SEARCH

SEARCH BY CITATION

Keywords:

  • companion cell;
  • macromolecular signalling;
  • mass flow;
  • phloem evolution;
  • phloem loading;
  • phloem-specific proteins;
  • phloem transport;
  • phloem unloading;
  • plasma-membrane-associated proteins;
  • plasmodesmata;
  • sieve-element plastids

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

This review deals with aspects of the cellular and molecular biology of the sieve element/companion cell complex, the functional unit of sieve tubes in angiosperms. It includes the following issues: (a) evolution of the sieve elements; (b) the specific structural outfit of sieve elements and its functional significance; (c) modes of cellular and molecular interaction between sieve element and companion cell; (d) plasmodesmal trafficking between sieve element and companion cell as the basis for macromolecular long-distance signalling in the phloem; (e) diversity of sieve element/companion cell complexes in the respective phloem zones (collection phloem, transport phloem, release phloem); (f) deployment of carriers, pumps and channels on the plasma membrane of sieve element/companion cell complexes in various phloem zones; and (g) implications of the molecular-cellular equipment of sieve element/companion cells complexes for mass flow of water and solutes in a whole-plant frame.


INTRODUCTION

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Phloem has been and still is a puzzling tissue due to its position and physiology. It is virtually out of experimental reach – it is deeply embedded into other tissues – and is very ‘touchy’– it shows a range of immediate defence reactions following any kind of manipulation. Because phloem is to be studied preferably in situ, much has been learned about phloem physiology by using non-invasive techniques such as the application of radioisotopes, nuclear magnetic resonance imaging, and confocal laser scanning microscopy and a range of molecular-biological approaches.

Novel techniques have revealed a number of unexpected cell- and molecular-biological properties of the sieve tubes. Thus far, the phloem system has been known for the translocation and distribution of photo-assimilate properties. It has been shown that mass flow in the sieve tubes transported numerous other solutes and undesired guests (plant viruses, bacteria). Over recent years, however, it was discovered that the phloem transmits alarm signals (systemic acquired resistance, SAR) from plant parts attacked by pests, chewing insects or herbivores to other unaffected parts. Exchange of macromolecules (proteins, RNA) between sieve element and companion cells may play a pivotal role in SAR and other forms of remote control. Even structural sieve-element proteins appear to be mobile in the phloem and can pass sieve plates. Molecular-biological studies unravelled a multitude of carriers, pumps and channels on the plasma membrane of the sieve tubes that explains the transport of solutes (mainly sugars) and corresponding amounts of water. Collectively these findings have brought about a different appreciation of the phloem system.

Basic to phloem functioning is a bizarre duo, the sieve element/companion cell complex which forms the functional unit of the sieve tubes in angiosperms. Whereas the sieve element is almost ‘clinically’ dead, the companion cell is a paragon of bubbling activity. The secret of their success lies in their concerted interaction while carrying out discrete tasks. The extraordinary structural and molecular equipment of sieve element and companion cell provides the fundament for the multifunctionality of phloem. In this review, phloem physiology is regarded as the result of the collective power of its single units, the unique sieve element/companion cell complexes.

ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

In the course of evolution, land plants have developed a system for long-distance transport of photo-assimilates from the autotrophic towards the heterotrophic parts (Schulz 1998). Such a translocation system could not exploit the physical forces produced by water potential differences in the soil–plant–air continuum as the xylem system does. Therefore, land plants ‘invented’ a water-driven system against the transpiration gradient. According to the brilliant concept of Ernst Münch (Münch 1930), the phloem system makes use of a turgor gradient along the sieve tubes as the driving force for mass flow. High turgor values resulting from massive photosynthate accumulation by collection phloem at the source ends propel the sieve tube sap toward sites of low turgor values caused by the escape of photosynthates and the corresponding loss of water from release phloem at the sink ends (see for a description of modifications to the original Münch concept, van Bel 1995). The sophisticated phloem system simultaneously met the increment in plant size, the division of labour between plant organs and the challenge imposed by the transpiration.

GENERAL FUNCTIONS OF THE PHLOEM

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

A pressure-driven mass flow system provides a good opportunity for all kinds of components to be transported over long distances in the plant. No wonder that one finds numerous classes of compounds in sieve tube sap (e.g. Ziegler 1975; Zimmermann & Ziegler 1975; Lohaus et al. 1995; Hayashi et al. 2000). Apart from a range of carbohydrates (e.g. monosaccharides, disaccharides, galactose oligosaccharides, sugar alcohols), amino acids and minerals occur in large quantities.

A number of signalling substances, transported in minor quantities, play a major role in the integration of functioning, growth and development of the plant. Among them are several phytohormones (Ziegler 1975) that are thought to effect gene expression in distant tissues. In many species, phloem translocation of secondary plant products was observed, some of which are engaged in defence against pests and predators (Murray & Christeller 1995; Christeller et al. 1998; Hartmann 1999; Dannenhoffer, Suhr & Thompson 2001). Pest-induced signals activating defence signals at locations remote from the site of attack are postulated to wander through sieve tubes (Ryals et al. 1996). RNA in the sieve tube sap is speculated to be involved in the control of gene expression in distant tissues (Ruiz-Medrano, Xoconostle-Cazares & Lucas 2001). Carbohydrates themselves too may exert a remote control on gene expression (Lalonde et al. 1999; Smeekens 2000). Even propagation of action potentials along sieve tubes may be part of the signalling outfit (Rhodes, Thain & Wildon 1996).

The ability of sieve tubes to translocate insecticides and herbicides within the plant are exploited in agriculture (Hsu & Kleier 1996; Lichtner 2000). Furthermore, sieve tubes provide an excellent vehicle for distribution of micro-organisms. In particular, plant viruses have adapted to the special structural/functional properties of sieve tubes and hitch-hike in the phloem system (Nelson & van Bel 1998; Oparka & Santa Cruz 2000).

SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Sieve tubes in angiosperms are arrays of sieve element modules, each of which is associated with one or a few companion cells. Sieve element (SE) and companion cell (CC) originate from the same precursor cell (Esau 1969; Behnke & Sjolund 1990). In view of the compulsory association between SE and CC, it may be more appropriate to define sieve tubes as arrays of SE/CC complexes (Fig. 1). The advantage of the modular construction is that channels can be extended ad libitum by meristematic activity and that damage can be readily repaired by replacement of injured elements (Aloni & Barnett 1996; Wang & Kollmann 1996).

image

Figure 1. In vivo structure of the sieve elements (SEs) in Vicia faba with companion cells (CCs), mostly in a staggered position (modified after Knoblanch & van Bel 1998). SEs and CCs are connected through numerous pore/plasmodesma units (PPUs). Sieve-element plastids (Pl), mitochondria (M), and ER stacks (ER) are parietally positioned and evenly distributed along the SE plasma membrane, whereas parietal proteins (PP) are locally aggregated. PPs and ER are sometimes located on the sieve plates (SPs) or at the margins of the sieve pores, but do not impede mass flow. A large spindle-shaped crystalline protein cluster (CP) rests close to the sieve plate; this massive protein is a peculiarity of the Fabaceae. C, callose; CW, cell wall; N, nucleus; P, plastids; V, vacuole.

Download figure to PowerPoint

PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

The presence of SEs in Phaeophytes and Plantae infers an analogous development of photosynthate transport systems probably imposed by division of labour and enlargement of the plant body. Most likely, the phylogenetic origin of SEs in Phaeophytes and in Plantae is not related. For instance, SEs are more advanced in Phaeophyta (Schmitz 1990) than in mosses (Scheirer 1990), which originate from the Chlorophyta, of which none of the present representatives possess SEs (Graham 1993; Niklas 1997).

Emergence of highly specialized cells such as sieve tube modules was a milestone in the evolution toward distinct ‘land plant phloem’. According to Taylor (1990), the diversion between sieve cells (sustained by Strasburger cells) and SEs (sustained by CCs) must have taken place in ancient cryptogams as early as the middle Devonian. Reliable documentation of SEs in the soft phloem tissues of fossil angiosperms is missing which renders an assessment of the origin of the accompanying cells impossible (Taylor 1990).

As result, the phylogeny of SEs is virtually obscure. As so often, ontogeny may serve here to trace the rough outlines of phylogenetic descent. The ontogenic transformation of plasmodesmata in the walls between SEs may reflect the phylogenetic conversion of plasmodesmal connections into sieve pores. Furthermore, CCs lack with certainty in root protophloem of pea (Schulz 1994) and in seed coat protophloem of bean (Offler & Patrick 1984) and with a high degree of probability in root protophloem of Arabidopsis (Linstead, Dolan & Roberts 1993). The protophloem configuration may represent the ancient state of SEs – without distinct CCs – as is found in mosses and cryptogams (Table 1). The lack of CCs may be compensated by support of adjacent parenchyma cells to SE metabolism. A possible interdependence between SEs and neighbouring parenchyma in sink protophloem reminds of the structural setting of SEs in mosses and ferns (Scheirer 1990; Evert 1990a).

