its cycling in plants, sulfate has to pass various membranes which requires influx / efflux transport systems across these barriers during the uptake into roots, loading (efflux) into the xylem, further uptake into parenchyma cells of the shoot, phloem loading and unloading and finally transport into the chloroplast for assimilatory reduction as well as into the vacuoles for storage (Hawkesford, 2003; Buchner et al., 2004b). Since the first reported cloning of a plant sulfate transporter from Stylosanthes hamata (Smith et al., 1995), it became apparent that sulfate transport in plants is carried out by a complex system of transport proteins encoded by a large gene family. In recent years, many genes encoding sulfate transporters from a variety of herbaceous dicotyledonous plant species like Stylosanthes hamata, 13 Introduction
Arabidopsis thaliana, Lycopersicon esculentum, Brassica oleracea and also from monocotyledonous like Hordeum vulgare, Oryza sativa and Zea mays have been isolated and characterized (Smith et al., 1995, 1997; Takahashi et al., 1996, 1997, 1999, 2000; Bolchi et al., 1999; Vidmar et al., 1999, 2000; Shibagaki et al., 2002; Yoshimoto et al., 2002, 2003; Godwin et al., 2003; Howarth et al., 2003; Buchner et al., 2004a; Kataoka et al., 2004a, 2004b; Hopkins et al., 2004; Nocito et al., 2006). The deduced amino acid sequences of the cloned cDNAs from these species are closely related to each other. Based on their sequence similarity the sulfate transporter gene family was subdivided into five groups (Figure 1.3) (Hawkesford, 2003) and, specific responsibilities were investigated for several isoforms.
Sulfate transporters of group 1 are mainly responsible for sulfate uptake from the soil into the roots. High affinity sulfate transporters with specific localization in root cells and sulfate uptake function have been identified from Arabidopsis thaliana AtSULTR1;1 / AtSULTR1;2, Hordeum vulgare HvST1, Lycopersicon esculentum LeST1;1 / LeST1;2, Zea mays ZmST1;1 and from the tropical forage legume Stylosanthes hamata ShST1 / ShST2 (Smith et al., 1995, 1997; Yoshimoto et al., 2002; Howarth et al., 2003; Nocito et al., 2006). Sulfate transporters belonging to group 2 are discussed to be involved in xylem loading and unloading. AtSULTR2;1, is localized specifically in xylem parenchyma cells of roots and leaves as well as in the phloem of leaves (Takahashi et al., 2000). This transporter and also sulfate transporter AtSULTR2;2 which is expressed in phloem cells of roots and in leaf vascular bundle sheath cells are supposed to be responsible for sulfate uptake from the apoplastic space of the xylem in Arabidopsis thaliana. Group 4 sulfate transporters are supposed to be responsible for vacuolar efflux (Kataoka et al., 2004a). Two vacuolar sulfate transporters have been identified for Arabidopsis thaliana, AtSULTR4;1 and AtSULTR4;2. Both are localized in the tonoplast membrane of xylem parenchyma cells in roots and hypocotyls where they facilitate the efflux of sulfate from the vacuoles into the cell cytoplasm (Kataoka et al., 2004a). BSULTR4;1 from Brassica oleracea exhibits a different expression pattern in leaves of different age (Parmar et al., 2007). This transporter was detected in old leaves that may be sources for sulfate, but not in young leaves.
14 Introduction
Figure 1.3. Phylogenetic analysis of plant sulfate transporter amino acid sequences. Accession numbers: Arabidopsis thaliana (see Table 3.1); Triticum tauschii, TtST1;1A (AJ238244) and TtST1;1B (AJ238245); Hordeum vulgare, HvST1 (X96431); Brassica napus / juncacea, BnST1;1 (AJ416460), BnST1;2 (AJ311388), BnST2;1 (AJ633705), BjST (AJ223495), BnST3;1 (AJ581745), BnST5;1 (AJ311388), BnST4;1 (AJ416461); Zea mays, ZmST1;1 (AF355602); Stylosanthes hamata, ShST1(X82255), ShST2 (X82256), ShST3 (X82454); Sporolobus stapfianus, SsST (X96761); Lycopersicum esculentum, LeST1 (AF347613), LeST2 (AF347614); Solanum tuberosum, StST1 (AF309643); Oryza sativa (see Table 3.2). For all sequences with an asterisk (*), see DuPont patent application No. WO 00/04154. (Image taken from Hawkesford, 2003).
15 Introduction
In comparison to the sulfate transporters of group 1, 2 and 4, the sequences clustering to group 3 are scarcely characterized and more variable in their transcript localization. Expression analyses of group 3 transporters in Brassica oleracea show some examples of transcript accumulation specific in the stem and the roots (Buchner et al., 2004a). Only one sulfate transporter BSULTR3;3 of Brassica oleracea, accumulates specifically in leaves (Buchner et al., 2004a), while three mRNA sequences from Arabidopsis thaliana (AtSULTR3;1, AtSULTR3;2, AtSULTR3;3) are expressed in this tissue (Takahashi et al., 2000). AtSULTR3;5 is co-localized together with AtSULTR2;1 in xylem root cells of Arabidopsis thaliana. As mentioned above these sulfate transporters mainly facilitate the retrieval of apoplastic sulfate into the xylem parenchyma cells. Sulfate uptake into a yeast expression system was almost not detectable when AtSULTR3;5 was expressed heterologously. However, in cells co-expressing both AtSULTR3;5 and AtSULTR2;1, sulfate uptake capacity was approximately 3 times higher than with AtSULTR2;1 alone. Thus, AtSULTR3;5 supports the sulfate uptake capacity of AtSULTR2;1 (Kataoka et al., 2004b). The only investigated sulfate transporter from Lotus japonicus SST1 which is phylogenetically close to AtSULTR3;5 from Arabidopsis thaliana (Figure 1.3) is crucial for symbiotic nitrogen fixation by intracellular rhizobia within root nodules (Krusell et al., 2005). SST1 is expressed in a nodule specific manner and is located in the symbiosome membrane of Lotus japonicus nodules. The symbiosome consists of bacteria which are surrounded by a membrane built by the plant in infected nodules (Roth and Stacey, 1989). SST1 is absent when the symbiont is missing. Therefore, it is assumed that SST1 transports sulfate from the plant cell cytoplasm to the intracellular rhizobia, where the nutrient is essential for protein synthesis. So far the numerous sulfate transporters analyzed in different plant species exhibit highly tissue specific expression pattern, and therefore permit a controlled allocation of sulfate within the plant. The tissue specificity of sulfate transporters constitutes a potential for plants to regulate the sulfate demand and supply of particular tissues and enables responses to changing environmental conditions.
In most organisms including higher plants absorption and assimilation of sulfate is highly regulated. This regulation is considered to be mediated by negative feedback control, in which sulfate or products of the sulfate assimilation such as cysteine or
16 Introduction glutathione serve as signal molecules (Rennenberg et al., 1988, 1989; Herschbach and Rennenberg, 1991, 1994; Lappartient et al, 1999; Herschbach et al., 2000, Hartmann et al., 2004; van der Zalm et al., 2005). Accordingly, transport proteins are important steps to regulate the rate of sulfate uptake into a certain cell or tissue. Regulation may be possible at the side of proteins by feedback activation / deactivation, post-transcriptional control such as phosphorylation (Maruyama- Nakashita et al., 2004c) or by transcriptional control (Maruyama-Nakashita et al., 2006). Down-regulation of sulfate uptake has been observed due to accumulation of sulfate assimilation products such as cysteine or glutathione (Rennenberg et al., 1988, 1989; Herschbach and Rennenberg, 1991, 1994; Lappartient et al, 1999; Bolchi et al., 1999; Vidmar et al., 1999, 2000; Hartmann et al., 2004; Maruyama- Nakashita et al., 2004a). High concentrations of cysteine and glutathione induce the repression of sulfate transporter transcripts in roots for instance of HvST1 in Hordeum vulgare, AtSULTR1;1 and AtSULTR1;2 in Arabidopsis thaliana and ZmST1 in Zea mays (Bolchi et al., 1999; Vidmar et al., 1999, 2000; Maruyama-Nakashita et al., 2004a). However, heavy metal exposure like Cd, Zn and Cu induces transcript accumulation of ZmST1;1 and the sulfate uptake in Zea mays roots (Nocito et al., 2006). The glutathione content is probably involved in the signalling pathway triggering increased sulfate transporter expression by heavy metal exposure. The resulted induction of sulfate uptake may be a reaction to decreasing glutathione content, because of its participation in heavy metal detoxification (Nocito et al., 2006). In contrast, an increased uptake of sulfate that correlates with an increased sulfate transporter expression was found in response to O-acetylserine (OAS) (Hirai et al., 2003; Maruyama-Nakashita et al., 2004a; Hopkins et al., 2005). OAS is the precursor of cysteine that connects the S-metabolism with the N-metabolism and is high when sulfate availability is limited or when nitrogen content is in surplus (Hesse et al., 2004). Many sulfate transporters are transcriptionally regulated in response to the sulfur availability (Bolchi et al., 1999; Vidmar et al., 1999, Takahashi et al., 2000; Buchner et al., 2004a; Hirai et al., 2003; Nikiforova et al., 2003; Kataoka et al., 2004a; Maruyama-Nakashita et al., 2004a; Parmar et al., 2007). The transcripts of AtSULTR1;1 and AtSULTR1;2 from Arabidopsis thaliana accumulate as a response to sulfur-starvation. As mentioned above, these sulfate transporters are responsible for the sulfate uptake into epidermis and cortex cells of the roots (Takahashi et al.,
17 Introduction
2000; Vidmar et al., 2000). The increased mRNA level correlates with increasing sulfate influx capacity in roots. Upon re-supply of sulfate, the uptake capacity is repressed within hours with parallel decreasing mRNA and protein levels. These results also indicate a rapid turnover of sulfate transporter proteins (Rennenberg et al., 1989; Takahashi et al., 2000). Comparable results were observed for ZmSULTR1 in Zea mays and HvST1 in Hordeum vulgare roots (Smith et al., 1997; Bolchi et al., 1999). All together these investigations reveal the control of sulfate uptake via sulfate transporter regulation at the transcriptional level.
Furthermore, sulfate loading into the xylem also responds to the availability of reduced sulfur or sulfate (Herschbach and Rennenberg, 1991). In general the distribution of sulfate through the vascular system seems to be transcriptionally regulated. The low affinity sulfate transporters AtSULTR2;1 and AtSULTR2;2 located in the xylem parenchyma, respond to changes in the external sulfate supply in Arabidopsis thaliana roots and leaves (Takahashi et al., 2000). Also the transcript of ZmST2;1 is up-regulated in leaves (Hirai et al., 2003; Hopkins et al., 2004). Limitation of external sulfate caused also the accumulation of the phloem specific sulfate transporter AtSULTR1;3 mRNA both, in leaves and roots (Yoshimoto et al., 2003). In addition, the transcript level of the tonoplast localized sulfate efflux transporter AtSULTR4;2 accumulates in Arabidopsis thaliana roots, predominantly in the xylem vessels during sulfur limitation (Kataoka et al., 2004b). This may indicate sulfate remobilization from vacuoles of xylem parenchyma cells under sulfur limitation. Altogether, the precious findings give evidence that the amount of sulfate transported via the xylem and phloem is probably regulated on the transcriptional level of transport proteins involved in loading and unloading of the vascular system.
Transcriptional control is associated with SLIM1, a transcription regulation factor identified together with Arabidopsis thaliana AtSULTR1;2. This transcription regulation factor is crucial for proper growth under sulfate limited conditions (Maruyama-Nakashita et al., 2006). SLIM1 seems to play an important role in activation of sulfate uptake. The up-regulation of four sulfate transporter gene transcripts (AtSULTR1;1, AtSULTR1;2, AtSULTR4;2 and AtSULTR3;4) depends on SLIM1 beside several proteins involved in sulfur metabolism like ATP sulfurylase, a putative thioglucosidase and serine O-acetyltransferase. Recently, a novel cis-
18 Introduction element has been described which is involved in transcriptional regulation when sulfate, cysteine or glutathione were re-supplied after S-deficient growth conditions to Arabidopsis thaliana plants (Maruyama-Nakashita et al., 2005). This cis-element or so called sulfur responsive element (SURE) was identified in the promoter region of the sulfate transporter AtSULTR1;1 and has been found in several other genes that are inducible by S-starvation (Maruyama-Nakashita et al., 2005). Within the signal transduction in response to sulfur limitation several transcription factors (Nikiforova et al., 2003; Kasajima et al., 2007) and phytohormones may be involved. S-starvation co-ordinately affects the metabolism of flavonoids, auxin and jasmonate, which are signalling compounds (Nikiforova et al., 2003; Jost et al., 2005). It has been shown that the phytohormone zeatin, which belongs to the compounds of cytokinins, down-regulates AtSULTR1;1 and AtSULTR1;2 transcript accumulation in Arabidopsis thaliana roots (Maruyama-Nakashita et al., 2004b). Expression profiling of Arabidopsis thaliana seedlings with methyl-jasmonate, an important signalling compound in host-pathogen interactions, revealed a high proportion of genes related to the sulfur metabolism among the induced genes (Jost et al., 2005). Recent investigations comparing the two sulfate uptake transporters AtSULTR1;1 and AtSULTR1;2 in Arabidopsis thaliana roots suggest that sulfate transporter expression depends on different regulation factors (Barberon et al., 2008; Rouached et al., 2008). Split root experiments showed that the expression of AtSULTR1;1 is locally regulated in response to sulfur starvation whereas accumulation of AtSULTR1;2 appeared to be mainly related to the metabolic demand and seems to be controlled by the photoperiod (Rouached et al., 2008). Therefore, it has been assumed that sulfate transporter expression depends on various complex regulation networks and local influences. Posttranscriptional regulations of sulfate transporters have been observed in Arabidopsis thaliana (Yoshimoto et al., 2007; Maruyama-Nakashita et al., 2004c). AtSULTR1;1 and AtSULTR1;2 proteins accumulated during sulfur deprivation while their corresponding mRNA level was constant in roots (Yoshimoto et al., 2007). An involvement of protein phosphorylation / dephosphorylation has been discussed in the signal transduction pathway (Maruyama-Nakashita et al., 2004c). It seems that up-regulation of AtSULTR1;1 during sulfur limitation requires protein phosphatase as a key factor. Furthermore, a conserved STAS (sulfate transporter anti sigma factor antagonist) domain has been identified at the carboxyl-terminal end of most sulfate
19 Introduction transporter protein sequences (Shibagaki and Grossman, 2004). The STAS domain is important to form a tertiary structure involved in nucleotide triphosphate binding and protein-protein interaction. This domain is also necessary for the correct localization in the membrane and influences kinetic characteristics of the sulfate transporter activity. The latter has been demonstrated for the Arabidopsis thaliana sulfate transporter AtSULTR1;2 (Rouached et al., 2005; Shibagaki and Grossman, 2006). The STAS domain shares significant similarity with bacterial anti-sigma factor antagonists such as SpoIIAA of Bacillus subtilis (Aravind and Koonin, 2000) which is a key component of the regulatory network involved in the induction of sporulation in response to nutrient deficiency. Therefore, this domain is possibly involved in the regulation of sulfate transporters due to the nutrient conditions. The STAS domain seems to be an important sequence area for the functionality of transporter and, thus, constitutes a potential contact point to influence protein activity.
Obviously, beside transcriptional control sulfate allocation in plants is coordinated at the protein level. A complex network of various factors is involved containing specific regulation factors in response to S-metabolism, phytohormones, posttranscriptional mechanisms and interactions with protein sequence domains. Because of the numerous isoform genes with similar sulfate transport function but highly tissue and rather cell specific transcript localizations, the transcriptional level seems to be the first important regulation step. The rapid turnover of these proteins enables a fast increase or decrease of sulfate uptake in a particular cell type. The transcription regulation factors like SLIM and SURE allow a synchronous regulation of sulfur related genes in the nucleus. In this context it is of great interest how a perennial plant coordinates the sulfate allocation over long transport distances. The transcriptional composition is supposed to be adapted to the tree specific requirements in particular due to changing environmental conditions.
1.5 Why research on poplar?
The most prominent characteristics that make trees different from annual herbaceous plants are their long distance between shoot and root, the stable supporting structure and long lifespan. Wood has both supporting and conductive structure allowing trees to achieve immense sizes and to compete with smaller plants for light, water and 20 Introduction nutrients. These prerequisites enable trees to live for centuries. Perennial growth characteristics that influence many aspects of morphology and physiology as well as the long-term permanent growth pattern are tree-specific areas of research. Trees in general produce most of the terrestrial biomass and dominate most terrestrial ecosystems (Brunner et al., 2004b). Poplar trees are widespread in North America as well as in Europe and are therefore of ecological importance. Many poplar species are adapted to conditions of abundant nutrient supply and unlimited water availability. Under such circumstances poplar species achieve very high relative growth rates and consequently rapid biomass accumulation (Cronk, 2005). For this purpose, poplars are good source of fibre for paper, lumber, plywood and a possible source of biofuel.
The use of Populus as research model system for trees and perennial plant biology increased when the complete genome sequence of Populus trichocarpa was published. For molecular biological studies poplar offers many advantages in comparison to other trees: (I) The known complete genome sequence allows the identification of gene families via blast search and further cloning of the genes of interest. (II) Poplar trees grow in relatively short time to tree size and therefore enable their use in functional genomic studies (Brunner et al., 2004b). (III) Furthermore, poplar is one of a few tree species that can be vegetatively propagated which allows the work with cloned plant material of the same genetic properties. Finally, (V) the phylogenetic relationship of Populus and Arabidopsis is useful for comparative functional and genomic studies, and facilitates research on genome and gene family evolution in eudicots (Jansson and Douglas 2007). Apparently, Populus offers many possibilities to study questions that cannot be addressed in Arabidopsis and rice, the two herbaceous prime model systems of plant biology and genomics (Jansson and Douglas, 2007).
1.6 Aims and research questions
As discussed, it is obvious that sulfate distribution within plants is based on the tissue specific expression of sulfate transporter isoforms. But, it is also obvious that trees have different requirements than herbaceous plants due to their long live span and due to the great distance between shoot and root that needs up to 100 meters long distance transport and more. The storage and remobilization of sulfur due to 21 Introduction seasonal changes, nutrient limitation and various environmental factors require a flexible adaptation of sulfate allocation in trees. In particular the ability to survive harsh winter conditions is a highly adaptive trait specific for perennial growth. Therefore, Populus is supposed to be suitable for investigations on the seasonal regulation of storage and mobilization of nutrients like sulfate.
The aim of the present study was to enhance our understanding about the molecular background of sulfate allocation via the phloem in the deciduous tree species Populus tremula x P. alba.
For this purpose the following issues have been addressed:
(1) Possesses Populus tremula x P. alba a sulfate transporter gene family?
(2) Are sulfate transporter genes in trees tissue specific expressed and allow the expression pattern first conclusions about their responsibilities in sulfate cycling?
(3) Are sulfate transporters involved in long distance transport by specifically localization in phloem tissue for loading and unloading of sulfate?
(4) If sulfate transporters are expressed in the phloem, do differences exist between collection phloem, transport phloem and release phloem?
(5) Does nutrient limitation like sulfur deficiency influence the transcriptional level of sulfate transporters in storage tissues like bark, wood and mature leaves to induce sulfate redistribution?
(6) Have changing environmental conditions during the season an impact on sulfate transporter expression to regulate the sulfate distribution via the phloem for mobilization and storage processes?
22 Materials and Methods
2 Materials and Methods
2.1 Plant material
All experiments were carried out with plants of the wildtype poplar hybrid Populus tremula x P. alba, clone 717 1-B4, originating from the Institute National de la Recherche Agronomique (INRA, France). This hybrid of trembling poplar and white poplar is also known as Populus x canescens.
2.1.1 Plant propagation
The poplar plants were micropropagated in vitro as described by Strohm et al. (1995). For this purpose, four weeks old sterile in vitro grown plants, circa 10 cm in length were used. After removing roots and leaves, the stem was cut into small pieces including one node from which a new adventitious shoot developed. The cuttings were cultivated in 20 ml of a sterile, modified MS culture medium (Murashige and Skoog,
1962) [5 mM NH4NO3, 4.7 mM KNO3, 0.75 mM CaCl2 x 2 H2O, 0.38 mM MgSO4 x 7
H2O, 0.3 mM KH2PO4, 0.1 mM H3BO3, 0.112 mM MnSO4 x H2O, 0.036 µM ZnSO4 x 7
H2O, 5 µM KJ, 0.1 µM CuSO4 x 5 H2O, 1 µM Na2MoO4 x 4 H2O, CoCl2 x 5 H2O, 50 µM -1 FeSO4 x 7 H2O, 50 µM Na-EDTA, 1 mg l nicotinic acid, pyridoxal-HCl, Ca- panthotenate, thiamine-HCl, L-cysteine-HCl, L-glutamine, indole-3-butyric acid each, 10 µg l-1 biotin, 0.1 g l-1 myo-inositol, 20 g l-1 sucrose, 1.2 g l-1 MES (chemicals from Sigma-Aldrich GmbH, München, Germany or Merck, Darmstadt, Germany)] containing 7 g l-1 agar (Bacteriological grade, ICN Inc. Biomedicals, Eschwege, Germany) in a glass culture flask (Sigma-Aldrich GmbH, München, Germany) closed with a plastic cap. The freshly prepared cuttings had to be kept in darkness over night to avoid degradation of the hormone indole-3-butyric acid. Subsequently, cuttings were grown four weeks in a culture room under controlled conditions, which comprehended long day conditions (16 h light / 8 h dark) at a photosynthetically active radiation of 100 µmol m-2 s-1 (Fluora L 58W/55, Osram, München, Germany), at 25°C room temperature.
23 Materials and Methods
2.1.2 Planting on soil
For growth on soil sterile plants were transferred into pots (circa 10 cm height, length and width) with substrate consisting of one part autoclaved commercial potting soil (Floradur, Floragard GmbH, Oldenburg, Germany), one part perlite (Perligran, Knauf Perlite GmbH, Dortmund, Germany) and one part quartz sand (grain size 0.2-0.7) thirty days after micropropagation. The soil was treated for 1 h with 200 ml 0.15% fungicide solution (Proplant, Dr. Stähler GmbH, Stade, Germany) per pot before planting the poplar seedlings. During the first two weeks, plantlets were protected with a glass cover. Pots were regularly watered with tap water and supplied with 200 ml commercial fertilizer (3 g l-1 Hakaphos blau; COMPO GmbH, Münster, Germany) every other week.
2.1.3 Growth conditions in the greenhouse
Poplar plants were cultivated in a greenhouse (26 ± 5°C) under long-day conditions with a 16 h photoperiod provided by natural daylight and mercury-vapour lamps (Osram HQL 400; Osram, München, Germany), which supplying plants with photosynthetically active radiation between 300-500 µmol m-2 s-1.
2.1.4 Field-grown poplar trees
For seasonal analyses poplar plants were cultivated from February 2005 in a greenhouse as described (Chapter 2.1.3). In autumn 2005 trees with a height of approximately 150 cm were trimmed, planted into bigger pots (20 cm width, length and 25 cm height) and transferred for acclimation during winter into the field near the institute. In April 2006 these poplar trees were planted in a field site close to the Institute of Forest Botany and Tree Physiology on the airport campus (University of Freiburg, Germany), each in a hole of one square meter containing nutrient rich humus. During planting each poplar tree was fertilized once with 120 g of a commercial long-time fertilizer (Basa cote Plus 12 M, COMPO, Münster, Germany). Trees were grown under natural conditions, except for very dry periods the plants were watered with tap water.
24 Materials and Methods
2.2 Plant treatments and sampling
2.2.1 Sulfate deprivation treatment on clay culture
Forty days old sterile poplar cuttings were transferred on moistened burned clay bales (2-6 mm in diameter; Leca Ton, Leca, Lamstedt, Germany) after micropropagation (Chapter 2.1.1). Plants were grown in a greenhouse as hydroponic cultures with 25% Hoagland nutrient solution for five months (Hoagland and Arnon, 1950). Subsequently, plants were treated with 25% Hoagland solution without any sulfate in the growth medium. In the sulfate free medium all sulfate salts were replaced by chloride salts [1.25 mM KNO3, 2.5 mM Ca(NO3)2 x 4 H2O, 5 mM MgCl2 x
6 H2O, 0.25 mM KH2PO4, 2.29 µM MnCl2 x 4 H2O, 1.16 µM H3BO3, 0.08 µM CuCl2 x
4 H2O, 0.19 µM ZnCl2, 0.13 µM Na2MoO4 x 4 H2O, 22.5 µM Fe-EDTA (chemicals from Merck, Darmstadt, Germany]. Fine roots of the poplar plants were harvested after 14 days of sulfur depletion and frozen in liquid nitrogen.
2.2.2 Sulfate deprivation treatment on sand culture
Thirty days old sterile poplar cuttings were directly transferred into pots (circa 10 cm height, length and width) filled with quartz sand (grain size 0.2-0.7 mm). Prior to planting, the sand of each pot was treated for 1 h with 200 ml 1/40 Hoagland solution (Hoagland and Arnon, 1950) including 0.15% fungicide solution (Proplant, Dr. Stähler GmbH, Stade, Germany). During the first two weeks young plantlets were covered with a glass to protect them from drought. Each pot was regularly watered with 600 ml 1/40 Hoagland solution and the plants were cultivated in a greenhouse as described for three months (Chapter 2.1.3). Subsequently, the medium was changed to full nutrient solution, that was modified according to 25% Hoagland solution [0.6 mM KNO3, 1.3 mM Ca(NO3)2 x 4 H2O, 0.3 mM MgSO4 x 7 H2O, 1 mM MgCl2 x 6
H2O, 0.25 mM KH2PO4, 2.3 µM MnCl2 x 4 H2O, 10 µM H3BO3, 0.08 µM CuCl2 x 4
H2O, 0.2 µM ZnCl2, 0.2 µM Na2MoO4 x 4 H2O, 0.04 µM CoCl2 x 6 H2O, 22.5 µM Na-
EDTA, 22.5 µM FeCl2, pH 5.5 (chemicals from Merck, Darmstadt, Germany], and poplar plants were grown for another three weeks. All sulfate containing micro- nutrient salts were replaced by chloride salts. Sulfate deprivation conditions were started by replacing the full nutrient solution by sulfate free medium [full nutrient solution with 0 mM MgSO4 but 1.3 mM MgCl2 x 6 25 Materials and Methods
H2O]. Three plants were treated for 2, 5, 9, 14, 20 and 26 days, respectively, with sulfate free medium. Three control plants were continuously watered with full nutrient solution. Each period of sulfate deprivation treatment was started the number of treatment days before sampling, so that all plants had the same developmental stage and could be harvested at the same day. To get an idea about sulfate availability during the treatment, sulfate concentrations were measured in the flow through after watering by ion chromatography (Chapter 2.5).
2.2.3 Plant protection
If required, pesticides were used against powdery mildew (member of the order Erysiphales; most likely Erysiphe adunca) and spider mites (acarian; member of the family Tetranychidae, most likely for greenhouse pest Tetranychus urticae). If symptoms of mildew infection were recognized plants were treated twice in one week intervals with 750 µl l-1 Baymat® (Bayer Vital GmbH, Leverkusen, Germany). Trees infested with spider mites were sprayed once a week with 250 µl l-1 Vertimec (Novartis Agro GmbH, Frankfurt, Germany). Lice (member of the family Lachnidae) on field-grown poplar trees were combated with 20 ml l-1 Neudosan® Neu Blattlausfrei (Neudorff GmbH, Emmerthal, Germany).
2.2.4 Sampling of plant material
Poplar plants grown in the greenhouse were harvested between 8 am and 10 am. During the harvest, young trees were separated into the apex and the 10th and 11th leaf counted from the first leaf about 1 cm in length. Leaves were separated into major leaf vein, which is equivalent to the midrib, and the leaf lamina. Stem sections between the 10th and 12th leaf were separated into bark and wood. Additionally, roots were separated into fine roots (up to 1 mm in diameter) and roots with secondary growth; the latter were separated into root wood and root bark. For further analyses plant material was frozen in liquid nitrogen and stored at -80°C.
For in situ hybridization analyses small pieces (approximately 3 mm in length) of stem sections from young stem parts between the 4th and 5th leaf, from stem parts between the 10th and 11th leaf and from basal stem parts between the 25th and 27th leaf counted from the first leaf of 1 cm in length were harvested. Additionally, the 11th 26 Materials and Methods leaf, separated into the major leaf vein and lamina, fine roots, small sections of roots with secondary growth and the apex were sampled. All plant material was immediately treated for fixation (Chapter 2.4.1).
Sampling of field-grown plants, which was started at the end of August 2006 and finished in September 2007, was performed between 10 am and 11 am. Twigs and leaves from sixteen trees were sampled randomly. Whenever applicable, two leaves between the 7th and the 9th leaf and the bark from twig sections between the 5th and 15th leaf, all counted from the apex of the twig, were harvested. A sampling overview is given in table 2.1. Tissues were collected from one appropriate twig of three trees at the same sampling date. Dormant buds were removed from twigs. During spring, six young developing leaves were collected. Bark tissue from one-year-old twigs and additionally pooled bark samples from three currently developed twigs were taken in spring (Table 2.1). For further analyses plant material was frozen in liquid nitrogen and stored at -80°C.
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Table 2.1. Sampling dates and tissues harvested from field-grown poplar trees.
Sampling date Leaves Bark tissue
th th August 2006 28th two between 7th and 9th between 5 and 15 leaf
th th September 2006 6th two between 7th and 9th between 5 and 15 leaf 13th two between 7th and 9th between 5th and 15th leaf 20th two between 7th and 9th between 5th and 15th leaf 26th two between 7th and 9th between 5th and 15th leaf
th th October 2006 5th two between 7th and 9th between 5 and 15 leaf 11th two between 7th and 9th between 5th and 15th leaf 18th two between 7th and 9th 25th two between 7th and 9th between 5th and 15th leaf
th th November 2006 8th two between 7th and 9th between 5 and 15 leaf 22nd between 5th and 15th leaf
th th December 2006 21st between 5 and 15 leaf
th th January 2007 22nd between 5 and 15 leaf
th th February 2007 15th between 5 and 15 leaf 27th between 5th and 15th leaf
th th March 2007 12th between 5 and 15 leaf 27th between 5th and 15th leaf
th th April 2007 3rd between 5 and 15 leaf 11th between 5th and 15th leaf 18th six new developing (Ø 2 cm) between 5th and 15th leaf 26th six new developing between 5th and 15th leaf
th th May 2007 3rd six new developing between 5 and 15 leaf 18th four from 5th to 8th (mature) between 5th and 15th leaf additional: between 5th and 11th leaf of three currently developing twigs
th th June 2007 4th four from 5th to 8th between 5 and 15 leaf additional: between 5th and 11th leaf of three currently developing twigs 29th two between 7th and 9th between 5th and 15th leaf of new twig
th th July 2007 31st two between 7th and 9th between 5 and 15 leaf of new twig
th th August 2007 14th two between 7th and 9th between 5 and 15 leaf of new twig 28th two between 7th and 9th between 5th and 15th leaf of new twig
September 2007 13th two between 7th and 9th
28 Materials and Methods
2.3 Molecular biological methods
The molecular biological methods were performed with absolute pure water (H2Odd) which was filtered through a water purification system (Micro-TKA, TKA Wasseraufbereitungssysteme, Niederelbert, Germany) after demineralization. Pipet tips, glass ware, reaction tubes and, if possible, the prepared solutions were autoclaved for 20 min at 120°C. Two different centrifuges were used depending on reaction tube size (5402 Centrifuge, Eppendorf, Hamburg, Germany; Rotina 48 R, Hettich, Tuttlingen, Germany). All plant materials were homogenized with a mortar and pestle in liquid nitrogen.
2.3.1 Preparation of RNA
Total RNA extraction
Total RNA for gene isolation (Chapter 2.3.13) from roots, leaves, bark, wood and apexes of soil grown plants (Chapter 2.1.2, 2.2.4) and from roots of sulfate starved plants (Chapter 2.2.1, 2.2.4) and for seasonal analysis from leaves of field-grown poplar trees (Chapter 2.1.4, 2.2.4) was extracted from 90 mg homogenized plant tissue. The extraction was performed using a commercial RNA isolation kit (RNeasy Plant Mini Kit; QIAGEN, Hilden, Germany) according to the manufacturer’s protocol with the following minor modifications. The RLC extraction buffer included in the kit was complemented with 10 µl β-mercaptoethanol (Sigma-Aldrich, München, Germany) and 20 mg polyethylenglycol (Sigma-Aldrich, München, Germany) per ml RLC extraction buffer and neither the buffer nor the samples were incubated at 56°C.
Total RNA isolation according to Kolosova et al. (2004)
For expression analyses with the Northern blot technique, total RNA was isolated from 1.2 g homogenized plant tissue applying the method described by Kolosova et al. (2004). Aliquots of 0.4 g tissue from three plants were pooled. RNA samples used for real-time RT PCR analyses (field-grown poplar trees; Chapter 2.1.4, 2.2.4) were prepared in a diminished approach. For this purpose approximately 120 mg bark tissue from each tree was used separately. Details in the following protocol given in square brackets represent the adjusted extraction protocol for small sample size.
29 Materials and Methods
The water used was treated with 0.1% diethylpyrocarbonate (DEPC, Sigma-Aldrich, München, Germany) for 1 h and subsequently autoclaved at 120°C for 20 min. All solutions except Tris-buffer were prepared with DEPC treated water. Immediately before use, the extraction buffer [200 mM Tris-HCL, pH 8.5, 1.5% (w/v) lithium dodecylsulfate, 300 mM LiCl, 10 mM disodium salt EDTA, 1% (w/v) sodium deoxycholate, 1% (w/v) Tergitol Nonidet® P-40 (indicated chemicals from Sigma- Aldrich, München, Germany)] was supplemented with 1 mM aurintricarboxylic acid, 10 mM dithiothreitol, 5 mM thiourea and 2% (w/v) polyvinylpolypyrrolidone 40 (PVPP) (indicated chemicals from Sigma-Aldrich, München, Germany). The buffer was incubated at 50°C in a water bath to dissolve these components. Aliquots of 1.2 g [120 mg] homogenized plant tissue were transferred to 50 ml polypropylene (PP) tubes [2 ml safe seal micro tubes] (Sarstedt, Nümbrecht, Germany) and homogenized in 20 ml [1.2 ml] extraction buffer by vigorous shaking. The suspensions were shock-frozen in liquid nitrogen. After thawing at 37°C, the samples were centrifuged at 3500 [5700] x g for 20 min at 4°C. Subsequently, the supernatants were transferred into new tubes. One-thirtieth volume of 3.3 M sodium acetate (pH 6.1) and 0.1 volume 100% ethanol of the current approach volume were added. After homogenization the solution was chilled on ice for 10 min to precipitate polysaccharides. To remove polysaccharides the extracts were centrifuged at 3500 [11400] x g for 30 min at 4°C. The supernatants were transferred into new tubes. For RNA precipitation, one-ninth volume of 3.3 M sodium acetate (pH 6.1) and 0.6 volume of ice-cold isopropanol were added and samples were stored at -20°C over night. The RNA pellets were received after centrifugation at 5000 [5700] x g for 30 min at 4°C and, subsequently, dried for 15 min on ice. The pellets were re- suspended in 8 ml [400 µl] TE-buffer [10 mM Tris-HCl, pH 8.0, 1 mM EDTA] plus 8 ml [400 µl] 5 M NaCl by vortex mixing between 15 and 30 min. To remove residual polysaccharides 4 ml [200 µl] 10% cetyltrimethylammonium bromide (CTAB, Sigma- Aldrich, München, Germany) was added. After homogenisation at room temperature samples were incubated for 5 min at 65°C in a water bath. RNA was then extracted twice with an equal volume of chloroform / isoamylalcohol (24:1, v/v). After centrifugation at 4000 [9140] x g for 30 min at room temperature the supernatants were supplemented with one-quarter volume 10 M LiCl, mixed and kept at 4°C over night. After centrifugation at 4000 [4570] x g for 30 min at 4°C, the supernatants were
30 Materials and Methods removed with a pipette. The RNA pellets were dissolved in 2 ml TE-buffer and transferred in 2 x 2 ml micro tubes (Sarstedt, Nümbrecht, Germany). 0.9 volume ice- could isopropanol and 0.1 volume 3.3 M sodium acetate were added to precipitate the RNA at -20°C during 1 h. The RNA pellets were received after centrifugation at 16000 x g for 30 min at 4°C, washed in 1 ml 70% ethanol and collected by centrifugation for 10 min. RNA samples from the small preparation approach were collected at 16000 x g directly after LiCl precipitation, without an additional isopropanol precipitation step, continued by washing with 70% ethanol. After a drying step for 10 min on ice the RNA pellets were re-suspended in 200 µl to 1000 µl [60 µl] water. RNA samples were stored at -80°C.
2.3.2 Isolation of genomic DNA
Genomic DNA was isolated from 90 mg homogenized leave tissue using the Nucleon Phytopure Genomic DNA Extraction Kit according to the manufacturer’s protocol (Amersham Biosciences, Freiburg, Germany).
2.3.3 Preparation of DNA free RNA
RNA extracted from plant tissue (Chapter 2.3.1) can be contaminated with genomic DNA, which leads to an overestimation when gene expression is quantified by real- time RT PCR. Therefore, genomic DNA was eliminated in RNA samples used for real-time RT PCR analyses. For this purpose, 30 µl (15-30 µg) of the extracted RNA was treated with 5 µl (5 U) deoxyribonuclease I (DNase I, Fermentas GmbH, St. Leon-Roth, Germany). Subsequently, 5 µl 10 x reaction buffer including 25 mM
MgCl2 (Fermentas GmbH, St. Leon-Roth, Germany) and 10 µl DEPC-treated water (see Chapter 2.3.1) were added. The reaction was carried out for 30 min at 37°C. Enzyme activity was stopped by adding 5 µl 25 mM EDTA solution (pH 8) and enzymes were degenerated at 65°C for 10 min. RNA was precipitated with 200 µl 100% ethanol for 30 min at -20°C. After centrifugation at 16000 x g at 4°C for 10 min, the RNA pellet was air dried at room temperature and dissolved in 40 µl DEPC- treated water.
31 Materials and Methods
2.3.4 Determination of nucleic acid concentration and quality
The concentration of RNA and DNA was determined by measuring the absorption at
260 nm (OD260) with a spectrophotometer (Beckman DU® 650, Beckman, Germany). For this purpose, aliquots of 200 µl of 1:100 diluted samples were filled in quartz glass cuvettes (Suprasil, Hellma, Mühlheim, Germany). An OD260 of 1 is equivalent to 40 µg ml-1 RNA (2) and 50 µg ml-1 DNA (1) (Sambrook and Russel, 2003). A first purity control was achieved by calculating the absorption ratio at 260 nm to absorption at 280 nm, each minus the absorption at 320 nm (3). The value obtained should range between 1.6 and 2 for appropriate purity (Sambrook and Russel, 2003).
-1 (1) DNA concentration (µg µl ) = (OD260 nm – OD320 nm ) * 5 -1 (2) RNA concentration (µg µl ) = (OD260 nm – OD320 nm ) * 4
(3) Nucleic acid purity = (OD260 nm – OD320 nm ) / (OD280 nm – OD320 nm)
RNA quality was further checked by separating 0.5 µg RNA on a 0.8% (w/v) agarose (Molecular Biology grade, MP Biomedicals, Illkirch, France) gel using horizontal electrophoresis (Chapter 2.3.5). The concentration of RNA used for real time RT- PCR analyses and the plasmid concentration with ligated cDNA for the calibration curve were measured undiluted in 1 µl sample with a nano-spectrophotometer (NanoDrop® ND-1000, Peqlab, Erlangen, Germany). DNA amplification quality was further checked by separating 2-5 µl PCR product on a 0.8-2% (w/v) agarose gel (Chapter 2.3.5) and comparing the amplified DNA segment with a commercial DNA standard (FastRuler™ DNA Ladder Low Range or GeneRuler™ 1 kb DNA Ladder, Fermentas GmbH, St. Leon-Roth, Germany). This method enabled an estimation of the concentration.
2.3.5 Separation of DNA segments by agarose gel-electrophoresis
Horizontal agarose gel-electrophoresis allows the separation of DNA and RNA molecules according to their size. The amount and length of DNA segments can be estimated by comparison with DNA standards of known length and concentration. Depending on the expected segment sizes, gels were prepared with agarose (Molecular Biology grade, MP Biomedicals, Illkirch, France) concentrations between 32 Materials and Methods
0.8% and 2% in 0.5 x TBE-buffer [5 mM Tris-HCl, pH 8, 50 mM borate, 1 mM EDTA]. Higher agarose concentrations are useful for smaller sizes. Agarose was dissolved in 0.5 x TBE-buffer by heating in a microwave. After the solution had been cooled down to circa 40°C, 1 µl (0.1% (v/v)) ethidium bromide (Sigma-Aldrich, München, Germany) was added per 100 ml TBE-buffer. This solution was poured into the gel tray of the electrophoresis system (PerfectBlue Gelsystem Mini L, peqlab, Erlangen, Germany) which was equipped with a comb.
Ethidium bromide intercalates between the DNA double strand and emits fluorescent light after exitation by UV-light (γ = 210-316). RNA and DNA samples were mixed with one-fifth volume loading dye [50% (v/v) glycerol, 100 mM EDTA, 0.04% (w/v) bromophenol blue, 0.04% (w/v) xylene cyanol FF, (Sigma-Aldrich, München, Germany)] and loaded into the gel slots. An aliquot of 4 µl of a DNA standard (FastRuler™ DNA Ladder Low Range or GeneRuler™ 1 kb DNA Ladder, Fermentas GmbH, St. Leon-Roth, Germany) was loaded in one slot for size comparison. Nucleic acid segments were separated electrophoretically at 100 volt in a horizontal chamber (PerfectBlue Gelsystem Mini L, peqlab, Erlangen, Germany) filled with 0.5 x TBE- buffer. Detection and documentation was performed using an UV-light imaging system (InGenius, Syngene Bio Imaging, Cambridge, UK). Digital pictures were saved and printed as needed for documentation.
2.3.6 Separation of RNA segments by agarose gel-electrophoresis
RNA separation was performed under denatured conditions to avoid RNA degradation. All solutions were prepared with water which was treated for 1 h with 0.1% DEPC followed by autoclaving at 120°C for 20 min. All used materials were autoclaved or cleaned with ethanol in order to remove possible contaminations by ribonuclease. RNA was separated on a 1% (w/v) agarose (Molecular Biology grade, MP Biomedicals, Illkirch, France) gel containing 6% (v/v) formaldehyde in 1 x MAE- buffer [0.02 M MOPS, 5 mM sodium acetate, 0.5 mM EDTA] by gel-electrophoresis. Formaldehyde was added after the agarose was dissolved in 1 x MAE-buffer by heating in a microwave and cooled down to 40°C. The gel was poured into the tray and polymerized within 1 h. 15 µg RNA was mixed with twice the volume of RNA loading buffer [100 µl 10 x MAE-buffer, 175 µl (37%) formaldehyde, 500 µl formamidedeion., 1 µl ethidium bromide (chemicals from Sigma-Aldrich, München, 33 Materials and Methods
Germany), 25 µl loading dye (see Chapter 2.3.5)]. Samples were incubated for 10 min at 65°C to denature RNA strands followed by chilling on ice for 5 min. This treatment prevents the building of tertiary structures. RNA segments were separated by gel-electrophoresis performed at 100 volt for 100 min in 1 x MAE-buffer. The RNA was immediately transferred and fixed on a nylon membrane (see Chapter 2.3.14).
2.3.7 Isolation of DNA segments from agarose gels and DNA purification
If necessary, designated DNA segments were cut out of the agarose gel after electrophoretical separation (Chapter 2.3.5) using a scalpel. DNA was extracted from the gel slices and purified using the QIAquick® Gel Extraction Kit (MinElute gel extraction kit, Qiagen, Hilden, Germany) according to the manufacturer’s instruction. DNA segments applied directly from PCR amplification were extracted using the QIAquick PCR purification kit according to manufacturer’s protocol (Qiagen GmbH, Düsseldorf, Germany). This procedure removes any left over PCR components such as non-incorporated deoxynucleotide triphosphates (dNTPs).
2.3.8 cDNA synthesis
Complementary DNA (cDNA) constitutes DNA synthesized from a mature mRNA template in a reaction catalyzed by the enzyme reverse transcriptase. Using a Poly T primer or a random oligonucleotide primer, the total cDNA is consistent with all transcribed genes in a particular sample. cDNA can be used for expression analysis in real time RT-PCR or rather to determine the DNA sequence, which encodes for the amino acid sequence of a gene.
First strand cDNA synthesis was performed with SuperScript II Reverse Transcriptase (Invitrogen, Karlsruhe, Germany) from 2 µg total RNA according to the manufacturer’s instruction. With the aim of gene isolation, 2 µl Poly T primer [5´-TTT
TTT TTT TTT TTT V-3´] and for quantitative real time RT-PCR analyses 2 µl R12 primer [5´-NNN NNN NNN NNN-3´] with a final concentration of 500 nM were added to a 20 µl standard reaction volume. The mixture of RNA, DEPC-treated water (see Chapter 2.3.1), primer and 1 µl dNTP mix (10 mM dATP, dTTP, dCTP, dGTP each) was incubated for 10 min at 65°C in order to denature the RNA. After shock cooling on ice, 4 µl 5 x First-Strand Buffer (Invitrogen, Karlsruhe, Germany), 1 µl 0.1 mM 34 Materials and Methods dithiothreitol (Sigma-Aldrich, München, Germany) and 1 µl (200 U) of SuperScript II Reverse Transcriptase were added and subsequently gently mixed with a pipette by pulling up and down. For primer annealing this mixture was incubated for 10 min at 25°C. The transcription reaction proceeded for 1 h at 42°C and was terminated by incubation at 70°C for 15 min. For quantitative gene expression analyses via real time RT-PCR (see Chapter 2.3.15) a ‘no RT’ control was prepared by the equal procedure except adding the transcriptase enzyme. This control was required to check whether genomic DNA contamination was still present in the sample.
2.3.9 Standart DNA amplification via PCR
The 20 µl standard reaction approach contained 2 µl 25 mM MgCl2 and 0.4 µl dNTP mix (10 mM dATP, dTTP, dCTP, dGTP each). 4 µl 5 x reaction buffer (Promega, Mannheim, Germany) and 0.2 µl (1 U) Go Taq Flexi DNA Polymerase (Promega, Mannheim, Germany) or 1 µl 10 x reaction buffer (Fermentas GmbH, St. Leon-Roth, Germany) and 0.05 µl (1 U) Taq DNA Polymerase (Fermentas GmbH, St. Leon-Roth, Germany) were added. The final concentration of each forward and reverse primer was 100 nM. DNA template concentrations ranged from 20-150 ng. Deionized water was used to adjust to the final volume of 20 µl. All components were added in 0.2 ml PCR Thermo-Strip tubes (Thermo scientific, Waltham, USA) standing on ice. When PCR approaches with a volume smaller or larger than 20µl were performed, the volumes of all components were added accordingly. DNA amplification was carried out using a light cycler system (Mastercycler® or Mastercycler® pro, Eppendorf, Hamburg, Germany). The PCR program was started with 3 min incubation at 94°C in order to denature the DNA template. Each cycle was initiated by the incubation for 30 s at 94°C to separate the newly synthesised DNA strands. The annealing temperature was applied for 30 s and ranged between 50°C and 56°C depending on the primers used. Subsequent time for polymerization at 72°C was dependent on the amplified DNA segment (1 min per 1000 bp). Number of cycles was between 32 and 38. A final polymerization step was performed for 5 min at 72°C. Subsequently, the reaction mixture was cooled down to 12°C until further analyses.
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2.3.10 DNA ligation into plasmid
The DNA segments amplified via PCR (Chapter 2.3.9) for gene isolation were ligated into the pCR2.1 vector (Invitrogen, Karlsruhe, Germany) via T4 DNA ligase (Invitrogen, Karlsruhe, Germany). The Taq DNA polymerase produces an adenine-overhang that fits into the pCR2.1 vector gap. The ligation reaction was performed immediately after DNA amplification by standard PCR and extraction from agarose gels (Chapter 3.2.5) because the adenine-overhang is not stable and can truncate during longer storage. 10 µl ligation reaction mixture contained 1 μl 10 x ligase buffer (Invitrogen, Karlsruhe, Germany), 1 μl (1 U) T4 DNA Ligase, 0.5 µl (12.5 ng) vector DNA and an adequate amount of the amplified DNA segment. Approximately, 12-20 ng of DNA amplicon was used. The reaction mixture was adjusted to a final volume of 10 µl with H2Odd and incubated at 14°C for 16 h.
2.3.11 Preparation of competent Escherichia coli cells
Bacteria cells are able to import foreign DNA. This process can be amplified with
CaCl2 or MgCl2 which increases the permeability of the membrane. For enhanced adsorption of plasmid DNA and subsequent amplification in great quantities the
Escherichia coli (E. coli) cells were treated with MgCl2. All materials for the generation of competent E. coli cells were autoclaved for 20 min at 120°C and all steps were performed under sterile conditions (Hera safe, Heraeus, Stuttgart, Germany). An over night culture of E. coli INVαF’ cells (Invitrogen, Karlsruhe, Germany) was prepared with 10 µl of a glycerol culture [500 µl E. coli INVαF’ over night cell culture plus 25% (v/v) glycerol shock frozen in liquid nitrogen, stored at -80°C] added to 5 ml LB medium [5 g l-1 yeast extract, 10 g l-1 tryptone (SERVA electrophoresis GmbH, Heidelberg, Germany), 5 g l-1 NaCl] in a 50 ml PP tube (Sarstedt, Nümbrecht, Germany). Bacteria culture was grown at 37°C and 160 rpm (Thermoschüttler, Infors AG, Bottmingen, Swiss) over night. Subsequently, 100 ml LB medium was inoculated with 1 ml E. coli INVαF’ over night culture and grown in 250 ml conical flasks (Erlenmeyer flask) closed with aluminium foil at 37°C and
160 rpm for circa 3 h. When bacteria cell suspension exhibited an OD600 (spectrophotometer, Beckman DU® 650, Beckman, Germany) of approximately 0.4, it was centrifuged at 2500 x g for 10 min at 4°C. After removing the supernatant, the 36 Materials and Methods cell pellet was carefully suspended in 10 ml ice cold TSS solution [LB medium including 10% (w/v) PEG 8000 (sterile filtrated) (Sigma-Aldrich, München, Germany),
5% (v/v) dimethylsulfoxide, 50 mM MgCl2] by pipetting up and down. Aliquots of 100 µl of these competent cells were immediately filled in 1.5 ml safe seal micro tubes (Sarstedt, Nümbrecht, Germany), shock frozen in liquid nitrogen and stored at -80°C.
2.3.12 Transformation of Escherichia coli strain INVαF’
The DNA segment, ligated in the plasmid (Chapter 2.3.10), was transformed into E. coli INVαF’ competent cells (Chapter 2.3.11) according to the manufacturer’s instruction (Invitrogen, Karlsruhe, Germany) (Sambrook et al., 2003). Aliquots of 200 µl of the transformed bacteria cell suspension culture were plated on LB medium [5 g l-1 yeast extract, 10 g l-1 tryptone, 5 g l-1 NaCl] including 50 µg ml-1 kanamycin (ICN Biomedicals Inc., Ohio, USA) and 2% (w/v) agar in petri dishes (Sarstedt, Nümbrecht, Germany). The surface of the LB medium in petri dishes was coated with 50 µl x-gal solution [20 mg x-gal (5-bromo-4-chloro-3-indolyl- beta-D-galactopyranoside; Duchefa, Haarlem, Netherlands) in 1 ml formamide (Sigma-Aldrich, München, Germany)] using a glass spatula for subsequent blue-white screening. E. coli clones containing the plasmid were selected via kanamycin resistance. The blue-white screening is a molecular technique that allows detection of successful ligation of DNA segments into the cloning side of the vector plasmid. If the ligation was successful, the foreign DNA is inserted within the lacZ gene, thus disrupting the production of functional β-galactosidase. Hydrolysis of colorless x-gal by the β- galactosidase causes the characteristic blue color of the colonies, which shows that the colonies contain an unligated vector. White colonies indicate insertion of the plasmid including ligated DNA segment. Positive white bacteria colonies were tested via ‘colony PCR’ to make sure that the DNA segment inserted had the expected length. For ‘colony PCR’ little amount of white E. coli colony was taken as template and the vector specific primers M13 reverse (5’-CAGGAAACAGCTATGAC-3’) and T7 (5’-
TAATACGACTCACTATAGGG-3’) were used for standard PCR (Chapter 2.3.9). Amplified segments were detected on an agarose gel (Chapter 2.3.5). Isolation of plasmids was performed using a commercial plasmid preparation kit according to the manufacturer’s instruction (QIAprep spin miniprep kit, Qiagen, Hilden, Germany).
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2.3.13 Isolation of sulfate transporter sequences
Comparison of the 14 sulfate transporter protein sequences from Arabidopsis thaliana with the Populus trichocarpa genome database DOE Joint Genomic Institute (jgi) version 1.0 (http://genome.jgi-psf.org/Poptr1/Poptr1.home.html) via BLAST (basic local alignment search tool, http://genome.jgi-psf.org/Poptr1/Poptr1.home.html, scoring matrix BLOSUM62) resulted in 18 putative open reading frames for putative sulfate transporter sequences in the poplar genome. The selected DNA sequences corresponding to the putative sulfate transporter genes were amplified by PCR from the poplar hybrid Populus tremula x P. alba. Gene specific primers were designed for each gene of interest based on the genomic sequence information of the Populus trichocarpa database (Table 2.2). Plants cultivated in the green house for gene isolation were harvested after four months as described in chapter 2.2.4. First strand cDNA was isolated (Chapter 2.3.8) from transcribed RNA of different poplar tissues (fine root, leaf, bark, wood and apex) (Table 2.2) and from roots of sulfur starved plants (Chapter 2.2.1, 2.3.1). Most sulfate transporter sequences were amplified via two nested PCRs. Only three genes could be amplified with one PCR using a specific forward (f) and reverse primer (r) (see Table 2.2).
All amplification reactions were performed with Taq DNA polymerase (Promega, Mannheim, Germany) according to standard protocol for a 20 µl reaction volume (Chapter 2.3.9). An aliquot of 1 µl (circa 150 ng) first strand cDNA (Chapter 2.3.8) was used as template for the amplification reaction. To amplify SULTR1;1, SULTR3;2a and SULTR3;3a by one PCR specific forward primer (f) and specific reverse primer (r) was added. For the remaining genes Poly T primer (r 1) and the specific primer 1 (f 1) (Table 2.2) were added for the first nested PCR. All primers had a final concentration of 100 nM. The second amplification reaction for nested PCR was performed with an aliquot of 1 µl solution of the first PCR as template and the forward 2 (f 2) and reverses 2 (r 2) primer (Table 2.2). Annealing temperatures ranged between 50°C and 55°C depending on the primer pairs used (Table 2.2). The elongation time for polymerase chain reaction was between 2 and 3 min (at least 1 min per 1000 bases).
38 Materials and Methods
Table 2.2. Oligonucleotide primers, annealing temperatures (T), amplified segment length in base pairs (bp) and the cDNA used for amplification of putative poplar sulfate transporter sequences. cDNA isolated from tissues of sulfur starved plants (S-), forward primers (f) and reverse primers (r) are indicated.
Gene and Segment Annealing Oligonucleotide primer sequences cDNA primer in bp T
SULTR1;1 2116 54°C root S- Sultr_1.1g 5´-CGATATGGATATAAGGAGCCTG-3´ f Sultr_1.1x 5´-TTTGGAGAGCATATCATAATGCG-3´ r
SULTR1;2 1885 50°C leaf Sultr_1.2a 5´-AATGAAATTGATCCTGTAGGCAAC-3´ f 1 Sultr_1.2b 5´-AGAATATCGGAGGCTGGCATTC-3´ f 2 PolyT mix 5´-TTTTTTTTTTTTTTT V-3´ r 2
SULTR2;1a 1854 54°C leaf Sultr_2.1a 5´-GCCTACAGAAATCTTGTTCTTACC-3´ f 1 Sultr_2.1b 5´-TGCATTTGGACTATTTAGGTGGC-3´ f 2 Sultr_2.1y 5´-AGACTCAGGGTTTGAAATTAGTC-3´ r 2
SULTR2;1b 2110 54°C root S- Sultr_2.2a 5´-TCATGCCAGGAGATTCCCAACTG-3´ f 1 Sultr_2.2b 5´-AAGACTGTAATTTCATTCCTGCAC-3´ f 2 Sultr_2.2x 5´-TTACACAGGTTATGGTTTATATTAC-3´ r 2
SULTR2;2 1904 55°C root Sultr_3.9a 5´-CTGAATTCTCCTGATCCACCAG-3´ f 1 Sultr_3.9b 5´-CCTATTGCAAGAGCTCGGTAG-3´ f 2 Sultr_3.9z 5´-CGGTTAATTAAGCAAGAGGCCG-3´ r 2
SULTR3;1a 1850 55°C apex Sultr_3.3a 5´-GACCACTTCACGAAGATTCGTAC-3´ f 1 Sultr_3.3b 5´-CTATATTCAAGCTTTATTCCACCTC-3´ f 2 Sultr_3.3z 5´-AAGATTCACACTGTAAAACCTGTAC-3´ r 2
SULTR3;1b 1989 54°C apex Sultr_3.2a 5´-TCTTTCCTGATGATCCTCTCAGG-3´ f 1 Sultr_3.2b 5´-ATTCGTCTTAGGCATAAAATATTTCC-3´ f 2 Sultr_3.2x 5´-TTCGAAGACTCATGTATGAAACG-3´ r 2
SULTR3;2a 1289 52°C wood Sultr_3.5g 5´-ATGGGTAACCCCTACTATGAATG-3´ f Sultr_3.5y 5´-GTTTGATATACGGAGATACAAATTG-3´ r
39 Materials and Methods
Gene and Segment Annealing Oligonucleotide primer sequences cDNA primer in bp T [°C]
SULTR3;2b 1546 54°C apex Sultr_3.1a 5´-TGCAACAATAGTGGGTTTCATGG-3´ f 1 Sultr_3.1b 5´-GCCTTCAGCAACTTAAAGGGATC-3´ f 2 Sultr_3.1z 5´-ATTCGATGTAAGGAGATACACAAA-3´ r 2
SULTR3;3a 2245 54°C bark Sultr_3.4g 5´-GCCATGGAGCCTAACGCCAGC-3´ f Sultr_3.4z 5´-CCGCTTGATCAAGCTGATGCATG-3´ r
SULTR3;3b 2034 55°C apex Sultr_3.10a 5´-CAGCAATAGCTTGCAACCTGATC-3´ f 1 Sultr_3.10b 5´-GCACAATACAGAAGCTCAAGAGC-3´ f 2 Sultr_3.10z 5´-CTGTGTTCGTGTTCCAAGTCATG-3´ r 2
SULTR 3;4a 2122 54°C apex Sultr_3.7a 5´-GGAAACCACTCTCAGAATCACCA-3´ f 1 Sultr_3.7b 5´-CTCAGAATCACCACTGAGTCAATA-3´ f 2 Sultr_3.7z 5´-AGTTCTCTTTCTAACGGTCACG-3´ r 2
SULTR3;4b 2024 55°C apex Sultr_3.6a 5´-CCAGTAATGCCAGGCATGGAGA-3´ f 1 Sultr_3.6b 5´-CTGAGATCTTCTTCCCTGATGAT-3´ f 2 Sultr_3.6z 5´-AACTTGTAATGGTCAGCACATGTC-3´ r 2
SULTR3;5 1970 55°C apex Sultr_3.8a 5´-CCAATTCCAGCATGAATCCTACC-3´ f 1 Sultr_3.8b 5´-ATCCTACCCAGGTGAATTTCAAC-3´ f 2 Sultr_3.8z 5´-GAGAACCTATCGGCTAACTAGAC-3´ r 2
SULTR4;1 2351 54°C root Sultr_4a 5´-GCACCACCGGTGCGCATGG-3´ f 1 Sultr_4.1b 5´-ATCTCCCCACAATCTCTGTACC-3´ f 2 Sultr_4.1z 5´-GCTCAATATAAACAGAGTGAAGG-3´ r 2
SULTR4;2 2262 54°C root S- Sultr_4a 5´-GCACCACCGGTGCGCATGG-3´ f 1 Sultr_4.2b 5´-GTTATCTTCCAACAATGTCTGCC-3´ f 2 Sultr_4.2v 5´-GTTTGGTCCTTTGAGGGAA-3´ r 2
SULTR5;1 1304 54°C apex Sultr_5.1a 5´-ATCTAGGTACGTACATACCAATC-3´ f 1 Sultr_5.1b 5´-TGGCCTTAACATTATCCGTTGAC-3´ f 2 PolyT 5´-TTT TTT TTT TTT TTT V-3´ r 2
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Gene and Segment Annealing Oligonucleotide primer sequences cDNA primer in bp T [°C]
SULTR5;2 1449 54°C root S- Sultr_5.2a 5´-TTACTGGGGCAATCTATGGCGTG-3´ f 1 Sultr_5.2b 5´-TTACTTTTAGGTGTCACGGGTC-3´ f 2 PolyT 5´-TTT TTT TTT TTT TTT V-3´ r 2 most genes PolyT 5´-TTT TTT TTT TTT TTT V-3´ r 1
Amplified DNA pieces were separated on a 0.8% agarose gel while comparing the size with a DNA standard (GeneRuler™ 1 kb DNA Ladder, Fermentas GmbH, St. Leon-Roth, Germany) (Chapter 2.3.5). DNA segments were extracted from the agarose gel and ligated into the pCR2.1 vector (Chapter 2.3.7, 2.3.10). The plasmid was transformed into E. coli competent bacteria cells (Chapter 2.3.12) for reproduction and subsequently isolated. The coding DNA sequences of the cloned and isolated gene segments were identified (MWG Biotech AG, Martinsried, Germany).
2.3.14 Northern blotting
Northern blotting is a method to determine the transcript level of specific genes. During the procedure RNA is transferred and fixed on a nylon membrane after the separation by gel-electrophoresis. mRNA accumulation on the membrane is detectable via hybridization with radioactively labeled DNA probes. For expression analyses by Northern blots Populus tremula x P. alba plants were cultivated for ten weeks in the greenhouse (Chapter 2.1.2, 2.1.3, 2.2.4) or under sulfate deprivation as described (Chapter 2.2.2).
Fixation of RNA on a membrane
After separation by denaturing agarose gel-electrophoresis (Chapter 2.3.6), the RNA was transferred from the gel to a Hybond-XL™ nylon membrane (Amersham Pharmacia, Freiburg, Germany) by capillary blotting with 20 x SSC buffer [3 M NaCl, 0.3 M sodium citrate] using a TURBOBLOTTER™ (Schleicher & Schuell, Dassel, 41 Materials and Methods
Germany). The blotting construction was started with 14 layers of dry Blotting-Paper- Sheets (thickness BF4, Sartorius, Göttingen, Germany), followed by four dry and one wet Blotting-Paper-Sheets (thickness BF2 Sartorius, Göttingen, Germany), followed by the Hybond-XL™ nylon membrane and the RNA agarose gel on top. Papers and the membrane were trimmed to the same size than the gel and the wet paper was soaked with 20 x SSC buffer. When setting up the blotting construction, it had to be ensured that no air bubbles were present between the gel and the membrane. Above the gel three thin, wet (20 x SSC buffer soaked) paper layers were added and one additional paper long enough to put its ends in the 20 x SSC solution of the buffer tank. The plastic lid of the apparatus was added on top of the blotting construction to avoid evaporation. After 18 h incubation, the RNA was transferred from the gel to the Hybond-XL™ nylon membrane. The RNA was fixed on the membrane by backing at 80°C for 2 h in a heating oven (T 6060, Heraeus Instruments, Stuttgart, Germany).
Probe synthesis and labeling for Northern blot hybridization
A gene specific DNA segment was synthesized for each sulfate transporter sequence from Populus tremula x P. alba by PCR (Chapter 2.3.9) using gene specific oligonucleotide primers (Table 2.3). If the reverse primer is missing in table 2.3, the reverse 2 primer shown in table 2.2 was used. DNA segments were between 150 and 360 bp long and partly localized in the 3´ untranslated region to enhance hybridization specificity and avoid cross-hybridization with closely related genes. The already cloned 5.8s rRNA segment (Accession number: AY781281) was 479 bp long and was amplified using the primer pair 5.8s rRNA_for [5´-GAACCCGTGGCATGACAAGCTG-3´] and
5.8s rRNA_rev [5´-AACCACCGCTCGTCGTGGCGA-3´]. For DNA probe amplification, a 40 µl standard PCR approach was performed with an annealing temperature of 52°C, one minute elongation time and 38 cycles. Approximately 20 ng of each cloned SULTR sequence ligated in the pCR2.1 vector (Invitrogen, Karlsruhe, Germany) was used as DNA template (Chapter 2.3.13). The amplification product was purified (Chapter 2.3.7) and dissolved in 35 µl H2Odd. An aliquot of 2 µl purified DNA product was separated on a 1% (w/v) agarose gel to estimate the concentration of the amplified DNA segment (Chapter 2.3.4).
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Table 2.3. Oligonucleotide primers used for DNA probe synthesis. Probe length in base pairs (bp), forward (f) and reverse primer (r) for each gene are indicated. If no reverse primer is listed for the respective gene the reverse 2 primer (r 2) given in table 2.2 was used.
Gene Primer Oligonucleotide primer sequences Probe in bp
SULTR1;1 Sultr_1.1n 5´-AGCTGATTGGAGAGGACAAG-3´ f 283 SULTR 1;2 Sultr_1.2n 5´-AATCTTCCTCACGGTTGCTG-3´ f 268 Sultr_1.2z 5´-CAATGCAGAGCATACAATGCA-3´ r SULTR 2;1a Sultr_2.1n 5´-CTAGCCAACTTTGCGACG-3´ f 360 SULTR 2;1b Sultr_2.2n 5´-GGTAGCCAACTTTGTGACC-3´ f 333 SULTR 2;2 Sultr_3.9n 5´-GACGTGTCCAATGTAATGAAC-3´ f 274 SULTR 3;1a Sultr_3.3n 5´-TTTCATGCTTCACACATGCAG-3´ f 282 SULTR 3;1b Sultr_3.2n 5´-ATACTCGCAAACCAGACCCC-3´ f 288 SULTR 3;2a Sultr_3.5n 5´-CTAGTAAAGTCGAAGTTCATC-3´ f 317 SULTR 3;2b Sultr_3.1n 5´-CTAGAGAAGTCAAATTTCATG-3´ f 339 SULTR 3;3a Sultr_3.4n 5´-ATCGAGCTATGTGTGATAATG-3´ f 284 SULTR 3;3b Sultr_3.10n 5´-TTTTCAGCAACTTGAATGTAATG-3´ f 277 SULTR 3;4a Sultr_3.7n 5´-GTTACATCAGTCTAAAACACTC-3´ f 308 SULTR 3;4b Sultr_3.6n 5´-AAATGCTGGACTCGTTTGGC-3´ f 296 SULTR 3;5 Sultr_3.8n 5´-AACTTTAGCTGAAGTGCTCAG-3´ f 292 SULTR 4;1 Sultr_4.1n 5´-TAAAACCTCTGCTCCCAAGC-3´ f 325 SULTR 4;2 Sultr_4.2n 5´-GGATAAACCAAGTTTCTTCC-3´ f 326 SULTR 5;1 Sultr_5.1n 5´-TGCTCTGCGGTTTCAATAAC-3´ f 324 Sultr_5.1z 5´-GTTTGACAGTAAGTACAAAGG-3´ r SULTR 5;2 Sultr_5.2n 5´-GCCATGTCCTGTTCTCTAG-3´ f 150 Sultr_5.2z 5´-GAAAGATTCTAACTGGTTGCC-3´ r
DNA segments were radioactively labeled using the random prime method (Feinberg and Vogelstein, 1983). During synthesis short oligonucleotide primers (mostly hexaoligonucleotides) of random sequences anneal on different sides of the denatured, single stranded DNA template. The Klenow-fragment-DNA-polymerase recognizes these double strands as start for the amplification reaction and incorporates labeled nucleotides. Because of the exonuclease activity of the Klenow- enzyme, which only functions in the opposite direction (3’ to 5’) than the synthesis, the 43 Materials and Methods amplification stops at the next bound primer. Therefore, the labeled probe is a mixture of DNA segments of variable length. DNA segments were labeled with [α-32P] dATP (3000 Ci mmol-1, 10 mCi ml-1, Hartmann Analytic, Braunschweig, Germany).
For the labeling reaction the Strip-EZ™ DNA Kit was used (Ambion, Cambridgeshire, UK) as described in the manufacturer’s instruction. Subsequent to the labeling reaction, probes were purified on a sephadex chromatography column (MicroSpin™ G-50 column, Amersham Pharmacia, Freiburg, Germany) in order to remove unincorporated radioactive nucleotides. Columns had to be centrifuged for 1 min at 9300 x g (Universal 30 RF, Hettich, Tuttlingen, Germany) to remove storage buffer. Subsequently, the labeling reaction approach was transferred to the column and centrifuged at 9300 x g for 2 min. Labeled DNA segments in the flow through were denatured for 5 min at 95°C. The amount of labeling was checked by measuring the radioactivity in 1 µl of the flow through. For this purpose, 3 ml scintillation-cocktail (Optiphase HiSafe 3, PerkinElmer, Massachusetts, USA) was added to 1 µl sample in a liquid scintillation vial (Packard, Groningen, Netherlands). Radioactivity was measured with a liquid scintillation counter (Wallac 1409, Wallac, Turcu, Finnland) implicating a quench correction about 90% - 95%.
Northern blot hybridization RNA sequences of interest can be visualized within total RNA fixed on a nylon membrane by hybridization with gene specific radioactively labeled probes. Northern blot hybridization of putative sulfate transporter transcripts was performed at 65°C in a hybridization oven (PersonalHyb, Stratagene, Amsterdam, Netherlands). For this purpose, nylon membranes were transferred into hybridization tubes (Stratagene, Amsterdam, Netherlands). Subsequently, pre-hybridized with denatured herring sperm DNA (Promega, Mannheim, Germany) avoids unspecific DNA probe binding on the nylon membrane because herring sperm DNA saturates unspecific binding sites. Herring sperm DNA (10 mg ml-1) was denatured for 5 min at 95°C. 10 ml FSB buffer
[0.05 M NaP2O7 x 10 H2O, 0.115 M NaH2PO4, 0.5 M EDTA] including 7% SDS and 40 µl herring sperm DNA were added to the membrane followed by an incubation for at least 1 h at 65°C. Aliquots of 60-180 x 106 Becquerel of the labeled probe per ml FSB buffer were added to the membrane and incubated over night. Excess DNA probe was removed after hybridization. For this purpose, the membranes were washed three times under high stringency conditions by low SDS concentration 44 Materials and Methods with 10 ml FSB buffer including 1% SDS at 65°C for 45 min for each step. In case of hybridization with an 5.8s rRNA probe two additional washing steps each with 10 ml 0.1 x SSC [15 mM NaCL, 1.5 mM sodium citrate] including 0.1% SDS were performed at 65°C for 45 min. The moist membranes were air tight wrapped and stored in plastic bags.
Specificity of DNA probes for sulfate transporters was checked by dot blot hybridization of closely related sequences. For this purpose, the pCR 2.1 plasmid (Invitrogen, Karlsruhe, Germany) containing the respective sulfate transporter sequence was denatured for 5 min at 95°C. 0.2-2 µl (12 ng) of the plasmid DNA was dripped on a Hybond-XL™ membrane (Amersham Pharmacia, Freiburg, Germany) and subsequently fixed by backing 1 h at 80°C. These membranes were enclosed in the hybridization treatments.
Membranes were exposed for one to six days to a Phosphor Imaging plate in dependence on the radioactive intensity (Bio-Rad Laboratories, München, Germany; or Fujifilm, Düsseldorf, Germany). A Phosphor Imaging plate consists of a thin layer of special crystals doped with lanthanides. Radioactive radiation excites the crystals and a latent image of the sample is formed on the plate. The image was readout by scanning the plate with a red laser beam using a dual optical fiber (Molecular Imager FX, Bio-Rad Laboratories, München, Germany). Red light further excites the pre- excited phosphor particles causing emission of blue light (Eraser, Bio-Rad Laboratories, München, Germany). The intensity of this emission is measured with a photomultiplier and used to construct an image of the detected transcript. Results were analysed with Quantity one® 1-D software (version 4.6.3, Bio-Rad Laboratories, München, Germany) and the images on the phosphor plates were cleared with light.