Table 1.  Ultrastructural features of the SEs in the respective groups of land plants reflecting their evolutionary development (from van Bel & Knoblauch 2000). Assessments are based on the data in Eleftheriou (1990), Evert (1990a,b), Scheirer (1990), and Schulz (1990). The overview shows a sequence of phylogenetic modifications in which the resistance of the conducting elements was increasingly reduced. As the nuclear control on metabolic processing in the SE was lost, assistance of the neighbouring parenchyma was invoked to sustain the deficient functioning of the SEs. The relationship between SEs and their associate cells became increasingly intimate, which culminated in the evolution of the SE/CC complex that originates from the same mother cell. The plasmodesmata on the end walls transformed into pores specialized in longitudinal transport, those on the walls between SE and CC into symplasmic corridors responsible for the exchange of metabolites and genetic and metabolic signals
 MossesVascular cryptogamsConifersAngiosperms
Ultrastructural features of sieve elements at maturity
Plasma membranePresentPresentPresentPresent
NucleusDegenerateNuclear remnantsNuclear remnantsNo
PlastidsInsignificant plastidsPresentP-type and S-type plastidsP- and S-type plastids
TonoplastSmall vacuoles filled with fibrillar materialAbsentAbsentAbsent
MitochondriaIntactIntactFew, dilated, functionality is matter of disputeFew, dilated, functionality is matter of dispute
RibosomesNot observedNoNoNo
Endoplasmic reticulumSmooth parietal nets and stacksSmooth parietal nets and stacksSmooth parietal nets and stacksSmooth parietal nets and stacks
Cytoskeleton no microfilamentsMicrotubules,NoNoNo
Golgi apparatusNoNoNoNo
Structural and physiological properties of sieve elements
Callose production?Present in some speciesPresentPresent
Specific inclusionsRefractive spherulesRefractive spherulesP- and S-type plastidsP- and S-type plastids, P- proteins with few exceptions
Symplasmic connectionsPlasmodesma-like on all wallsSmall pores equally distributed over the wallsSame degree of pore specialization on all wallsLarge pores on end walls, much smaller on side walls
Associated cellsParenchyma with high metabolic activityParenchyma with high metabolic activityStrasburger cellscompanion cells
Connectivity with associated cellsModerately symplasmically connectedMany single plasmodesmata with adjacent parenchymaPore at the SE-side, branched pd at the Strasburger cell sidePore at the SE-side, branched pd at the companion cell side

On the premise that present concepts on plant evolution are correct, the (ultra)structural trends in the extant plant groups may contribute to the understanding of the phylogeny of SEs and accompanying cells (Behnke & Sjolund 1990; van Bel 1999). Of course, this evolutionary reconstruction is only useful if plant groups have largely retained the conducting elements of their ancestors.

Optimization of longitudinal flow was apparently achieved by increasing the porosity of the end walls through enlargement of the symplasmic windows (Table 1). The SEs elongated and developed thicker cell walls, possibly to sustain increasing turgor pressures. In addition, most of the cytoplasmic structure was dismantled to diminish the resistance to mass flow (Table 1). As the cytoplasmic outfit degenerated finally leading to full nuclear disintegration, the adjacent parenchyma cell gradually took over command over the events in the SE (Table 1).

As nuclear control of metabolic processing vanished in SEs, the metabolic input of the neighbouring cell was invoked to sustain the deficient functioning of the SE (Table 1). An absolute prerequisite for fine-tuning the activities of SE and CC are intimate symplasmic contacts. Consequently, the highly specialized plasmodesmata between CC and SE became responsible for permanent and selective exchange of micro- and macromolecules (e.g. Ruiz-Medrano et al. 2001).

ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

In general, the meristematic mother cell of a SE/CC complex divides unequally in the longitudinal direction (see Behnke & Sjolund 1990). One daughter cell develops into one or several metabolically hyperactive CCs possessing a dense cytoplasm and numerous mitochondria with well-developed cristae (Esau 1969; Evert et al. 1969; van der Schoot & van Bel 1989; van Bel 1993). The other daughter cell goes through a controlled disintegration process which may be designated loosely as ‘programmed cell semi-death’ (e.g. Esau 1969; Wergin, Palevitz & Newcomb 1975; Eleftheriou 1990; Evert 1990b).

The ontogeny may largely reflect the successive phylogenetic stages (Table 1). The nucleus gradually disintegrates, the vacuolar membrane disappears just as cytoskeletal elements, ribosomes, Golgi bodies and the mitochondria are reduced in number (Esau 1969; Behnke & Sjolund 1990). What finally remains is the plasma membrane, a thin layer of parietal cytoplasm, composed of mostly stacked endoplasmic reticulum (ER) or fenestrated ER (Thorsch & Esau 1981a,b; Sjolund & Shih 1983) and a few often dilated mitochondria (Evert 1990b). Conspicuous elements of the SE cytoplasm are phloem-specific plastids (Behnke 1991a) and sieve element P-proteins (Eleftheriou 1990; Evert 1990b; Cronshaw & Sabnis 1990; Behnke 1991b; Iqbal 1995).

The ontogenic development of symplasmic contacts between SE/CC complex and adjacent parenchyma were studied in detail in the collection phloem of a few dicotyledonous leaves (Gamalei 1990; Beebe & Russin 1999). Apart from one attempt (Turgeon & Hepler 1989), however, the actual connectivity between SE/CC complex and surrounding cells in minor vein phloem has not been tested yet. As for transport phloem, intracellular injection of Lucifer Yellow into the phloem cells of Lupinus stems gave some insight into the symplasmic connectivity in the course of the SE/CC ontogeny (van Bel & van Rijen 1994).

The precursors of SEs have similar plasmodesmal contacts at the respective interfaces with adjoining meristematic cells. All plasmodesmata shut off prior to differentiation into SE/CC complexes (van Bel & van Rijen 1994) which may be a general event in meristematic cells going into differentiation (Ehlers, Binding & Kollmann 1999; Ruan, Llewellyn & Furbank 2001). Plasmodesmal closure is considered to be a prerequisite for autonomous development without interference by neighbouring cells (Pfluger & Zambryski 2001).

The plasmodesmata undergo a disparate development at the respective interfaces of SEs in transport phloem.

  • 1
    Plasmodesmata at the outer surface of the SE/CC complex, responsible for communication between CCs and phloem parenchyma cells (PPs), can change in frequency and become branched during ontogeny (Kempers, Ammerlaan & van Bel 1998; K. Ehlers, personal comm.). At the CC/PP interface, the symplasmic connectivity is limited; at the SE/PP interface, symplasmic connectivity is absent (Kempers et al. 1998).
  • 2
    Plasmodesmata located in the transverse walls are transformed into large pores up to 1 µm. In the early stages of SE ontogeny, callose platelets are deposited around the plasmodesmata which are largely dissolved at a later stage (Evert 1990b). The plasma membranes of SEs are continuous through the sieve pores, but a net-like ER resides at their margins (Evert 1990b)
  • 3
    Plasmodesmata between SE and CC develop into special symplasmic connections, termed pore-plasmodesm units (PPUs, van Bel & Kempers 1997). They are branched at the CC side with up to 100 branches (Evert 1990b) coalescing in a central cavity which connects to a somewhat wider symplasmic channel at the SE side. As in sieve pores of gymnosperms, the ER tubules seem to traverse the PPUs (Ding et al. 1993). As for the central cavity, the PPU ultrastructure is as yet unresolved (van Bel & Kempers 1997).

STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

The power of the minimal outfit of the sieve element

The secret of the evolutionary success of the SEs seems to be simple. It combines the suitability for mass flow with the capacity to manage turgor control and, in doing so, to direct photo-assimilate fluxes. It is tempting to speculate why SEs have retained exactly this structural and metabolic equipment. An attempt will be therefore made to understand the cell biology of SEs and their interaction with the CCs.

Necessity of a plasma membrane

In the course of evolution long-distance mass flow conduits became necessary to meet the increasing demands of the plant body. In angiosperms, the vessel members undergo a programmed cell death resulting in the construction of dead channels through which mass flow is driven by transpiration. However, translocation in a direction opposite to the transpiration required retention of a SE plasma membrane necessary to generate osmotic gradients. The associated water fluxes were regarded as being responsible for mass flow (Münch 1930) until the advent of electron microscopy cast doubts on this concept.