The Ambion-EZ technology allows multiple stripping and re-hybridization of blotted membranes. Modified CTPs incorporated in the probe predetermine breaking points and are therefore relatively easy to remove at mild stripping conditions. In order to use each blotted membrane several times, probes were removed by stripping according to the manufacturer's instruction (Strip-EZ™ DNA Kit, Ambion, Cambridgeshire, UK).
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2.3.15 Quantitative real-time RT PCR
Real-time RT PCR is a method which allows quantification of gene expression at a particular time and in a particular tissue. PCR in the presence of a fluorescent dye enables the real time measurement of amplified DNA templates during the reaction. Because of previous reverse transcription of RNA into cDNA, the measured amount of a particular DNA sequence can be used to refer of the transcript level. The fluorescent dye SYBER Green, which intercalates in double stranded DNA increases proportional with the amplified DNA product. The amount is measured at the end of each cycle. Quantification is performed within the exponential DNA amplification phase because optimal reaction conditions are present during that time (Figure 2.1). DNA doubling is not limited due to the template amount and all components like primers and dNTPs are available in sufficient amounts. The measured crossing point (CP) value is corresponding to the number of PCR cycles necessary to achieve a constant fluorescence level at the exponential phase.
At CP value, each reaction tube has the same amount of new synthesized DNA. Absolute quantification is carried out with a calibration curve based on continuous dilutions of the DNA segment of interest ligated into a plasmid. The measured CP values of known concentrated DNA templates enable the quantification of the number of mRNA copies in the samples. Compared with the Northern blot analysis, real-time RT PCR is more sensitive. Therefore, genes expressed at low levels can be detected and slight differences in transcript amounts can be determined. It is also possible to measure large sample numbers simultaneously. In contrast, Northern blot technique is more useful to screen the expression of several genes in a small amount of samples.
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Figure 2.1. Example for real-time RT PCR amplification curves.
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Primer design and pre-experiments
The program Primer3Plus (http://www.bioinformatics.nl/cgi-bin/primer3plus/ primer3plus.cgi) (Rozen and Skaletsky 2000) was used to design and optimize primers for real-time RT PCR. All primers had similar lengths, the same GC content of approximately 40%, and an annealing temperature of approximately 60°C. Primers were designed for SULTR1;1, SULTR3;3a, SULTR4;1 and SULTR4;2 (Table 2.4). A sequence belonging to the elongation factor gene family (elongation factor 1-beta; EF1beta) was selected as reference gene, because it was shown that these genes undergo stable expression in plants (Brunner et al., 2004a; Nicot et al., 2005). EF1beta is involved in the translation process of mRNA into protein sequences and, within this process, elongation factors are responsible for the enzymatic delivery of aminoacyl tRNAs to the ribosome (Brands et al., 1986; Lauer et al., 1984). Primers for EF1beta used in the present study were designed by Henning Wildhagen (personal communication; Accession number: FJ372570).
Table 2.4. Oligonucleotide primers used for real-time RT PCR and amplified sequence lengths in base pairs (pb).
Gene Primer Oligonucleotide primer sequences Segment in bp
SULTR1;1 Sultr_1.1ss 5´-TTTATAACCCGTGCAGATAAGGAC-3´ 113 bp
Sultr_1.1uu 5´-CCTTTTAGCAAATGGTCACCAC -3´ SULTR3;3a Sultr_3.4rr 5´-GCCCCTCTTGTGTCTGTGATC-3´ 115 bp
Sultr_3.4tt 5´-TCCACGAGGGAGGATTTAGC-3´ SULTR4;1 Sultr_4.1r 5´-GGCACTGCGTATATATGATATCTGTC-3´ 114bp Sultr_4.1t 5´-AAACCTTACGACAAGTATTGCATTG-3´ SULTR4;2 Sultr_4.2r 5´- GAGGCAGGGCGTAGATTG -3´ 132bp
Sultr_4.2t 5´- GGAAGCAAGCCTTACAATGC -3´ EF1beta Elonga_for 5´-TGAGGATCTCTGGTGTCGAAG-3´ 100 bp Elonga_rev 5´-GTCTCAGCAGATGGAGGAGTG -3´
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To make sure that the designed primers are specific for the respective sulfate transporter sequence and, that the reverse transcriptase reaction was successful, the quality of primers and cDNAs were tested by standard PCR using Taq polymerase (Fermentas GmbH, St. Leon-Roth, Germany) (Chapter 2.3.9). The final concentration of each primer was 500 nM in a 10 µl reaction approach. 2.5 µl (25 ng) cDNA (ten fold diluted) from bark tissue was added as amplification template. In order to check if the cDNA sample is still contaminated with genomic DNA, 2.5 µl of either the 'no RT' (Chapter 2.3.8) reaction or genomic DNA was added instead of cDNA as template (Figure 2.2). The reactions were performed using a light cycler system and a standard program with 55°C annealing temperature and 15 s elongation time during 38 cycles. Amplified DNA segments were separated using a 1% or 2% agarose gel and visualized via UV absorption (Chapter 2.3.5) (Figure 2.2).
Figure 2.2. Quality test of primers and cDNA. Amplified DNA segments achieved by using primers for SULTR1;1, SULTR3;3a, SULTR4;1 and SULTR4;2 (Table 2.4) and different templates were separated on 1% or 2% agarose gels. As templates, either 25 ng bark cDNA (line 1), 'no RT' reaction (line 2), genomic DNA (line 3) or 50 ng bark cDNA (line 4) were used. A commercial DNA standard was applied (S) (FastRuler™ DNA Ladder Low Range, Fermentas GmbH, St. Leon-Roth, Germany).
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Resulting PCR fragments were 113 bp for SULTR1;1, 115 bp for SULTR3;3a, 114bp for SULTR4;1 and 132 bp for SULTR4;2. Primer specificity for all four sulfate transporter sequences were verified by PCR with bark cDNA as template because only one specific DNA segment was detected on the agarose gel with the expected size (Figure 2.2, line 1, 4). The primers designed for the elongation factor EF1beta were already established (Henning Wildhagen, personal communication). For SULTR1;1 and SULTR3;3a it was possible to chose a segment that enclosed an intron. These primers would amplify a longer PCR product if cDNA templates were contaminated with genomic DNA. The sequence length including an intron was 256 bp for SULTR1;1 and 195 bp for SULTR3;3a. Both fragments were not detectable when cDNA templates were used (Figure 2.2, line 1), but were amplified when genomic DNA was used as control template (Figure 2.2, line 3). Also the ‘no RT’ control did not show any amplified DNA segment (Figure 2.2, line 2). These results indicated that the designed primers are specific for the respective gene of interest and the RNA was not contaminated with genomic DNA.
Optimal primer concentrations for the real-time amplification reaction were established by testing primer concentrations of 0.6, 0.8, 1.2 and 1.8 µM using the LightCycler® 480 system (Roche GmbH, Mannheim, Germany) and the LightCycler® 480 SYBR Green I Master solution (Roche GmbH, Mannheim, Germany) in 10 µl reaction volume (see following chapter). Also different cDNA template concentrations (25 ng, 50 ng) were tested.
Real-time RT PCR mRNA contents were determined by real-time RT PCR using the LightCycler® 480 system (Roche GmbH, Mannheim, Germany), the LightCycler® 480 SYBR Green I Master solution (Roche GmbH, Mannheim, Germany) and the Multiwell Plate 384 (Roche GmbH, Mannheim, Germany). Specific amplifications of SULTR1;1, SULTR3;3a, SULTR4;1, SULTR4;2 and of EF1beta were performed using gene specific primers (Table 2.4). Total RNA was isolated (Chapter 2.3.1) and transcribed in cDNA after genomic DNA digestion (Chapter 2.3.3, 2.3.8).
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Figure 2.3. Example of melting peaks of amplified DNA. After real-time RT PCR amplification, samples were heated for denaturation. The melting peak exhibits the temperature which is necessary for DNA double strand separation in each sample. This temperature is dependent on the amplified segment length.
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10 µl reaction volume contained 2.5 µl (25 ng) cDNA (diluted ten times), 5 µl of LightCycler® 480 SYBR Green I Master solution, 1 µl containing both primers each with a final concentration of either 0.8 µM or 1.2 µM. Two technical replicates were always measured. The ‘no RT’ control of each cDNA used was included once in real- time RT PCR analysis. In addition, one reaction without primers was included to make sure that no unspecific amplification was measured. All solutions were stored on ice and intensive light exposure was avoided, because the SYBER Green dye is light sensitive. The multiwell plates were centrifuged at 3500 x g for 10 min and were adjusted in the LightCycler® 480 system. After a ‘hot start’ for 2 min at 95°C, 45 PCR cycles were performed with a 15 s melting step at 95°C, 15 s annealing time at 55°C and 15 s elongation at 72°C. For final polymerization, the reaction mixture was incubated 2 min at 72°C. After amplification, samples were heated to 95°C with simultaneous measurement of the decreasing fluorescent signal due to DNA denaturation. The resulting melting curve and the melting temperature are dependent on the length of the amplified DNA segment (Figure 2.3). Therefore, the melting temperature has to be similar for each reaction with the same primer. The detected DNA segments were analyzed once on a 2% (w/v) agarose gel (Chapter 2.3.5), gel eluted (Chapter 2.3.7) and subsequently sequenced (MWG Biotech AG, Martinsried, Germany) to demonstrate that the expected DNA was amplified.
Calculation of transcript quantity
A standard calibration curve for each gene was generated using 1 ng µl-1 (nano- spectrophotometer NanoDrop® ND-1000, Peqlab, Erlangen, Germany) of linearized plasmid containing the respective gene. In order to linearize the plasmids containing SULTR1;1, SULTR3;3a or SULTR4;2, they were digested with the restriction enzyme SpeI (Fermentas GmbH, St. Leon-Roth, Germany). The plasmid which contained the SULTR4;1 sequence was digested with XbaI (Fermentas GmbH, St. Leon-Roth, Germany) and the plasmid containing EF1beta was digested with EcoRV. For digestion, a 50 µl reaction approach with 1 µg plasmid DNA, 5 µl 1 x Tango™ buffer (Fermentas GmbH, St. Leon-Roth, Germany) and 1 µl (10 U) restriction enzyme was prepared and incubated over night at 37°C. Series of dilutions, 1, 0.1, 0.01, 0.001, 0.0001 pg µl-1, were prepared from the linearized plasmid.
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During real-time RT PCR the CP values of the standard calibration curve were measured and automatically correlated to the plasmid concentrations. The system calculates a calibration curve with the LightCycler® 480 software (version 1.5.0, Roche GmbH, Mannheim, Germany). Efficiency of the reaction was automatically calculated during each real-time RT PCR run corresponding to the amplification of the calibration curve (Figure 2.4). The amount of DNA product per PCR cycle should theoretically double in case of 100% PCR efficiency. Therefore, the optimal efficiency value is two. Efficiency of the performed real-time reactions varied between 1.77 and 2.51 depending on the gene, which is in the expected range (Table 2.5). The error values of the calibration curves were between 0.015 and 0.096 (Table 2.5, Figure 2.4).
Error: 0.0148 Efficiency: 1.84
Figure 2.4. Example for a real-time RT PCR standard calibration curve. Correlations between CP values and DNA template concentrations are presented. The trend line was calculated automatically by LightCycler® 480 software (version 1.5.0, Roche GmbH, Mannheim, Germany). The calculated reaction efficiency and error values are indicated (n = 2).
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Table 2.5. Efficiency and error values of standard calibration curves. Error values were calculated for measurement points of the standard calibration curve for the sulfate transporter sequences and the elongation factor 1-beta during the bark and leaf sample real-time RT PCR measurement. Values were calculated from two replicates for each run. Reaction efficiency was calculated due to the amount of DNA product after each cycle and should be around two.
SULTR1;1 SULTR3;3a SULTR4;1 SULTR4;2 EF1beta
1.84 2.21 1.89 1.89 1.98 Efficiency Bark 0.015 0.030 0.019 0.043 0.034 Error 1.88 2.45 1.84 1.77 2.51 Efficiency Leaf 0.027 0.096 0.021 0.039 0.071 Error
In order to convert DNA concentrations into copy numbers, the known size of the plasmid DNA that contains the gene of interest, which was used for the standard calibration curve, was determined and had to be multiplied with the average Dalton value of two nucleotides (616 Da). This value had to be divided by the Avogadro's number to get gram [g] molecules-1 also known as copy numbers (4). This copy numbers were then related to the quantity of RNA used for reverse transcription to get copies per µg RNA.
(4) Copy number [gram molecules-1] = (bp of plasmid + insert) * (308 Da * 2 nucleotides pb-1) / (Avogadro's number)
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2.4 Histochemistry – in situ hybridization
In situ hybridization is a method to localize a specific DNA or RNA sequence in tissues or single cells by using a labeled nucleotide strand complementary to the analyzed sequence (Figure 2.5). In this study a labeled RNA probe was applied to detect the two sulfate transporter transcripts SULTR1;1 and SULTR3;3a in poplar tissues at the cellular level. In situ hybridization experiments were carried out according to Cnops et al. (2006). All steps were performed with absolute pure water
(H2Odd), which was filtered with a water purification system (Micro-TKA, TKA Wasser- aufbereitungssysteme, Niederelbert, Germany) after demineralization. Water was autoclaved for 20 min at 120°C after 0.1% DEPC treatment for one hour.
Figure 2.5. Probe synthesis and in situ hybridization reaction scheme. Digoxigenin (DIG) incorporation in RNA strands during in vitro transcription (A), DIG-labeled probe binding during hybridization reaction (B) and anti-DIG-alkaline phosphatase (AP) binding with color reaction (C) are shown.
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2.4.1 Tissue fixation and embedding in paraffin
For in situ hybridization analysis plants were cultivated for ten weeks in the greenhouse under the described conditions (Chapter 2.1.3). Freshly harvested 3 mm plant tissue pieces (Chapter 2.2.4) were transferred into 10 ml glass vials with plastic lids (VWR, Darmstadt, Germany). Tissues were immediately covered with 4% paraformaldehyd (Sigma-Aldrich, München, Germany) in phosphate buffer saline
[PBS: 140 mM NaCl, 10 mM KCl, 6.4 mM Na2HPO4, 2 mM NaH2PO4, pH 7] including 0.01 % Triton X-100 (Sigma-Aldrich, München, Germany). Paraformaldehyd was used to fix RNA transcripts in place and to increase access for probes during hybridization. Samples were vacuum infiltrated using an exsiccator (Kartell, Noviglio, Italy) and a vacuum pump. The negative pressure during vacuum treatment removed the air from intercellular spaces. Samples were kept at 4°C over night. Subsequently, tissues were washed twice in PBS for 5 min. Tissue samples were exposed for adjacent dehydration to an increasing ethanol series [30%, 50%, 70% ethanol in PBS] and finally three times to 100% ethanol. For each step, samples were kept on ice for 30 min. Finally, samples were incubated in 100% ethanol over night at 4°C. Ethanol was replaced by Roticlear® (Carl Roth GmbH, Karlsruhe, Germany). Rising Roticlear® series consisting of 35%, 50% and 70% Roticlear in ethanol were added by following two final steps in 100% Roticlear®. Each step was kept for 30 min at room temperature. Subsequently, plant material was incubated over night in 42°C Roticlear® solution saturated with paraffin (Paraplast® Plus, Sigma-Aldrich GmbH, München, Germany). Paraplast® Plus was melted at 60°C. All further steps were performed at 60°C and all materials used in the following steps were pre-heated to 60°C, because Paraplast® solidifies immediately at lower temperatures. Replacement of Roticlear was performed by exchanging melted Paraplast® four times. This procedure was carried out over two days. For each step, vacuum infiltration was repeated several times in a 60°C aluminium block exsiccator. Negative pressure made sure that paraffin was distributed within the entire tissue. The glass vials were kept open during this procedure so that Roticlear® could evaporate. Samples were transferred to 19 x 16 mm Petri dishes (Sarstedt, Nümbrecht, Germany) and embedded in paraffin by cooling down to room temperature. The embedded plant pieces were stored at 4°C.
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2.4.2 Preparation of tissue slices
Embedded tissues (Chapter 2.4.1) were cut into thin slices. Depending on the tissue 7- 10 µm thick sections of embedded plant material were cut with a microtome (HM355, Microm, Walldorf, Germany). This work was done in an air-conditioned room. Paraffin blocks were stored on ice to avoid folding of paraffin slides during cutting.
Subsequently, the cuttings were expanded while swimming on H2Odd at 37°C in a water bath. Cuttings were transferred on coated microscope glass slides (SuperFrost® Plus, Menzel GmbH, Germany). Due to the coating, the slide surface is positive charged. This results in a better adhesion of the mostly negative charged protein rich cutting surface. The slides were dried on a heating plate at 37°C for 2 h and stored in a dust free box at 4°C.
2.4.3 Synthesis of DIG-labeled RNA probes
Specific RNA probes for SULTR1;1 and SULTR3;3a sequences were generated by standard PCR using Taq polymerase (Fermentas GmbH, St. Leon-Roth, Germany) (Chapter 2.3.9) and the primer pairs given in table 2.3. The sense primers for control hybridization (sense probe) were tagged with the promotor sequence for T3 polymerase [5´-AATTAACCCTCACTAAAGGGAGA-3´] and the antisense primers for probe synthesis (antisense probe) were tagged with the promotor sequence for T7 polymerase [5´-GCTTCTAATACGACTCACTATAGGGAGA-3´]. The sense probe has the same sequence than the target mRNA and therefore, it constitutes the control for specific signals caused by the target mRNA complementary antisense probe. As DNA template 20 ng of either SULTR1;1 or SULTR3;3a sequences ligated in the pCR2.1 vector (Invitrogen, Karlsruhe, Germany) were used. The standard PCR program was performed with minor modifications. For the first two cycles the annealing temperature was 52°C, for the following two cycles 60°C and for the last 32 cycles 64°C each with 30 s incubation.
After PCR amplification the solution was transferred on a 1% agarose gel to separate PCR products (Chapter 2.3.5). The segments received were eluted from the gel (Chapter 2.3.7) and used as template for a second PCR. This reaction was aimed to increase the amount of DNA. A 200 µl approach that contained the same primer pairs 57 Materials and Methods as before was applied to a standard amplification protocol with 64°C annealing temperature and 1 min elongation time. The PCR product was purified (Chapter 2.3.7) and suspended in 40 µl H2Odd. 2 µl of PCR product was used to calculate the concentration on a 1% agarose gel by comparison with a DNA standard (GeneRuler™ 1 kb DNA Ladder, Fermentas GmbH, St. Leon-Roth, Germany) (Chapter 2.3.4).
Amplified DNA sequences were transcribed in RNA strands and labeled with digoxigenin (DIG) during in vitro (Figure 2.5 A). The plant steroid DIG from Digitalis lanata is linked to uridintriphosphate (UTP) and can only be detected upon binding. The reaction approach for a 25 µl volume contained 0.8-1 µg template DNA, 2.5 µl DIG RNA labeling mix (Roche GmbH, Mannheim, Germany), 0.5 µl RNase inhibitor (40 U µl-1, Fermentas GmbH, St. Leon-Roth, Germany), 1 µl T7 or T3 RNA Polymerase (10-20 U µl-1, Fermentas GmbH, St. Leon-Roth, Germany) and 5 µl 5 x transcription buffer (Fermentas GmbH, St. Leon-Roth, Germany). The reaction volume was adjusted with water to a final volume of 25 µl. The transcription reaction was incubated for 2-3 h at 37°C. Thereafter, DNA templates were digested with 2 µl DNase I (1 U µl-1 RQ1 RNase free, Promega, Mannheim, Germany) at 37°C for 15 min. 4 µl yeast tRNA (10 mg ml-1, Ambion, Applied Biosystems, Germany) was added before digestion was started to avoid unspecific RNA segment digestion. RNA was precipitated after adding 2 µl EDTA [0.2 M, pH 8], 2.5 µl LiCl [4 M] and 75 µl ethanol within 1 h at -80°C. Subsequently, RNA was separated by centrifugation at 16000 x g for 30 min at 4°C (5402 Centrifuge, Eppendorf, Hamburg, Germany). The pellet was washed twice with 500 µl 80% (v/v) ethanol. After each washing step RNA was centrifuged for 5 min at 16000 x g. After drying the pellet, the DIG labelled RNA probe was dissolved in 100 µl water and probe quality was tested by separating 5 µl electrophoretically on a 1% (w/v) agarose gel (Chapter 2.3.5). The concentration was calculated upon measurement of the absorption at 260 nm with a spectrophotometer (Chapter 2.3.4).
2.4.4 In situ hybridization with DIG labeled probes
Pre-hybridization treatment was started by removing paraffin from tissue slices which were fixed on coated microscope glass slides (Chapter 2.4.2) by treating the samples three times with 100% Histo-Clear® (Biozym scientific, Oldendorf, Germany) for 10 min. Histo-Clear® was further replaced by a decreasing series of Histo-Clear® in 58 Materials and Methods ethanol. The percentage of Histo-Clear® was 60% (v/v) than 30% (v/v) followed by three times 100% ethanol. During each step, samples were incubated for 3 min at room temperature. Tissue slices on the microscope slides were re-hydrated in decreasing ethanol series [95%, 70%, 50%, 30%, 0% (v/v) in H2Odd including 0.85% (w/v) NaCl]. Subsequently, samples were immersed in 0.2 M HCl for 20 min at room temperature and washed again with 0.85 % (w/v) NaCl. Proteins were digested by proteinase treatment [125 µg ml-1 Pronase (Sigma-Aldrich GmbH, München,
Germany) in PBS: 140 mM NaCl, 10 mM KCl, 6.4 mM Na2HPO4, 2 mM NaH2PO4, pH 7] for 10 min at 37°C, which enhanced accessibility for the probe within the tissue slices during hybridization. This enzymatic reaction was stopped with 0.2% (v/v) glycerol followed by 5 min post-fixation with 4% (v/v) paraformaldehyd in PBS. Thereafter, slides were washed three times with PBS and one time with 0.85% (w/v) NaCl at room temperature to remove paraformaldehyd.
Before hybridization tissue slices on microscope glass slides were treated with 100 µl pre-hybridization buffer [50% (v/v) formamide (37%) (Sigma-Aldrich, München, Germany), 2 x SSC [0.3 M NaCL, 30 mM sodium citrate] including 40 U ml-1 RNase inhibitor (40 U µl-1 RiboLock™, Fermentas GmbH, St. Leon-Roth, Germany)] for 1 h at 50°C. Each glass slide was separately covered with a parafilm® strip (Brand, Wertheim, Germany) and slides were stored in a humid chamber during incubation. The humid chamber contained 4 x SSC [0.6 M NaCL, 60 mM sodium citrate] soaked paper towel. The solution on the slides was replaced by 100 µl hybridization solution [50% (v/v) formamide (37%), 4 x SSC, 10 x Denhardt solution [2 g l-1 Ficoll 400, 2 g l-1 polyvinylpyrrolidone, 2 g l-1 BSA (chemicals from Sigma-Aldrich GmbH, München, Germany)] 0.5 mg ml-1 yeast t-RNA (Ambion, Cambridgeshire, UK), 10% (v/v) dextran -1 sulfate and 40 U ml RNase inhibitor in H2Odd] including equal concentrations (20 ng µl-1) of either an antisense or a sense DIG labelled probe (Figure 2.5 B). The hybridization solution was heated to 65°C for 5 min to denature nucleotide sequences. Prepared slides were covered with parafilm® strips and incubated for 16 h at 50°C in a humid chamber kept in darkness.
Washing steps with twice 3 x SSC, once 1.5 x SSC and once 0.75 x SSC were performed in glass boxes at 50°C each step for 30 min to remove excess probe. Not hybridized single stranded RNA probe was eliminated with ribonuclease (20 µg ml-1
59 Materials and Methods
RNase A, Carl Roth GmbH, Karlsruhe, Germany) in TE-buffer [10 mM Tris-HCl, pH 8.0, 1mM EDTA] including 0.5 mM NaCl to reduce the background signal. After the second washing step, ribonuclease was added for 20 min at 37°C. Slides were equilibrated in TE-buffer for 20 min at 37°C before and after ribonuclease treatment. Further washing steps were performed with 0.3 x SSC and 0.1 x SSC each for 15 min. Subsequently, slides were incubated in buffer 1 [100mM Tris-HCl, pH 7.5, 150 mM NaCl]. Unspecific binding sites within the tissue were blocked with blocking solution (BS) consisting of 0.5% (w/v) bovine serum albumin (BSA, Sigma-Aldrich GmbH, München, Germany) and 0.02% (v/v) Tween-20 (Sigma-Aldrich, München, Germany) added in buffer 1 which improves antibody treatment against DIG (anti-digoxigenin- UTP-alkaline phosphatase, FAB fragments, Roche GmbH, Mannheim, Germany). The antibody reaction was performed in a humid chamber for 1 h at room temperature in darkness. For this purpose, each slide was treated with 100 µl antibody diluted 1:2000 in BS and covered with a parafilm® strip. Slides were washed once in blocking solution followed by washing five times for 20 min with buffer 1. Samples were equilibrated in detection buffer [100 mM Tris-HCl, pH 9.5, 100 mM NaCl, 50 mM
MgCl2] for subsequent immunological detection.
2.4.5 Immunological detection of hybridized DIG probes
After probe hybridization (Chapter 2.4.4) the fixed tissue on the slides was prepared for immunological detection of DIG, incorporated in the probe, with an antibody (Figure 2.5 C). Visualization of the antibody was based on the enzymatic alkaline phosphatase (AP) reaction with 5-bromo-4-chloro-3-indolyl phosphate (BCIP, Roche GmbH, Mannheim, Germany). For the AP colour reaction, samples were incubated over night with two substrates, which were nitroblue tetrazolium chloride (NBT, Roche GmbH, Mannheim, Germany) and BCIP each at a final concentration of 0.2 mM. The reaction was performed in 70 ml detection buffer [100 mM Tris-HCl, pH 9.5,100 mM NaCl, 50 mM MgCl2] at room temperature in darkness. AP is linked to the antibody against DIG, cuts off a phosphate from BCIP that starts the redox reaction of dephosphorylated BCIP and NBT (Figure 2.6). This reaction results in a blue / violet insoluble precipitate. Slides were washed twice for 5 min with water to stop the reaction, mounted with 50% (v/v) glycerol and covered with a coverslide. The coverslides were surrounded with nail polish for permanent fixation. Visual detection was performed with inverted light microscopes (Axioplan 2 imaging and Axiovert 200 M MAT, Zeiss GmbH, Göttingen, 60 Materials and Methods
Germany) and a dissecting microscope (Stemi SV 11 Apo, Zeiss GmbH, Göttingen, Germany). Images were taken with a digital camera (AxioCam MRc, Zeiss GmbH, Göttingen, Germany) and were documented with the AxioVision 4.6 software (Zeiss GmbH, Göttingen, Germany).
Figure 2.6. Redox reaction of BCIP and NBT. The alkaline phosphatase cuts a phosphate residue from the substrate 5-bromo-4-chloro-3-indolyl phosphate (BCIP) which results in a blue / violet precipitate by redox reaction with nitroblue tetrazolium chloride (NBT).
2.5 Determination of sulfate
Sulfate contents in tissue samples were determined by ion exchange chromatography, which allows the separation of different anions depending on the charge properties of the molecules. During chromatography the charged molecules interact with the column matrix. Anion exchange chromatography retains ions using positively charged functional groups which are bound to the stationary phase. The target anions are eluted by increasing the concentration of a similarly charged species that displaces the analytical ions from the stationary phase. The ions discharged and eluted can be measured by conductivity detected. 61 Materials and Methods
2.5.1 Sample preparation
100 mg PVP was added to 1 ml H2Odd in a 2 ml PP micro tube (Sarstedt, Nümbrecht, Germany), closed with a screw cap and soaked over night at 4°C. 50 mg homogenized tissue (Chapter 2.2.2, 2.2.4), was added and strongly mixed. After 1 h shaking at 4°C the samples were boiled in a water bath for 10 min to denaturize proteins followed by cooling down on ice. Samples were centrifuged at 16000 x g (5402 Centrifuge, Eppendorf, Hamburg, Germany) and 4°C for 10 min. The supernatants were used for ion exchange chromatography measurements. The flow through of the nutrient solution from sulfate deprivation treatments of poplar trees was used directly for sulfate concentration measurement (Chapter 2.2.2). Aliquots of 80 µl either undiluted or 1:3 diluted with H2Odd were filled into 200 µl glass micro-inserts (0.1 mm x 15 mm, VWR, Darmstadt, Germany). These tubes were placed into clear 2 ml liquid chromatography glass vials (VWR, Darmstadt, Germany) and closed with rubber coated septa. 50 µl of the samples were injected automatically into the ion exchange chromatography system.
2.5.2 Ion chromatographic measurements
Sulfate concentrations were quantified using the ion chromatograph system DX-120 (Dionex, Idstein, Germany) which operates on the principle of isocratic ion analysis and conductivity detection. The system was controlled by a workstation running the PeakNet software package (Version 4. 3, Dionex, Idstein, Germany). Samples were injected with an autosampler (AS3500, Thermo Separation Products, Piscataway, USA) into a sample loop of 50 µl volume. The chromatograph system was equipped with a first guard column (IonPac® NG1, 4 x 35 mm, Dionex, Idstein, Germany) which absorbs phenolic contaminations followed by a second guard column (RFIC™ IonPac® AG9-SC, 4 x 50 mm, Dionex, Idstein, Germany). The second guard column accumulates ions in the sample which reduces the retention time. The analytical column (RFIC™ IonPac® AS9-SC, 4 x 250 mm, Dionex, Idstein, Germany) separates anions via an isocratic gradient built by sodium bicarbonate buffer of 0.75 -1 mM NaHCO3 and 2 mM Na2CO3. The flow rate was 0.8 ml min . The separation of eluted anions was enhanced with a self-regenerating anion suppressor (ASRS® ULTRA II 4-mm, Dionex, Idstein, Germany) for improved detection. The suppressor 62 Materials and Methods enhances analyte conductivity while suppressing eluent conductivity (background). The suppressor is packed with an ion-exchange material that allows ions to migrate into and out-of a capillary. For anion suppressors the capillary wall is a cation- exchange material in the acid form. The ions required for eluent suppression are generated by continuous electrolysis of water. This results in a dramatic improvement in analyte detection limits. Anions were detected in a conductivity module and data were transmitted to the work station. Standard calibration curves for sulfate were generated using 1 mM Na2SO4 to prepare serial dilutions (25, 50, 100, 200, 400 and 800 µM) (Figure 2.7).
8x106 r = 0.999 7x106 p < 0.0001
6x106
5x106
6 4x10
peak area 3x106
2x106
1x106
0 0 100 200 300 400 sufate concentration (µM)
Figure 2.7. Standard calibration curve for sulfate. The standard calibration curve for sulfate was generated using 1 mM Na2SO4 to prepare serial dilutions (25, 50, 100, 200, 400 and 800 µM). Sulfate was measured using ion chromatography with conductivity detection. Data points are means of three individual measurements. Error bars indicate standard deviations. The solid line indicates the linear regression fit. The regression coefficient (r) is given in the graph.
63 Materials and Methods
2.6 Bioinformatic analysis
Sequence analyses of the isolated sulfate transporter amino acid sequences were performed with the open available software blast search (blastp; http://blast.ncbi.nlm.nih.gov/Blast.cgi), using the score matrix BLOSUM62 and the open gap penalty of 11 and gap extension penalty of 1 to create alignment. The statistical significance threshold for reporting matches against sequences of the database was a default value of 10, such that 10 matches are expected to be found merely by chance, according to the stochastic model of Karlin and Altschul (1990). If the statistical significance ascribed to a match is lower than the expect threshold, the match is significant (E < 0.01; Pearson, 2000). The multiple sequence alignments presented were obtained using ClustalW (http://www.ebi.ac.uk/Tools/clustalw2/ index.html, Thompson et al., 1994) with the same parameters described before. Sequence identity and similarity were analyzed by local alignment (http://www.ncbi.nlm.nih.gov/blast/bl2seq/wblast2.cgi, Tatusova and Madden, 1999) with the same parameters for gap and the same score matrix. Multiple sequence alignments and phylogenetic analyses were carried out using the molecular evolutionary genetics analysis program (MEGA 4; Tamura et al., 2007) including ClustalW2 (Thompson et al., 1994) with gap open penalty of 50 and gap extension penalty of 0.2 with the score matrix BLOSUM62.
Computed distances within and between indicated groups (MEGA 4; Tamura et al., 2007) constitute sequence diversity based on amino acid substitutions of the compared sequence sites. Poisson correction was chosen as distance estimation model and gaps inserted during alignment were treated by pairwise deletion. Poisson correction calculates the mean pairwise distance for the set of sequences under study. This distance (number of amino acid substitutions per site) is the proportion of amino acid sites at which the two compared sequences are different. The Poisson correction distance assumes equality of substitution rates among sites and equal amino acid frequencies while correcting for multiple substitutions at the same site. The distance within groups are arithmetic mean values of all pairwise alignment distances between taxa of one group. Distances between groups computed the average distances. Secondary structures within amino acid sequences were 64 Materials and Methods determined based on a combination of PSIPRED (Jones, 1999) and JNET (Cuff and Barton, 1999) available with the Quick2D bioinformatic toolkit (http://toolkit.tuebingen.mpg.de/quick2_d).
2.7 Statistical analyses and calculations
All graphs were plotted with the program package Origin® (version 6.1, OriginLab Corporation, Northampton, USA). Quantification of Northern blot results were evaluated using the Quantity one® 1-D software (version 4.6.3, Bio-Rad Laboratories, München, Germany). For standard mathematical calculations such as mean values and standard deviations, the program Microsoft Office Excel 2003 was used. All statistical analyses were performed using a statistic software package (SPSS® GmbH software, version 16, München, Germany).
The transcript abundance data in copy numbers per µg RNA and sulfate contents from seasonal analysis (Chapter 2.1.4, 2.3.15) were transformed (ln) to improve normality (Gaussian distribution). Data transformation resulted in normality and homogeneity of variances of data in conformity with assumptions of conventional statistical methods. Differences in transcript abundances as well as sulfate contents among dates were tested by analysis of variance (ANOVA) using a significance level of p < 0.05. Post-hoc tests were used to determine significant differences between adjacent values as indicated by ANOVA. Turkey test was applied, if the prerequisite for Turkey test application, i.e. that sample exhibited homogeneity of variances, was achieved. If sample variances were not homogeneous distributed, the Games- Howell-Test was applied. Bivariate data correlations were performed with transformed (ln) transcript (copy numbers per µg RNA) mean values and sulfate content mean values of the three tree samples of the respective dates using Pearson’s linear correlation function. The weather data (maximum, minimum and mean temperature, relative humidity and sun shine term) were available from a meteorological station in close vicinity (Station 10803 of the Deutscher Wetterdienst, Airport Freiburg, Germany). Meteorological variables were mean values of measured data in the period of 24 h before sampling.
65 Materials and Methods
Principal components analysis (PCA) was applied to summarize the variation of 5 variables (mRNA levels) and sulfate contents as well as the sampling dates. Principal components analysis is a multivariate technique of data reduction that analyzes the correlation matrix to compute a few independent factors that are linear combinations of the variables and that explain most of the variation in the data (Sokal and Rohlf 1995). Used data were standardized (Standardized value = value – mean value / standard deviation) which is recommended, if variables with different dimensions like sulfate content and transcript amount are compared. The results are shown in bivariate diagrams corresponding to the main factors extracted. The usefulness of the PCA was assessed by Kaiser-Meyer-Olkin’s measure of sampling adequacy (KMO). The KMO should be above 0.5 if variables are interdependent and a PCA is useful. The computed PCA score values of the two factor loadings were shown in a bivariate diagram and correlated with meteorological data (maximum, minimum and mean temperature, relative humidity and sun shine term day length).
66 Results
3 Results
3.1 Populus tremula x P. alba sulfate transporter gene family
3.1.1 Characterization of cDNAs encoding putative Populus tremula x P. alba sulfate transporters
In the present study, the gene family of sulfate transporters have been investigated in poplar trees. The known 14 sulfate transporter sequences from Arabidopsis thaliana (Table 3.1) were compared with the Populus trichocarpa genome database (http://genome.jgi-psf.org/Poptr1/Poptr1.home.html). This search indicated that there are 18 putative open reading frames located in the poplar genome. Predicted sequences encoding the putative 18 sulfate transporters were isolated from the poplar hybrid, Populus tremula x P. alba (Table 3.1, Chapter 2.3.13). The cloned cDNAs of SULTR1;1, SULTR3;3a, SULTR3;2a, SULTR4;1, and SULTR4;2 were full length, while the remaining 13 sequences were partial. The cloned cDNA segments were between 1304 bp and 2351 bp long (Table 2.2).
16 of the deduced polypeptides encoded by putative poplar sulfate transporter genes had significant identities (E < 0.01; Pearson, 2000) with sequences of proton-coupled sulfate transporters from Arabidopsis thaliana (http://blast.ncbi.nlm.nih.gov/Blast.cgi) that have been isolated and functionally characterized (Takahashi et al., 1997, 2000; Yoshimoto et al., 2002, 2003). Only the amino acid sequences of SULTR5;1 and SULTR5;2 did not exhibit a high level of statistical significance (E > 0.01) for the homology to a sequence in the Arabidopsis thaliana database TAIR (The Arabidopsis Information Resource; http://www.arabidopsis.org). But still high percentage of sequence identity were found by local alignment of SULTR5;1 and SULTR5;2 with the most similar sequence from Arabidopsis thaliana (Table 3.1). Sequence comparison and phylogenetic analysis of predicted poplar protein sequences to the known sulfate transporter families from Arabidopsis and rice indicated that the 18 sequences from poplar could be classified into at least five groups (Hawkesford, 2003) (Figure 3.1). 67 Results
100 PtaSULTR1.1
98 PtaSULTR1.2 AtSULTR1.2 63 100 AtSULTR1.3 40 AtSULTR1.1 100 OsSULTR1.3 OsSULTR1.1 100 OsSULTR1.2
93 100 PtaSULTR2.1a 99 PtaSULTR2.1b 97 AtSULTR2.1 PtaSULTR2.2 89 AtSULTR2.2 100 OsSULTR2.1 100 OsSULTR2.2
100 PtaSULTR3.1a
52 PtaSULTR3.1b 98 PtaSULTR3.2a 98 100 PtaSULTR3.2b 98 AtSULTR3.1 100 AtSULTR3.2 OsSULTR3.1 82 90 OsSULTR3.2
100 PtaSULTR3.5 AtSULTR3.5 97 OsSULTR3.5 96 OsSULTR3.6 92 100 PtaSULTR3.3a 100 PtaSULTR3.3b AtSULTR3.3
100 PtaSULTR3.4a 100 100 PtaSULTR3.4b AtSULTR3.4 98 OsSULTR3.4
100 PtaSULTR4.1 97 PtaSULTR4.2 AtSULTR4.1
100 100 AtSULTR4.2 OsSULTR4.1
94 PtaSULTR5.1 69 PtaSULTR5.2 98 AtSULTR5.2 OsSULTR5.2 100 AtSULTR5.1 95 OsSULTR5.1
0.2 substitutions per site
Figure 3.1. Phylogenetic analysis of sulfate transporter sequences. Neighbor-joining tree (MEGA 4; Tamura et al., 2007) from the multiple alignment (ClustalW; Thompson et al., 1994) of the amino acid sequences of Arabidopsis thaliana (Accession numbers, Table 3.1), rice (Accession numbers, Table 3.2) and the deduced amino acid sequences of the cloned Populus tremula x P. alba sulfate transporter gene family (Accession numbers, Table 3.1) are shown. The bootstrap values, expressed as a percentage, were obtained from 1000 replicate trees. Poisson correction was used as substitution model. Number of amino acid substitutions per site indicates the relation distance.
68 Results
The names of the cloned sequences were adjusted according to existing denotations from other plant species (Hawkesford, 2003). Putative sulfate transporters within one group exhibited closely related sequences and, therefore, clustered at the same branch. Groups exhibit relations based on phylogenetic distances (Figure 3.1).
Table 3.1. Populus tremula x P. alba and Arabidopsis thaliana sulfate transporter gene family. Sequence names and Accession numbers of the cloned poplar sequences and the known Arabidopsis genes are indicated. Putative full length poplar sequences were marked in bold letters. Similarity and identity of homologous amino acid sequences between Populus tremula x P. alba and Arabidopsis thaliana and between closely related poplar sequences are given. These comparative analyses were carried out by local sequence alignments (http://www.ncbi.nlm.nih.gov/blast/bl2seq/wblast2.cgi) (Tatusova and Madden, 1999). Groups reflect cluster built by phylogenetic analysis (see Figure 3.1).
Group Populus tremula x P. alba poplar P. to A. Arabidopsis thaliana Accession Identity / Identity / Accession Gene name Gene name number Similarity Similarity number 1 PtaSULTR1;1 DQ906929 75 / 87% AtSULTR1;3 AB049624 89 / 94% PtaSULTR1;2 DQ174472 79 / 89% AtSULTR1;2 AB042322 AtSULTR1;1 AB018695 2 PtaSULTR2;1a DQ906931 68 / 83% 87 / 93% AtSULTR2;1 AB003591 PtaSULTR2;1b DQ906933 71 / 85% PtaSULTR2;2 DQ174473 69 / 83% AtSULTR2;2 D85416 3 PtaSULTR3;1a DQ174470 77 / 89% AtSULTR3;1 D89631 90 / 96% PtaSULTR3;1b DQ906928 78 / 89% PtaSULTR3;2a DQ174469 63 / 80% 83 / 89% AtSULTR3;2 AB004060 PtaSULTR3;2b DQ906934 66 / 84% PtaSULTR3;3a DQ906924 78 / 89% 90 / 94% AtSULTR3;3 AB023423 PtaSULTR3;3b DQ906926 78 / 88% PtaSULTR3;4a DQ174467 79 / 90% 89 / 94% AtSULTR3;4 AB054645 PtaSULTR3;4b DQ174466 79 / 90% PtaSULTR3;5 DQ906927 66 / 81% AtSULTR3;5 AB061739 4 PtaSULTR4;1 DQ906930 77 / 88% 89 / 93% AtSULTR4;1 AB008782 PtaSULTR4;2 DQ906935 74 / 86% AtSULTR4;2 AB052775 5 PtaSULTR5;2 DQ174477 66 / 84% 89 / 94% AtSULTR5;2 NP_180139 PtaSULTR5;1 DQ174475 71 / 86% AtSULTR5;1 NP_178147
69 Results
PtaSULTR1;1 and PtaSULTR1;2 belonged to group 1 including high affinity sulfate uptake transporters (Smith et al., 1997; Vidmar et al., 2000; Shibagaki et al., 2002, Yoshimoto et al., 2002, 2003) (Figure 3.1). Three sequences, PtaSULTR2;1a, PtaSULTR2;1b and PtaSULTR2;2 clustered to group 2 of low affinity transporters (Takahashi et al., 2000) and nine, PtaSULTR3;1a, PtaSULTR3;1b, PtaSULTR3;2a, PtaSULTR3;2b, PtaSULTR3;3a, PtaSULTR3;3b, PtaSULTR3;4a, PtaSULTR3;4b, PtaSULTR3;5 sequences clustered to group 3 (Figure 3.1). Little is known about the function of the numerous sulfate transporters of group 3 except a demonstrated ability of SULTR3;5 to increase the rate of root-to-shoot sulfate translocation in Arabidopsis thaliana (Kataoka et al., 2004b). Both group 4 and 5 contained two poplar sulfate transporter proteins. PtaSULTR4;1 and PtaSULTR4;2 associated to group 4, while PtaSULTR5;1 and PtaSULTR5;2 associated to group 5 (Figure 3.1). Group 4 transporters from Arabidopsis thaliana are localized in the tonoplast and facilitate sulfate efflux from the vacuole (Kataoka et al., 2004a). Current investigations raised the discussion that members of group 5 transport molybdate instead of sulfate (Tomatsu et al., 2007). The poplar genome often harboured two sequences with high similarity around 93% on the amino acid level (Table 3.1 line poplar). These sequences were related to only one sulfate transporter from the Arabidopsis genome (Figure 3.1). Within group 2 and 3 the respective proteins were designated with the same number, plus indices “a” or “b”; for instance PtaSULTR3,3a and PtaSULTR3;3b. Rice sulfate transporter genes are represented within each group but always at a separated branch, distant from the two dicotyledonous plants (Figure 3.1).
Local sequence alignment analyses exhibited the relations of the respective poplar and known Arabidopsis thaliana sequences based on highest similarity (http://www.ncbi.nlm.nih.gov/blast/bl2seq/wblast2.cgi) (Tatusova et al., 1999). The sequence similarity of Populus tremula x P. alba sulfate transporters to the homologous Arabidopsis thaliana proteins was in the range of 80% to 90% and the identity ranged between 63% and 79% on the amino acid level (Table 3.1). Determined identities of the predicted amino acid sequences from the poplar hybrid and the respective denoted sequences from the Populus trichocarpa database were between 93% and 99%. Rice is a monocotyledonous plant which exhibit 13 predicted sulfate transporters and is phylogenetic more distant from poplar than Arabidopsis.
70 Results
Local alignments of the predicted amino acid sequences from rice with homologous sulfate transporters from Populus tremula x P. alba revealed similarities between 77% and 85%, while the identity was in the range of 57% and 75% (Table 3.2).
Table 3.2. Populus tremula x P. alba and Oryza sativa sulfate transporter gene family. Sequence names and accession numbers of the cloned poplar sequences and the known rice genes are indicated. Putative full length poplar sequences were marked in bold letters. Similarity and identity of homologous amino acid sequences between Populus tremula x P. alba and Oryza sativa are given. These comparative analyses were carried out by local sequence alignments (http://www.ncbi.nlm.nih.gov/blast/bl2seq/wblast2.cgi) (Tatusova and Madden, 1999). Groups reflect cluster built by phylogenetic analysis (see Figure 3.1).
Group Populus tremula x P. alba P. to O. Oryza sativa Accession Identity / Accession Gene name Gene name number Similarity number 1 PtaSULTR1;1 DQ906929 73 / 85% OsSULTR1;3 AF493790 PtaSULTR1;2 DQ174472 75 / 85% OsSULTR1;1 AF493790 OsSULTR1;2 XP_470587 2 PtaSULTR2;1a DQ906931 60 / 78% OsSULTR2;1 AAN59769 PtaSULTR2;1b DQ906933 63 / 80% PtaSULTR2;2 DQ174473 64 / 79% OsSULTR2;2 AAN59770 3 PtaSULTR3;1a DQ174470 65 / 80% OsSULTR3;1 NP_921514 PtaSULTR3;1b DQ906928 67 / 83% PtaSULTR3;2a DQ174469 65 / 80% OsSULTR3;2 AAN06871 PtaSULTR3;2b DQ906934 68 / 83% PtaSULTR3;3a DQ906924 63 / 78% OsSULTR3;4 BAD68396 PtaSULTR3;3b DQ906926 64 / 78% PtaSULTR3;4a DQ174467 70 / 85% PtaSULTR3;4b DQ174466 69 / 84% OsSULTR3;5 NM_192602 PtaSULTR3;5 DQ906927 57 / 77% OsSULTR3;6 NM_191791 4 PtaSULTR4;1 DQ906930 73 / 84% OsSULTR4;1 AF493793 PtaSULTR4;2 DQ906935 74 / 85% 5 PtaSULTR5;2 DQ174477 63 / 76% OsSULTR5;2 BAD03554 PtaSULTR5;1 DQ174475 63 / 77% OsSULTR5;1 BAC05530
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Additionally, comparison of further known sulfate transporter sequences indicated identities of SULTR1;2 to tomato LeST1-1 (AF347613) and potato StST1 (AF309643) of about 79%, while the identity to Stylosanthes hamata ShST1 / ShST2 (X82255 / X82256), Hordeum vulgare HvST1 (X96431) and Zea mays ZmST1;1 (AAK35215) was in between 75% and 77%. Distances within and between indicated groups (MEGA 4; Tamura et al., 2007) were specified based on their phylogenetic relatedness (Figure 3.1). Distance calculation compute sequence diversity based on amino acid substitutions of the compared sequence sites. This distance (number of amino acid substitutions per site) is the proportion of amino acid sites at which the two compared sequences are different. Distance calculation within groups demonstrated that the relation between sequences ranged between 0.33 substitutions per site for the most homogenous group 1 and 0.59 for the biggest most variable group 3 (Table 3.3). Among group 1, 2 and 3 distances ranged between 0.69 and 0.81. These sequences clustered within one big branch which separated group 4 and 5 with a calculated distance above 1 (Figure 3.1). The most distant branch consisted of the more diverse group 5, which was reflected by values above 2 (Table 3.3).
Table 3.3. Phylogenetic distances between amino acid sequences. Sequence groups built by phylogenetic analysis (see Figure 3.1) were examined regarding the distance within groups and between group means (MEGA 4; Tamura et al., 2007). Distance values indicate the number of amino acid substitutions per site.
Distance Distance between groups within group
Group 1 2 3 4
1 0.33 2 0.45 0.69 3 0.59 0.75 0.84 4 0.33 1.10 1.19 1.26 5 0.51 2.36 2.35 2.22 2.30
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3.1.2 Tissue specific expression analyses of the putative Populus tremula x P. alba sulfate transporter gene family
To get a general idea about the role of individual sulfate transporters in Populus tremula x P. alba, tissue specific expression analyses were performed by the Northern blot technique (Chapter 2.3.14, Figure 3.3). Fine roots, wood, bark, leaves, leaf veins, apex, root bark and root wood were examined separately to get an overview over the whole tree. In order to discriminate the expression level of different sulfate transporter isoforms with specific probes, it was necessary to identify isoform specific parts of the cDNA sequences. The coding region of closely related genes was up to 92% identical at the nucleotide level. Therefore, the selected sequence segments included parts of the more diverse 3’ untranslated region (3’ UTR). The 3’ UTR is situated between the stop codon of the mRNA and the end of each gene transcript (poly-A tail). The optimized probes were between 150 and 360 nucleotides long to enhance the respective gene specificity (Table 2.3). Unspecific binding of the probes during hybridization reaction was tested for each closely related gene pair on the cDNA integrated in the plasmid pCR2.1 (Invitrogen, Karlsruhe, Germany) (Figure 3.2). The transcript level of the 18s rRNA was detected as an equal loading control of the RNA amount used. Unspecific hybridization between close related sequences were not observed for numerous probes (SULTR1;1, SULTR 2;1a, SULTR 2;1b, SULTR 3;1b, SULTR 3;2a, SULTR 3;2b, SULTR 3;3b, SULTR 3;4b, SULTR 4;1) (Figure 3.2). While probes for SULTR1;2, SULTR5;1, SULTR5;2, SULTR3;3a and SULTR3;4a exhibited a slight signal (less than 20%) with their closely related SULTR, only two probes for SULTR3;1a and SULTR4;2 exceeded the critical amount of approximately 40% for unspecific binding. Tissue specific expression analyses revealed that the gene pairs with critical probes did not exhibit the same expression pattern except for both SULTR4 (Figure 3.3). Therefore, unspecific binding of probes can be mainly neglected and has to be implicated just for the results of group 4.
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Figure 3.2. Dot blots for cross-hybridization tests of sulfate transporter probes of closely related sequences. Unspecific binding capacity of each sulfate transporter probe with closely related sequences was tested. Therefore 12 ng of the respective cDNA ligated in the plasmid pCR 2.1 (Invitrogen, Karlsruhe, Germany) were fixed on a nylon membrane and hybridized with the indicated 32P-labeled probe (Chapter 2.3.14). Indicated percentage value of cross hybridization was calculated using Quantity one® analysis software (Quantity one® 1-D version 4.6.3, Bio-Rad Laboratories, München, Germany).
The family of sulfate transporter genes from Populus tremula x P. alba exhibited strong tissue specific expression patterns (Figure 3.3). Only one gene, namely SULTR1;2 was exclusively expressed in fine roots (Figure 3.3 a), while two genes were detected in bark tissue (Figure 3.3 b). These two genes, SULTR1;1 and SULTR3;3a were not only detected in bark of stem and roots, but additionally in leaf veins. While SULTR3;3a exhibited constant transcript abundances within the detected tissues, the expression of SULTR1;1 was approximately three fold stronger in bark and leaf veins compared to root bark. SULTR1;1 indicated additionally a weak transcript abundance in stem wood. Noticeable for these two genes was their abundant expression in leaf veins and the simultaneously absence of any signal in the associated leaf lamina.
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Figure 3.3. Expression analysis of the putative Populus tremula x P. alba sulfate transporter gene family. Transcript levels were analysed from 15 µg total RNA by the Northern blot technique (Chapter 2.3.14). Hybridization was performed with 32P-labeled gene specific probes. To quantify RNA loading, transcript accumulation of the 5.8s rRNA was detected. Plants were grown on soil in a greenhouse for ten weeks. Total RNA was extracted from the indicated tissues: fine roots, leaves, major leaf veins, the apex, bark, wood, root wood and root bark from the poplar hybrid Populus tremula x P. alba (n = 3). Grouping reflects similar expression pattern (a, b, c, d ,e).
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Three gene transcripts were detected in leaves, with SULTR3;3b being the only gene exclusively expressed in leaf tissue and the apex (Figure 3.3 c). Another two putative sulfate transporters, SULTR2;2 and SULTR3;1b, exhibited strong transcript accumulation in leaf tissue (Figure 3.3 c). In addition, SULTR2;2 mRNA was detectable in stem wood with comparable abundance than in leaves. However, the transcript level observed in the apex, leaf veins, root bark and stem bark was more than threefold lower compared to leaves and stem wood. SULTR3;1b mRNA expression was lower in stem wood as well as in the apex and root wood compared to the transcript abundance within leaves (Figure 3.3 c). Very slight SULTR3;1b transcript levels were also observed in fine roots and leaf veins. For the two genes SULTR3;3b and SULTR3;1b it is notable that transcript accumulation was observed in leaves without any detectable signal in leaf veins, which was opposite to the pattern of SULTR1;1 and SULTR 3;3a.
The biggest number of genes was specifically expressed in wood tissues (Figure 3.3 d). Within this group SULTR2;1b, SULTR3;2a, SULTR3;2b and SULTR3;4a exhibited equal mRNA accumulation in stem wood and root wood. SULTR3;5 was the only sequence exclusively detected in root wood, while SULTR3;1a was solely expressed in stem wood. Beside the accumulation of SULTR2;1a and SULTR3;4a mRNA in wood tissues, very weak mRNA accumulation of these sulfate transporter genes was also detected in root bark and leaf veins (Figure 3.3 d). The sequences of SULTR4;1, SULTR4;2, SULTR5;1 and SULTR5;2 exhibited a consistent expression through all analysed tissues with variations of intensity (Figure 3.3 e). No expression was detected for SULTR3;4b under the growth conditions applied to the poplar plants (Figure 3.3 d). For this study, the two genes SULTR1;1 and SULTR 3;3a were most noticeable, because they were detected in bark of stem and roots as well as in leaf veins (Figure 3.3 b). Because the phloem vascular system is localized within these tissues, the subsequent analyses were focused on these putative sulfate transporters. Additionally, the two sequences of group 4 as putative vacuolar efflux sulfate transporters were included in further analyses, because in Arabidopsis thaliana tonoplast localization of AtSULTR4;1 and AtSULTR4;2 and vacuolar sulfate distribution were demonstrated (Kataoka et al., 2004a).
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3.1.3 Structure analyses of predicted sulfate transporter amino acid sequences
Analyses of predicted amino acid structures were done with the two sulfate transporter sequences PtaSULTR1;1 and PtaSULTR3;3a according to their specific expression in bark and leaf veins (Figure 3.3 b). Due to the fact that sulfate redistribution is supposed to be connected with sulfate transport out of vacuoles, both sequences that belong to group 4, PtaSULTR4;1 and PtaSULTR4;2, were also included. The beginning of the open reading frame of these full length isolated genes were based on the identification of the most likely ATG, which encodes the translation start using the TIS Miner grogram (http://dnafsminer.bic.nus.edu.sg/) (Liu et al., 2004). An amino acid sequence alignment was carried out with deduced full length protein sequences from poplar including one representative Arabidopsis transporter AtSULTR1;1 (Figure 3.4). Predicted polypeptides of the cloned SULTR sequences had a length of 646 amino acids (aa) for PtaSULTR1;1, 652 aa for PtaSULTR3;3a, 678 aa for PtaSULTR4;1 and 676 aa for PtaSULTR4;2 (Figure 3.4). The hydrophobic profile of predicted polypeptides encoded by the cDNA sequences of PtaSULTR1;1, PtaSULTR3;3a, PtaSULTR4;1 and PtaSULTR4;2 indicated twelve membrane spanning domains (MSDs) (Figure 3.4). Such membrane spanning helices are characteristic for integral membrane proteins. A combined analysis with MEMSAT (http://saier-144-37.ucsd.edu/memsat.html) (Jones et al., 1994) and TMAP (http://mobyle.pasteur.fr/cgi-bin/MobylePortal/portal.py) (Persson and Argos, 1994) programs enabled the most probable locations of MSDs within the amino acid sequences. The received topology with twelve MSDs fits to the model that has been proposed for the sulfate transporter isolated from Stylosanthes hamata (Smith et al., 1995). According to this model, several basic amino acid residues (Lys, Arg) are located on both sides of the membrane. Among them, Arg-(R-391/390/363/361), which is located between membrane spanning domain (MSD) nine and ten, was a basic residue that is identical to the published sulfate transporter amino acid sequences from Arabidopsis thaliana, Stylosanthes hamata and Hordeum vulgare (Takahashi, 1997) (Figure 3.4).
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AtSULTR1.1 MSGTINPPDGGGSGARNPPVVRQRVLAPPKAGLLKDIKSVVEETFFHDAPLRDFKGQTPA 60 PtaSULTR1.1 --MDIRSLSSSHRHPQDAPYVHK-VGLPPKQNLFKEFKDTVKETFFADDPLRSFKDQPRS 57 PtaSULTR3.3a MEPNASNNMQPDHCLGITPMEVHKVVPPPHRSTIQKLKSKLKETFFPDDPLLQFKRQPLG 60 PtaSULTR4.1 ------MPTRPVKIIPLQHP------NTTTSSSLNPLPGALFSRWTAKVKR 39 PtaSULTR4.2 ------MPNRPVKTIPLQHP------NTTSSSSS-PLAQAMFSRWTAKIKR 38 1 AtSULTR1.1 KKALLGIQAVFPIIGWAREYTLRKFRG-DLIAGLTIASLCIPQDIGYAKLANVDPKYGLY 119 PtaSULTR1.1 KKFILGLQAIFPILEWGRSYSFAKFRG-DLIAGQTIASLCIPQDIGYAKLANLDPQYGLY 116 PtaSULTR3.3a KKWILAAQYVFPILQWGPNYSFKLFKS-DIVSGLTIASLAIPQGISYAKLASLPPIVGLY 119 PtaSULTR4.1 ITLVQWIDTFLPCCRWIRTYKWREYFQPDLMAGLTVGVMLVPQAMSYAKLAGLHPIYGLY 99 PtaSULTR4.2 TTPSQWIDTFLPCYRWIRTYKWREYLQPDLTAGLTVGIMLVPQAMSYAKLAGLHPIYGLY 98 2 3 4 AtSULTR1.1 SSFVPPLIYAGMGSSRDIAIGPVAVVSLLVGTLCQ-AVIDPKKNPEDYLRLVFTATFFAG 178 PtaSULTR1.1 TSFVPPLIYAFMGSSRDIAIGPVAVVPLLLGTLLQSEIADPVANAAEYRRLAFTATFFAG 176 PtaSULTR3.3a SSFVPPLVYAVLGSSRDLAVGPVSIASLILGSMLR-QKVSPINDPLLFLQLAFSSTFFAG 178 PtaSULTR4.1 TGFIPIFVYAIFGSSRQLAIGPVALVSLLVSNVLGGIVNSSDE---LYTELAILLAFMVG 156 PtaSULTR4.2 IGFIPIFVYAIFGSSRQLAIGPVALVSLLVSNVLGG-MDLSDE---LYTELAILLAFMVG 154 5 AtSULTR1.1 IFQAGLGFLRLGFLIDFLSHAAVVGFMGGAAITIALQQLKGFLGIKTFTKKTDIVSVMHS 238 PtaSULTR1.1 ITQVTLGFLRLGFLIDFLSHAAIVGFMGGAAITIALQQLKGFLGIKKFTKKTDIVSVMHS 236 PtaSULTR3.3a LFQASLGLLRLGFIIDFLSKAILIGFMAGAAVIVSLQQLKSLLGITHFTKQMGLVPVLSS 238 PtaSULTR4.1 ILECIMALLRLGWLIRFISHSVISGFTSASAIVIALSQAKYFLGYD-IVRSSKIVPLIKS 215 PtaSULTR4.2 IMECIMAFLRLGWLIRFISHSVISGFTTASAIVIALSQAKYFLGYD-VVRSSKIVPLIKS 213 6 7 AtSULTR1.1 VFKNAEHGWNWQTIVIGASFLTFLLVTKFIGKRNRKLFWVPAIAPLISVIISTFFVFIFR 298 PtaSULTR1.1 VFASARHGWNWQTIVIGVSLLSFLLFAKYIGKKNKRLFWVPAIGPLISVILSTFFVFITR 296 PtaSULTR3.3a AFHNIN-EWSWQTILMGFCFLVFLPLARHVSMRKPKLFWVSAGAPLVSVILSTILVFAFK 297 PtaSULTR4.1 IISGAH-KFSWPPFVMGSCILAILLVMKHLGKSRKQFTFLRAAGPLTAVVLGTLFVKMFH 274 PtaSULTR4.2 IISGAH-KFSWPPFVMGSCILAILLVMKHLGKSRKQFRFLRPAGPFTAVVLGTVFVKMFH 272 8 AtSULTR1.1 ADKQGVQIVKHIDQGINPISVHKIFFSGKYFTEGIRIGGIAGMVALTEAVAIARTFAAMK 358 PtaSULTR1.1 ADKDGVQIVKHMEKGINPSSVNQIYFSGDHLLKGVRIGIVAAMIALTEAIAIGRTFAAMK 356 PtaSULTR3.3a AQHHGISVIGKLQEGLNPPSWNMLHFHGSNLGLVIKTGLVTGIISLTEGIAVGRTFAALK 357 PtaSULTR4.1 P--SSISLVGEILQGLPSFSFPKKFEYAKSLIP---TAMLITGVAILESVGIAKALAAKN 329 PtaSULTR4.2 P--SSISLVGDIPQGLPSFSIPKKFEYAKSLIP---SAMLITGVAILESVGIAKALAAKN 327 9 10 AtSULTR1.1 DYQIDGNKEMIALGTMNVVGSMTSCYIATGSFSRSAVNFMAGVETAVSNIVMAIVVALTL 418 PtaSULTR1.1 DYQLDGNKEMVALGTMNIVGSMTSCYVATGSFSRSAVNFMSGCQTAVSNIVMSIVVFLTL 416 PtaSULTR3.3a NYQVDGNKEMMAIGLMNVIGSATSCYVTTGAFSRSAVNHNAGAKTAVSNVVMSVTVMVTL 417 PtaSULTR4.1 GYELDSSQELFGLGLANIMGSLFSAYPSTGSFSRSAVNNESGAKTGLSGVVAGIIMCCSL 389 PtaSULTR4.2 GYELDSSQELFGLGLANILGSFFSAYPSTGSFSRSAVNDDSGAKTGLAGIVAGTIMGCSL 387 11 12 AtSULTR1.1 EFITPLFKYTPNAILAAIIISAVLGLIDIDAAILIWRIDKLDFLACMGAFLGVIFISVEI 478 PtaSULTR1.1 QFITPLFKYTPNAVLSAIIISAVIGLVDFDAAYLIWKIDKFDFVACMGAFFGVVFASVEI 476 PtaSULTR3.3a LFLMPLFQYTPNVVLGAIIVTAVIGLIDFPAACQIWKIDKFDFVVMLCAFFGVVFISVQD 477 PtaSULTR4.1 LFLTPLFEYIPQCALAAIVISAVMGLVDYDEAIFLWHVDKKDFVLWIITSATTLFLGIEI 449 PtaSULTR4.2 LFLTPLFEYIPQCGLAAIAISAVMGLVDYDEAIFLWHVDKKDFVLWIITSTTTLFLGIEI 447
AtSULTR1.1 GLLIAVVISFAKILLQVTRPRTTVLGKLPNSNVYRNTLQYPDAAQIPGILIIRVDSAIYF 538 PtaSULTR1.1 GLLIAVSISFFKLLLQVTRPRTAILGKLPRTAVYRNILQYPEATKVPGVLIVRVDSAIYF 536 PtaSULTR3.3a GLAIAVAISIFKILLQVTRPKTLVLGNIPGTDIFRNLHHYKDATRIPGFLILSIEAPINF 537 PtaSULTR4.1 GVLVGVGASLAFVIHESANPHIAVLGRLPGTTVYRNIEQYPEAYTYNGIVIVRIDAPIYF 509 PtaSULTR4.2 GVLVGVGVSLAFVIHESANPHIAVLGRLPGTTVYRNIQQYPEAYTYNGIVIVRIDAPIYF 507 STAS AtSULTR1.1 SNSNYVRERASRWVREEQENAKE-YGMPAIRFVIIEMSPVTDIDTSGIHSIEELLKSLEK 597 PtaSULTR1.1 SNSNYIKERILRWLIDEEELVNK-SSQPKIQFLVVEMSPVTDIDTSGIHALEELYRSLQK 595 PtaSULTR3.3a ANTTYLKERILRWINEYETEEDI-KKQSSIHFLILDLSAVSAIDTSGVSLFKDLKKAVEN 596 PtaSULTR4.1 ANISSIKDRLREYEVDADKSSRRGPEVEKIYFVILEMSPITYIDSSAVQALKDLHQEYKS 569 PtaSULTR4.2 ANISFIKDRLREYEADVDKSARHGPEVERIHFLILEMSPITYIDSSAVQALKDLHQEYKS 567
AtSULTR1.1 QEIQLILANPGPVVIEKLY-ASKFVEEIGEKNIFLTVGDAVAVCSTEVAEQQT------649 PtaSULTR1.1 REIQLILANPGPVVIDKLH-ASDFAQLIGEDKIFLTVANAVAACSPKLVEEV------646 PtaSULTR3.3a KGVELVLVNPVGEVLEKLIRADDARDIMGPDTLYLTVGEAVAALSPTMKGQSSSYV---- 652 PtaSULTR4.1 RDIQICISNPNRDVLLTLT-KAGIVELLGKERYFVRVHDAVQVCLQHVQSSTQSPKKPDP 628 PtaSULTR4.2 RDIEICIANPNQDVLLTLT-KAGIVELIGKEWYFVRVHDAVQVCLQHVQSLNQTPKNPDS 626
AtSULTR1.1 ------PtaSULTR1.1 ------PtaSULTR3.3a ------PtaSULTR4.1 SAEEKPRIFKRLSKQREEDLSIAELESGDNKTSAPKHTKPHLEPLLSRRS 678 PtaSULTR4.2 FAEDKPSFFQRLSKQREEDLSIAELESGDKKTSVPKFTEPHLEPLLSRKS 676
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Figure 3.4. Amino acid sequence alignment of putative sulfate transporters from Populus tremula x P. alba and Arabisopsis thaliana. Full length amino acid sequences of PtaSULTR1;1, PtaSULTR3;3a, PtaSULTR4;1 and PtaSULTR4;2 were aligned with the Arabidopsis AtSULTR1;1 sulfate transporter protein (Accession numbers, Table 3.1). The alignment was performed using ClustalW2 (http://www.ebi.ac.uk/Tools/clustalw2/index.html). Consensus amino acids are yellow highlighted while gray highlighted amino acids have similar properties. Gaps in the sequence are inserted to obtain the best alignment and are indicated by dashes. Twelve membrane spanning domains (MSDs) were predicted by a combination of hydrophobic analysis with the MEMSAT (Jones et al., 1994) and TMAP (Persson and Argos, 1994) programs and accounted with black bars. A conserved basic amino acid residue (Arg) between MSD 9 and 10 is indicated by a solid dot. A conserved STAS (sulfate transporter anti sigma factor antagonist) domain was identified and indicated by a dark blue box. Predicted transit peptide regions were calculated using the ChloroP 1.1 program (http://www.cbs.dtu.dk/services/ChloroP/) (Emanuelsson et al., 1999) and are highlighted with bold letters.
At the carboxyl-terminal end a conserved STAS (sulfate transporter anti sigma factor antagonist) domain was identified. The carboxyl-terminal end of predicted Populus tremula x P. alba sulfate transporter sequences were compared with the STAS domain from the Arabidopsis thaliana SULTR1;2 protein sequence characterized by Shibagaki et al. (2004) (Figure 3.4, 3.5). Except the members of group 5, the remaining sixteen putative sulfate transporters from Populus tremula x P. alba exhibited this STAS domain at their carboxyl-terminus like in Arabidopsis (Figure 3.5). Secondary structures within this area which are likely built due to the amino acid sequences, were predicted with the Quick2D Bioinformatic toolkit (http://toolkit.tuebingen.mpg.de/ quick2 d). The predicted α-helices and β-strands are conserved within the 16 poplar sulfate transporters (Figure 3.5). Predicted transit peptide sequences were identified with the TargetP 1.1 program (http://www.cbs.dtu.dk/services/TargetP/) (Emmanuelsson et al., 2000). Only the full length sequences were examined because transit peptides are supposed to be situated at the N-terminal end of proteins, which was absent in the partial cloned sulfate transporter sequences (Keegstra and Cline 1999; Bruce, 2001). This analysis revealed the transit peptide characteristics of chloroplast localization for sulfate transporters of group 4, while for the other sequences a clear localization was not found. Also additional analyses of the predicted sulfate transporter sequences from the Populus trichocarpa database did not reveal a clear localization for the remaining proteins from group 1, 2, 3 and 5. The program ChloroP 1.1 (http://www.cbs.dtu.dk/ services/ChloroP/) (Emanuelsson et al., 1999) indicated a transit peptide length of 32 aa for PtaSULTR4;1 and of 24 aa for PtaSULTR4;2 at the N-terminal site (Figure 3.4).