Massive deposits on the sieve plate pores (e.g. Robidoux et al. 1973; Johnson, Freundlich & Barclay 1976) were considered to be incompatible with mass flow. In order to meet the presumptive sieve pore occlusion several alternatives to mass flow were advanced. In the end, however, the physiological and mathematical evidence in favour of mass flow became overwhelming (e.g. Goeschl & Magnuson 1986; Magnuson, Goeschl & Fares 1986) despite apparent sieve plate occlusion. Over the course of the dispute, several authors argued that the occlusion was an artefact triggered by cutting and fixation procedures (Evert et al. 1969; Kollmann 1973; Johnson et al. 1976). If extreme care is taken in preparation, sieve plates are found to be essentially open (Kollmann 1973; Johnson et al. 1976; Sjolund & Shih 1983). Gentle in situ fixation (fixation was done in situ in intact plants or stems) of broad bean stem phloem (Fig. 2) showed that protein deposits do occur in vivo on sieve plates. Narrow corridors are present in the protein clots so that mass flow through the sieve pores is unhindered (Ehlers, Knoblauch & van Bel 2000).

image

Figure 2. Corridors through the protein deposits onto the sieve pores (modified after Ehlers et al. 2000). EM picture of the sieve-plate area between Vicia faba sieve elements (SEs) fixed in situ. Deposits of sieve-element proteins and stacked ER cisternae occur on the sieve-plate walls (arrowheads), but corridors (arrow) in the front of the sieve pores leave an unhindered passageway for mass flow. Sectioned portions of the large crystalline protein body and sieve-element organelles are visible. CC companion cell. (×4150)

Download figure to PowerPoint

Deposits on sieve plates (Fig. 2) were also observed in confocal laser scanning microscopy (CLSM) studies which provided unequivocal visual proof of mass flow in sieve tubes. Despite presence of protein clots and ER on sieve plates, dye accumulation or retardation of dye movement was not observed (Knoblauch & van Bel 1998). Real-time recordings of the water flow through sieve tubes have also been made using nuclear magnetic resonance (NMR) techniques (Köckenberger et al. 1997; Rokitta et al. 1999). These recordings showed flow rates similar to those calculated earlier from the displacement of radiolabelled substances (Goeschi & Magnuson 1986; Magnuson et al. 1986). Provided that the resolution of the method increases, NMR promises to be an excellent tool for in vivo visualization of water mass flow as well as for identification of the solutes translocated (Köckenberger 2001).

Compatibility of mass flow with parietal stability of sieve-element organelles

Given the mass flow in sieve tubes, the stable parietal position of structural SE elements is remarkable. Organelles and parietal proteins are not dragged along with the turbulent mass flow. Recently, about 7 nm long macromolecular extensions were discovered (Fig. 3) that anchor ER, mitochondria and plastids to each other and to the plasma membrane (Ehlers et al. 2000). It appears that SEs express more than any other cell type such an anchoring system keeping organelles and proteins together as a propoplasmic sheet (Ehlers et al. 2000).

image

Figure 3. Macromolecular anchors between the organelles themselves and between the organelles and the SE plasma membrane (modified after Ehlers et al. 2000). Macromolecular anchors in the cytoplasmic lining of a Lycopersicon esculentum sieve element visible in a digitized false-colour micrograph (bar, 50 nm; insert bar, 200 nm). Single, straight clamp-like structures (arrows), about 7 nm long and 4 nm thick, occur between the plasma membrane and the outer membrane of the sieve-element plastids (P) and the parietal ER-cisternae (ER). The clamps are mostly twice as long (15 nm long, 4 nm thick) between the membranes of the parietal ER-cisternae and the outer plastid membrane (arrowheads). The long clamps are often H-shaped or Y-shaped.

Download figure to PowerPoint

Endoplasmic reticulum: ion reservoir and conveyer belt?

In comparison with other organelles, ER in SEs is well-preserved in the form of nets or stacks (Thorsch & Esau 1981a,b; Sjolund & Shih 1983). The parietal fenestrated ER which may share some traits with the cortical ER (Staehelin 1997) was postulated to be a structural necessity for ATP-fuelled retrieval (Sjolund & Shih 1983). Recently, ER in SEs has been proposed to function as a rail system for the trafficking of proteins between SEs and CCs (Oparka & Turgeon 1999) which may extend into the SEs serving as a means for protein sorting (van Bel & Knoblauch 2000).

Mysterious sieve-element plastids

Occurrence of sieve-element plastids is universal among spermatophytes. Sieve-element plastids are very diverse in appearance and are strongly family specific (Behnke 1991a). According to the nomenclature of Behnke (1991a), sieve-element plastids are classified as S-type (starch inclusions only) or P-type (proteinaceous inclusions together or without starch bodies) plastids. Of 382 dicotyledonous families investigated, 320 had exclusively S-type plastids, 48 families had representatives with only P-type plastids, whereas 14 families included species with S-type or P-type plastids (Behnke 1991a).

Despite their ubiquitous presence, the function of sieve-element plastids is obscure. It was suspected that starch grains of disrupted sieve-element plastids were engaged in the wound-induced occlusion of sieve plates (Barclay, Oparka & Johnson 1977). In this context, it is worth noting that several types of sieve-element plastids enclose crystalline proteins (Behnke 1991a). CLSM-aided observation has shed some light on the in vivo behaviour of sieve-element plastids in Vicia faba (Knoblauch & van Bel 1998). Upon insertion of micropipettes with a tip diameter of 1 µm, sieve-element plastids exploded instantly. In contrast, sieve-element plastids remained intact in reaction to being impaled by a 0·1 µm electrode tip. After explosion, membranes of ruptured plastids stayed attached to the plasma membrane which demonstrates inadvertently the effectiveness of the macromolecular anchors (Ehlers et al. 2000). Sieve-element plastids seem to be less sensitive to injury and to changes in turgor potential than phloem-specific proteins which clog around the tip of even the smallest microcapillaries (S. Günther, unpubl. results). This conclusion is corroborated by the fact that electron microscopic pictures always show intact sieve-element plastids after gentle fixation (Kollmann 1973; Ehlers et al. 2000). All in all, there is no strong evidence that sieve-element plastids are involved primarily in sealing of sieve plates and their in vivo function remains a mystery to date. Sieve-element plastids might well function as storage units, provided that the sets of enzymes needed for synthesis and breakdown of macromolecules is present. In that case, sieve-element plastids should contain fewer inclusions in times of food shortage.

Phloem-specific proteins, more than remnants of a cellular past?

In sieve-tube exudate, at least 150 proteins occur with a molecular mass mostly in the order of 20–60 kDa (Fisher, Wu & Ku 1992; Nakamura et al. 1993; Sakuth et al. 1993; Schobert et al. 1995, 1998). These sieve-tube exudate proteins (STEPs; Schobert et al. 1995) may include all classes of phloem-specific proteins, namely, soluble proteins, traces of detached or expanded structural proteins and proteins from exploded sieve-element plastids. Possibly, many others will be found with improvement in detection methods (Kehr et al. 1999; Haebel & Kehr 2001).

The major part of the STEPs are water-soluble proteins. Some metabolic and other processes in which they are engaged are beginning to emerge (Sjolund 1997; Thompson & Schulz 1999; Hayashi et al. 2000). For example (see for review, Hayashi et al. 2000), STEPs may be engaged in sugar metabolism (sucrose synthase, mannitol dehydrogenase), transmembrane sugar transport (ATPases, sucrose carriers), membrane water permeability (aquaporins), protein degradation (ubiquitin) and detoxification of reactive oxygen species (glutaredoxin, glutathione reductase). A trypsin inhibitor (Murray & Christeller 1995), that prevents the proteolytic action of trypsin or chymotrypsin, and aspartic acid protease inhibitor (Christeller et al. 1998) were collected from transport phloem of Cucurbita. The protease inhibitors play a role in the defence against insect herbivory (Christeller et al. 1998; Dannenhoffer et al. 2001). Further properties of phloem-specific proteins in long-distance signalling are discussed in one of the sections below.

Structural phloem-specific originate from protein bodies which develop during SE ontogeny, except in many palms and grasses (Eleftheriou 1990). These structural P-proteins disperse at maturity (Behnke 1991b) and reside parietally in mature SEs (e.g. Kollmann 1973; Evert 1990b; Knoblauch & van Bel 1998). In about 10% of dicotyledonous species investigated, non-dispersive P-protein bodies have been observed (Behnke 1991b).