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β1 Sultr1.1 VEIGLLIAVSISFFKLLLQVTRPRTAILGKLPRTAVYRNILQYPEATKVPGVLIVRVDS- 532 Sultr1.2 VEIGLLIAVSISFAKILLQVTRPRTAILGNLPRTTVYRNILQYPEAAKVPGVLIVRVDS- 370 Sultr2.1a VEIGLLAAVIISFVKIIIISIRPGTEELGRLPGTDIFCDVNQYPMAVKNSKALIIRVKSG 354 Sultr2.1b VEIGLLAAVTISFVKILIISIRPGAEVLGRLPETDIFCDVDQYPMAAKNPQVLVIRVKSG 475 Sultr2.2 VEIGLLAAVTISFARILLNAIRPGIEALGRLPRADVYCDMNQYPMAVKTPGILAVRINSA 505 Sultr3.1a VEIGLVIAVAISVLRLLLFVARPKTFILGNIPNSMIYRNVEQYLNTSSVPGVLILEIDA- 416 Sultr3.1b VEIGLVVAVAISLLRVLLFVARPKTFILGNIPNSMIYRNVEQYTNTSSVPGVLILEIDA- 456 Sultr3.2a VEIGLVIAVTISLLRMILSVARPRTFLLGNIPNSMIFRSIEQYPVANNIPGVLILQIDA- 508 Sultr3.2b VEIGLVIAVAISLLRMLMSVARPRTFLLGNIPNSMIYRSIDQYPIANTVPGVLILQIDA- 320 Sultr3.3a VQDGLAIAVAISIFKILLQVTRPKTLVLGNIPGTDIFRNLHHYKDATRIPGFLILSIEA- 533 Sultr3.3b VQDGLAIAVGISIFKILLQVTRPKTVVLGDIPGTDIFRNFHHYKEAMRIPGFLILSIEA- 501 Sultr3.4a VPSGLGIAVGVSVFKILLHVTRPNTLIMGNIRGTNVYQCLGRYKEASRIPSFLVLAIES- 525 Sultr3.4b VPLGLGIAVGVSVFKILLHVTRPNSSIMGNIKGTQIYHSLSRYKEASRVPSFLILAIES- 491 Sultr3.5 MDMGLMISVGLALLRALLYVARPAACKLGKLPDSTLYRDTEQYAEASGPPGILAIQLGS- 512 Sultr4.1 IEIGVLVGVGASLAFVIHESANPHIAVLGRLPGTTVYRNIEQYPEAYTYNGIVIVRIDA- 505 Sultr4.2 IEIGVLVGVGVSLAFVIHESANPHIAVLGRLPGTTVYRNIQQYPEAYTYNGIVIVRIDA- 503 Sultr5.1 GRSGGCVALLG------AAKLVLGLVLGSSLVMVLNQFPVG------329 Sultr5.2 GRSGGCVALLG------AAKMLLGLVLGSSLVMVLKQFPVG------262 * .: . :* : : : ::
β2 α1 β3 α2 STAS Sultr1.1 AIYFSNSNYIKERILRWLIDEEELVNK-SSQPKIQFLVVEMSPVTDIDTSGIHALEELYR 591 Sultr1.2 AIYFSNSNYIKERILRWLIDEEELVNK-SGQTKIQFLIVELSPVTDIDTSGIHAMEELLR 429 Sultr2.1a LLCFANANFVKEKIMKWATEEEENDSK-GKR-TVQVVILDMSNLMNIDMSGIASLLELQN 412 Sultr2.1b LLCFANANFVKEKIMKLATEEEEG-RK-GKR-TVQVVILDMSNLMNIDVSGITSLVELHK 532 Sultr2.2 LPCFANANFIRERILRWVTEEVNEIKE-STEGGIKAVILDVSNVMNIDTAGILALEELHK 564 Sultr3.1a PIYYANSGYLRERIARWVDDEEDKLKS-SGETSLQYVILNMGAVGTIDTSGISMLEEVKK 475 Sultr3.1b PIYFANASYLRERIARWVDEEEDKLKS-SGETSLQYVILDMGAVGNIDTSGISMLEEVKK 515 Sultr3.2a PVNFANANYLRERISRWIYEEEEKLKS-TGGSSLQYVILDLSAVGSTDTSGISMFKEVKK 567 Sultr3.2b PVYFANANYLRERISRWIYEEEEKVKS-TGGSSLQYVILDLSAVGSLDTSGISMLEEVKK 379 Sultr3.3a PINFANTTYLKERILRWINEYETEEDI-KKQSSIHFLILDLSAVSAIDTSGVSLFKDLKK 592 Sultr3.3b PINFANTTYLKVRILRWIDEYETEEDT-KRQSSIHFLILDLSAVSSIDTSGVSLLKDLKK 560 Sultr3.4a PIYFANSTYLQERILRWIREEEDWIKA-NNEDTLKCVILDMTAVTAIDTSGIDLVCELRK 584 Sultr3.4b PIYFANSTYLQERVLRWIREEDEWIKA-NNGSPLKCIILDMTAVTAIDTSGIDLLCELRK 550 Sultr3.5 PIYYAYGNYIRERILRWIRNDE------GNGKAVKHVLLDLTGVTSIDTTGIETLAEVLR 566 Sultr4.1 PIYFANISSIKDRLREYEVDADKSSRRGPEVEKIYFVILEMSPITYIDSSAVQALKDLHQ 565 Sultr4.2 PIYFANISFIKDRLREYEADVDKSARHGPEVERIHFLILEMSPITYIDSSAVQALKDLHQ 563 Sultr5.1 ------VLGVLLLFAGIELAMASRDMNTKEEAFV 357 Sultr5.2 ------VLGVLLLFAGIELALASRDMNTKEEAFV 290 : ::: : :
β4 α3 β5 α4
Sultr1.1 SLQKREIQLILANPGPVVIDKLHAS-DFAQLIGEDKIFLTVANAVAACSPKLVEEV---- 646 Sultr1.2 SLQKREIQLILANPGPAVIDKLHAS-GSAQLIGEDKIFLTVADAVASCCPKSVGEV---- 484 Sultr2.1a NLASGGMELAITNPKWQVIHKLRLA-NFATKMG-GRVFLTAGEAVDACLGAKMAAV---- 466 Sultr2.1b NLASSGMELAITNPKWQVIHKLRVA-NFVTKIG-GRVFLTIGEAMDACLGAKMAAV---- 586 Sultr2.2 ELLVHEAQLAIANPKWQVIHKLRLA-KFIDRIGRGWIFLTVSEAVDACVSSKLTALANC- 622 Sultr3.1a VMDRRGLKLVMANPGAEVMKKLNKA-KFIEKIGQEWIHLTVGEAVEACDFMLHTCSPG-P 533 Sultr3.1b VMDRRELQLVLANPGAEVVKKLNKS-KLIEKIGQEWMYLTVGEAVGACNFMLHTRKPD-P 573 Sultr3.2a NIYSRGLKLVLANPRSEVIKKLVKS-KFIESIGQEWIYLTVGEAVAACNFMLHASKSNNQ 626 Sultr3.2b NIDRRDFKLVLANPRSEVIKKLEKT-KFMESIGQEWIYLTVGEAVAACNFMLHRSKSNNP 438 Sultr3.3a AVENKGVELVLVNPVGEVLEKLIRADDARDIMGPDTLYLTVGEAVAALSPTMKGQSSSYV 652 Sultr3.3b ALENTGAELVLVNPVGEVLEKLQRADDVRDVMSPDALYLTVGEAVAALSSTVKGRSSSHV 620 Sultr3.4a MLEKRSFQLVLANPVGSVMEKLHQS-KTLDSFGLNGIYLTVGEAVADISALWKSQP---- 639 Sultr3.4b MLEKRSLKLVLTNPVGSVMEKLHQS-KMLDSFGLNGIYLAVGEAVADISALWKSQPDFPE 609 Sultr3.5 MLEVKHIKMKIVNPRLEVFEKMMKS-KFVDKIGEESIFLCMEDADEASYDFSVTTEKQGF 625 Sultr4.1 EYKSRDIQICISNPNRDVLLTLTKA-GIVELLGKERYFVRVHDAVQVCLQHVQSSTQSPK 624 Sultr4.2 EYKSRDIEICIANPNQDVLLTLTKA-GIVELIGKEWYFVRVHDAVQVCLQHVQSLNQTPK 622 Sultr5.1 MLICSAVSITGSSAALGFLCGIAVH------LLLKVRNWHNDQPCSTV------399 Sultr5.2 MLICAAVSLVGSSAALGFVCGIIVH------VLLYLRNWQKEQPCPVL------332 .: .. .. : : .
Figure 3.5. Amino acid sequence alignment of the carboxyl-terminal end of putative sulfate transporters. Amino acid sequences of the 18 sulfate transporters from Populus tremula x P. alba (Table 3.1) were aligned using ClustalW2 program (http://www.ebi.ac.uk/Tools/clustalw2/index.html). Excluding group 5 sulfate transporters, consensus amino acids are yellow highlighted while gray highlighted are amino acids with similar properties. Consensus amino acids of all sequences are indicated by asterisk while dots highlight amino acids with similar properties. Gaps in the sequences are inserted to obtain best alignment and are indicated by dashes. A conserved STAS (sulfate transporter anti sigma factor antagonist) domain was identified and indicated by a dark blue box with predicted α-helices (red banners) and β-strands (black arrows) using the bioinformatics tool Quick2D (http://toolkit.tuebingen.mpg.de/quick2_d). 80 Results
3.2 Cellular transcript localization of SULTR1;1 and SULTR3;3a
According to the transcript accumulation in bark and leaf vein tissue (see Figure 3.3b) the two putative Populus tremula x P. alba sulfate transporter sequences SULTR1;1 and SULTR3;3a were selected to gain more detailed insight in the organization of sulfate allocation through the phloem vascular system. In situ hybridization was performed for further observations at the cellular level (Chapter 2.4). Differences of the expression patterns between the two genes were of interest as well as variations of the respective putative sulfate transporters within different tissues. The phloem connects source leaves with sink organs. Under full nutrient growth conditions sulfate is loaded into the phloem of mature leaves for distribution throughout the plant in different directions and subsequent supply via unloading into sink tissues. To get an overview of the cellular expression at the whole plant level, transcript accumulation of SULTR1;1 and SULTR3;3a was investigated in leaves assumed as sulfate source, stem and roots with secondary growth as organs where sulfate unloading for storage takes place and additionally fine roots and the shoot apex as expected sink organs regarding sulfate.
3.2.1 Cellular transcript localization of SULTR1;1 in leaves
SULTR1;1 mRNA accumulation was investigated in transverse sections of leaf vein tissue (midrib) where transport phloem is expected and furthermore in the leaf lamina where collection phloem is localized (Figure 3.6). As a control, comparable sections were treated with the sense probe, which did not result in any staining in leaf veins or lamina cross sections (Figure 3.6 Ia). Violet staining by binding of the antisense probe exhibited accumulation of sulfate transporter SULTR1;1 mRNA predominately in the phloem area of major leaf veins (Figure 3.6 I). The enlarged view exhibited strong transcript accumulation exclusively in companion cells (Figure 3.6 Ib). Neither sieve elements nor phloem parenchyma cells exhibited SULTR1;1 expression. A comparatively weak signal was detected in xylem parenchyma cells associated with xylem vessels (Figure 3.6 Ic).
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Figure 3.6. In situ localization of SULTR1;1 mRNA in leaf tissue. Images shown are transverse sections through the lamina of the 10th leaf and of a major leaf vein from Populus tremula x P. alba. Cuttings were treated with DIG-labeled antisense probe for SULTR1;1 (I, Ib-If). As control, sections were treated with DIG-labeled sense probe for SULTR1;1 (Ia). Magnification of the phloem (Ib) and xylem (Ic) are indicated with black boxes (I). Enlargement of the vascular bundle within minor leaf veins (Ie) are accounted with a black box (Id). Palisade parenchyma is presented (If). Bars indicate real µm. Abbreviations: companion cell (cc); lower epidermis (le); palisade parenchyma (pp); phloem (ph); sieve element (se); spongy parenchyma (sp); upper epidermis (ue); xylem (x); xylem parenchyma (xp); xylem vessel (v).
SULTR1;1 transcript was detectable in minor leaf veins of the lamina where cellular mRNA localization was observed in phloem cells (Figure 3.6 Id, Ie). In minor leaf veins the phloem revealed no clear separation into sieve tubes and companion cells. Transverse lamina sections exhibited transcript accumulation additionally in the palisade parenchyma cells, mainly in the cell layer directly adjacent to the upper epidermis (Figure 3.6 Id, If). Slight transcript accumulation was also detected in the spongy mesophyll and in the upper epidermis cell layer.
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3.2.2 Cellular transcript localization of SULTR1;1 in stem and roots with secondary growth
Poplar stems were divided into three sections of different ages, which feature variable physiological stages. Young stem sections (between 4th and 5th leaf) were scrutinized because in this area the vascular tissue is at an early developmental stage. Middle stem sections with young mature leaves (between 10th and 11th leaf) are supposed to be mature while older basal stem parts (between 25th and 27th leaf) were taken because increased storage of sulfate can be expected in this area of poplar stem (Hartmann et al., 2000). The control-hybridization with SULTR1;1 sense probe did not result in a signal in the three investigated stem sections (Figure 3.7 Ia, IIa, IIIa). Violet staining by the antisense probe exhibited accumulation of sulfate transporter SULTR1;1 transcripts predominantly in the vascular cambium and phloem area (Figure 3.7 I). In addition, mRNA transcripts were detectable in the periderm and protoxylem. Different physiological stages of stems exhibited similar transcript pattern for SULTR1;1 (Figure 3.7 I, II, III).
When stem sections were observed from the outside to the inside, epidermis cells of young stems revealed strong SULTR1;1 expression. However, in the middle and basal stem sections SULTR1;1 accumulation was noticeable in the cell layer directly below the epidermis. Comparing the periderm area of the three stem sections of different ages with a higher magnification, only young stem sections exhibited SULTR1;1 transcript accumulation in the external epidermis cells (Figure 3.7 Ib, IIb, IIIb). Additionally, transcript accumulation was observed in the epidermal region of young stems where a lenticel was localized (Figure 3.7 I, Ig). A lenticel is an area within the cortex with big intercellular spaces to allow gas exchange between the internal tissue and the atmosphere. Beside SULTR1;1 accumulation, more detailed observations indicated dark brown suberic cells between epidermis and parenchyma cells, probably caused by the accumulation of phenolic components (Figure 3.7 Ih).
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Figure 3.7. In situ localization of SULTR1;1 in stem tissues. Transverse stem sections of Populus tremula x P. alba of three different ages, young stem between the 4th and 5th leaf (I), middle stem between the 10th and 11th leaf (II) and basal stem between the 25th and 27th leaf (III) are presented. Cuttings were treated either with DIG-labeled sense probe (Ia,IIa,IIIa) or with antisense probe (all except Ia,IIa,IIIa). b-f/h show magnifications of accounted sectors which were labeled with black boxes (I, II, III.) Higher magnification of the phloem area (If, IIf, IIIf), xylem (Id, IId, IIId), protoxylem facing pith cells (Ic, IIc, IIIc), lenticel (Ig, Ih) and periderm (Ib, IIb, IIIb) are presented. Bars indicate real µm. Abbreviations: cambium (c); companion cell (cc); cortex (co); cuticula (cu); epidermis (e); lenticel (l); periderm (pe); phloem (ph); pith (p); protoxylem (pr); ray initials (ri); sclerenchymatous phloem fibers (s); sieve element (se); xylem (x); xylem parenchyma (xp); xylem ray (r); xylem vessel (v).
Compared to the other investigated stem sections only basal stem cuttings exhibited SULTR1;1 transcript accumulation in bark parenchyma cells (cortex), which are localized between the sclerenchymatous phloem fibers and extend to the periderm (Figure 3.7 III). Below the sclerenchymatous phloem fibers, considerable mRNA accumulation was especially found in the phloem cells. A higher magnification of phloem tissue indicated that SULTR1;1 expression is concentrated exclusively in companion cells, as also observed for the leaf veins (Figure 3.7 If, IIf, IIIf). The transporter transcript was neither detected in sieve cells nor in phloem parenchyma cells. This cell specific expression pattern was almost identical in all analyzed stem sections. Within the cambium, the developmental precursors of pith ray cells, namely the ray initials, exhibited a stronger signal than the surrounding cambium cells in particular the fusiform cells (Figure 3.7 Ie, IIe, IIIe). Again this pattern was detected in all stem sections.
Cells of wood tissue including cells of the xylem did not exhibited staining within any cell type. Only xylem ray cells and mainly the xylem parenchyma cells associated with xylem vessels exhibited substantial SULTR1;1 transcript accumulation (Figure 3.7 Id, IId, IIId). This was again independent of the age of the stem section investigated. Similarly, parenchyma cells associated with the protoxylem that is localized adjacent to pith tissue showed mRNA accumulation especially in cells close to xylem vessels. While in old basal stem sections this particular staining was weak, transporter transcript accumulation was substantial in young and mature stem sections (Figure 3.7 Ic, IIc, IIIc).
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Figure 3.8. In situ localization of SULTR1;1 in roots with secondary growth. Transverse sections of Populus tremula x P. alba roots with secondary growth were treated either with DIG-labeled sense probe (Ia) or with antisense probe (I, lb-d). Ib-d are magnifications of sectors labelled with black boxes (l). Magnification of the phloem area (Ic), xylem (Ib), and periderm (Id) are presented. Bars indicate real µm. Abbreviations: companion cell (cc); epidermis (e); periderm (pe); phloem (ph); sclerenchymatous phloem fibers (s); sieve element (se); xylem (x); xylem ray (r); xylem vessel (v).
Additionally, roots with secondary growth were included in the expression analyses, because of their function as sink organs for sulfate delivered from leaves (Hartmann et al., 2000). Similar to the results from stem sections, no signal was detected in roots with secondary growth when treated with the sense probe as control (Figure 3.8 Ia). Only a slight unspecific staining was observed in sclerenchymatous phloem fibers and in young xylem. In general a comparable expression pattern of SULTR1;1 transcript was detected in roots with secondary growth than in stem sections. Comparing the epidermal area of middle stem sections and roots, SULTR1;1 transcript accumulation was equally localized in cells below the suberized epidermis (Figure 3.8 Id, 3.7 llb). All cells of the cortex and the cambium showed slight mRNA accumulation (Figure 3.8 I). As observed in stem sections, strong mRNA accumulation was especially detected in the phloem and in xylem parenchyma cells (Figure 3.8 I). A stronger magnification of the phloem indicated again that SULTR1;1 transcripts were concentrated exclusively in companion cells (Figure 3.8 Ic). Also, only xylem ray cells associated with xylem vessels exhibited considerable mRNA accumulation within the wood (Figure 3.8 Ib). Due to the developmental pattern of roots with secondary growth, neither pith tissue nor protoxylem was present.
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3.2.3 Cellular transcript localization of SULTR3;3a in leaves
In addition, the sulfate transporter SULTR3;3a as a candidate for phloem specific expression was selected for further examinations at the cellular level. These observations were supposed to exhibit both differences and similarities in the expression patterns of the two transporters, SULTR3;3a and SULTR1;1, in various tissues. As control, treatment with the sense probe for SULTR3;3a did not exhibit staining within leaves (Figure 3.9 Ia), whereas violet staining was achieved by the antisense probe exhibiting accumulation of sulfate transporter SULTR3;3a transcripts. Throughout all experiments SULTR3;3a hybridization reactions were less intensely stained than SULTR1;1 probe hybridizations (Figure 3.6, 3.7, 3.8, 3.9, 3.10, 3.11, 3.12, 3.13).
Figure 3.9. In situ localization of SULTR3;3a in leaf tissues. Transverse sections through the lamina of the 10th leaf and through a major leaf vein from Populus tremula x P. alba were investigated. Cuttings were treated with DIG-labeled transcript complementary antisense probe for SULTR3;3a (I, Ib-Ie) or as a control with DIG-labeled sense probe for SULTR3;3a (Ia). Magnification of the phloem (Ib) and xylem (Ic) are indicated with black boxes (I). Enlargement of the vascular bundle of minor leaf veins (Ie) are accounted with a black box (Id). Bars indicate real µm. Abbreviations: companion cell (cc); lower epidermis (le); palisade parenchyma (pp); phloem (ph); sieve element (se); swamp parenchyma (sp); upper epidermis (ue); xylem (x); xylem parenchyma (xp); xylem vessel (v).
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The overview of major leaf veins exhibited SULTR3;3a transcript accumulation mainly in phloem tissue (Figure 3.9 I). A higher magnification of major leaf veins indicated SULTR3;3a mRNA accumulation clearly in companion cells and with weaker intensity in sieve tubes (Figure 3.9 Ib). Comparable transcript accumulation was detected in xylem parenchyma cells surrounding xylem vessels (Figure 3.9 Ic). Several consensuses were noticeable, but also several variations of the expression pattern comparing the two genes SULTR1;1 and SULTR3;3a in leaves (Figure 3.6, 3.9). Both are expressed in companion cells and xylem parenchyma cells surrounding xylem vessels. But, in contrast to SULTR1;1 no clear signal of SULTR3;3a mRNA was detected in minor leaf veins or in any other cells of the lamina like palisade parenchyma or epidermis cells (Figure 3.9 Id, Ie). The slight expression of SULTR3;3a in sieve elements was not examined for SULTR1;1.
3.2.4 Cellular transcript localization of SULTR3;3a in stem tissues and roots with secondary growth
Transverse sections of three different stem parts with different age were examined to compare variable physiological stages of the poplar stem (see Chapter 3.2.2). Roots with secondary growth were also included because of their function as sink organs for sulfate delivered from leaves (Hartmann et al., 2000). Neither in stem sections nor in roots a signal was detected with the sense probe of SULTR3;3a used as control (Figure 3.10 Ia, IIa, IIIa, 3.11 Ia). Violet staining by the treatment with antisense probe exhibited the accumulation of sulfate transporter SULTR3;3a transcripts. SULTR3;3a mRNA was predominantly observed in phloem tissue, in xylem tissue and in parenchyma cells of the protoxylem (Figure 3.10 I, II, III). This expression pattern was almost identical in all three stem sections. A higher magnification indicated that within the phloem SULTR3;3a mRNA was accumulated in the companion cells and slightly in sieve elements (Figure 3.10 If, IIf, IIIf). Similar transcript accumulation was observed in the major leaf vein (Figure 3.7 Ib). SULTR3;3a transcript accumulation was also noticed in the periderm but only of basal and middle stem sections (Figure 3.10 Ib, IIb, IIIb). The wood exhibited staining exclusively in xylem ray cells and within these cells mainly the parenchyma cells adjacent to xylem vessels (Figure 3.10 Id, IId, IIId). In addition, SULTR3;3a mRNA was detected in parenchyma cells associated to xylem vessels of the protoxylem (Figure 3.10 Ic, IIc, IIIc).
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Figure 3.10. In situ localization of SULTR3;3a in stem sections. Transverse sections of Populus tremula x P. alba stems of three different developmental stages were investigated. Young stem sections were collected between the 4th and 5th leaf (I), middle stem sections between the 10th and 11th leaf (II) and basal stem between the 25th and 27th leaf (III). Cuttings were treated either with DIG- labeled sense probe (Ia, IIa, IIIa) or with transcript complementary antisense probe (all except Ia, IIa, IIIa). I/II/IIIb-f/g showed sectors labelled with a white or black box (I, II, III). Higher magnification of the selected sections show phloem area (If, IIf, IIIf), xylem (Id, IId, IIId), protoxylem adjacent to pith (Ic, IIc, IIIc) and periderm (Ib, IIb, IIIb). Bars indicate real µm. Abbreviations: cambium (c); companion cell (cc); cortex (co); epidermis (e); lenticel (l); periderm (pe); phloem (ph); pith (p); protoxylem (pr); ray initials (ri); sclerenchymatous phloem fibers (s); sieve element (se); xylem (x); xylem parenchyma (xp);. xylem ray (r);xylem vessel (v);
The particular expression pattern in the wood was very similar to SULTR1;1 transcript accumulation (Figure 3.7 Id, IId, IIId). But, clear differences were observed when comparing the expression of SULTR3;3a and SULTR1;1 in the epidermal area. While sulfate transporter SULTR1;1 exhibited considerable transcript accumulation in epidermis cells of young stem sections and lenticels, no mRNA accumulation was observed for SULTR3;3a in these tissues (Figure 3.7 Ib, Ig, 3.10 Ib, Ig). Transcript of SULTR1;1 accumulated in the cell layer directly below the epidermis in old and mature stem, which was also not observed for SULTR3;3a (Figure 3.7 llb, lllb, 3.10 llb, lllb). However, transcript abundance of SULTR3;3a was visible in sieve tubes in contrast to SULTR1;1 results. Contrary to SULTR1;1 transcript accumulation, SULTR3;3a mRNA accumulation was completely absent in roots with secondary growth (Figure 3.11) and in the vascular cambium of all three stem sections examined (Figure 3.10 Ie, IIe, IIIe).
Figure 3.11. In situ localization of SULTR3;3a in roots with secondary growth. Transverse sections of Populus tremula x P. alba roots with secondary growth were treated either with DIG- labeled sense probe (Ia) or with antisense probe (I). Bars indicate real µm. Abbreviations: periderm (pe); phloem (ph); sclerenchymatous phloem fibers (s); xylem (x).
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3.2.5 Transcript localization of sulfate transporter SULTR1;1 and SULTR3;3a in sink tissues, i.e. fine roots and the shoot apex
Cellular transcript accumulation of SULTR1;1 and SULTR3;3a was investigated in fine roots and the shoot apex as expected sink organs regarding sulfate distribution in plants. As control, neither treatment with sense probe for SULTR1;1 nor the sense probe for SULTR3;3a resulted in a violet staining (Figure 3.12 Ia, IIa). The brown spots, which were detected in the endodermis and also in the root cortex of both sense and antisense probe treated sections, originated from phenolic components. In situ hybridization analysis of fine roots exhibited cellular transcript accumulation of SULTR1;1 within the stele, solely (Figure 3.12 I, Id).
Figure 3.12. In situ localization of SULTR1;1 and SULTR3;3a in fine roots. Transverse sections and longitudinal sections (If, Ig) through fine roots from Populus tremula x P. alba were treated either with DIG-labeled antisense probe for SULTR1;1 (I, Ib-g) or with antisense probe for SULTR3;3a (II, IIb). As control, cross sections were treated with DIG-labeled sense probe for SULTR1;1 (Ia) and SULTR3;3a (IIa). Tetrarch (I, II) and diarch (Id) organized xylem, enlargement of the phloem (IIb, Ic, Ie), the stele (Ib) and lateral roots (Ib) are shown. Longitudinal sections of developing lateral roots (If, Ig) are indicated. Bars indicated real µm. Abbreviations: companion cell (cc); endodermis (en); lateral root (lr); phloem (ph); xylem (x);.
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SULTR1;1 mRNA signals were observed in vicinity of the xylem predominantly in cells that were assigned as phloem. This cellular specific expression pattern was obvious through all developmental stages of fine roots from the early diarch xylem up to the tetrarch organization (Figure 3.12 Ib, Ic, Ie). The same cells were stained when root primordia were developed (Figure 3.12 If). Longitudinal sections of fine roots which had developed root primordia revealed additional SULTR1;1 transcript abundance in the meristematic tissue, while strongest intensity was detected in vicinity of vascular cells (Figure 3.12 Ig). In transversal sections of fine roots where lateral roots were developed, transcript accumulation occurred mainly in the cells associated with xylem vessels (Figure 3.12 Ib). No detectable expression was found for SULTR3;3a neither in the early developmental stage nor in roots already forming tetrarch xylem (Figure 3.12 II, IIb).
Early developmental stages of the shoot were analyzed as further sink organ for sulfate. Both sense probes did not reveal any staining due to transcript accumulation (Figure 3.13 Ia, IIa). The dark brown color of leaf margin cells of very young developing leaves was caused by accumulation of phenolic compounds and was visible in both sense and antisense probe treated sections (Figure 3.13 I, II, Ia, IIa). Longitudinal shoot apex sections exhibited for both SULTR1;1 and SULTR3;3a mRNA accumulation in the shoot apex (Figure 3.13 I, II). Strong staining was observed caused by SULTR1;1 expression in leaf primordia, the shoot apical meristem and in provascular strands (Figure 3.13 I, Ib, Ic). The provascular strands arise at the leaf primordia cells and continue longitudinally throughout the apex. These cells have a long and narrow shape compared to parenchyma cells (Figure 3.13 Ic). Weak signals were detected for SULTR3;3a in provascular strands and the shoot apical meristem (Figure 3.13 II). Young developing leaves showed structures that seem to be the precursor of the leaf petiole without transcript accumulation of SULTR1;1 (Figure 3.13 Id). At an early development stage pre-lamina cells, which are equivalent to the leaf margin meristem, exhibited strong SULTR1;1 accumulation (Figure 3.13 Ie) especially in long-shaped cells (Figure 3.13 If). Corresponding expression patterns were not detected for SULTR3;3a.
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Figure 3.13. In situ localization of SULTR1;1 and SULTR3;3a in the shoot apex and developing leaves. Longitudinal sections through the shoot apex from Populus tremula x P. alba were treated either with DIG-labeled antisense probe for SULTR1;1 (I, Ib-f) and for SULTR3;3a (II). As control, longitudinal sections were treated with DIG-labeled sense probe for SULTR1;1 (Ia) and SULTR3;3a (IIa). Shoot apical meristem (Ib) and provascular cells (Ic) are indicated. Early developing leaves (Id) with an enlargement of the leaf margin meristem (Ie, If) accounted in the black box (Id), are shown. Bars indicate real µm. Abbreviations: epidermis (ep), leaf margin meristem (lm); leaf primordia (lp); provascular strands (ps); shoot apical meristem (sam).
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3.3 Influence of sulfur deprivation on sulfate transporter mRNA expression
It is assumed that sulfate distribution within plants in response to the sulfur statues is associated with the transcript abundance of sulfate transporter genes. Previous studies demonstrated that the transcript level of various sulfate transporters in many plant species changed due to the sulfate concentration in the nutrient solution (Takahashi et al., 1997; Yoshimoto et al., 2003; Buchner et al., 2004a; Hopkins et al., 2004; Kataoka et al., 2004a). In this study, the effect of sulfur deprivation on the expression of genes belonging to the sulfate transporter gene family of Populus tremula x P. alba was investigated. mRNA accumulation was analyzed in major leaf veins, bark and wood of stems by Northern blot technique (Figure 3.16-3.18). Therefore, poplar trees were grown on sand and were watered for 2, 5, 9, 14, 20 or 26 days with nutrient solution without sulfate (Chapter 2.2.2). Due to this treatment the sulfate content available for uptake through poplar roots decreased exponentially (Figure 3.14).
600
500 sulfate deprivation 400 full nutrient solution
300
sulfate (µM) 200
100
0 0 5 10 15 20 25 days of sulfate deprivation
Figure 3.14. Sulfate concentrations in the flow through during the sulfate deprivation experiment. Data exhibit sulfate concentrations in the flow through of plant pots during the sulfate deprivation treatment (black squares) and of control plant pots with full nutrient solution (black dots) (Chapter 2.2.2). The sulfate concentration in the flow through was measured by ion chromatography after watering each plant with 600 ml solution. Fits of decay indicated by solid lines were calculated using the software Origin® version 6.1 (OriginLab Corporation, Northampton, USA). 94 Results
Sulfate concentrations within the flow through of the nutrient solution were 200 µM after 5 days, 100 µM after 9 days and approximately 50 µM after 11 days of sulfur depletion. Finally, after 17 days sulfate was not detected any more in the flow through of plant pots after watering. The control plants were continuously watered with sulfur- rich nutrient solution containing 300 µM sulfate (Figure 3.14). The plants did not show any visible sign of sulfur deficiency. Neither the growth nor the constitution seemed to be affected after 26 days of sulfur deprivation. However, analyses of sulfate accumulation in major leaf veins, bark and wood tissue indicated sulfate depletion in the respective tissues (Figure 3.15).
4 c leaf vein bc abc bc abc
3 ab*
2 a
1
0 2.0 b b bark FW) -1 1.5 a a a a
1.0 a 0.5
0.0 0.8
sulfate (nmolmg wood d d d 0.6 cd
0.4 bc b 0.2 a
0.0 0 5 10 15 20 25 30 days of sulfate deprivation
Figure 3.15. Sulfate content in major leaf vein, bark and wood of stems. Four months old Populus tremula x P. alba trees grown on sand in a greenhouse were treated for 2, 5, 9, 14, 20 and 26 days with sulfate free nutrient solution as indicated (Chapter 2.2.2). Control plants were continuously grown with nutrient solution containing 0.3 mM sulfate (0 days of sulfate deprivation). The data presented are means including standard deviations of three individual plants (n = 3). (*) Different indices indicate significant differences for p< 0.05 between different time points of sulfate deprivation within one tissue.
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The three investigated tissues exhibited a slight increase in sulfate content after two or five days of sulfur depletion. Highest sulfate contents in leaf veins were observed after two days which corresponded to a significant increase of about 43%. After 26 days of sulfur deprivation sulfate content in leaf veins was only half compared to control plants. Sulfate contents in bark and wood increased by 64% and 57% after five days of sulfur depletion. Subsequently, sulfate content decreased in both tissues (Figure 3.15). Lowest sulfate levels were measured after 26 days of sulfur deprivation in all three tissues investigated. In bark (0.56 ± 0.07 nmol mg-1 FW) and leaf vein (1.29 ± 0.3 nmol mg-1 FW) sulfate contents were approximately 40% less compared to control plants and a significantly diminished sulfate content of about 80% was observed in wood (0.08 ± 0.02 nmol mg-1 FW) in comparison to all other time points.
3.3.1 Influence of sulfate deprivation on sulfate transporter transcript accumulation in bark and leaf vein
To compensate for decreased sulfate availability plants are supposed to mobilize sulfate out of vacuoles from storage tissues like bark, wood or mature leaf mesophyll cells. Upon mobilization sulfate would be loaded into the sieve tubes of storage tissue, transported via the phloem vascular system and unloaded in sink tissues, in order to supply the growing as well as developing tissues with sulfate. Mature poplar leaves are considered source tissues that supply sinks like young developing leaves with sulfate (Hartmann et al., 2000). Due to this assumption the phloem specific sulfate transporters SULTR1;1 and SULTR3;3a were investigated in major leaf veins and in bark because these tissues include the phloem vascular system. To take sulfate re-mobilization from vacuoles into account, the two sulfate transporters of group 4 were analyzed. Finally, transcript abundance of sulfate transporter SULTR2;2, which was detected in the leaf lamina, bark and wood (Figure 3.3 c), has been included in these analyzes.
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Figure 3.16. Effects of sulfur deprivation on sulfate transporter mRNA expression in leaf vein tissue. Transcript levels were analysed from 15 µg total RNA of three pooled plants by Northern blot technique (Chapter 2.3.14). Hybridization was performed with 32P-labeled, sequence specific probes for SULTR1;1, SULTR3;3a, SULTR4;1, SULTR4;2 and SULTR2;2 of Populus tremula x P. alba. Sulfate deprivation was carried out with four months old plants grown in a green house. Three plants were treated with sulfate free nutrient solution for 2, 5, 9, 14, 20 and 26 days, respectively. Control plants (c) were continuously grown with nutrient solution containing 0.3 mM sulfate. To qualify RNA loading and calculate the relative expression level transcript accumulation of the 5.8s rRNA was detected.
Growth of poplar trees under sulfate deficient conditions resulted in a two- to threefold increased expression of SULTR1;1 in major leaf veins exclusively after 26 days of sulfate deprivation (Figure 3.16), when also diminished sulfate contents were observed in leaf veins. In contrast to SULTR1;1 expression, the transcript accumulation of SULTR3;3a was slightly decreased about one fourth after two and five days of sulfur deprivation in leaf vein tissue (Figure 3.16). Transcript abundance stayed constant at this level until the end of the experiment. There was no substantial 97 Results impact on both SULTR4.1 and SULTR4.2 expression in major leaf vein tissue due to sulfate deprivation. Also SULTR2;2 transcript abundance was constant except for the expression level after nine days of sulfate deprivation which was about one-third higher compared to control plants (Figure 3.16).
Figure 3.17. Effects of sulfur deprivation on sulfate transporter mRNA expression in bark tissue. Transcript levels were analysed from 15 µg total RNA of three pooled plants by Northern blot technique (Chapter 2.3.14). Hybridization was performed with 32P-labeled, sequence specific probes for SULTR1;1, SULTR3;3a, SULTR4;1, SULTR4;2 and SULTR2;2 of Populus tremula x P. alba. Sulfate deprivation was carried out with four months old plants grown in a green house. Three plants were treated with sulfate free nutrient solution for 2, 5, 9, 14, 20 and 26 days, respectively. Control plants (c) were continuously grown with nutrient solution containing 0.3 mM sulfate. To qualify RNA loading and calculate the relative expression level transcript accumulation of the 5.8s rRNA was detected.
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Poplar trees grown under sulfate deprivation exhibited an increased expression of SULTR1;1 in bark (Figure 3.17). Already after two days transcript abundance was increased and was approximately doubled after 5 days compared to control plants. Between 9 and 20 days the transcript level remained constant except minor fluctuations. A fourfold increase of SULTR1;1 transcript accumulation in bark took place after 26 days of sulfate deprivation (Figure 3.17). Comparable to SULTR1;1 both, SULTR4.1 and SULTR4.2, exhibited a substantial increase of transcript accumulation in bark after 26 days. Transcript amount increased threefold for SULTR4.2 and one and a half fold for SULTR4.1, compared to control plants. The sulfate transporter SULTR3;3a, which was also specifically expressed in phloem cells (Figure 3.17), exhibited a different expression pattern in comparison to SULTR1;1. In bark tissue mRNA abundance was halved after two days of sulfur deprivation followed by an increase of the transcript level at day five (Figure 3.17). A diminished SULTR3;3a expression was observed again after 20 days. SULTR2;2 exhibited an increased transcript accumulation in bark which was 80% after five days of sulfur deprivation compared to the control trees (Figure 3.17). Transcript abundance stayed at this high level except minor fluctuations until day 20. Further increased expression, approximately threefold, after 26 days of sulfur deprivation was observed compared to control plants.
3.3.2 Influence of sulfate deprivation on sulfate transporter transcript accumulation in stem wood
Sulfate transporter mRNA expression in poplar stem wood was analyzed under sulfur deficiency because living cells of wood tissue are involved in sulfur storage (Herschbach and Rennenberg, 1996). Seven sulfate transporter genes transcribed in woody tissues (see Figure 3.3 d) were investigated in addition to SULTR4;2 and SULTR2;2. The strongest increase of transcript accumulation was exhibited for SULTR4.2, which was 5-fold after 26 days of sulfur deprivation (Figure 3.18). Slightly increased transcript accumulations of SULTR2;2 of approximately 30% were detected after two, 14 and 20 days of sulfur depletion in stem wood.
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Figure 3.18. Effects of sulfur deprivation on sulfate transporter mRNA expression in stem wood. Transcript levels were analyzed from 15 µg total RNA of three pooled plants by Northern blot technique (Chapter 2.3.14). Hybridization was performed with 32P-labeled, sequence specific probes for SULTR4;2, SULTR2;2, SULTR2;1a, SULTR2;1b, SULTR3;2a, SULTR3;2b, SULTR3;1a, SULTR3;1b and SULTR3;4a of Populus tremula x P. alba. Sulfate deprivation was carried out with four months old plants grown in a green house. Three plants were treated with sulfate free nutrient solution for 2, 5, 9, 14, 20 and 26 days, respectively. Control plants (c) were continuously grown with nutrient solution containing 0.3 mM sulfate. To qualify RNA loading and calculate the relative expression level transcript accumulation of the 5.8s rRNA was detected.
Three of the selected wood specific transporters, SULTR2;1a, SULTR2;1b, and SULTR3;2b exhibited similar expression patterns. These genes showed diminished mRNA abundances after 26 days of sulfate deficiency (Figure 3.18). The mRNA amount of SULTR2;1a declined more than 6-fold whereas the amounts of both SULTR2;1b and SULTR3;2b were approximately 3-fold diminished compared to the control plants. While both SULTR2;1 genes exhibited an abrupt decrease in transcript abundances, the level of SULTR3;2b decreased continuously. Three of the remaining four sulfate transporters, namely SULTR3;1a, SULTR3;2a and SULTR3;4a showed mainly constant expression levels during the treatment, except slight fluctuations (Figure 3.18). SULTR3;2a exhibited exactly the same expression pattern in stem wood tissue than SULTR2;2, which was an approximately 30% increase at day two, 14 and 20. Sulfate transporter SULTR3;1b exhibited substantially decreased mRNA accumulation after two days. Afterwards, mRNA abundance increased again peaking after 14 days and subsequently decreasing successively (Figure 3.18).
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3.4 Seasonal influences on sulfate transporter gene expression
Seasons are characterized by a combination of variations in temperature, day length, rainfall, humidity and solar radiation (Figure 3.19). At the temperate climate in Central Europe, seasonal influences on poplar development and growth cycles are supposed to be mediated mainly by temperature and day length (Jansson and Douglas, 2007). Especially the transition state between the growing season in summer and the dormant season during winter and vice versa are of high significance. In most parts of Europe the growing season is defined by the average number of days per year with a 24-hour average temperature of at least 5°C, which is typically from April until October or November. This pattern, however, varies considerably with latitude and altitude. In 2006, mean temperatures below 5°C were first observed at October 2nd and the last day below 5°C in 2007 was on March 31st. The latter date coincided with bud break (Figure 3.20 C).
100 30
25 90 Relative humidity(%) Relative 20 80
15 70 10 60 5 Temperature (°C) Temperature
0 50
5 40 Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun Jul Aug
Date
Figure 3.19. Weather conditions in Freiburg from August 2006 to September 2007. Maximum temperature (black circles), minimum temperature (black triangles) and mean temperature (black squares) are average values calculated from the 24 h period before sampling. Grey bars exhibit the average relative humidity of the 24 h period before sampling. The weather data were available from the meteorological station 10803 (Deutscher Wetterdienst , Airport Freiburg, Germany).
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Plant material was collected on 29 dates distributed over the year to analyze seasonal expression patterns of sulfate transporter genes. 16 field grown Populus tremula x P. alba trees were used for this study and for each date samples were taken from three independent trees between end of August 2006 and September 2007 (Table 2.1, Figure 3.20 A). In order to investigate if phloem loading and unloading of sulfate is correlated with environmental parameters during spring and fall, sampling of plant material was carried out weekly during spring (end March until start May) and autumn (start September until end October) (Table 2.1). During summer and winter sampling was less frequent. Twig bark and leaves were analyzed, because these tissues switch within their function as source and sink organs for sulfate during annual growth (Herschbach and Rennenberg, 1996).
Figure 3.20. Populus tremula x P. alba trees during the season. Full habitus (A) and foliage (B) of the trees investigated in August 2006. Beginning of bud break on April 3rd 2007 (C), open buds on April 11th 2007 (D) and young developed leaves at the end of April 2007 (E) are shown. Senescent leaves still attached were observed at November 15th 2006 (F).
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Gene expression of the two phloem specific sulfate transporters SULTR1;1 and SULTR3;3a was investigated to analyze their contribution on sulfate distribution within the plant via the phloem vascular system. Further analyses were performed to investigate group 4 sulfate transporters from Populus tremula x P. alba, which are probably localized in the tonoplast of vacuoles, a major site of sulfate storage. The amounts of sulfate transporter mRNA were analyzed by quantitative real-time RT PCR (Chapter 2.3.15). A sequence belonging to the elongation factor gene family (elongation factor 1-beta; EF1beta) was selected as reference gene, because it was shown that these genes are stable expressed during biotic and abiotic stresses in plants (Brunner et al., 2004a; Nicot et al., 2005).
3.4.1 Seasonal influences on sulfate transporter gene expression in bark tissue
The expression patterns of the sulfate transporter genes SULTR1;1, SULTR3;3a, SULTR4;1 and SULTR4;2 in the bark exhibited significant variation in mRNA abundance during the seasons. All four transporters showed mainly long term trends with partly comparable expression patterns. For instance, continuously diminishing transcript levels were observed in the bark of young, developing twigs from mid May until July (Figure 3.21). This decreasing transcript abundance was noticeable for all investigated genes including EF1beta from May 18th until July 2007 (p < 0.05; May 18th, June 5th > July 31th, August 14th, August 28th). All genes examined except SULTR3;3a showed higher transcript abundance in bark of new developing twigs (2007) in May and June compared to bark tissue of twigs that developed the year before (2006). The most noticeable influence on SULTR1;1 mRNA abundance in bark tissue was detected during spring 2007. Bud break took place between April 3rd and 11th and the first leaves were observed around mid April (Figure 3.20 E). Coinciding with bud break (Figure 3.20 C) SULTR1;1 transcript amount increased in bark significantly 2- fold during each week between April 3rd and April 18th (Figure 3.21) (p < 0.05; April 3rd < April 11th < April 18th). On April 18th, the transcript level reached its maximum for the entire sampling period. After the first leaves were developed, a significant decrease of SULTR1;1 transcript amount was observed within one week from April 18th to the 26th (Figure 3.21).
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6 FW)
-1 4
2 sulfate sulfate
(nmol mg 0 SULTR1;1 15 10
5 0 30 SULTR3;3a 20
total RNA) total 10 -1 0 20 SULTR4;1 15 10
5 0 20 SULTR4;2 15 mRNA (copies * 100000 µg
10 5 0 200 EF1beta 150
100
50 0 Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun Jul Aug Sep 2006 - 2007
Figure 3.21. Transcript levels and sulfate contents during the season in bark tissue. Transcript abundances from field grown Populus tremula x P. alba trees were determined with quantitative real- time RT PCR using gene specific primers. SULTR1;1, SULTR3;3a, SULTR4;1, SULTR4;2 and the reference gene EF1beta were investigated (Chapter 2.3.15). Samples were harvested from August 2006 to August 2007 at the dates given in table 2.1. Dashes indicate the 15th of the respective month. Bark developed in 2006 (grey) was differentiated from bark developed in 2007 (black). Sulfate contents were measured by ion chromatography (Chapter 2.5). Values presented are means ± SD of three individual trees. The arrow indicates bud break. 105 Results
Another significant diminishing of mRNA abundance was detected during autumn within one week between October 5th and 11th. Within this time period in 2006 the growing season ended and switched into the dormant period, as indicated by mean temperatures below 5°C. The lower expression level during the dormant period was interrupted by a significant increase in December, when the transcript abundance about doubled compared to October. This increase coincided with the lowest temperature observed in December for the entire season with a mean value around 0°C and a minimum value around -5°C (Figure 3.19).
The second phloem specific sulfate transporter SULTR3;3a showed a different expression pattern compared to SULTR1;1. Most noticeable was the absence of transcript abundance in bark tissue during dormancy from October 2006 until March 2007 (Figure 3.21). Strong changes were visible during spring with a weekly significant increase of mRNA accumulation from March 12th until the April 18th (p < 0.05; March 12th < April 3rd, p < 0.068; April 3rd < April 18th). This considerable increase occurred simultaneously with the start of the growing season and was similar for both phloem specific sulfate transporters. The total increase of SULTR3;3a mRNA between March and April was 35-fold and compared to the lowest expression level in December the transcript abundance on April 18th was 250-fold higher. While the transcript abundance of SULTR1;1 was constant during late summer and autumn 2006 (end August until start October) strong fluctuations were detected for SULTR3;3a with peak transcript accumulations at August 28th and September 26th (Figure 3.21). Seasonal expression patterns of the two sulfate transporters SULTR4;1 and SULTR4;2 in bark tissue were different during most periods (Figure 3.21), but comparably similar during late summer 2006, when a significant decrease in mRNA abundance (p < 0.05; Aug 28th > Sep 13th) was followed by higher mRNA amounts (late September). A further decrease was observed in autumn at October 5th when the growing season turned into the dormant period (Figure 3.21). However, strong differences were noticed in the expression pattern of SULTR4;1 and SULTR4;2 in bark tissue during winter and spring. While the expression of SULTR4;2 was constant during dormancy at a very low level, the transcript amount of SULTR4;1 increased significantly during winter with the highest expression level at December 21st (Figure 3.21) (p < 0.05; Oct 25th < Nov 22nd < Dec 21st > Jan 22nd). The mRNA amount on
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December 21st was 15-fold higher than on October 25th. In contrast the mRNA abundance of SULTR4;2 was almost not detectable during winter which was comparable to the expression pattern of SULTR3.3a. The transcript abundance of SULTR4;2 increased again in spring with a significant threefold increase between March 12th and March 27th just before bud break (Figure 3.20 C, 3.21). The subsequent slight increase lasted until May 18th and resulted in a 23-fold mRNA abundance compared to the lowest level in winter. At this date young leaves were fully developed. By contrast, SULTR4;1 exhibited a constant expression level from January until April 11th (Figure 3.21). The average mRNA level of all sulfate transporter genes investigated was comparable in stem bark tissue. The reference gene EF1beta exhibited almost constant transcript amounts in poplar stem bark with slight fluctuations, for instance in November, when mRNA amounts increased. A significant decrease was observed during the switch from the growing season into dormancy in October. Additionally, mRNA abundance declined within developing bark from end of June until August (Figure 3.21) (May 18th, June 4th > July 31st Aug 14th, Aug 28th). The sulfate content in bark tissue exhibited a strong seasonal fluctuation. The content was high during dormancy and low during the growing season (Figure 3.21). During leaf senescence in autumn, the sulfate content in bark tissue increased 4-fold from 1.2 ± 0.3 nmol mg-1 FW at the end of October to 4.8 ± 0.8 nmol mg-1 FW in December (p < 0.05; Oct 5th < Oct 25th < Nov 8th < Dec 21st). This high sulfate content was observed during the entire dormant period. Simultaneously with bud break, sulfate content declined suddenly and significantly after April 3rd within two weeks. Subsequently, a slight decrease was observed until May 18th when leaves were fully developed (p < 0.05; April 18th > May 3rd, May 18th).
3.4.2 Seasonal influences on sulfate transporter gene expression in leaves
The seasonal expression pattern in leaves of the two phloem specific sulfate transporters SULTR1;1 and SULTR3;3a exhibited different mRNA abundances during leaf development in spring 2007 (Figure 3.22). For both, a successive increase led to a peak followed by continuously diminishing transcript accumulation. This expression pattern was temporally different for the two sulfate transporters.
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SULTR1;1 transcript abundance increased continuously from the first foliage harvested on the 18th of April and reached peak amounts on the 18th of May when leaves were fully developed. The following decrease lasted until June. Both, the increase and decrease were significant (p < 0.05; April 18th < May 18th > Aug 14th, Aug 28th, Sep 13th) and caused approximately 10-fold differences when comparing the highest and the lowest transcript amount. During summer and autumn, SULTR1;1 expression remained at a low level, except a considerable increase of mRNA abundance in senescent, but still attached yellow leaves (Figure 3.20 F, 3.22). This increase was observed from October 25th to November 8th and was 6-fold. SULTR3;3a also exhibited an increase followed by a decrease of transcript amount in the leaves developed in 2007. However, peak expression occurred approximately 1.5 months latter than observed for SULTR1;1 (Figure 3.22). The increase of SULTR3;3a mRNA started after May 18th and reached a peak at the end of June. Thereafter, SULTR3;3a mRNA levels declined until September 13th 2007, when the lowest expression level was observed. At least the highest and the lowest expression values were significantly different. The same low expression level was detected from late summer 2006 until leaf fall in autumn.
The expression pattern of both SULTR4 sulfate transporters exhibited continuously increasing transcript abundances during leaf development in spring which were followed by declining transcript abundances after leaf maturation (Figure 3.22). SULTR4;2 showed the highest peak expression level on May 18th like SULTR1;1, while SULTR4;1 showed the highest peak value at the 4th of June. The decrease in mRNA abundance visible for both genes was significant between the highest peak and the transcript amounts observed from August until September 2007. Transcript accumulation in young leaves on April 18th was significantly lower compared to the transcript accumulation in mature leaves in mid May. This expression pattern was comparable to the observation for SULTR1;1. Another analogy to SULTR1;1 was noticeable for SULTR4;1 in autumn 2006 (Figure 3.22). The transcript abundance of both genes increased in senescent leaves between the 25th of October and 8th of November before leaf fall. During late summer and autumn 2006 expression of group 4 sulfate transporters exhibited constant levels except minor fluctuations which was also found for SULTR1;1 and SULTR3;3a.
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8
FW] 6 -1 4
sulfate 2
[nmol mg 0 40 SULTR1;1 30
20
10 0 3 SULTR3;3a
2
1
total RNA) 0 -1 80 SULTR4;1 60
40
20 0 80 SULTR4;2 60 40
mRNA (copies * 100000 µg 20 0 3000 EF1beta
2000
1000
0 Aug Sep Oct Nov Dec Jan Feb Mar Apr May Jun Jul Aug Sep 2006 2007
Figure 3.22. Transcript levels and sulfate contents during the season in leaves. Transcript abundances in leaves from field grown Populus tremula x P. alba trees were determined with quantitative real-time RT PCR using gene specific primers. SULTR1;1, SULTR3;3a, SULTR4;1 SULTR4;2 and the reference gene EF1beta were investigated (Chapter 2.3.15). Leaves were harvested from August 2006 (grey bars) until September 2007 (black bars) at the dates given in table 2.1. Dashes indicate the 15th of each month. Sulfate contents were measured by ion chromatography (Chapter 2.5). Data presented are means ± SD from three individual trees.
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EF1beta exhibited a rather constant mRNA level in leaves during the whole season usually with only minor fluctuations, but with one exception (Figure 3.22): Significantly elevated transcript amounts were observed after bud break in young developing leaves in April. The sulfate content in leaves showed no significant differences between senescent leaves collected in autumn 2006 and the newly developed leaves collected in spring 2007 (Figure 3.22). Sulfate content decreased significantly during leaf development from April to May 2007 followed by a continuous increase until June (p < 0 .05; April 18th > May 18th < June 29th). In both years highest sulfate contents were found in summer on the 28th of August 2006 and 13th of September 2007. In late summer 2006, the sulfate content declined significantly by about 70% from 6.8 ± 1.4 nmol mg- 1 FW to approximately 2 ± 0.3 nmol mg-1 FW and remained constant at this low level until leaves became senescent (p < 0.05; Aug 28th, Sep 6th > Sep 20th).
3.4.3 Principle component, correlation and regression analysis
Principle component analysis (PCA) was used to extract systematic trends from the large data set of seasonal investigations. This method is a multivariate technique that analyzes a correlation matrix to compute few independent factors that are linear combinations of the variables and explain most of the variation in the data (Sokal and Rohlf, 1995; Soler et al., 2008). Data of gene expression levels and sulfate contents were reduced to a few principal components characterizing the strongest systematic effects according to the variance in the data (Sjödin et al., 2008). Each of these principal components describes independent effects in the data, which relate the samples by means of score vectors and the corresponding features using loading vectors. In the present study, PCA was applied to summarize the variations within the variables of the four investigated sulfate transporter genes SULTR3;3a, SULTR4;2, SULTR1;1, SULTR4;1 and the sulfate contents as well as within the sampling dates. The results are shown in bivariate diagrams corresponding to the two main factors extracted. The Kaiser-Mayer-Olkin’s (KMO) measure of sample adequacy assessed that many transcript and sulfate content variables were interdependent (leaves: KMO = 0.675; bark: KMO = 0.610). The two axes of the principle component analysis (PCA) explained for bark 80% and for leaves 77% of the variations. PCA factor loadings for the four sulfate transporter gene transcript profiles and sulfate contents
110 Results and the scores of the sampling dates for the two PCA components were separately investigated for bark and leaves (Figure 3.23, 3.27).
Analysis of bark tissue
The first principle component of bark analysis explained 48% of the variations and separated the sulfate content from the gene expression data, while the transporter transcripts SULTR3;3a, SULTR4;2, SULTR1;1 and to a lesser extend SULTR4;1 grouped together (Figure 3.23 A). Because the dates indicated mainly high transcript amounts clustered right in the score plot around 2 and the mainly low values cluster left around -1, these factor score reflect individual variations in the transcript or sulfate content amounts. The transcript abundance of all four sulfate transporter genes were particularly high at August 2006 28th, April 18th, May 18th and June 4th and low at January 22nd, February 15th, March 12th, November 22nd, November 8th, October 25th and October 11th (Figure 3.23 B).
loading score
A B
Figure 3.23. Principle component analysis (PCA) of transcript accumulations and sulfate contents in bark. PCA of the transcript abundances of the four sulfate transporter genes SULTR1;1, SULTR3;3a, SULTR4;1, SULTR4;2 and of sulfate contents were analyzed. 26 dates during August 2006 until August 2007 plus young twigs (written italic) at two particular dates were implicated (Table 2.1). Factor loadings of the four genes and sulfate content for the two PCA axes (A) and mean PCA scores of the sampling dates for the two axes extracted (B) are presented.
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The second principle component explained 32% of the variations and highlighted one group of two sulfate transporters consisting of SULTR3;3a and SULTR4;2. Both were separated from SULTR1;1, SULTR4;1 and the sulfate content (Figure 3.23 A). Second factor score separated samples taken on December 21st from the remaining samples. At this date main differences within gene expression of the grouped sulfate transporters compared to the investigated mRNA levels of SULTR1;1 and SULTR4;1 occurred (Figure 3.23 B). Transcript abundances of SULTR3;3a and SULTR4;2 were almost not detectable during dormancy, while SULTR1;1 and SULTR4;1 and sulfate content peaked in December. Furthermore, the samples early in the year 2007 from January 22nd until April 3rd grouped. At bud break and during leaf development the gene transcripts particularly of SULTR1;1, SULTR3;3a and SULTR4;2 increased while the sulfate contents decreased (Figure 3.21). The second factor score was significantly negative correlated to maximum, minimum, mean temperature and additionally to day length (r = -0.611**, -0.717**, -0.721**; 0.497** at p < 0.01), but unrelated to other meteorological parameters like relative humidity, wind, rain and sun hours.
Table 3.4. Correlation matrix among the transcript abundance in bark, with sulfate contents and weather parameters. Values presented are (Pearson-) correlation coefficients of the transcript abundances of the sulfate transporter genes indicated (**, p < 0.01). Correlations of expression levels with sulfate content and weather parameters (temperature (T), day length) are presented. The coefficient can reach values between -1 and 1. Values of approximately 1 signify a distinct increasing linear trend while values of approximately -1 indicate a distinct decreasing linear trend. Values of approximately 0 verify no statistical linear correlation.
SULTR1;1 SULTR3;3a SULTR4;1 SULTR4;2 sulfate
SULTR3;3a 0.301 SULTR4;1 0.546** 0.013 SULTR4;2 0.363 0.881** 0.238 sulfate - 0.010 - 0.621** 0.375 - 0.521** T mean - 0.068 0.749** - 0.454 0.596 ** - 0.682** T max - 0.100 0.703** - 0.421 0.583** - 0.529** T min 0.008 0.691** - 0.399 0.491** - 0.750** day length - 0.067 0.814** - 0.138 0.866** - 0.639**
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15 AB14.5 r = 0.881 14 p < 0.0001 14.0 y = 0.518x + 6.45 n = 29 13 13.5 mRNA mRNA 12 13.0
12.5 11 SULTR4;2 r = 0.546 12.0 SULTR4;1 10 p = 0.002 y = 0.664x + 4.40 n = 29 11.5 9 8 9 10 11 12 13 14 15 11.5 12.0 12.5 13.0 13.5 14.0 14.5 SULTR3;3a mRNA SULTR1;1 mRNA
C D r = -0.521 2.5 2.0 p = 0.004 y = -0.332x + 4.83 2.0 n = 28 1.5 1.5 1.0 1.0
0.5
0.5 sulfate sulfate
0.0 0.0 r = -0.621 -0.5 p < 0.001 y = -0.238x + 3.47 -0.5 n = 28 -1.0 -1.0 8 9 10 11 12 13 14 15 9 101112131415 SULTR3;3a mRNA SULTR4;2 mRNA
Figure 3.24. Regression analysis of sulfate transporter expression levels and sulfate contents in bark. Regression analysis between correlated gene transcript accumulations of SULTR3;3a with SULTR4;2 (A) and SULTR1;1 with SULTR4;1 (B) are shown. SULTR3;3a (C) and SULTR4;2 (D) transcripts correlated with sulfate contents are indicated. 26 or 27 dates during August 2006 until August 2007 plus young twigs at two particular dates were analyzed (Table 2.1). Correlation coefficient (r), probability that r is cero (p), samples (n) and the linear equation (y) are presented.
To analyse linear correlations between the expression pattern of two genes and the expression pattern of a particular gene with weather conditions, bivariate Pearson- correlation analyses were performed and the linear regression was calculated. The grouped transcript abundances of SULTR3;3a and SULTR4;2 according to PCA second factor score were significantly positive correlated in bark tissue (r = 0.881; p < 0.01) (Figure 3.24 A) and both mRNA levels were negatively correlated to the sulfate content in bark tissue (r = -0.621; -0.521; p < 0.01) (Table 3.4, Figure 3.24 C, D).
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300 250 r = 0.583 p < 0.001 250 y = 31.20x -211.46 200 n = 29
200 150
150 100 T max T mean 100 50 r = 0.596 p < 0.001 y = 29.02x -234.26 50 0 n = 29 200 1000 r = 0.491 150 p = 0.007 900 y = 24.22x -224.47 n = 29 100 800
T min 50 700 length day 0 r = 0.866 600 p < 0.0001 y = 101.35x -523.17 -50 n = 29 500
9 1011121314159 101112131415 SULTR4;2 mRNA SULTR4;2 mRNA
300 250 r = 0.703 p < 0.0001 250 y = 22.13x - 81.32 200 n = 29
200 150
150 100 T max T mean 100 50 r = 0.749 p < 0.0001 y = 21.43x - 123.55 50 n = 29 0 200
r = 0.691 150 p < 0.0001 900 y = 20.07x - 158.17 n = 29 100 800
50 700 T min
daylength 0 600 r = 0.814 p < 0.0001 -50 y = 56.05x + 88.05 500 n = 29 8 9 10 11 12 13 14 15 8 9 10 11 12 13 14 15 SULTR3;3a mRNA SULTR3;3a mRNA
Figure 3.25. Regression analysis of SULTR3;3a and SULTR4;2 mRNA expression levels of bark tissue with weather parameters. Regression analysis between SULTR3;3a mRN (bottom graphs) or SULTR4;2 mRN (top graphs) abundances with mean temperature (T mean), maximum temperature (T max), minimum temperature (T min) and the day lengths (one day before sampling date) are indicated. Weather data are means of the 24 h period before sampling. 27 dates during August 2006 until August 2007 plus young twigs at two particular dates were analyzed (Table 2.1). Correlation coefficient (r), probability that r is cero (p), samples (n) and the linear equation (y) are presented.
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Additionally, SULTR1;1 and SULTR4;1 mRNA levels were positively correlated with each other (Figure 3.24 B). SULTR3;3a and SULTR4;2 transcripts were positively and the sulfate content was negatively correlated with mean temperature, maximum temperature, minimum temperature and the day lengths (Table 3.4, Figure 3.25, 3.26), which agreed with the effect extracted by the second PCR factor score (Figure 3.23).
300 r = -0.682 250 p < 0.0001 y = -50.95x + 166.67 250 n = 28 200
200 150
150 100 T max T mean
100 50 r = -0.529 p = 0.004 50 y = -42.69x + 213.01 0 n = 28
200 r = -0.639 1000 p < 0.001 y = -166.77x +835.23 150 n = 28 900
100 800
50 700 daylength T min
0 600 r = -0.750 p < 0.0001 -50 y = -58.22x + 118.67 500 n = 28
-1.0 -0.5 0.0 0.5 1.0 1.5-1.0 2.0 -0.5 0.0 0.5 1.0 1.5 2.0 sulfate sulfate
Figure 3.26. Regression analysis of sulfate content in bark tissue with weather parameters. Regression analysis between sulfate contents and mean temperature (T mean), maximum temperature (T max), minimum temperature (T min) and the day length (one day before sampling date) are indicated. Weather data are means of 24 h before sampling. 26 dates during August 2006 until August 2007 plus young twigs at two particular dates were analyzed (Table 2.1). Correlation coefficient (r), probability that r is cero (p), samples (n) and the linear equation (y) are presented.
115 Results
Analyses of leaf tissue
In the data set of leaf samples, the first principle component explained 56% of the variations and separated SULTR3;3a transcript abundances and the sulfate content from SULTR1;1, SULTR4;1, SULTR4;2 mRNA levels, which grouped (Figure 3.27 A). The first factor score mostly separated November 8th, May 18th and June 4th (Figure 3.27 B) from the remaining dates, which were generally the dates with highest transcript levels during the season. The expression patterns of the three sulfate transporters SULTR1;1, SULTR4;1 and SULTR4;2 were positively correlated to each other in leaves (Table 3.5; Figure 3.28 A, B, D) and compared to SULTR3;3a highly expressed on November 8th, May 18th and June 4th. The first axis score was significantly negative correlated to maximum temperature (r = -0.477; p < 0.05) but unrelated to other meteorological parameters. Bivariate correlation analyses exhibited only for SULTR4;2 a negative linear correlation with maximum temperature and a positive correlation with relative humidity (Table 3.5, Figure 3.29).
loading score
A B
Figure 3.27. Principle component analysis (PCA) of transcript accumulations and sulfate contents in leaf. PCA of the transcript abundances of the four sulfate transporter genes SULTR1;1, SULTR3;3a, SULTR4;1, SULTR4;2 and the sulfate content were analyzed on 20 dates during August 2006 until September 2007 (Table 2.1). Factor loadings of the four genes and sulfate content for the two PCA axes (A) and mean PCA scores of the sampling dates for the two axes extracted (B) are presented.
116 Results
Table 3.5. Correlation matrix among the transcript abundance in leaves, with sulfate contents and weather parameters. Values presented are (Pearson-) correlation coefficients of the transcript abundances of the sulfate transporter genes indicated (**, p < 0.01). Correlations of expression levels with sulfate content and weather parameters (temperature (T), day length, humidity) are presented. The coefficient can reach values between -1 and 1. Values of approximately 1 signify a distinct increasing linear trend while values of approximately -1 indicate a distinct decreasing linear trend. Values of approximately 0 verify no statistical linear correlation.
SULTR1;1 SULTR3;3a SULTR4;1 SULTR4;2 sulfate
SULTR3;3a - 0.002 SULTR4;1 0.587** 0.303 SULTR4;2 0.725** 0.076 0.589** sulfate - 0.645** 0.160 - 0.275 - 0.529 T mean - 0.296 - 0.057 - 0.224 - 0.506 0.383 T max - 0.381 - 0.057 - 0.259 - 0.638** 0.328 T min - 0.266 - 0.194 - 0.328 - 0.204 0.301 day length 0.112 0.299 0.185 - 0.187 - 0.021 humidity 0.342 0.067 - 0.011 0.564** - 0.238
The second component explained 23% of the variations in expression and grouped the two sulfate transporters SULTR4;2 and SULTR1;1, while SULTR3;3a was separated. The second factor score separated most summer dates (June 4th, June 29th, July 31st, Aug 14th Aug 28th 2006) from the remaining dates and additionally on the opposite scale side both May dates (Figure 27 B). The transcript abundances of SULTR4;2 and SULTR1;1 increased from April 18th until May 18th followed by an additional slight increase until August (Figure 3.22). SULTR1;1 was the only sulfate transporter which exhibited linear negative correlations between the transcript abundances and the sulfate content in leaf tissues (Table 3.5; Figure 3.28 C).
117 Results
15.6 15.6 A B 15.4 r = 0.589 15.4 p = 0.006 y = 0.606x + 5.75 15.2 15.2 n = 20
15.0 15.0 mRNA mRNA
14.8 14.8
14.6 14.6 SULTR4;2 r = 0.725 SULTR4;2 14.4 14.4 p < 0.001 y = 0.335x + 10.48 14.2 n = 20 14.2
14.4 14.6 14.8 15.0 15.2 15.4 15.6 15.8 12.0 12.5 13.0 13.5 14.0 14.5 15.0 SULTR4;1 mRNA SULTR1;1 mRNA
C D 15.8 2.0
15.6
1.5 15.4
15.2 1.0 mRNA
15.0 sulfate 0.5 14.8 SULTR4;1 r = 0.587 r = -0.645 p = 0.007 p = 0.002 y = 0.264x + 11.68 14.6 0.0 y = -0.464x + 7.16 n = 20 n = 20 14.4 12.0 12.5 13.0 13.5 14.0 14.5 15.0 12.0 12.5 13.0 13.5 14.0 14.5 15.0 SULTR1;1 mRNA SULTR1;1 mRNA
Figure 3.28. Regression analysis of sulfate transporter expression levels and sulfate contents in leaf. Regression analysis between correlated gene transcript accumulations of SULTR4;1 with SULTR4;2 (A), SULTR1;1 with SULTR4;2 (B) and SULTR4;1 with SULTR1;1 (C) are shown. SULTR1;1 transcripts correlated with sulfate contents (D) are indicated. 20 dates during August 2006 until September 2007 were analyzed (Table 2.1). Correlation coefficient (r), probability that r is cero (p), samples (n) and the linear equation (y) are presented
118 Results
100 28
26 90
24 80 22
70
20 T mean 18 60 r = -0.638 r = 0.564 16 humidity relative p = 0.002 p = 0.01 y = -9.16x + 158.16 50 14 y = 20.81x - 238.93 n = 20 n = 20 12 40 14.2 14.4 14.6 14.8 15.0 15.2 15.4 15.6 14.2 14.4 14.6 14.8 15.0 15.2 15.4 15.6 SULTR4;2 mRNA SULTR4;2 mRNA
Figure 3.29. Regression analysis of SULTR4;2 transcript abundances in leave tissue with weather parameters. Regression analysis between SULTR4;2 transcript abundances and mean temperature (T mean) and relative humidity are indicated. Weather data are means of 24 h before sampling. Correlation coefficient (r), probability that r is cero (p), samples (n) and the linear equation (y) are presented.
In summary, the observed correlations between sulfate transporter gene expressions were exclusively positive. Correlations for leaf and bark were rarely equal. The SULTR1;1 transcript pattern correlated with the mRNA level of both sulfate transporters from group 4 in leaves and with SULTR4;1 in bark tissues. In contrast SULTR3;3a correlated with SULTR4;1 transcript abundance in bark only. The mRNA levels of the two sulfate transporter belonging to group 4 were only correlated in leaves. The expression patterns of the two phloem specific sulfate transporters were not correlated to each other neither in bark nor in leaves. Transcript abundances of SULTR3;3a and SULTR4;2 were negatively correlated with the sulfate content in bark, but not in leaves. Influences of temperature and day length were mainly noticeable for the expression of SULTR3;3a, SULTR4;2 and the sulfate content in bark.
119 Discussion
4 Discussion
4.1 Populus tremula x P. alba sulfate transporter gene family
In the present study, the gene family of sulfate transporters were investigated in poplar trees. Poplar showed a larger number of putative open reading frames for predicted sulfate transporters compared to the appropriate gene family of herbaceous plant species. For Populus trichocarpa via database analysis as well as for Populus tremula x P. alba via sequence cloning 18 putative sulfate transporter genes were identified. Like Arabidopsis thaliana, Populus belongs phylogenetically to the rosids and therefore it is well positioned for comparative analysis (Brunner et al., 2004b). Comparative mapping has revealed extensive genome co-linearity between species of closely related plants (Schmidt, 2002). Phylogenetic analyses of gene families can identify putative orthologs of genes that have been characterized in Arabidopsis thaliana and can reveal lineage-specific duplications. Compared to the Arabidopsis thaliana sulfate transporter gene family which contains 14 sequences and rice with 13 members, poplar seems to exhibit gene duplications in this family. In terms of genome evolution, analyses of Arabidopsis thaliana and poplar ESTs have revealed whole poplar genome duplication (salicoid duplication) (Tuskan et al., 2006). Thus, a single- copy gene in Arabidopsis thaliana is typically represented by two copies in Populus species (Jansson and Douglas, 2007). This could be the reason for the observation that two highly homolog SULTR sequences were related to only one sulfate transporter of the Arabidopsis thaliana gene family in the present study. It can be excluded that the closely related genes are derived from the two combined genomes of the used hybrid Populus tremula x P. alba because the sequences found are also represented in the Populus trichocarpa genome. In many cases the two closely related SULTR sequences from poplar exhibited comparable tissue specific transcript accumulation (SULTR2;1a/b, SULTR3;2a/b, SULTR4;1/2, SULTR5;1/2) (Figure 3.3). This was not observed for both SULTR3;1a/b and SULTR3;3a/b. It was shown that many duplicate genes in the Populus genome have undergone subfunctionalization in expression (Tuskan et al., 2006) suggesting genome reorganization during evolution. Additionally, not each gene sequence is presented twice in poplar because a loss of 120 Discussion genes during evolution can minimize this effect.
Sequence comparison including accurate statistical estimation allows the characterization of sequence relationships (Pearson, 2000). The significant homologies of deduced amino acid sequences of polypeptides encoded by 16 of the 18 putative Populus tremula x P. alba sulfate transporter genes with sequences of proton / sulfate co-transporters that were isolated and functionally characterized from Arabidopsis thaliana (Takahashi et al. 2000, Yoshimoto et al., 2002, 2003; Kataoka et al., 2004b) make it highly likely that the poplar genes have the expected sulfate transport function. Only the partial sequences of Populus tremula x P. alba putative sulfate transporters of group 5 did exhibit less high homologies. For these a sulfate transport function is questionable because contrary to previous suggestions, the ortholog protein AtSULTR5;2 from Arabidopsis thaliana was recently identified as molybdate transporter (Tomatsu et al., 2007).