The spindle-shaped crystalline P-protein bodies in SEs of Fabaceae (Palevitz & Newcomb 1971; Wergin et al. 1975; Lawton 1978a,b) display a range of astonishing properties (Knoblauch et al. 2001). Insertion of a microcapillary into a SE of intact Vicia faba main veins caused dispersion of the 10–30 µm long crystalloid (Fig. 4). The original conformation was restored spontaneously after some time (Knoblauch et al. 2001). Dispersion and re-integration must therefore be ordered processes. The same expansive effect had the replacement of pure water in which the SEs were bathed, by 800 mOsmol solutions and, most strikingly, by the reverse treatment as well. Analogous treatments with 400 mOsmol solutions had no effect (Knoblauch et al. 2001). Application of Ca2+ also triggered disintegration of the crystalloid. The effects of microcapillary insertion and Ca2+ could be prevented or reverted by ethylenedaiminetetraacetic acid (Knoblauch et al. 2001). Similar reversible reactions were obtained with crystalloids of other legumes, but not with non-dispersive proteins of other families such as Urticacea and Rosaceae (Knoblauch et al. 2001). Perhaps, evolution has ‘frozen’ P-proteins of Fabaceae in the transition from non-dispersive to dispersive proteins.

image

Figure 4. Dynamics of crystalline P-protein bodies in Fabaceae. Non-dispersed and dispersed crystalline P-proteins in the sieve tubes of Vicia faba observed by confocal laser scanning microscopy in ploem tissue stained with 5(6)-carboxy-4′,5′-dimethylfluorescein diacetate (modified after Knoblauch et al. 2001). The crystalloid P-protein body before (A) and after (B) insertion of a micropipette (not visible; tip diameter 2 µm). Micropipette insertion triggers the transformation of the dense, elongate crystalloid into a roundish sieve tube plug. SE, sieve element.

Download figure to PowerPoint

According to a provisional model, Ca2+ is responsible for rearrangement of water molecules within the crystalloid leading to a shift in protein conformation. It is speculated that a turgor shock activates membrane elastiticity-dependent channels to release Ca2+ into the SE lumen (Knoblauch et al. 2001). Calcium influx from the apoplast may be mediated by channels in the SE plasma membrane which have recently been found (Volk & Franceschi 2000). Alternatively, ER stacks in the SEs may serve as a calcium reservoir used for dispersion of crystalloids (Sjolund & Shih 1983; Arsanto 1986).

Sealing of sieve tubes

Decisive for the evolutionary success of the phloem in higher plants must have been the development of efficient sealing systems. More than any other cell type the SE is in danger of acting as a massive leak for plant saps. The internal pressure in the SEs is high, the pores between the SEs are huge and rapid down-regulation of the turgor pressure is difficult given the inadequate cellular machinery.

It must be realized in this context that phloem exudation experiments using detached leaves are artefactual. In nature, mechanical damage to plants is mostly incurred due to removal of leaves or twigs by feeding animals or wind. It is unlikely that under natural conditions leaf detachment causes loss of phloem sap in the absence of mass flow by phloem loading in leaves. Thus, there may be a second compelling reason for comprehensive sealing devices. Phloem wounding may be hazardous as it provides easy access for phytopathogens. The lectin character of PP2 in Cucurbita (Read & Northcote 1983) suggests that sealing also acts as a barrier against bacteria, viruses and fungi.

CLSM studies indicated that structural phloem-specific proteins play a prominent role in sealing sieve tubes independent of the nature of disturbance. Parietal proteins detach from the plasma membrane in response to laser radiation or turgor loss (Knoblauch & van Bel 1998). In the latter instance, the parietal proteins are drawn away as threads like chewing gum (Knoblauch & van Bel 1998), which are probably identical to the ‘transcellular strands’ reported by Thaine (1962, 1969). The detached protein particles move with the mass flow to the next sieve plate, where the sieve pores become occluded. In legumes, parietal proteins detach simultaneously with the dispersion of crystalloids providing supplementary sealing capacity. Turgor disturbance seems to be a principal activator of conformation changes in structural phloem-specific proteins. A potential role of ER-contents in sieve plate occlusion deserves our attention given the pronounced effect of turgor disturbance on the ER structure (Ehlers et al. 2000).

The prevailing view is that callose deposition plays a central part in sealing of SEs, as callose is found to be associated with sieve plates of wounded sieve tubes. A dominant role of callose in sieve pore occlusion makes evolutionary sense because it is presumed to be involved in plasmodesmal closure (Lucas, Ding & van der Schoot 1993; Botha & Cross 2000, 2001; but Radford, Vesk & Overall 1998); sealing of sieve pores being transformed plasmodesmata would be functionally homologous. Yet, the abundant presence of callose in sieve pore regions may have a different ground. Callose plays an important part in the genesis of the sieve pores (Evert 1990b) and may be still present as collars around the sieve pores in intact sieve tubes (Ehlers et al. 2000).

Undoubtedly, callose is responsible for long-term plugging of sieve plates. The crucial question regarding the importance of callose in sieve tube sealing is whether callose synthesis is rapid enough for full and instant occlusion of the sieve plates. In other words, how much of the callose seen on electron microscope pictures is due to permanent callose or to preparative treatments such as cutting and fixation (Radford et al. 1998)? Older experiments seemingly showed that phloem transport is reduced by callose-inducing treatments (Scott et al. 1967; McNairn & Currier 1968), whereas others called the effect of callose deposition on phloem transport into question (Eschrich et al. 1965; Webster & Currier 1965). It remains actually doubtful if the massive amounts of callose observed can be produced enzymatically within seconds in cell walls, but more thorough work is needed for a definitive conclusion.

Strong involvement of proteins in instant sieve pore occlusion raises questions on the physiological fitness of sieve tube sealing. Logically, only species survived that invested efforts in evolving adequate sealing mechanisms. Not all angiosperms possess the same plugging tools. Phloem-specific proteins are absent in the SEs of several monocotyledons (Eleftheriou 1990); crystalline P-protein bodies only occur in members of the Fabaceae (Palevitz & Newcomb 1971; Wergin et al. 1975; Lawton 1978a, b; Behnke 1991b). There may be a inverse relationship between the size of the sieve pores and the efficiency and number of sealing devices. In the very narrow SEs of grasses (Eleftheriou 1990), sieve-element plastids may be sufficient to plug the tiny sieve pores. In the majority of dicot species, parietal phloem-specific proteins may be sufficient for occlusion. More sealing systems in one species suggests that a single one was insufficient as sieve pores widened during evolution. As a final speculation, sieve pore occlusion by phloem-specific proteins may present a temporary line of defence, sealing by callose may have a more definitive nature (‘definitive callose’, Esau 1969; ‘dormancy callose’, Davis & Evert 1970).

INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Plasmodesmal gateways between between sieve element and companion cell

Decisive for the evolutionary success of the phloem in angiosperms was the interaction between SE and CC. That SEs were dependent on CCs was suspected for a long time (e.g. Esau 1969). Owing to the support of CCs, SEs would survive over many years (in palms as many as 30 years) despite the absence of a nucleus and a deficit of mitochondria (Raven 1991). Expectedly, such an intimate relationship requires that many tasks of the SE are actually executed by the CC and implies that, for instance, energy-carrying substances and macromolecules are channelled from CC to SE (Raven 1991).

Macromolecular transfer was substantiated by showing protein trafficking between CC and SE (Bostwick et al. 1992; Fisher et al. 1992; Sakuth et al. 1993; Clark et al. 1997; Dannenhoffer et al. 1997). Most likely, many, if not all, of the 150–200 proteins detected in sieve tube sap to date (Fisher et al. 1992; Nakamura et al. 1993; Sakuth et al. 1993; Schobert et al. 1995, 1998; Hayashi et al. 2000), are synthesized in CCs (Clark et al. 1997; Dannenhoffer et al. 1997) and therefore must be trafficked to SEs via PPUs (Ruiz-Medrano et al. 2001). Turnover of phloem-specific proteins, the production of which cannot possibly take place in the enucleate SEs, was demonstrated by strong incorporation of 35S-methionine into phloem-specific proteins collected from phloem exudate (Fisher et al. 1992; Sakuth et al. 1993). Moreover, gene expression of phloem-specific proteins takes place in the CCs (Bostwick et al. 1992). The PP2 mRNA of the 42 kDa dimeric PP2 lectin was localized only within CCs by in situ hybridization of a digoxigenin-labelled antisense probe. As no trace of PP2 mRNA was found in SEs, PP2 molecules must have passed through the PPU corridor. In a similar way, sucrose carriers operating at the SE plasma membrane are thought to be produced in CCs and trafficked to SEs of Solanum (Kühn et al. 1997).

In view of macromolecular trafficking, a large diameter of PPUs was predicted (Fisher et al. 1992). A molecular exclusion limit of PPUs in the order of 20–30 kDa was detected using fluorescence-tagged macromolecules (Kempers & van Bel 1997). The estimated exclusion limit range matched the observation that the 27 kDa green fluorescent protein, expressed in CCs of tobacco and Arabidopsis, moves to the SEs and migrates within the sieve tubes (Imlau, Truernit & Sauer 1999). Phloem-specific proteins were even able to enlarge the molecular exclusion diameter of mature plasmodesmata between mesophyll cells (usually 1 kDa) up to 30 kDa (Balachandran et al. 1997, Ishiwatari et al. 1998). Six weight classes of STEPS ranging from 10 to over 100 kDa, extracted from sieve tube exudates of Ricinus communis and Cucurbita maxima, were able to widen plasmodesmata in Cucurbita leaves. Likewise, RPP13-1 (a thioredoxin h protein) from rice phloem sap had the capacity to increase the molecular exclusion limit of plasmodesmata between tobacco mesophyll cells to a size between 9 and 20 kDa (Ishiwatari et al. 1998). The findings suggest that the unusually large size of PPUs is due to the permanent presence of gating effectors in sieve tube sap

Mechanisms for trafficking macromolecules through PPUs

Although macromolecular trafficking through PPUs seems to be beyond doubt, the mode of transfer is a matter of debate. The multitude of mechanistic proposals shows that experimental evidence is far from conclusive. In sieve-tube sap, heat-shock proteins have been detected that may be involved in the unfolding of proteins to be trafficked through PPUs (Schobert et al. 1995). In a relatively simple model (Oparka & Turgeon 1999; Oparka & Santa Cruz 2000), some phloem-specific proteins act as chaperones which dock onto receptor proteins in the ER membrane. The chaperones pick up and carry proteins to be transferred along the ER rod through the PPU while ‘widening’ the PPU diameter in an unknown fashion. Recently, specific peptides which might operate at the orifice of the PPU, were reported to interfere with plasmodesmal dilation (Kragler et al. 2000). In a more sophisticated model therefore a proteinaceous complex (docking protein, chaperone protein and the protein to be transferred) docks onto two receptor proteins at the entry of the PPU before being channelled (Lucas, Yoo & Kragler 2001).

Some sieve-element proteins are also proposed to be engaged in CC-to-SE trafficking of mRNA present in the SE lumen (Ruiz-Medrano et al. 2001). The transfer mechanism may bear resemblance to protein trafficking via PPUs (Ruiz-Medrano et al. 2001). Movement proteins of phloem-limited viruses are regarded as functional analogues to plasmodesmal chaperones. For instance, potato leaf roll virus (PLRV) may mimic transfer of plant-borne mRNA in order to traffic its genome through PPUs (Sokolova et al. 1997; Prüfer et al. 1997). PLRV is a 5·6 kb RNA virus that produces a 17 kDa movement protein (Prüfer et al. 1997). The acidic terminus of the movement protein is phosphorylated by a membrane-bound kinase located in the vicinity of PPUs (Sokolova et al. 1997). Phosphorylation may be involved in a conformational change of viral RNA prior to PPU passage. Collectively, these models render a distinct function to the ER as a protein-sorting device in SEs in keeping with the observation that movement protein of cucumber mosaic virus crept over a blanket of parietal ER in SEs (Blackman et al. 1998).

Companion cell as the power house of the sieve element

In addition to macromolecular support, the SE also needs permanent input of energy to ensure its viability. Sugar metabolism and the associate energy production of the CC have a dramatic impact on the physiological fitness of the SE/CC complex (Fig. 5). Sugar uptake or retrieval by the SE/CC complex is inhibited by blocking sugar breakdown in CCs as was shown in a number of elegant experiments using transgenic tobacco plants (Lerchl et al. 1995; Geigenberger et al. 1996). After insertion of a phosphorylase gene from Escherichia coli behind a rolC promoter, pyrophosphate necessary for conversion from UDP glucose to glucose 1-phosphate was hydrolysed. As result, the glycolysis was blocked and, hence, breakdown of sucrose and production of ATP were minimized (Fig. 5). As a result, phloem loading was impaired to a high extent and massive assimilate loss along the translocation pathway was observed (Geigenberger et al. 1996). Apparently, insufficient energy was available to sustain proton extrusion, responsible for the energization of carrier-mediated uptake and retrieval by SE/CC complexes (Lerchl et al. 1995). As a matter of fact, sugar breakdown was restored in phosphorylase transgenic plants in whose genome an invertase gene from Saccharomyces cerevisiae was inserted (Fig. 5) (Lerchl et al. 1995). In double-transformants, phloem loading and sucrose loss reached the same levels as in wild-type plants (Lerchl et al. 1995; Geigenberger et al. 1996).

image

Figure 5. Carbohydrate as cargo and fuel; the impact of an impaired glucose hydrolysis on the uptake and retrieval of sucrose (from van Bel & Knoblauch 2000). Effect of some enzymes involved in the sugar metabolism of Nicotiana tabaccum companion cells on phloem loading and phloem translocation (from Lerchl et al. 1995 and Geigenberger et al. 1996). In transgenic plants with an Escherichia coli pyrophosphatase gene (PPa) behind a phloem-specific rol C promoter, glycolysis in the companion cells is impaired by pyrophosphatase. As a result, phloem loading is strongly inhibited – probably by the lack of proton pump fueling – and intense photosynthate loss along the pathway takes place due to the loss of retrieval capacity. The situation is restored to the wild-type level in transgenic plants supplemented with a Saccharomyces cerevisiae cytosolic invertase (Suc2).

Download figure to PowerPoint

The importance of sugar metabolism for phloem action is illustrated by the specific location of sucrose synthase to CCs (Nolte & Koch 1993). The CC specificity of sucrose synthase has been related to the high respiratory demand and need for readily available UDPGlc during callose biosynthesis (Koch & Nolte 1995). The former idea is consistent with the constitutive expression of of Asus 1 sucrose synthase in phloem of Arabidopsis and its induction elsewhere only when ATP supplies are limited (Martin et al. 1993). So far, localization of invertase to the phloem is less precise; vacuolar Ivr2 invertase has been detected in vascular bundles of maize leaves, but the resolution is too low to determine its exact location (Kim J-Y et al. 2000).

Three alternatives have been proposed for the energy supply of SEs of virtually symplasmically isolated SE/CC complexes (van Bel 1993, 1996). Basic to all models is the assumption that CCs produce substances required for energization of transport events at the SE plasma membrane. In the first model, sugar is accumulated by CCs and transported to SEs through PPUs. Such a mechanism seems to be highly inefficient in transport phloem, where CCs only partly cover the SEs (Zahur 1959; Fig. 6A). The interface between SE and CC amounts approx. 25% of the SE surface in transport phloem of advanced herbs (A.J.E. van Bel, unpubl. results). There, sugars would easily escape from SE areas uncovered by CCs.

image

Figure 6. Potential energy transfer and metabolic interactions between sieve element (SE) and companion cell (CC) (modified after van Bel 1993). The model presents three possible ways to maintain the sugar concentration in the SE. (A) sucrose is retrieved by sucrose/H+-symporters energised by the proton-motive force of the CC and transferred to the SE via unilaterally branched pore-plasmodesm units in the cell wall (CW) between CC and SE. (B) ATP produced by the abundantly occurring mitochondria in the CC move via the pore-plasmodesma units to the SE and energize the proton pumps in the energy-deficient SE. The proton-motive force drives the sucrose-retrieving H+-symporters in the SE plasma membrane. (C) The electrogenic potential generated by the proton pumps in the plasma membrane of the CC partly propagates via the plasma membrane lining of the pore-plasmodesma units to the plasma membrane of the SE. There, the electrogenic potential contributes to the proton-motive force that fuels sucrose/proton retrieval mediated by H+-symporters in the SE plasma membrane.

Download figure to PowerPoint

Two models provide a tight and direct control of the sugar content by SEs themselves. In one of them, ATP produced in the CC diffuses to SEs, where it is used to fuel H+-ATPases in the plasma membrane of the CC (Fig. 6B). Incidently, this model is consistent with the high ATP content of sieve tube sap (Lehmann 1979). In a more speculative model, the electrogenic membrane potential generated by the CC propagates via the plasma membrane lining of the PPUs to SEs (Fig. 6C). Circumstantial evidence in support of this model is the fractionally more negative membrane potential of the CC in tomato stem phloem (van der Schoot & van Bel 1989).

Finally, it should be underscored that sugars accumulated by phloem parenchyma for temporary storage are brought back into SE/CC complexes. Thus, phloem parenchyma cells are probably distantly involved in the energy management of SEs

Further consequences of the positioning of carriers, pumps and channels for the interaction between SE and CC (and phloem parenchyma) will be discussed below.