Substantial blocks of high sequence conservation support possible functional motifs of proteins. Therefore, strong consensuses allow description or rather the functional demonstration of the cloned poplar genes because the compared proteins are well characterized (Takahashi et al., 1997; Shibagaki et al., 2004; Rouached et al., 2005). The topology of the sulfate transporter sequences PtaSULTR1;1, PtaSULTR3;3a, PtaSULTR4;1 and PtaSULTR4;2 fits the twelve membrane spanning domain (MSD) model that has been proposed for the sulfate transporters isolated from Stylosanthes hamata and Arabidopsis thaliana (Smith et al., 1995; Takahashi et al., 1997; Hawkesford, 2003). Such membrane spanning helices are wide spread structures for sulfate transporters (Hästbacka et al., 1994; Smith et al., 2000) and characteristic for integral membrane proteins in plants (Chen et al., 2008). According to this model for sulfate transporters, several basic amino acid residues (lysine, arginine) were located on both sides of the membrane. One basic arginine residue, which is located between MSD 9 and 10, is identical in most SULTR sequences from poplar, Arabidopsis thaliana, Stylosanthes hamata and Hordeum vulgare (Takahashi et al., 1997). It is found for all SULTR sequences from poplar except for PtaSULTR3;2a and the members of group 5. As previously indicated, this residue may have some functional significance for binding or channelling the sulfate anion (Smith et al., 1995, 2000). The absence of this basic residue in both PtaSULTR5 sequences agrees with the
121 Discussion mentioned before studies revealing that the Arabidopsis thaliana AtSULTR5;2 sequence encodes a molybdate transporter instead of a sulfate transporter (Tomatsu et al., 2007). Correspondingly, PtaSULTR5;1 and PtaSULTR5;2 exhibited the farthest phylogenetical distance to all other sequences of this gene family (Table 3.3) and did not reveal the STAS domain. Therefore, both PtaSULTR5 are supposed to fulfil other functions possibly the transport of molybdenum.
The STAS domain is an approximately 130 amino acid long, conserved element at the carboxyl-terminal end of the predicted sulfate transporter proteins that extends into the cell cytoplasm. This sequence shares significant similarity with bacterial anti-sigma factor antagonists such as SpoIIAA of Bacillus subtilis (Aravind and Koonin, 2000). The bacterial SpoIIAA protein is a key component of the regulatory network involved in the induction of sporulation in response to nutrient deficiency. Possibly this element is somehow involved in the regulation of the nutrient status in plants, but such a function has not been shown until now for the STAS domain. The predicted α-helices and β- strands in the STAS region are conserved within 16 poplar sulfate transporters and agree with the structural analysis available for Arabidopsis thaliana and Stylosanthes hamata (Rouached et al., 2005). This particular domain is necessary for a correct localization in the membrane and influences the kinetic characteristics of the AtSULTR1;1 and AtSULTR1;2 sulfate transporters from Arabidopsis thaliana (Shibagaki and Grossman, 2004, 2006; Rouached et al., 2005). Because the 16 putative PtaSULTR sequences from group 1 to 4 reveal all the characteristics of sulfate transporters it is concluded that these poplar proteins feature comparable functions.
The sulfate transport function was investigated for PtaSULTR1;1 and PtaSULTR3;3a by heterologous expression in Xenopus oocytes and electrophysiological two- electrode voltage clamp technique according to Carpaneto et al. (2005) with different sulfate concentrations in the bathing solution. Because it is assumed that sulfate transport is a symport mechanism and one sulfate molecule is transported together with three protons, it was expected that sulfate uptake into the Xenopus cell would result in an inward (negative) current. However, such a negative current was not measured. One reason could be that sulfate transport is, contrary than assumed, an electrically neutral symport with 2 protons or an exchange mechanism with 2 HCO- or
122 Discussion chloride ions which was demonstrated in rabbit membrane vesicles for human sulfate transporters (Kuo and Aronson, 1988; Hästbacka et al., 1994; Haila et al., 2001). Then, sulfate transport across the membrane is not detectable with the two-electrode voltage clamp method. Due to this result further investigations were done by heterologous expression of PtaSULTR1;1 and PtaSULTR3;3a in the yeast mutant CP 154-7A lacking the function of both own sulfate transporter genes (Cherest et al., 1997). This yeast mutant can only grow with an alternative sulfur source like cysteine or methionine. It has been shown that many plant sulfate transporters can replace the yeast protein function which results in the growth of the yeast mutant with sulfate as solely sulfur source (Smith et al., 1995; Takahashi et al., 2000; Howarth et al., 2003; Kataoka et al., 2004b; Krusell et al., 2005; Nocito et al., 2006). This was not achieved for the poplar proteins investigated. One reason could be that the protein sequences of PtaSULTR1;1 and PtaSULTR3;3a start more upstream in the DNA transcript because the start codon (ATG) for translation was only theoretically determined with the TIS Miner program (Liu et al., 2004) and comparison with the orthologs from Arabidopsis thaliana. This case would result in a too short protein sequence which could be functionally uncapable. Also post-translational folding and other factors like phosphorylation or methylation processes could be the reason for non-functional proteins because the translational apparatus of both an eucaryal organism like yeast and an animal like Xenopus can fail to perform correct translation of a plant protein. There are several reasons for the failed results of the functional analyses which possibly reflect differences between trees and herbaceous plants and therefore require further extensive investigations.
4.2 Tissue specific sulfate transporter gene expression
In many plants like Arabidopsis thaliana, Stylosanthes hamata, Zea mays, Lycopersicon esculentum and Hordeum vulgare members of group 1 sulfate transporters were verified for sulfate uptake into roots (Smith et al., 1997; Takahashi et al., 2000; Yoshimoto et al., 2002; Howarth et al., 2003; Hopkins et al., 2005; Nocito et al., 2006). The exclusive expression of the poplar PtaSULTR1;2 sequence in fine roots supports the view of a sulfate uptake function from the soil for this protein. The three sequences of Populus tremula x P. alba, PtaSULTR2;1a, PtaSULTR2;1b and PtaSULTR2;2 belong phylogenetically to group 2, which contains low affinity 123 Discussion sulfate transporters (Takahashi et al., 2000). Both PtaSULTR2;1 genes are comparably expressed in wood of stem and roots while PtaSULTR2;2 was mainly localized in leaves and stem wood. The two members of Arabidopsis thaliana AtSULTR2;1 and AtSULTR2;2 are localized within the vascular system (Takahashi et al., 2000). Sulfate transporter AtSULTR2;1 is specifically expressed within xylem parenchyma cells of roots and leaves as well as in the phloem of leaves, while sulfate transporter AtSULTR2;2 transcripts are localized in phloem cells of roots and leaf vascular bundle sheath cells (Takahashi et al., 2000). In Arabidopsis thaliana group 2 sulfate transporters are supposed to be responsible for the uptake of sulfate from the apoplastic solution within the vascular bundle and are possibly involved in optimizing long distance root-to shoot transport (Takahashi et al., 2000). This function is indicated by an apparent movement of sulfate from roots to leaves with increase of the mRNA level of AtSULTR2;1 in roots by selenate treatment. Selenate cause sulfate deficiency because it compete for the uptake into root cells. In leaves AtSULTR2;2 was expressed in bundle sheath cells surrounding the vascular tissue of leaf veins. This suggests a role in the uptake of sulfate released from xylem vessels for transfer to the primary sites of assimilation in leaf pallisade and mesophyll cells. The functions could be similar for poplar due to similar transcript localization. Both PtaSULTR2;1 genes are expressed mainly in the wood of roots and stem where retrieval of sulfate into xylem parenchyma cells, which leaked into intercellular space, probably takes place. However, PtaSULTR2;2 is expressed highly in leaves and stem wood where sulfate uptake released from xylem vessels can be assumed. Also ShST3 (X82454) from Stylosanthes hamata which clustered into group 2 is expressed in leaf tissue (Smith et al.,1995) and exhibits 73% identity to PtaSULTR2;2. BSULTR2;1 from Brassica oleracea is expressed in stem and leaves while BSULTR2;2 is localized in roots (Smith et al., 1995; Buchner et al., 2004). All these results from different plant species, which are supplemented by the results from poplar in the present study, show comparable expression patterns.
The biggest number of 7 putative SULTR transcripts was abundant in the wood, which includes the xylem. Therefore, these proteins are possibly involved in nutrient distribution along the stem from the roots to the shoot and in the allocation into storage tissues. The high quantity of wood specific localized sulfate transporters in poplar in comparison to herbaceous plants, indicate a tree specific adaptation.
124 Discussion
Members from both, group 2 and 3 sulfate transporters, exhibit this wood specific expression indicating that sequence similarities are not obligatory for the same transcriptional localization. Interestingly, PtaSULTR3;5 was the only sulfate transporter exclusively expressed in root wood. This result is consistent with results from two other plant species. LjSST1 from Lotus japonicus was found within nodules of plants performing nitrogen fixation by bacterial symbiosis with rhizobia (Krusell et al., 2005). This sulfate transporter is expressed in a nodule specific manner, which means it is exclusively located in the symbiosome membrane of Lotus nodules (Krusell et al., 2005). LjSST1 is absent under none symbiotic growth conditions. However, AtSULTR3;5 from Arabidopsis thaliana is transcriptionally co-localized together with AtSULTR2;1 in vascular xylem cells of the root. AtSULTR3;5 reinforces the sulfate uptake capacity of AtSULTR2;1 from the apoplastic space into the xylem parenchyma of roots under sulfur limited conditions. Thus, this transporter may improve the root to shoot transport (Kataoka et al., 2004b). Both functions coincide with the specific localization in root wood which is not contradictory but indicate variations between species due to environmental adaptation. The exclusive expression of PtaSULTR3;5 in root wood without rhizobia interaction supports the idea of an uptake function in xylem parenchyma cells as described for AtSULTR3;5 in Arabidopsis thaliana.
The remaining group 3 sulfate transporters are scarcely investigated, but several observations support the assumption that these transporters are more variable. Expression analyses of the whole group 3 SULTR members in Brassica oleracea show some genes with transcript accumulation specific in stem, some in roots and only one BSULTR3;3 in leaves (Buchner et al., 2004). These results are consistent with the presented results from Populus tremula x P. alba where only PtaSULTR3;3b was exclusively expressed in leaves and the apex. PtaSULTR3;1b was similarly expressed in leaves and the apex, but additionally in wood. Three investigated Arabidopsis thaliana genes (AtSULTR3;1, AtSULTR3;2, AtSULTR3;3) are expressed in leaves but analyses in the stem have not been reported. These results allow no further interpretation of the respective sulfate transporter expression in poplar regarding specific functions. It can just be summarized that compared to wood tissue only a few SULTR sequences were detected in the leaf lamina of poplar trees, but the functional significance of these sulfate transporters has to be further investigated.
125 Discussion
The function of sulfate transporters which cluster in group 4 is much clearer because several investigations were done in Arabidopsis thaliana. The two SULTR4 protein sequences from poplar are closest related to AtSULTR4;1 and AtSULTR4;2 from Arabidopsis thaliana, which are localized in the tonoplast membrane predominantly in xylem parenchyma cells of roots and hypocotyls. Both transporters are facilitating the sulfate efflux from vacuoles (Kataoka et al., 2004a). Because vacuoles are present in nearly all cells from all tissues, the ubiquitous expression in numerous tissues of both poplar group 4 sulfate transporters supports the assumed localization within the vacuolar membrane. Both SULTR4 proteins from poplar revealed a chloroplastic transit peptide sequence that was also found for the Arabidopsis thaliana transporters from group 4 (Takahashi et al., 1999). Still, these sulfate transporters are located within the tonoplast membrane and function as efflux transporters (Kataoka et al., 2004a). Therefore, it can be assumed that also PtaSULTR4;1 and PtaSULTR4;2 facilitate sulfate efflux from vacuoles. In summary, the highly tissue specific expression pattern of 16 sulfate transporter sequences indicates a transcriptional regulation and organization of sulfate distribution within poplar trees. Regarding sulfate transport in the phloem it was most interesting that the two sulfate transporters PtaSULTR1;1 and PtaSULTR3;3a were specifically expressed in the bark and leaf veins. Due to this result these two sequences were selected for more precise localization analyses.
4.3 Cellular transcript localization of two sulfate transporters in minor leaf veins and the lamina
The two sulfate transporter sequences PtaSULTR1;1 and PtaSULTR3;3a from Populus tremula x P. alba exhibiting transcript accumulation in bark and leaf vein tissue (Figure 3.3b) were analyzed by in situ hybridization. This method was applied to gain more detailed insight into the organization of sulfate allocation through the phloem vascular system at the cellular level. During vegetative growth sulfate is transported into poplar leaves via the xylem system and sulfate is further removed out of mature leaves through the phloem (Hartmann et al., 2000). The phloem
126 Discussion connects source leaves with sink organs. At full nutrient supply, mature poplar leaves are mainly sources for carbohydrates (Dickson, 1991) and therefore, the phloem of minor leaf veins may function as collection phloem (van Bel, 2003) also for nutrients such as sulfate. PtaSULTR1;1 transcript was abundant in minor leaf veins of the leaf lamina where mRNA localization was observed in phloem cells. This indicates that phloem loading with sulfate might be carried out by PtaSULTR1,1 for transport to different sink organs. In contrast PtaSULTR3;3a was not detectable in minor leaf veins. Apoplastic phloem loading of carbohydrates is mediated by sucrose transporters (Hellmann et al., 2000; Patrick et al., 2001; Carpaneto et al., 2005; Schmitt et al., 2008). Two polyol transporters are discussed to be involved in phloem loading with sorbitol in Plantago major leaves (Ramsperger-Gleixner et al., 2004) and sucrose transporters have been identified in phloem cells of source leaves of Plantago major, potato and Arabidopsis thaliana (Stadler et al., 1995; Stadler and Sauer 1996; Kühn et al., 1997; Ramsperger-Gleixner et al., 2004). However, also symplastic sucrose loading has been shown for cucurbits where sugars enter the sieve element-companion cell (SE- CC) complex directly via plasmodesmata (Turgeon et al., 1993; Turgeon, 1996). Also many deciduous trees are discussed to be symplastic loaders due to their high plasmodesmata frequency between companion cells and surrounding parenchyma cells (Gamalei, 1989). Still apoplastic phloem loading is assumed because on this way sulfate can accumulate in companion cells and it is superior for regulation. Such a regulation may be required for a controlled sulfate allocation from source to sink.
Previously, it has been shown that the sulfate transporter AtSULTR1;3 from Arabidopsis thaliana is essential for sulfate loading into the phloem of cotyledons to supply the young growing shoot with sulfate during early development. It is assumed that sulfate transporters are dependent (driven) on the proton motive force and function by a proton-symport mechanism. Because the H+ concentration at the plasma membrane is generally higher outside the cell relative to the cytosol (Lass and Ullrich-Eberius, 1984), the specific expression of PtaSULTR1;1 within phloem cells adverts to an uptake function from the apoplast into the cytoplasm. EM- immunocytochemistry revealed the predominant cellular location of proton pumps in companion cells of the collection phloem of Vicia faba (Bouché-Pillon et al., 1994). Plasma membrane H+-ATPase 4 (PM4) was also notably expressed in companion
127 Discussion cells of minor veins of source leaves in Nicotiana plumbaginifolia (Moriau et al. 1999). Suppression of PM4 led to an increase in the leaf sugar content and a decrease of sucrose in phloem exudates of Nicotiana plumbaginifolia (Zhao et al., 2000). Together, these results indicate the involvement of H+-ATPase in apoplastic phloem loading for sucrose by proton / sucrose symporters and support the idea of an apoplastic loading also for sulfate by proton / sulfate co-transport. Due to the specific expression of PtaSULTR1;1 in the phloem of minor leaf veins, this transporter is supposed to be involved in sulfate uptake into the collection phloem.
Beside the described localization in the phloem, PtaSULTR1;1 but again not PtaSULTR3;3a transcript was located in mesophyll cells of the leaf lamina in particular in the upper palisade parenchyma. This may indicate the contribution of PtaSULTR1;1 protein in sulfate uptake into these cells. In green beech leaves sulfur is mainly located in the cytoplasm of palisade parenchyma cells, bundle sheath cells and in sieve elements (Eschrich et al., 1988). The palisade parenchyma cells are photosynthetically active and provide protons via ferredoxin for the sulfate reduction process (Leustek and Saito, 1999). Therefore, high amounts of sulfate are supposed to be transported into these cells which might be the requirement of PtaSULTR1;1. The slight expression of PtaSULTR1;1 in the epidermis and spongy mesophyll cells also indicates relevance of this transporter for sulfate uptake into various cell types of the leaf but with less priority.
4.4 Sulfate transporter expression in the transport phloem and in bark
Transcript accumulation of PtaSULTR1;1 and PtaSULTR3;3a was investigated in major leaf veins, along the stem and in roots with secondary growth to get an overview of the cellular expression at different functional zones of the phloem. Major leaf veins, the stem and roots with secondary growth are organs where sulfate is mainly transported over long distances but also released and retrieved. In major leaf veins and along the stem highest PtaSULTR1;1 transcript signals were detected in companion cells while PtaSULTR3;3a was consistently expressed in phloem cells. In the transport phloem the sieve element-companion cell complex is symplastically
128 Discussion isolated from the surrounding parenchyma cells in Salix alba and Ricinus (van Bel and Kempers, 1991). In Populus trichocarpa, companion cells but not sieve elements possess plasmamembrane H+-ATPases (Arend et al., 2002). This coincides with the specific expression of PtaSULTR1;1 in companion cells. Thus, PtaSULTR1;1 may be involved in retrieval of sulfate leaked out from the phloem through companion cells. A comparable function for return into companion cells has been discussed for sucrose transporters that are expressed along the transport phloem (Stadler and Sauer 1996; Williams et al., 2000, van Bel, 2003). Therefore, both sulfate transporters are supposed to be responsible for retrieval of apoplastic sulfate into companion cells. Another function of sulfate transporters in companion cells could be the transport out of these cells into the sieve elements which is less likely because companion cells and sieve elements built a complex with plasmodesmata connections and sulfate can flow passively in the direction of the concentration gradient (van Bel et al., 1996). Phloem unloading from companion cells into the apoplastic space via PtaSULTR1;1 and PtaSULTR3;3a seems also rather unrealistic because sulfate transporters depend on the proton motive force (Lass and Ullrich-Eberius, 1984; Hawkesford et al., 1993). However, mature leaves export 35S-sulfate that is taken up via flap-feeding mainly in basipetal direction to the poplar stem where it is stored (Hartmann et al., 2000). Sulfate must therefore be transported out of the sieve element-companion cell complex into the apoplastic space. This unloading may be mediated by voltage dependent anion channels (Frachisse et al., 1999, Roberts 2006), because plasma membrane anion channels of hypocotyl cells in Arabidopsis thaliana can generate large currents with sulfate. In plants, anion channels regulate efflux from the cytoplasm into the extracellular space, which is driven by the anion-gradient across the plasma membrane and the negative membrane potential of plant cells (Czempinski et al., 1999). In summary, it is most likely that sulfate uptake from the outside into the sieve element-companion cell complex is carried out by PtaSULTR1;1 and PtaSULTR3;3a. Still, it can not be completely excluded that PtaSULTR1;1 and PtaSULTR3;3a are involved in phloem unloading because it has been shown that ZmSUT1 mediates sucrose efflux depending on the sucrose gradient and the membrane potential (Carpaneto et al., 2005). During heterologous ZmSUT1 expression in Xenopus oocytes sucrose efflux was measured when the sucrose concentration was higher in
129 Discussion the cytosol than extra cellular. Regarding these results the sucrose transport direction is invertible but if this is the case under in vivo conditions in plants is questionable and was not shown for sulfate transporters until now. Nevertheless, accumulation of sulfur in companion cells of the stem has been observed after feeding 35S-sulfate via flap-feeding to mature poplar leaves (Herschbach, unpublished results). This accumulation can probably be achieved by active transport via PtaSULTR1;1 and PtaSULTR3;3a from sieve elements or by anion channels. However, passive transport through plasmodesmata depends on high sulfate concentrations in sieve elements to constitute a concentration gradient that triggers sulfate uptake into companion cells. This contradicts the observed 35S accumulation in companion cells. Another accumulation process could be a direct transfer of sulfate into vacuoles of companion cells which would reduce the cytosolic sulfate concentration. In plants cytoplasmic concentrations of sulfate are kept relatively constant and the surplus is stored in the vacuoles (Hawkesford, 2000). On this way, an incident sulfate gradient from sieve elements into companion cells is probably induced. For this purpose, it is possible but not obligatory that PtaSULTR1;1 and PtaSULTR3;3a are involved in the transport of sulfate out of the sieve elements to enforce unloading of sulfate and further its accumulation in companion cells.
Beside the strong expression in companion cells PtaSULTR3;3a was additionally detected in sieve elements. The angiosperm sieve elements are lacking any apparent capacity for transcription or translation (van Bel et al., 2002). Therefore, detection of PtaSULTR3;3a in the sieve tubes is an indication of mRNA leakage out of companion cells for subsequent movement by phloem mass flow. Evidences for plant transcript delivery from companion cells into sieve elements has been claimed for the sucrose carrier SUT1 in potato by in situ hybridization (Kühn et al., 1997), thioredoxin h and actin in rice (Sasaki et al., 1998). Additionally, mRNAs of several proteins, including transporters, have been found in phloem exudates (Doering-Saad et al., 2002, Lough and Lucas, 2006, Omid et al., 2007, Le Hir et al., 2008, Kehr and Buhtz, 2008). 36 ESTs have been identified in the phloem sap of melon plants belonging to transporters and channels including 7 putative sugar transporters (Omid et al., 2007). mRNA translocation has been shown also by grafting experiments in tomato (Kim et al., 2001), cucumber (Ruiz-Medrano et al., 1999) and potato (Banerjee et al., 2006). Apparently, plants have evolved a system to transport mRNA for signalling purposes
130 Discussion at the whole plant level and thereby coordinating developmental processes (Laugh and Lucas, 2006). Transportation of PtaSULTR3;3a mRNA within the phloem is in principle possible from the present results, but has to be demonstrated in further experiments. Analyses of different stem sections exhibited that the expression pattern of both sulfate transporters was independent from the developmental state of the stem. Different from the stem, roots with secondary growth exhibited strong PtaSULTR1;1 transcript accumulation in companion cells but no expression of PtaSULTR3;3a. These roots fulfil storage functions like the stem and therefore, contribution of PtaSULTR1;1 in phloem unloading seems suitable. Still, as discussed above, both PtaSULTR proteins seem to be involved in re-loading of sulfate into the phloem but, with varying responsibilities in different tree sections. It seems possible that only PtaSULTR1;1 transports sulfate from the apoplastic space into companion cells of roots with secondary growth. While retrieval by PtaSULTR1;1 and PtaSULTR3;3a may be important along the entire trunk.
The further transport of sulfate from the phloem into storage tissues of the wood may be released via its uptake into ray initials. In principle, en exchange between phloem and xylem can be assumed for poplar trees (Hartmann et al., 2000). Sulfur from just expanded poplar leaves is exported mainly into apical tree parts and only to a minor extent into basipetal direction (Hartmann et al., 2000). Because 35S-sulfate was detected only apical to the fed leaf, phloem to xylem exchange must be assumed for 35S-sulfate (Hartmann et al., 2000). PtaSULTR1;1 mRNA was detected in the vascular cambium and expression was strongest in ray initials which was not found for PtaSULTR3;3a. An antibody against maize plasmamembrane H+-ATPase has been used to detect ATPases in transversal twig sections from Populus trichocarpa (Arend et al., 2002). In spring during cambial growth strong immunoreactivity against H+-ATPase was observed in cambial cells and expanding xylem cells. Thus, PtaSULTR1;1 seems responsible for the uptake of sulfate in growing and dividing tissue in the cambium. Previously it was found that 35S-sulfate which was fed to beech leaves was mainly translocated into ray, medullary sheath and pith cells (Ziegler, 1965). The enrichment of PtaSULTR1;1 in ray initials possibly indicates the way of sulfate uptake into ray cells while the further transport within the ray pith may take place by cell to cell transport via plasmodesmata. Therefore, it can be assumed
131 Discussion that the exchange from phloem to xylem is carried out by the uptake of sulfate into ray initials with a subsequent symplastic transport in ray cells and a further transport into xylem vessels.
Localization of PtaSULTR1;1 was also observed in the epidermis of the young stem and in the cell layer adjacent the epidermis, the periderm, in older stem as well as in roots with secondary growth. It is feasible that these cells are supplied with sulfate from the apoplastic solution by PtaSULTR1;1. The nutrient supply could be important especially for the cell layer adjacent the epidermis during development because in older stem parts the periderm builds the cork cambium (phellogen) where cell division and cork growth takes place. During cell development many sulfur containing compounds like proteins and sulfolipides have to be synthesized which requires high sulfate content and uptake in these cells. Compared to the other investigated stem sections only basal stem sections exhibited PtaSULTR1;1 transcript accumulation in cells of the cortex, which are localized between the sclerenchymatous fibers and extend from the phloem up to the periderm. Probably these cells built a connection between the vascular system and the outer cell layers for supply. The expression in the epidermis may be associated with the protection against environmental influences like pathogen defence and detoxification. Sulfur rich compounds like glutathione have been discussed for protection against oxidative damage and pathogen defence (Rausch and Wachter, 2005). This could be an explanation for sulfate uptake into outer cell layers for further assimilation and synthesis of defence compounds. In this context, high glutathione synthesis and high expression of genes involved in cysteine and glutathione biosynthesis have been observed in trichomes of Arabidopsis thaliana (Gutiérrez-Alcalá et al., 2000). Strong transcript accumulation of PtaSULTR1;1 in lenticels supports this assumption and allows further speculations about sulfate uptake into these bark localized areas. Accordingly, AtSULTR1;1 in Arabidopsis thaliana was specifically expressed in hydathode cells of cotyledons (Takahashi et al., 2000). A lenticel is an area within the cortex with big intercellular spaces to allow gas exchange between the internal tissue and the atmosphere while hydathodes deliver water from leaves. Therefore, both tissues are in contact to the aerial space and share similar premises like epidermis cells where PtaSULTR1;1 also accumulated.
132 Discussion
4.5 Sulfate transporter expression in wood
PtaSULTR1;1 and PtaSULTR3;3a expression is not only obvious in bark tissues. Both transporters are present in pith ray cells predominantly adjacent to xylem vessels and in parenchyma cells associated to the primary xylem located at the border between the pith and wood tissue. Because sulfate was found as the predominant sulfur compound in xylem sap of deciduous trees (Herschbach and Rennenberg, 2001) it can be assumed that these sulfate transporters are involved in the uptake of sulfate into storage cells of the wood from xylem vessels. In walnut trees a putative sucrose transporter JrSUT1 has been detected in vessel-associated parenchyma cells (Decourteix et al., 2006). Immunolocalization studies showed that JrSUT1 was co-localized with plasma membrane H+-ATPase (JrAHA) in these cells. Because the presence of the proton motive force is a prerequisite for sucrose and also for sulfate uptake the presence of ATPase in ray cells adjacent to xylem vessels in poplar (Arend et al., 2002) supports the assumption that PtaSULTR1;1 and PtaSULTR3;3a could be involved in sulfate uptake from xylem vessels into the symplast of pith ray cells for storage. Uptake of sulfate from the xylem into xylem parenchyma cells of leaf veins by both transporters, PtaSULTR1;1 and PtaSULTR3;3a, seems also relevant. The observed expression pattern in the xylem was independent from the developmental stage of stem sections. Therefore, sulfate is supposed to be taken up from the xylem along the whole stem. Only in old basal stem sections PtaSULTR1;1 mRNA accumulation close to the primary xylem was weak compared to the staining in other cells, while transporter transcript accumulation was substantial in young and mature stem sections. This observation indicates a decreasing importance with increasing stem age to supply older wood tissues and pith cells with sulfate. It may be speculated that the sulfate pool of older storage tissues of the wood is already filled. The only accumulation of PtaSULTR1;1 transcript in xylem parenchyma cells surrounding xylem vessels in roots was an obvious difference between PtaSULTR1;1 and PtaSULTR3;3a expression. Uptake of sulfate into these cells is concluded to be exclusively carried out by PtaSULTR1;1.
133 Discussion
4.6 Sulfate transporter expression in sink tissues
To get an overview of the cellular expression at the whole plant level, transcript accumulation of PtaSULTR1;1 and PtaSULTR3;3a was investigated in fine roots and the shoot apex as expected sink organs regarding the dependence on sugar supply from source organs. It has been shown that sugar transporters are expressed in sink organs like tobacco pollen (Nicotiana tabacum, Lemoine et al. 1999), broad bean seeds (Vicia faba, Weber et al., 1997) and plantain young ovules (Plantago major, Gahrtz et al., 1996). In Populus tremula x P. alba sulfate is transported to the shoot apex and to fine roots (Hartmann et al., 2000) which indicate sulfate transport to expected sink organs. PtaSULTR1,1 transcript is highly abundant in the shoot apical meristem (SAM), in leaf primordia, leaf margin meristems (LMM) and provascular strands. It is discussed that within typical sink tissues companion cells are very small or missing and nutrients and metabolites may leak directly from sieve elements into the apoplastic space. From the apoplast nutrients have to be taken up by the surrounding cells. Therefore, PtaSULTR1,1 could be important for the uptake of sulfate into dividing and growing cells of the shoot for further reduction and assimilation. Symplastic unloading via plasmodesmata from sieve elements directly to sink cells is assumed for various plants and different tissues (van Bel, 2003). Such a mechanism would be opposed to the involvement of PtaSULTR1,1. New developing leaves of poplar trees are not dependent on the supply of reduced sulfur from mature leaves, because all enzymes of the assimilatory sulfate reduction are highly active in young developing leaves (Hartmann et al., 2000). Even the apex of poplar trees is capable to reduce and assimilate sulfate (Herschbach, 2003). These findings support the uptake function of PtaSULTR1;1 in sink tissues, but the extent of sulfate assimilation in heterothrophic and dividing tissues is still unknown (Hartmann et al., 2000). High PtaSULTR1;1 abundance in the root apical meristem of developing side roots with strongest intensity close to xylem vessels again supports its relevance for sulfate uptake into dividing and growing cells. Very weak signals were detected also for PtaSULTR3;3a in provascular strands of the shoot apex but no signal was detected in fine roots or any other sink tissue analyzed. PtaSULTR1;1 but not PtaSULTR3;3a was abundant in the protophloem of fine roots. Sulfate reduction probably takes 134 Discussion place in fine roots because the enzymes involved in assimilatory sulfate reduction are also present in roots. In maize strongest APS reductase activity, the key enzyme of the sulfate assimilation pathway (Vauclare et al., 2002), has been detected in root tips (Brunold, 1990; Kopriva et al., 2001; Kopriva and Koprivova, 2004). These results support the idea that sulfate is loaded into the phloem by PtaSULTR1;1 for further allocation to the root tip where it can be taken up and used after reduction for protein synthesis.
4.7 Influence of sulfate deprivation on the sulfate transporter transcript level
In higher plants sulfate transport is intensively regulated. This regulation is considered to be mediated by negative feedback control, in which sulfate or products of sulfate assimilation such as cysteine, methionine or glutathione serve as effector molecules (Herschbach and Rennenberg, 1991, 1994; Lappartient, 1999). To compensate for decreased availability of sulfate, the most significant sulfur form internally mobilized is supposed to be sulfate if it is present in the particular tissue (Hawkesford, 2000). Bark and wood are preferential storage tissues for sulfur (Herschbach and Rennenberg, 1996) and sulfate can also be transported out of mature leaf mesophyll cells to supply sink tissues in poplar trees with sulfate (Hartmann et al., 2000). If sulfate from the vacuole of leaves is mobilized, it must be loaded into the phloem and / or xylem for further transport to expected sink organs. However, it has been shown that the endogenous sulfur pool in some herbaceous plants is relatively immobile, but without distinguishing between reduced and oxidized sulfur (Larsson et al., 1991; Sunarpi and Anderson, 1996; Eriksen et al., 2001). Sulfate efflux from the vacuole of leaf mesophyll and root cells of Macroptilium atropurpureum was slow and did not significantly contribute to sulfur efflux from cells during sulfur deficiency (Bell et al., 1994, 1995). Therefore, it is questionable if sulfate is mobilized from vacuoles during sulfur deficiency in appreciable amounts. The up-regulation of the phloem specific transporter PtaSULTR1;1 in both, mature leaf veins and bark tissues, supports the expectation that loading of sulfate into the phloem of source tissues during sulfate deprivation increased. This result corresponds to decreasing sulfate contents in both bark and leaf vein tissues.
135 Discussion
Accordingly, in Arabidopsis thaliana sulfur deprivation caused the accumulation of AtSULTR1;3 mRNA both in leaves and roots (Yoshimoto et al., 2003), which gives evidence that sulfate loading into the phloem may increase and is regulated at the gene expression level due to the sulfur status in the plant. In contrast, PtaSULTR3;3a was not clearly affected on the transcriptional level by sulfur deficiency which indicates differences in the regulation of these two phloem specific sulfate transporters in poplar. The up-regulation of both putative vacuolar sulfate efflux transporters in the bark and additionally the substantial up-regulation of PtaSULTR4;2 in stem wood supports the hypothesis that sulfate is mobilized form vacuoles of storage tissues of the trunk like bark and wood. Contradictory to the previous described immobility of the vacuolar sulfate pool in Macroptilium atropurpureum (Bell et al., 1994, 1995), the efflux function of AtSULTR4;2 in Arabidopsis thaliana promotes the transport of sulfate from vacuoles into the cytoplasm particularly in the xylem parenchyma cells under conditions of low sulfur supply (Kataoka et al., 2004a). This movement is supposed to optimize the flux of sulfate towards the xylem vessels and confirms the assumption, that sulfate is mobilized from vacuoles. The increase of PtaSULTR4;1 and PtaSULTR4;2 transcript abundance in poplar coincides with diminished sulfate concentrations in bark. The parallel increase of PtaSULTR1;1 mRNA support the idea that sulfate is mobilized from vacuoles of storage tissues, and is subsequently taken up via PtaSULTR1;1 into the companion cell-sieve element complex for further transport into sink tissues. A parallel increase of PtaSULTR4 and PtaSULTR1;1 mRNA was found in bark after 26 days of sulfate deficiency, while the expression of both sulfate transporters from group 4 was not affected in leaf veins. Probably the bark is more important for sulfate storage and remobilization than mature leaves in trees or sulfate mobilization takes place from the cytoplasm of mesophyll cells of the leaf lamina. These results in combination with the previous results mentioned from herbaceous plants indicate differences in the vacuolar sulfate mobilization between different tissues and between different plant species. It can be discussed that trees utilize the stem as nutrient reservoir while some herbaceous plants are supposed to either not mobilize vacuolar sulfate or utilize also the stele preferentially then roots and leaves for storage. Because living cells of the wood are involved in sulfur storage (Herschbach and Rennenberg, 1996), sulfate mobilization from the wood may also be relevant during
136 Discussion sulfur starvation. The strongly decreased sulfate content in wood tissue upon sulfur deprivation reflects either a decreased support from the leaves by phloem transport or an insufficient sulfate supply via the transpiration stream in the xylem. To address this question sulfate transporter expression in stem wood was investigated. Three wood specific sulfate transporters SULTR2;1a, SULTR2;1b and SULTR3;2b seemed to be down-regulated due to sulfur limitation. This observation led to the suggestion that in poplar trees transport out of or into the xylem is diminished under sulfate deficiency. If these transporters are involved in unloading of sulfate from the xylem vessels, decreased transcript accumulation would make sense because decreased uptake into storage tissue of the wood could improve the sulfate supply of growing tissues such as young developing leaves or the apex. However, most sulfate transporters expressed in the wood which cluster into group 3 did not exhibit a clear influence in response to sulfur starvation. Comparable results were found for Brassica oleracea where the whole group 3 did not show any effects in response to the sulfur depletion treatment in stem, leaf and root (Buchner et al., 2004). The sulfate transporter PtaSULTR2;2 transcript which was abundant in all shoot tissues revealed increasing transcript accumulation only in bark, while in leaf vein and wood no clear effect was found. This indicates different regulation mechanisms of the same sulfate transporter in different tissues. Therefore, further investigations are required to identify the cellular localization and potential function of this sulfate transporter.