LONG-DISTANCE SIGNALLING IN THE PHLOEM

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Physiological impact of macromolecules in the sieve tubes

Currently, the function of macromolecules in sieve tubes is subject of extensive research, because the presence of proteins and mRNA has been linked with long-distance signalling (see for recent reviews, e.g. Citovsky & Zambryski 2000; Ruiz-Medrano et al. 2001; Lucas et al. 2001). However, caution must be exercised as to this hot topic as critical uncertainties regarding experimental validity have been identified (Oparka & Santa Cruz 2000). It is, for instance, not entirely sure whether the release of proteins and transcripts into the SEs represents the unselective loss of superfluous products or whether it is a selective process for signalling purposes (Oparka & Santa Cruz 2000). The pertinent macromolecules must be demonstrated to move from source to sink in the sieve tubes. In this respect, contamination of collected phloem sap by nuclear neighbour cells has to be excluded in view of the low amount of transcript in SEs (Oparka & Santa Cruz 2000). There is also a debate about the mechanisms of macromolecular transfer through PPUs and the macromolecular entrance in sink tissues (Santa Cruz 1999; Oparka & Santa Cruz 2000; Kragler et al. 2000). It must therefore be shown unequivocally that macromolecules can be unloaded into the sink cells where the function of specific cells is effected (Oparka & Santa Cruz 2000). In spite of the criticism, evidence seems to be mounting in favour of involvement of phloem-borne macromolecules in long-distance signalling. Conceptually, long-distance signalling may be more acceptable when one realizes that sieve tubes are giant, elongate enucleate syncytia through which materials flow at a high speed.

Translocation of phloem-specific proteins

It has been suggested that several phloem-specific proteins have not only a significance for maintenance of the SEs. Several proteins manufactured in CCs probably exert a distant effect on cellular mechanisms. After systemic transport, the 18 amino acid polypeptide systemin produced in CCs triggers production of proteinase in distant organs as a weapon against damage by chewing insects (Narvaez-Vasquez et al. 1995). It has not yet been unequivocally excluded whether systemin is produced in the mesophyll and loaded into the phloem. However, it seems to be more likely that systemin is manufactured in CCs prior to translocation in the phloem. In transformants expressing the GUS reporter gene under the control of the prosystemin promoter, activity was only detected in CCs and adjoining phloem parenchyma cells (Jacinto et al. 1997).

In intergeneric grafts of Cucurbitaceae, stock-specific proteins were present in sieve-tube exudates collected from the scion (Golecki, Schulz & Thompson 1999). Having crossed the graft border these proteins were immunolocalized in the CCs of the scion (Golecki et al. 1998). RNA gel blot analysis or reverse transcription polymerase chain reaction indicate that proteins rather than their transcripts are translocated across the graft union (Golecki et al. 1999). Even structural proteins can cross intergeneric graft borders of curcurbit species (Golecki et al. 1998, 1999). The filamentous protein (PP1) of Cucurbita maxima can undergo conformational changes and is translocated in a 88 kDa globular form (Leineweber, Schulz & Thompson 2000) which can easily pass sieve pores. The observations raise a multitude of questions concerning conformational changes of phloem-specific proteins (Leineweber et al. 2000; Knoblauch et al. 2001), the turnover site of these proteins along the pathway, their effects on CC physiology along the path, and their impact on events in sinks, to which some phloem-specific proteins may be transported (Oparka & Santa Cruz 2000) as is the case for green fluorescent protein (GFP, Imlau et al. 1999).

Remote control by mRNA translocated in sieve tubes

It has been argued that mRNA occurring in the SE could not possibly have a local effect because SEs are devoid of ribosomes. Consequently, it is postulated that plant mRNA or RNA-protein complexes exert remote control on gene expression, for example, in sinks (Jorgensen et al. 1998; Lucas 1999). Evidence for plant transcript delivery from CCs into SEs has been claimed for the sucrose carrier SUT1 in potato (Kühn et al. 1997), thioredoxin h, oryzacystatin and actin in rice (Sasaki et al. 1998), and CmNACP, a plant paralog to a virus movement protein, in Cucurbita (Ruiz-Medrano, Xoconostle-Cazares & Lucas 1999; Xoconostle-Cazares et al. 1999). Thus far, remote control by mRNA is mainly inferred by long-distance transmission of signals inducing post-transcriptional gene silencing (PGTS). In PGTS, excess or foreign RNA is degraded by a process including several steps such as detection of transcribed target RNA, copying to complementary RNA (cRNA) via a RNA-dependent RNA-polymerase, formation of an mRNA-cRNA duplex or double-stranded RNA (dsDNA) and degradation through a dsRNAase to short dsRNA-fragments (Fagard & Vaucheret 2000; Lucas et al. 2001). dsRNA-fragments, their dsRNA precursors or cRNA are candidates for phloem translocation and gene silencing in distant organs (Lucas et al. 2001).

There is some support for such a control signalling. Transgenic tobacco lines in which nitrite and nitrate reductase are over-expressed showed PGTS of these genes which initiated within small leaf patches, expanded and propagated to other plant parts (Palauqui et al. 1996). In a subsequent set of experiments, a sequence-specific signal moved from a silenced stock into a non-silenced scion where exclusively expression of nitrite/nitrate reductase was suppressed (Palauqui et al. 1997).

The systemic signal conferring breakdown of GFP moved through three-way grafts indicating that the signal was actually phloem-transmitted (Voinnet et al. 1998). The progression of silence resembled the pattern of phloem unloading of GFP unloading in sink leaves (Voinnet et al. 1998; Imlau et al. 1999). Other evidence has been obtained with heterografts of pumpkin stocks and cucumber scions (Ruiz-Medrano et al. 1999, Xoconostle-Cazares et al. 1999). In phloem sap collected from pumpkin plants, an RNA transcript (CmNACP) was found possibly involved in PGTS of the gene encoding the cucumber homologue of CmPP16 (a phloem-specific protein of Cucurbita maxima) in the scion.

Phenotypic proof for long-distance transmission of PGTS signals has been provided using grafts of tomato plants. Xa(xanthophyllic)-scions (yellowish, wild-type unipinnate leaves having leaflets with acute lobes) were grafted onto Me(mouse ear)-stocks (octopinnate leaves with unlobed leaflets) (Kim, M et al. 2001. The phenotype of the Me-mutant is caused by gene fusion between pyrophosphate-dependent phosphofructokinas (PFP) and LeT6, a tomato knotted-1 like homeobox gene. Grafting induced changes in the leaf morphology of Xa-scions; the leaves had a higher order of pinnation and the leaflet lobes were more rounded as compared to Xa-plants (Kim, M et al. 2001).

Signalling by sugars and sensing of sugars

In analogy to sugar sensing by animal and yeast cells, sieve tube sugars may function as signals reporting on the sugar status (Lalonde et al. 1999; Smeekens 2000; Hellmann et al. 2000). As sucrose is the predominant transport sugar in many species, sucrose sensors are likely to be positioned on the SE plasma membrane. Three potential sucrose sensors have been detected so far.

In Arabidopsis, translation of a leucine zipper ATB2 gene is repressed specifically by sucrose at physiological concentrations. ATB2 is expressed in the funiculus and in association with vascular tissues of developing leaves (Rook, Weisbeek & Smeekens 1998). In sugar beet, high sucrose concentrations lead to a decline of transmembrane sucrose transport correlating with a reduction in steady-state mRNA levels of BvSUT1 (Chiou & Bush 1998). Both sensors may be responsible for down-regulation of (the expression of) sucrose/H+-symporters.

A SUT2 protein in tomato contains extended intracellular domains resembling the SNF3/RTG2 sucrose sensors in yeast (Weise et al. 2000). In contrast to the structurally related sucrose/H+-symporters SUT1 and SUT4, SUT2 is hardly found in source tissue and does not show membrane transport activity (Weise et al. 2000). SUT2 is an obvious candidate for sucrose sensing: it colocalizes with SUT1 and SUT4 on the SE plasma membrane of transport and release phloem and its expression is induced by high sucrose concentrations. Thus SUT2 may be rather involved in the up-regulation of (the expression of) sucrose/H+-symporters.

As for glucose sensors, the situation is more complex, as sink activity is not only dependent on the amount of glucose sensed extra- and intracellularly. Hexokinase activity may play a key role in intracellular glucose sensing (Jang et al. 1997). Extracellular glucose sensing may be executed by membrane-bound systems resembling the SNF3/RTG2 sensors in yeast (Öczan, Dover & Johnston 1998).

DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Different functional zones of the phloem

The phloem system in angiosperms comprises three sequential sectors each of which executes a specific task (van Bel 1996). The phloem collects photosynthate in the minor veins of mature leaves (collection phloem), distributes photosynthate between the centres of growth and storage via the phloem in main veins, petioles, stems, peduncles, main roots, etc. (transport phloem), where photosynthate is released into expanding or accumulating cells of various sinks (release phloem) (Fig. 7). Co-ordination and integration of these tasks result in an orchestrated machinery distributing photo-assimilates in a way that is appropriate for the stage of growth in keeping with the environmental conditions.

image

Figure 7. The proportional volumes of SE and CC in the respective phloem zones and the local fluxes of photo-assimilate and water (modified after van Bel & Ehlers 2000). A dynamic version of the Münch mass flow model. Photo-assimilates are translocated via the phloem through essentially leaky instead of hermetically sealed pipes. The solute content, and implicitly the turgor, are controlled by release/retrieval mechanisms in the sieve element/companion cell complexes (SE/CC complexes). Differential release/retrieval balances control the influx/efflux of sugars (closed arrows) and water (open arrows) in the various phloem zones. In the collection phloem (phloem loading), the influx will dominate, in the release phloem (phloem unloading) the efflux. In the transport phloem having a dual task (nourishment of axial and terminal sinks), the balance between influx and efflux varies with the requirements of the plant. The gradual loss of solute and commensurate amounts of water towards the sink, where massive delivery of water and solutes takes place, has been ascribed to a proton-motive force gradient in the SE/CC complexes from source to sink (van Bel 1995). Alternatively, the relative size reduction of companion cells along the source-to-sink path may explain a decreasing retrieval capacity of the SE/CC complexes in the direction of the sink.