In many plants the first response to sulfur deficiency is the increase of mRNA levels of specific sulfate transporters in roots which results in an increased sulfate uptake capacity (Takahashi et al., 2000). Thus, the fastest regulated genes under sulfate deprivation seem to be sulfate uptake transporters in roots (Smith et al., 1997; Takahashi et al., 1997). Also in Populus tremula x P. alba, the expression level of sulfate transporter PtaSULTR1;2 probably responsible for sulfate uptake by the roots starts to increase fastest, which was after 14 days of sulfate deprivation (Honsel et al., 2009). This increased transcript abundance was accompanied by a reduced sulfate content inside the roots. For Arabidopsis thaliana and Nicotiana tabacum beside uptake of sulfate by the roots also its distribution to the shoot was significantly increased by sulfur deprivation (Herschbach and Rennenberg, 1991; Kataoka 2004b). AtSULTR2;1 mRNA accumulation was increased, particularly in roots and
137 Discussion
AtSULTR2;2 increased in leaves of sulfur starved plants (Takahashi et al., 1999, 2000). Because AtSULTR2;1 is localized in xylem parenchyma cells of roots and AtSULTR2;2 in bundle sheath cells of leaves, their increased expression supports the idea that these transporters are involved in the uptake of sulfate from the xylem into the surrounding cells. Therefore, also the distribution of sulfate through the vascular system is regulated in response to sulfur starvation in herbaceous plants (Herschbach and Rennenberg, 1991; Shibagaki et al., 2003; Kataoka et al., 2004b). However, in contrast to the sulfate uptake transporters of the root, the low affinity sulfate transporters possibly responsible for xylem unloading respond, if at all, more slowly to changes in the external sulfate supply (Kataoka et al., 2004b). Comparing these results from Arabidopsis thaliana with the results from Populus tremula x P. alba it is noticeable that AtSULTR2;1 is up-regulated in roots while both PtaSULTR2;1a and PtaSULTR2;1b mRNA levels in poplar seemed to decrease in wood due to sulfur deprivation. However, sulfate transporter expression was not analysed in Arabidopsis thaliana stems. Therefore, the present result either reflects different regulations between tissues or different regulation mechanisms between poplar and Arabidopsis thaliana. It also has to be considered that during the presented experiments with poplar trees low sulfate amounts were still available to the roots until day 17 in the sand used as soil substrate. Therefore, direct comparison between results from the experiments with Arabidopsis thaliana performed in solutions with zero sulfate and the presented experiments with poplar on sand substrate has to be taken with care. Together the presented results support the idea that sulfate remobilization is a secondary response to prolonged sulfur deficiency and is more significant from bark and wood compared to mobilization of the vacuolar sulfate from mature leaves. The first reaction of plants to sulfate deficiency is the up-regulation of root uptake sulfate transporters to increase the internal sulfate amount. It seems that only if this reaction is insufficient to meet the plants sulfur demand, sulfate is mobilized from vacuoles of storage tissues.
4.8 Seasonal variation in sulfate transporter expression
Storage of starch and proteins is well established in the seasonal growth pattern during late summer and autumn in deciduous trees (Zimmermann, 1974; Bonicel et 138 Discussion al., 1987; Dickson, 1991; Sauter and van Cleve, 1994; Cooke and Weih, 2005). Studies with several tree species showed that storage proteins synthesized during autumn are localized in the vacuoles of pith rays and phloem parenchyma cells (Sauter and Wellenkamp, 1988; Sauter et al., 1988). Also sulfur compounds are stored in trees during winter. When 35S-sulfate was fed to a mature beech leaf in late summer it was allocated through the phloem to basipetal sections of the trunk to storage tissues (Herschbach and Rennenberg, 1996). Export of sulfate and of reduced sulfur from mature leaves and further storage in bark and wood tissues has been shown for several tree species (Herschbach and Rennenberg 1995, 1996; Hartmann et al., 2000). Sulfate accumulation in the bark of field-grown Populus tremula x alba trees started in autumn when the mRNA of both phloem specific sulfate transporters and both putative vacuolar efflux transporters decreased during the beginning of October. This indicates reduced phloem reloading and sulfate efflux from the vacuoles in bark. During that time the growing season switched into the dormant period. Parallel with a steep increase of the sulfate content in bark the PtaSULTR1;1 transcript accumulation increased in senescent leaves. Because PtaSULTR1;1 transcript is abundant in the collection phloem of minor leaf veins, the enhanced transcript accumulation in leaves during abscission may be interpreted as an increase of phloem loading with sulfate. Enhanced phloem loading might implicate sulfate recycling out of the leaves and further transport into storage tissues such as the bark. This result is consistent with the observation that during leaf senescence the sulfur fraction in beech leaves increases in the apoplastic compartments at the expense of the cytoplasmic sulfate of palisade cells, bundle sheath cells and sieve elements (Eschrich et al., 1988). From the apoplastic space sulfate might be taken up via transport proteins into the phloem. The finding that 35S-sulfate fed to a mature beech leaf in late summer was allocated through the phloem to storage tissues in the stem (Herschbach and Rennenberg, 1996) and the increased sulfate content in bark during leaf senescence in poplar confirm the supposed sulfate reloading. Because both PtaSULTR4 transcripts increased also in senescent poplar leaves and due to their putative vacuolar sulfate efflux function, they possibly empty the vacuoles before leaves fall. The observation that ESTs encoding for 5 cysteine proteases were most abundant and significantly enriched in a Populus library from senescent leaves collected in autumn, in comparison to a young mature leaf library (Bhalerao et al.,
139 Discussion
2003) adverts to protein degradation processes. These proteases were apparent orthologs of a class of vacuolar located cysteine proteases (Hayashi et al., 2001). Therefore, reduced sulfur in the form of degraded amino acids is possibly converted into sulfate and further available via vacuolar sulfate efflux transporters. Such degradation processes can be discussed because 35S-cysteine fed to a mature beech leaf in early summer was partially metabolized to sulfate which was further transported by the phloem to storage tissues of the trunk (Herschbach and Rennenberg, 1995). Whether cysteine, methionine or proteins are mobilized and converted to sulfate has to be investigated in further studies. The sulfate, cysteine and protein content of fallen leaves could give important information in this context.
In the further annual course sulfate content was high in the bark during the whole dormant period when PtaSULTR3;3a and PtaSULTR4;2 expression was almost not detectable. It seems that retrieval of sulfate in the transport phloem and vacuolar export comes to rest. The sulfate content reached its maximum during December, while PtaSULTR1;1 and PtaSULTR4;1 expression in the bark revealed significant peak values. During that time lowest temperatures below zero degree Celsius were observed. Under the assumption that PtaSULTR4;1 removes sulfate out of the vacuole and PtaSULTR1;1 transports sulfate into the phloem their transcript increase with a concomitant high sulfate level in bark seems first to be contrary. But some previous investigations indicate compound accumulation and transport events in correlation to cold temperature. In Populus x canadensis glycerol and sucrose contents increase sharply in twig wood during winter (Sauter and van Cleve, 1994). Also an increase of fat was found in the bark of poplar trees during the cold period. Furthermore, the putative xylem sucrose transporter JrSUT1 is up-regulated by freezing–thawing cycles over the autumn and winter period in walnut trees (Juglans regia) (Decourteix et al., 2006). It has been discussed whether the increase in glucose and sucrose during dormancy in the wood of poplar twigs (Sauter and van Cleve, 1994; Sauter et al., 1998) correlates with cool temperature to protect cells from freezing injury. In this context the present results implement that temperatures below zero degree Celsius might be a signal for sulfate redistribution. The induced upregulation of PtaSULTR4;1 possibly removes sulfate out of the vacuole and PtaSULTR1;1 transports sulfate into the phloem. Whether sulfate is involved in frost protection or metabolized to frost protecting compounds has to be investigated in
140 Discussion further studies. It seems that dormancy and associated cold hardiness that has been developed in over-wintering aboveground plant parts require more adaptation and protection than known until now in trees.
Starch and proteins which are stored during late summer and over winter are mobilized during spring when current year flushes are developed (Sauter and Wellenkamp, 1988, 1998; Sauter et al., 1994, 1998; Cooke and Weih, 2005). During bud break also sulfur is mobilized from soluble and insoluble compounds in bark and wood tissues of beech (Herschbach and Rennenberg, 1996). During early spring the increased transcript level in bark of the putative vacuolar sulfate efflux transporter PtaSULTR4;2 coincides with bud swelling and early leaf development. This expression pattern probably indicates sulfate mobilization from the vacuole of bark cells. Simultaneously, PtaSULTR1;1 and PtaSULTR3;3a mRNA accumulation increased during early leaf development in bark and the sulfate content decreased drastically within two weeks. Apparently, both sulfate transporters, PtaSULTR1;1 and PtaSULTR3;3a, are supposed to be responsible for enhanced sulfate loading into the phloem during spring for further allocation to developing sink tissue. Most likely transport within the phloem support swelling buds during spring. In spring during cambial reactivation and growth H+-ATPase protein increased strongly in cambium and phloem cells of Populus trichocarpa (Arend et al., 2002). At the same time also high PtaSULTR1;1 and PtaSULTR3;3a expression was observed. Correlation between the appearance of ATPase protein and both PtaSULTR1;1 and PtaSULTR3;3a increment is probably required because plasmamembrane H+- ATPase generates the proton motive force for sulfate loading into the cell cytoplasm.
In spring sulfate content decreased first in expanding leaves after bud break. This decrease probably results from dilution because of leaf expansion or is caused by high sulfate consumption for assimilation and incorporation into proteins. Simultaneously, PtaSULTR1;1 expression in leaves increased during development and reached a maximum when leaves were fully expanded. Thus, increasing sulfate transport out of leaves might also reduce sulfate content during expansion. When the leaf lamina has the required size to generate an appreciable transpiration stream in the xylem the growing tissues might be increasingly supplied with sulfate from the soil. During this time in April PtaSULTR1;1 transcript decreased rapidly in the bark
141 Discussion and it seems that developing tissues are less dependent on the supply from stored sulfur. In early May simultaneous peak expression of PtaSULTR1;1 and PtaSULTR4;2 mRNA in mature leaves coincides with lowest sulfate contents in leaves. At this developmental stage sulfate export from mature leaves is supposed to be carried out by PtaSULTR1;1. Under the assumption that PtaSULTR4;2 is a vacuolar efflux transporter (Kataoka et al., 2004a), sulfate efflux from the vacuole increases during leaf development and possibly gets relevant in fully expanded leaves. During summer sulfate content increased in mature leaves when PtaSULTR1;1 expression decreased. Accordingly, reduced phloem loading of sulfate enables sulfate accumulation in leaves. Simultaneously, expression of the putative vacuolar sulfate efflux transporter decreased during summer and this enables sulfate accumulation in the vacuole. The expression of the second phloem specific sulfate transporter PtaSULTR3;3a which is obvious in major leaf veins only increased later in the season when expression of PtaSULTR1;1 was again decreased. Therefore, phloem reloading in leaves may be relevant later in the season when storage processes along the stem are possibly of increasing significance.
The expression of the analyzed sulfate transporter genes varied obviously during the season, indicating that they are regulated on the transcriptional level due to environmental factors. In bark tissue the seasonal transcript accumulations of the phloem specific sulfate transporter PtaSULTR3;3a and the putative vacuolar efflux transporter PtaSULTR4;2 are positively correlated. Therefore, the expression of these genes seems similar regulated. Both are negatively correlated with the sulfate content in bark. This inverse correlation between sulfate content and sulfate transporter transcript accumulation supports the assumed vacuolar export function of PtaSULTR4;2 and phloem loading function of PtaSULTR3;3a. The combined function of both transporters results in sulfate flow out of the tissue. A similar transcriptional regulation of both genes is supported by the positive correlation of mRNA accumulation with weather parameters. Beside the influence of temperature the expression patterns are strongest correlated with day length which is a decisive parameter for seasonality (Jansson and Douglas, 2007). The increasingly shorter days in autumn are considered the most important input signals in this context (Jansson and Douglas, 2007). Studies with Populus have shown that once endodormancy has been induced by short day length, extended periods of chilling
142 Discussion temperatures are needed to condition trees to break dormancy (Rhode et al., 2000) and put the tree in a ‘standby’ position, waiting for spring conditions to grow. Increasing temperatures and day lengths correlate with increasing transcript accumulations of PtaSULTR3;3a and PtaSULTR4;2 during spring. The opposite effect is reflected during autumn. Autumn senescence has clear adaptive values, because remobilization of nitrogen, in particular, is important in forest ecosystems where nitrogen is often a limiting factor for plant growth (Jansson and Douglas, 2007) and uptake from the soil is minute in early spring due to low soil temperatures (Millard et al., 1998). Hence, it seems that PtaSULTR3;3a and PtaSULTR4;2 transcript abundance is regulated by seasonality in bark tissue. Furthermore, it can be assumed that temperature and light in the form of day length plays an important role in triggering transcriptional changes.
In contrast, the phloem specific sulfate transporter PtaSULTR1;1 and the putative vacuolar localized sulfate transporter PtaSULTR4;1 which are positively correlated in bark did not show any correlations to the sulfate content or weather conditions. Nevertheless, the transcript abundance increased during bud break and leaf development in spring. One possible reason of disconnecting PtaSULTR1;1 and PtaSULTR4;1 expression in bark from seasonal influences could be a plant internal regulation mechanism in the context of a developmental program. Another reason for regulatory disconnection from regular seasonal influences could be the accounted punctual high transcript abundance of PtaSULTR1;1 and PtaSULTR4;1 in December in connection with temperatures below zero degree Celcius. This increase is contradictory to dormancy during winter and seems to be induced by extreme temperature and the associated freezing conditions. Still, as PtaSULTR1;1 and PtaSULTR4;1 did not correlate to sulfate content or weather parameters transcription of these sulfate transporters seems to be regulated by different factors or additional influences overlay the seasonal adaptation.
An interesting observation regarding regulation is the missing appreciable influence of weather parameters on sulfate transporter expression in leaves. Still, PtaSULTR1;1, PtaSULTR4;1 and PtaSULTR4;2 are positively correlated to each other and PtaSULTR1;1 correlates negatively to the leaf sulfate content which corresponds to the described export and phloem loading functions. But, the first
143 Discussion increase and later decrease of PtaSULTR1;1 and PtaSULTR4;2 mRNA during leaf development appear to be regulated mainly by internal developmental processes. Environmental influences on gene expression during onset of leaf development in spring can largely be separated from developmental and leaf-age dependent influences in Populus (Wissel et al., 2003). This was shown by Wissel et al. (2003) analyzing the mRNA levels of 11 nuclear genes in leaves of a free-growing poplar tree throughout the growing season. Multivariate statistics allowed the differentiation between influences of environmental factors and developmental responses. Gene expression in leaves up to one month after bud flush seemed to be mainly determined by a developmental program, but later in summer environmental factors were much more important (Wissel et al., 2003). Developmental implication might also be the reason for highest transcript accumulation of EF1beta detected in the first small leaves in spring. As EF1beta is involved in the translation process of mRNA into new proteins (Brands et al., 1986; Lauer et al., 1984) it is convincing that during early leaf development the amount of an elongation factor might be high. Also the continuously diminishing transcript abundance in developing bark of young twigs from mid May until the end of July which was observed for all genes including the reference gene EF1beta might be regulated by internal developmental processes.
In summery, the influences of seasonality reflected by day length and temperature on sulfate transporter expression in bark may be interpreted as transcriptional regulation to control and adjust sulfate distribution. For this purpose sulfate transporters in the phloem are involved in the regulation of storage and mobilization processes in deciduous trees. Indeed leaves underlie a developmental process but whether and how the transcript abundances of sulfate transporters are regulated and which factors are mainly involved in leaves has to be investigated in further experiments.
4.9 Conclusion
Previous research gave information about sulfate distribution in trees but excluded genetic aspects. The current study has highlighted for the first time many details facilitating the assumption that a family of membrane transporters are involved in sulfate distribution within deciduous trees. Inherently, the amount of 16 determined genes in Populus tremula x P. alba is remarkable because comparison to the 144 Discussion consistently lower number of orthologs in herbaceous plants seems to be a first hint to a tree specific adaptation in sulfate allocation. In particular detailed knowledge about the regulation of long distance transport regarding storage and mobilization is of great interest in trees due to their perennial growth. In this context the cell specific identification of two sulfate transporter transcripts, PtaSULTR1;1 and PtaSULTR3;3a, in the phloem support the hypothesis that phloem loading is carried out via transport proteins and include an apoplastic step in poplar. The concluded contribution of both genes in sulfate retrieval into transport phloem and the assumed contribution of PtaSULTR1;1 in sulfate loading into the collection phloem of leaves constitute the first insight in elucidating the largely unknown control of phloem transport. But it has to be taken in account that the presented results were restricted on the transcriptional level. The definitive transport direction has to be verified in further studies and localization analyses on the protein level should follow to confirm the supposed functions.
The detailed expression overview of the whole sulfate transporter gene family in a tree in this thesis including the separation of bark and wood in stem and roots, separation of leaf vein, lamina and the apex is unique. The so identified tissue specific expression of sulfate transporters constitutes a potential for plants to regulate the sulfate supply of particular tissues and enables responses to environmental conditions.
The presented results allow for the first time insight in the transcriptional regulation of sulfate transporters due to seasonal changing. The correlation between weather parameters and the transcript amount of phloem specific and putative vacuolar sulfate transporters as well as the inverse correlation between expression und sulfate content in bark supports the assumption that these genes participate in sulfate mobilization. Further seasonal investigations of wood and roots might complete our knowledge of sulfate mobilization and estimate the importance of different tissues for storage. Therefore, the identification of wood specific sulfate transporters on the cellular level and their changes due to the season might be of great interest. In this context a complex unsolved area for future research is the trigger pathway between seasonal input signals and their conversion into mRNA accumulation of sulfate transporter genes.
145 Summary
5 Summary
Sulfur is one of the six macronutrients which plants require for growth and development. Sulfur is available to plants primarily in the form of sulfate in the soil where it is taken up by the roots. Beside short distance cell-to-cell transfer, sulfate is distributed within higher plants by long distances through the vascular system. The long distance transport from roots to the leaves takes place in the xylem while the transport from the so called ‘source’ to ‘sink’ tissues takes place in the phloem. Within herbaceous plants it seems that sulfate distribution is regulated by the tissue specific expression of numerous sulfate transporter isoforms. The aim of this thesis was the determination of molecular biological knowledge in poplar (Populus tremula x P. alba) to enhance the understanding of sulfate transport processes in trees in particular by the phloem.
In the present study, the gene family of sulfate transporters was investigated in poplar trees. 18 potential sulfate transporter genes were identified. Compared to the Arabidopsis thaliana sulfate transporter gene family which contains 14 sequences and to rice with 13 orthologs, poplar exhibited gene duplications. The significant homologies of deduced amino acid sequences encoded by 16 putative Populus tremula x P. alba sulfate transporters to the functionally characterized orthologs from Arabidopsis thaliana, indicated that the poplar sequences are highly likely to possess the expected sulfate transport function.
Northern blot analyses exhibited highly tissue specific expression patterns of the 16 sulfate transporter genes from Populus tremula x P. alba. This result adverts to a transcriptional regulation and organization of sulfate distribution within poplar trees. The solely expression of PtaSULTR1;2 in fine roots supports its significance in sulfate uptake from the soil. Seven poplar sequences were abundant in stem wood and root wood which coincides with the tree specific importance of nutrient distribution and storage along the stem. The constant expression of both PtaSULTR4;1 and PtaSULTR4;2 transporters in every tissue was consistent with the putative localization within the vacuolar membrane controlling cell internal sulfate distribution. Most 146 Summary interesting with regard to phloem transport of sulfate was the observation that the two sulfate transporters PtaSULTR1;1 and PtaSULTR3;3a were specifically expressed in the bark and leaf veins.
Due to these results in situ hybridization analyses were performed for PtaSULTR1;1 and PtaSULTR3;3a to localize the transcripts at the cellular level within different tissues. In the transport phloem along the stem and major leaf veins, PtaSULTR1;1 and PtaSULTR3;3a transcript accumulation was highest in companion cells. This specific expression suggests an uptake function for sulfate from the apoplast into the phloem. Furthermore, both transporters were present in pith ray cells predominantly adjacent to xylem vessels and in parenchyma cells around the protoxylem. This result indicates sulfate uptake into the cytosol of ray cells for storage in the wood. Exclusively PtaSULTR1;1 was present in the collection phloem of minor leaf veins which suggests that phloem loading with sulfate in leaves is achieved by PtaSULTR1,1 for further distribution through the sieve elements to sink tissues. Strong expression of PtaSULTR1;1 was also abundant in developing tissues like the shoot apical meristem, leaf margin meristem, side root meristem, provascular strands and ray initials. Therefore, this sulfate transporter seems to be important for the uptake of sulfate into developing cells for further assimilation. In summary, for the first time two phloem specific sulfate transporters were identified within a plant species which are with great feasibility involved in sulfate distribution from ‘source’ to ‘sink’ tissues.
To investigate the regulation of sulfate distribution in response to nutrition Populus tremula x P. alba trees were grown under sulfate deficient conditions. While the transcript of the phloem specific sulfate transporter PtaSULTR1;1 increased in both, mature leaf veins and bark tissue, simultaneously the sulfate content decreased in these tissues. Simultaneous up-regulation of both putative vacuolar sulfate efflux transporters of group 4 in bark and the up-regulation of PtaSULTR4;2 in stem wood supports the assumption that sulfate is mobilized from vacuoles of storage tissues under these conditions. This mobilized sulfate is supposed to be taken up via PtaSULTR1;1 into the companion cell-sieve element complex for subsequent transport to developing sink tissues. However, PtaSULTR3;3a was not clearly affected at the transcript level by sulfur deficiency, indicating regulatory differences between the two phloem specific sulfate transporters. In contrast to bark and wood,
147 Summary the expression of both SULTR4 transporters was not influenced in leaf veins by sulfur deficiency. Apparently, bark tissue is more important for sulfate remobilization than mature poplar leaves. The down-regulation of the three wood specific sulfate transporters, PtaSULTR2;1a, PtaSULTR2;1b and PtaSULTR3;2b, indicates that in poplar also xylem transport is influenced under sulfur limitation. In summary, sulfate deficiency induced the transcriptional regulation of sulfate transporters in trees which probably improve the supply of growing tissue.
Seasonal expression patterns of phloem specific sulfate transporters and of putative vacuolar sulfate efflux transporters were analyzed from August 2006 until September 2007 using field grown poplar trees. Sulfate accumulation in the bark started in late autumn when PtaSULTR1;1 expression in senescent leaves was high. Because PtaSULTR1;1 transcript is abundant in the phloem of minor leaf veins, the increase of its mRNA during abscission suggests an increase in phloem loading of sulfate for further transport out of the leaves into storage tissues. Sulfate content in the bark was high during the entire dormant period when PtaSULTR3;3a and PtaSULTR4;2 expression was nearly not detectable. The transcript abundance of the putative vacuolar sulfate efflux transporter PtaSULTR4;2 and both phloem specific sulfate transporters increased in the bark during bud swelling and early leaf development. This expression pattern with simultaneously decreasing sulfate contents in bark indicates sulfate mobilization from the vacuoles of bark cells and further loading into the phloem during leaf development in spring. Statistical analyses revealed that the accumulation of these sulfate transporter transcripts was exclusively positively correlated with each other. PtaSULTR3;3a and PtaSULTR4;2 were in addition negatively correlated to the sulfate content in bark. Apparently, sulfate remobilization from the vacuoles correlates with further uptake into the phloem which contributes to a loss of sulfate in bark tissue. A seasonal influence of temperature and day length was mainly found for the sulfate content, PtaSULTR3;3a and PtaSULTR4;2 transcript abundances in bark. Thus, the seasonal analyses of sulfate transporter expression and sulfate content in the present study provide for the first time insight into the transcriptional regulation of sulfate transporters during changing environmental conditions within the annual life-cycle.
148 Summary
6 Zusammenfassung
Schwefel zählt zu den sechs Makro-Nährelementen, die essentiell sind für das Wachstum und die Entwicklung von Pflanzen. Pflanzen nehmen Schwefel hauptsächlich in Form von Sulfat aus dem Boden über die Wurzeln auf. Neben dem Kurzstreckentransport von Zelle zu Zelle über Plasmodesmen wird Sulfat in höheren Pflanzen auch über längere Strecken in den Leitbündeln transportiert. Der Langstreckentransport von den Wurzeln zu den Blättern findet im Xylem statt während der Transport von den so genannten ’source’ Geweben zu den so genannten ’sink’ Geweben im Phloem stattfindet. Die Verteilung von Sulfat wird in krautigen Pflanzen durch zahlreiche, gewebespezifisch expremierte Sulfattransporter Gene reguliert. Ziel dieser Arbeit ist die Ermittlung molekularbiologischer Grundlagen in der Pappel (Populus tremula x P. alba) zum besseren Verständnis der Sulfattransportprozesse in Bäumen insbesondere über das Phloem.
In der vorliegenden Arbeit wurde die Gen-Familie der Sulfattransporter in der Pappel untersucht. Anhand der klonierten cDNA Sequenzen wurden 18 potentielle Sulfattransporter Gene identifiziert, welche zahlenmäßig die größte Familie im Vergleich zu den bisher untersuchten krautigen Pflanzen darstellt. Im Vergleich zu Arabidopsis thaliana mit 14 Sequenzen und Reis mit 13 wies die Pappel Gen- Verdopplungen auf. Aufgrund der signifikanten Homologien zwischen den abgeleiteten Aminosäuresequenzen von 16 der potentiellen Sulfattransporter aus der Pappel mit funktionell charakterisierten Sulfattransportern aus Arabidopsis thaliana, kann davon ausgegangen werden, dass die Pappel-Proteine entsprechend Sulfat transportieren.
Die durchgeführten Northern-Blot Analysen zeigten stark gewebespezifische Expressionsmuster der 16 Sulfattransporter Sequenzen aus Populus tremula x P. alba. Diese Ergebnisse weisen auf eine transkriptionelle Regulation und Organisation der Transporter-Gene und somit der Sulfatverteilung in Bäumen hin. Die ausschließliche Expression von PtaSULTR1;2 in Feinwurzeln deutet auf eine Beteiligung an der Sulfataufnahme aus dem Boden hin. Insgesamt 7 Sequenzen der 149 Summary
Gen-Familie waren im Holz von Wurzel und Stamm nachweisbar. Diese große Anzahl von Transportern im Holz kann mit der baumspezifischen Verteilung von Sulfat entlang des Stammes mehrjähriger Pflanzen in Zusammenhang gebracht werden. Des weiteren stimmt die konstante Expression von PtaSULTR4;1 und PtaSULTR4;2 in allen Geweben mit deren potentiellen Lokalisation in der Vacuolen- Membran überein. Ein zentrales Ergebnis der vorliegenden Arbeit war die spezifische Detektion der zwei Sulfattransporter PtaSULTR1;1 und PtaSULTR3;3a in Rinde und Blattader. Dieses Expressionsmuster deutet auf eine Lokalisation im Phloem hin.
Aufgrund dieser Hinweise wurde mittels in situ Hybridisierung die zelluläre Expression der Sulfattransporter PtaSULTR1;1 und PtaSULTR3;3a in verschiedenen Geweben nachgewiesen. Im Transport-Phloem der Blattadern und entlang des Stammes zeigte sich die stärkste Transkript Akkumulation in den Geleitzellen. Dies ist ein Hinweis auf eine Beteiligung der Transporter an der Sulfataufnahme ins Phloem. Beide Sulfattransporter waren zudem in den Markstrahlenzellen angrenzend an Tracheen sowie im Protoxylem nachweisbar. Demzufolge ist eine Beteiligung an der Sulfataufnahme ins Cytosol der Markstrahlenzellen für die Speicherung im Holz zu erwarten. Die ausschließliche Expression von PtaSULTR1;1 im Phloem der Blattadern höherer Ordnung deutet darauf hin, dass die Beladung des Phloems mit Sulfat in den Blättern für den Langstreckentransport zu den Sink-Geweben vornehmlich durch PtaSULTR1,1 realisiert wird. Zudem konnte eine starke Expression dieses Sulfattransporters in sich entwickelndem Gewebe wie dem Apical-, Blattrand- und Seitenwurzelmeristem, den provasculären Strängen und den Vorläuferzellen der Markstrahlen im Kambium nachgewiesen werden. Diese Lokalisation legt eine Beteiligung von PtaSULTR1,1 an der Sulfataufnahme in wachsendes Gewebe nahe. Damit wurden in dieser Arbeit erstmals zwei Phloem spezifische Sulfattransporter in einer Pflanzenart nachgewiesen, welche mit großer Wahrscheinlich an der Verteilung von Sulfat von ’source’ zu ’sink’ Geweben beteiligt sind.
Um die Regulation der Sulfatverteilung aufgrund der Nährstoffverhältnisse zu untersuchen, wurden Populus tremula x P. alba Bäume unter Schwefelmangel Bedingungen angezogen. Während die Expression des Phloem-spezifischen Sulfattransporters PtaSULTR1;1 in Rinde und Blattader anstieg, nahm die Sulfatkonzentration in diesen Geweben ab. Der gleichzeitige Anstieg der mRNA
150 Summary potentieller vacuolärer Sulfat-Efflux-Transporter aus Gruppe 4 in der Rinde und zusätzlich im Holz unterstützt die Annahme, dass Sulfat aus den Vacuolen der Speichegewebe mobilisiert wird. Dieses Sulfat könnte über PtaSULTR1;1 in die Geleitzellen des Phloems transportiert und anschließend zu den wachsenden Sink- Geweben geleitet werden. Hingegen zeigte PtaSULTR3;3a keine Expressions- unterschiede durch Sulfatmangel, was auf einen offensichtlichen Unterschied in der Regulation der beiden Phloem-spezifischen Sulfattransporter schließen lässt. Im Gegensatz zur Expression in Rinde und Holz wurde der Transkript-Gehalt der beiden SULTR4 Sequenzen in Blattadern durch Schwefelmangel nicht beeinflusst. Diese gewebespezifische Ausprägung lässt den Schluss zu, dass der Stamm in Bäumen das primäre Sulfat-Reservoir darstellt. Die Abnahme der Expression der drei Holz- spezifischen Sulfattransporter PtaSULTR2;1a, PtaSULTR2;1b und PtaSULTR3;2b weist zudem darauf hin, dass in Pappeln auch der Transport aus dem Xylemsaftstrom entlang des Stammes durch Sulfatmangel beeinflusst wird. Diese Ergebnisse zeigen, dass Schwefelmangel in Bäumen eine transkriptionelle Regulation der Sulfattransporter in Speichergeweben hervorruft, um vermutlich die Versorgung der wachsenden Gewebe zu gewährleisten.
Um die saisonale Expression der Phloem-spezifischen Sulfattransporter und der potentiellen Vacuolen-Efflux-Transporter zu untersuchen, wurden von August 2006 bis September 2007 Proben von Populus tremula x P. alba Bäumen genommen, die im Freiland wuchsen. Im Herbst erfolgte eine Akkumulation von Sulfat in der Rinde während die Expression von PtaSULTR1;1 in welken Blättern anstieg. Da PtaSULTR1;1 im Phloem der feinen Blattadern vorliegt, deutet dessen transkriptioneller Anstieg während der Blatt-Seneszens auf eine Beladung des Phloems mit Sulfat hin. Im Phloem kann Sulfat anschließenden aus den Blättern in die Speichegewebe des Stammes transportiert werden. Während der gesamten Ruheperiode im Winter war die Sulfatkonzentration in der Rinde hoch, während die Expression der Sulfattransporter PtaSULTR3;3a und PtaSULTR4;2 kaum nachweisbar war. Im Frühling stieg der Transkriptgehalt des potentiellen Vacuolen- Efflux-Transporters PtaSULTR4;2 und der beiden Phloem-spezifischen Sulfattransporter in der Rinde an. Dieser Anstieg erfolgte parallel mit dem Knospenschwellen und der frühen Blattentwicklung. Die gesteigerte Expression und der gleichzeitige Sulfatverlust in der Rinde weisen auf eine Sulfat Mobilisierung und
151 Summary
Beladung des Phloems in der Rinde hin während im Frühjahr die Blätter austreiben. Statistische Analysen zeigten, dass die Transkript Akkumulation der Phloem- spezifischen und potentiellen vacuolären Sulfattransporter ausschließlich positiv miteinander in Beziehung stehen. Zusätzlich korrelieren PtaSULTR3;3a und PtaSULTR4;2 negativ mit der Sulfatmenge in der Rinde. Dies unterstützt die Annahme, dass die Sulfat Mobilisierung aus den Vacuolen und die anschließende Aufnahme ins Phloem zusammenhängen und zum Sulfatverlust in der Rinde führen. Ein saisonaler Einfluss von Temperatur und Tageslänge auf die Expression von PtaSULTR3;3a, PtaSULTR4;2 und auf die Sulfatmenge war in der Rinde zudem erkennbar. Diese Untersuchungen zeigen, dass die saisonale Sulfatverteilung innerhalb der Pappel auf der transkriptionellen Ebene der Sulfattransporter reguliert wird.
152 References
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Acknowledgement
Für die Bereitstellung des interessanten Forschungsthemas, für die Betreuung und Diskussionsbereitschaft während der Arbeit bedanke ich mich herzlich bei Prof. Cornelia Herschbach.
Durch die finanzielle Unterstützung der Deutschen Forschungsgemeinschaft (DFG; Project: He 3003/2) und der Albert-Ludwigs-Universität Freiburg (Promotionsstipendium der Landesgraduiertenförderung) wurde diese Arbeit ermöglicht.
Ich bedanke mich bei Prof. Heinz Rennenberg, der mir die Durchführung der Arbeit am Institut für Baumphysiologie der Albert-Ludwigs-Universität Freiburg ermöglichte und mich finanziell und durch seine wissenschaftliche Erfahrung unterstützte.
Dr. Frank Ditengou danke ich für die umfangreiche Einweisung in die Technik der In situ Hybridisierung, die Betreuung während der Versuchsdurchführung und die gute Zusammenarbeit. In diesem Zusammenhang möchte ich Prof. Klaus Palme danken der mir diese Arbeiten am Institut für Botanik II der Universität Freiburg ermöglichte.
Vielen Dank an Prof. Siegfried Fink, der die Aufgabe des Korreferenten übernommen hat und mich beim Antrag des Promotionsstipendiums durch ein Gutachten unterstützte.
Henning Wildhagen möchte ich für die außerordentlich gute Zusammenarbeit im Besonderen bei der Durchführung des saisonalen Freilandversuchs danken.
Ganz herzlich danken möchte ich Anne Honsel, Gunda Stölken, Gertraud Michl, Barbara Ehlting, Cristiane Steinki-Schwarz, Sebastian Pfautsch und Henriette Dietrich für die außerordentlich gute Zusammenarbeit im Labor, für die fachliche Diskussionsbereitschaft, die unermüdliche Hilfsbereitschaft, die gute Atmosphäre und die freundschaftliche Unterstützung während der Arbeit. Barbara Ehlting danke ich zudem für das Korrekturlesen dieser Arbeit.
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Für die schweißtreibende Hilfe bei der Säuberung der Freilandfläche und Pflanzung der Pappeln möchte ich besonders Michael Rienks, Gertraud Michl, Sebastian Pfautsch, Peter Escher und den restlichen Helfern des Instituts für Baumphysiologie danken. Michael Rienks danke ich zudem für die unermüdliche Hilfe bei zahlreichen EDV-Problemen.
Der Wissenschaftlichen Gesellschaft in Freiburg danke ich für die finanzielle Unterstützung, die es mir ermöglichte meine Arbeit im Rahmen der internationalen Tagung „Plant Vascular Biology 2007“ in Taiwan vorzustellen.
Jeanne-Claude Davidian, INRA Montpellier in Frankreich, danke ich für die Bereitstellung der Hefemutante CP154-7A zur funktionellen Untersuchung der Sulfat Transporter. Zudem danke ich Ihm und Emmanuelle Cabannes für die Bereitschaft meine Fragen zu diesem Thema zu beantworten.
Prof. Rainer Hedrich, Institut für Botanik I der Universität Würzburg, danke ich für die Möglichkeit in seiner Arbeitsgruppe Versuche mit der elektrophysiologischen Spannungsklemmen-Technik durchzuführen. Für die umfangreiche theoretische und praktische Betreuung bei dieser Arbeit Bedanken ich mich herzlich bei Dr. Dietmar Geiger.
Ganz besonders danke ich meiner Familie und Freunden für ihre Unterstützung, ihr Interesse und Anteilnahme an meiner Arbeit. Ganz herzlich danken möchte ich Natalie Dürr und Peter Leciejewski für den Laptop der mir die Arbeit sehr erleichtert hat.
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