Download figure to PowerPoint

Incidentally, it should be noted that most of the experimental evidence for interaction between SE and CC was obtained using transport phloem. There, SEs are the largest and therefore the most amenable to manipulation or accessible to experimentation (e.g. Daie 1987; Grimm et al. 1990; van Bel & van Rijen 1994; Kempers & van Bel 1997).

Size ratio between sieve element and companion cell

The structure of the SE/CC complex seems to reflect the local phloem function (Fig. 7). The decreasing volume ratio between CCs and SEs along the phloem stretch (Fig. 7) may be related with a decreasing energy requirement for photosynthate retention in collection, transport, and release phloem, respectively (van Bel 1996). Although the CCs are much larger than SEs in collection phloem, where they often enclose the SEs, CCs only partly cover the SE surface in transport phloem (at least in some evolutionarily advanced dicotyledons; van Bel 1996), and are again smaller or missing in release phloem (Offler & Patrick 1984; Schulz 1994). The SE : CC ratios indicate that transport phloem is a functional hybrid between collection and release phloem which is consistent with balanced release/retrieval processes along the transport path (Minchin & Thorpe 1987).

Ultrastructure of the companion cell

Particularly in collection phloem, the ultrastructure of CCs is diverse (Fig. 8). Three classes of CCs having a dense cytoplasm in common are distinguished: intermediary cells (ICs) with numerous vesicles of unknown nature, transfer cells (TCs) with numerous cell wall invaginations and ordinary or smooth-walled companion cells (SCs) without special characteristics (Gamalei 1989; Turgeon 1996). The differences between CCs in collection phloem continue in a less conspicuous manner in transport phloem (Kempers et al. 1998).

image

Figure 8. Diagrams of the sieve element (SE)/companion cell complexes in the minor veins of angiosperm leaves (modified after van Bel 1999). Three types of companion cells are distinguished on the basis of subcellular structure and plasmodesmal density at the interface with the adjacent parenchyma cells. A, Intermediary cells, IC (type 1 according to the classification of Gamalei 1989) contain fragmented vacuoles of vesicles of unknown identity. B, Smooth-walled or ordinary companion cells, SC (types 1–2a, 2a according to the classification of Gamalei 1989) show the basic features of a companion cell (Turgeon 1996; Turgeon & Medville 1998). C, Transfer cells, TC (type 2b according to the classification of Gamalei 1989) possess conspicuous cell wall protrusions. During evolution, a functional and structural uncoupling of the sieve element/companion cell complexes may have taken place as is speculated here. Originally, sugar transport from the mesophyll to the sieve elements occurred by diffusion along a sugar concentration gradient through a symplasmic continuum in the species with SCs (B). In the next stage, sugar metabolism in production (mesophyll) compartment was functionally uncoupled from the transport compartment in species with ICs (A). In species with TCs, the sugar production compartment and the transport compartment were functionally and structurally uncoupled (C). The latter development enabled phloem loading against a steep sugar concentration gradient. The evolution towards sugar-concentrating species probably occurred several times in parallel (cf. Turgeon et al. 2001)

Download figure to PowerPoint

Plasmodesmal frequency between companion cells and adjacent parenchyma

In collection phloem, plasmodesmal frequencies between CCs and adjacent leaf parenchyma (LP) vary by a factor 1000 (Fig. 8). At the IC/LP interface (Fig. 8A) the density varies between 10 and 60 plasmodesmata µm−2, at the SC/LP interface (Fig. 8B) between 10 and 0·1 plasmodesmata µm−2; the density is mostly lower than 0·01 plasmodesmata µm−2 at the TC/PP interface (Fig. 8C) (Gamalei 1989, 1991). The SE/IC complexes (type 1, Gamalei 1989) are mostly associated with symplasmic phloem loading against a sugar gradient, the SE/TC complexes (type 2b, Gamalei 1989) mostly with apoplasmic phloem loading against a sugar gradient (Turgeon & Wimmers 1988; van Bel et al. 1992, van Bel, Ammerlaan & van Dijk 1994). It seems that SE/SC complexes (types 1–2a and 2a, Gamalei 1989) are associated alternatively with symplasmic phloem loading along a sugar gradient or with apoplasmic phloem loading (van Bel et al. 1992; van Bel 1999; Schrier 2001), but more detailed work is required. It appears that plasmodesmal frequency at the interface between CCs and adjoining parenchyma cells is not the only determinant of the mode of phloem loading. The identity of transport sugar(s) are just as important as the ultrastructure of the SEs for assessment of the mode of phloem loading (cf. Schrier 2001; Lalonde et al., this issue).

On the basis of plasmodesmal frequencies and the nature of the sugars translocated (Fig. 8), one may speculate on three evolutionary steps in phloem loading (van Bel 1999; Schrier 2001). The cornerstone of this speculation is that phloem loading occurred symplasmically in primitive angiosperms as seems to be the case in ferns and gymnosperms, possibly along a concentration gradient being the simplest solution. According to this concept, the photosynthesizing compartment was originally tightly linked to the SE/SC complex (Fig. 8B) and loading (or better ‘filling’) of sieve tubes happened along a concentration gradient (cf. Turgeon & Medville 1998). In the next stage (Fig. 8A), the sugar metabolism of mesophyll and SE/IC complex was uncoupled (Turgeon 1991; Haritatos, Ayre & Turgeon 2000), so that sugars can be accumulated by the SE/IC complexes against a concentration gradient. In the ultimate evolutionary stage (Fig. 8C, mesophyll and SE/TC complex operate virtually uncoupled as the symplasmic connectivity between both domains is largely reduced. The uphill sugar gradient may be steeper than in the previous type. It seems that the three types of phloem loading are all represented abundantly among the present dicotyledonous families (Gamalei 1989; Turgeon, Medville & Nixon 2001).

Projection of the phloem loading types on a phylogenetic tree of dicotyledon evolution (Soltis et al. 2000) does not seem to support this speculation. That evolution of phloem loading was not a single, but a multiple event in evolution does not need to be inconsistent with the above speculation (Turgeon et al. 2001). More seriously, however, phloem loading by Liriodendron of the ancient Magnoliaceae (Soltis et al. 2000) was shown to be apoplasmic (Goggin, Medville & Turgeon 2001). Moreover, symplasmic phloem loading along a concentration gradient was observed in Salix (Turgeon & Medville 1998) which is not at all ancestral in any case (Soltis et al. 2000). Obviously this issue requires further research as only a few species of a minor part of the dicotyledonous families have been investigated. To reach conclusive concepts, ultratructural, physiological and sugar-analytical data of dicots and monocots has to be compared systematically in the framework of evolution.

In comparison with collection phloem, few data on the connectivity between SE/CC complex and adjacent phloem parenchyma cells (PPs) are available for transport and release phloem. The interspecific variety in plasmodesmal connectivity in transport phloem is expected to be smaller than in collection phloem given the higher environmental strain upon leaf tissues. In transport phloem, plasmodesmal density at the interface between SE/CC complex and adjacent parenchyma varies by a factor of 10 in the few species investigated (Kempers et al. 1998) contrasting a difference by a factor of 1000 in their collection phloem (Gamalei 1989). Further, plasmodesmal frequencies between CCs and PPs are low in comparison with those at the other interfaces (Kempers et al. 1998) with virtually no plasmodesmata between SEs and PPs.

As for release phloem, plasmodesmal connectivity between SE and PP is relatively high as for instance in cereal caryopses (Oparka & Gates 1981) and in root apices (Warmbrodt 1985, 1986). However, symplasmic unloading in terminal sinks may not be universal given the symplasmic isolation of SEs in developing maize leaves (Evert & Russin 1993). The gating behaviour of plasmodesmata in collection, transport and release phloem may be different and significant for directing the nutrients to growing organs (Lalonde et al. in press). Symplasmic control via plasmodesmata is complemented by a control over solute exchange between apoplast and symplast by means of plasma membrane proteins (see Lalonde et al. in press).

PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Single units doing a collective job

As has been set out before, the presence of a plasma membrane is an absolute prerequisite for functioning of the SEs. In order to generate and control the osmotic potentials necessary for mass flow, the SE plasma membrane must be equipped with the appropriate substrate carriers, ion pumps, ion channels and aquaporins. Over the past decennium, several phloem-specific plasma-membrane-bound proteins have been identified (see also Patrick et al. 2001; Lalonde et al. in press). Among them are proton pumps (Bouché-Pillon et al. 1994; DeWitt & Sussman 1995; Moriau et al. 1999; Zhao et al. 2000; Langhans et al. 2001), sucrose carriers (Riesmeier, Hirner & Frommer 1993, Riesmeier, Willmitzer & Fromer 1994; Truernit & Sauer 1995; Stadler et al. 1995; Stadler & Sauer 1996; Lemoine et al. 1996; Kühn et al. 1996, 1997; Barker et al. 2000; Gottwald et al. 2000; Noiraud, Delrot & Lemoine 2000; Weise et al. 2000), mannitol carriers (Noiraud, Maurousset & Lemoine 2001), potassium channels (Marten et al. 1999; Phillipar et al. 1999; Deeken et al. 2000; Lacombe et al. 2000), calcium channels (Volk & Franceschi 2000), and aquaporins (Robinson et al. 1996; Schäffner 1998; Tyerman et al. 1999).

Deployment of carriers, pumps and channels in collection phloem

So far, molecular studies on phloem loading have been carried out using exclusively apoplasmically loading species. Hence, studies have focused on sucrose carriers and proton pumps. Diverse molecular strategies (see also Lalonde et al. in press) have revealed a set of sucrose/H+-symporters involved in loading and export. Insertion of an apoplasmic yeast invertase gene or by antisense repression of sucrose/H+-symporters impaired sucrose uptake by collection phloem. Blockage of phloem loading led to a massive increase of soluble sugars and starch in the mesophyll (e.g. Riesmeier et al. 1994; Kühn et al. 1996; Lemoine et al. 1996; Schulz et al. 1998; Gottwald et al. 2000). In studies focused on the location, sucrose carriers turned out to be associated with the veins (Noiraud et al. 2000) or were shown to occur in the minor veins (Riesmeier et al. 1993; Truernit & Sauer 1995). A higher spatial resolution of the distribution of sucrose carriers between cell types has not been reached for collection phloem.

As for the cellular location of proton pumps, EM-immunocytochemistry revealed their overwhelming presence in CCs of collection phloem of Vicia (Bouché-Pillon et al. 1994). At the interface between TC and SE, the amount of H+-ATPases was five times higher at the CC-side. Similarly, pm4 (for plasma membrane H+-ATPase 4) was notably expressed in CCs of minor veins of source leaves of Nicotiana plumbaginifolia (Moriau et al. 1999). Co-suppression of pm4 led to an increase in the leaf sugar content and a decrease of sucrose in phloem exudate of Nicotiana plumbaginifolia (Zhao et al. 2000). Co-suppression was much stronger in source than in sink leaves. Collectively, the results indicate the involvement of H+-ATPase in phloem loading by fuelling H+/sucrose symporters.

To my knowledge, no convincing molecular evidence has been presented thus far for the cellular localization of amino acid transporters in phloem tissues (Fischer et al. 1998).

Uptake of photo-assimilates by SE/CC complexes coincides with compensatory ion fluxes and mass movement of water (Smith & Milburn 1980a, b). The steep osmotic gradient at the SE/CC-border in the collection phloem induces water fluxes that require very high permeability constants which can only be accommodated by high densities of aquaporins (Tyerman et al. 1999). Indeed, aquaporins have been found to be associated with minor veins CCs of Arabidopsis (Robinson et al. 1996; Schäffner 1998).

Deployment of carriers, pumps and channels in transport phloem

Contrasting observations regarding the deployment of sucrose carriers have been obtained for transport phloem that is more suitable for a spatial resolution. Using strategies in which GUS was co-expressed with a SUC2 carrier, the carriers appeared to be deployed on the CC plasma membrane of Arabidopsis and Plantago (Stadler et al. 1995; Stadler & Sauer 1996). Recently, a weakly expressed SUC1 carrier was discovered on the SE membrane of Arabidopsis (R. Stadler et al. unpubl. results). In solanaceaous species, the sucrose carriers SUT1 – equivalent to SUC2 – (Kühn et al. 1997), SUT2 (Barker et al. 2000), and SUT4 (Weise et al. 2000) are positioned onto the SE plasma membrane as evidenced by immunofluorescence. Localization of sucrose carriers on the CC plasma membrane (Stadler et al. 1995; Stadler & Sauer 1996) lends support to a model in which sucrose is mainly retrieved by the CC (Fig. 6A). Predominant occurrence of the sucrose carriers on the SE plasma membrane (Kühn et al. 1997; Barker et al. 2000; Weise et al. 2000) rather favours an energy supply by CCs and photo-assimilate retrieval by SEs (Fig. 6B & C). It should be noted that several authors make confusing claims with regard to phloem cells engaged in ‘phloem loading’. Not only SEs in petioles and stems, but also those in main veins belong to the transport phloem (van Bel 1993). Evidence obtained with transport phloem may not be representative for events in collection phloem, the site of phloem loading. Unequivocal assessment of the vein order is required as results obtained with lower-order veins rather pertain to retrieval by the phloem.

The functional assessment of a PMF-driven mannitol carrier (AgMaT1) in celery (Noiraud et al. 2001) illustrates the difficulties in interpreting molecular-biological results if the exact location of the carriers is not known. Although the mannitol carriers reside unmistakably in petiole phloem, it is uncertain if the carrier is responsible for mannitol retrieval along the pathway or for storage by phloem parenchyma. It is not excluded hat AgMaT1 is located in phloem parenchyma cells given the PMF-dependence of mannitol uptake by plasma membrane vesicles of storage parenchyma (Salmon et al. 1995).

As for transport phloem, immunolocalization showed the presence of an epitope-tagged H+-ATPase (AHA3) localized to CCs in stems of Arabidopsis (DeWitt & Sussman 1995). Similarly, members of the anti-CIF2 immunoreactive H+-ATPases subfamily were detected on the CC plasma membrane by immunogold labelling (DeWitt et al. 1996). However, polyclonal antibodies raised against a conserved H+-ATPase sequence were not able to detect H+-ATPase in SEs (DeWitt & Sussman 1995). In contrast, Langhans et al. (2001) demonstrated different H+-ATPase isoforms on the plasma membrane of both CCs and SEs in transport phloem of Ricinus communis and Cucurbita pepo.

A detailed positioning of potassium channels has not been achieved yet. The ak2/ak3 gene (for the AK2/AK3 potassium channel) is expressed in collection and transport phloem of Arabidopsis thaliana (Marten et al. 1999; Deeken et al. 2000, Lacombe et al. 2000). This channel may responsible for potassium uptake in collection phloem and potassium release/retrieval along the phloem pathway (Lacombe et al. 2000).

Remarkably, phloem-specific (DHP)-type calcium channels reside exclusively on the SE plasma membrane in minor and major veins of Nicotiana tabacum and Pistia stratiotes (Volk & Franceschi 2000). Using immofluorescence or immunocytochemistry no labelling was observed to be associated with the CCs.

Deployment of carriers, pumps and channels in release phloem

Transcripts of sucrose and monosaccharide transporters have been detected in vascular tissues of sinks such as root tips (Truernit et al. 1996), sink leaves (Barker et al. 2000; Noiraud et al. 2000, 2001; Weise et al. 2000), and flowers (Truernit et al. 1996; Barker et al. 2000; Büttner et al. 2000). Although reporter gene expression or in situ hybridization has demonstrated localization of transcripts to protophloem (Stadler et al. 1995), no clear cellular localization has been achieved for release phloem. As for proton pumps, H+-ATPase was found to be mainly associated with the phloem in seedling roots (DeWitt, Harper & Sussman 1991).

Necessity of cell-specific localization of membrane-bound proteins

The spatial arrangement of SE/CC complexes along the phloem path and the increasing SE/CC volume ratios from sources to sinks predict that CCs become less dominant towards the sinks. It is likely therefore that the molecular outfit of the SE/CC plasma membrane will shift along the phloem pathway. This contention can only be verified by systematic investigation of cell-specific occurrence of the respective plasma-membrane proteins. Thus far, most investigations on membrane-bound proteins in the phloem have focused on showing their association with the phloem – hardly any with specific cell types within the phloem of various zones – in apoplasmically loading species only.

DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

Photosynthate is fuel and cargo at the same time

According to the classic Münch concept, photo-assimilates amass in the sieve tubes of sources and escape from the sieve tubes in sinks. The resultant turgor difference drives the mass flow (Fig. 5). The ingenuity of the phloem system thus is that fuel and cargo are identical! Over the past 10 years, the original Münch model has been modified marginally (e.g. van Bel 1995) with important implications, though, for water management in the phloem.

Generation of the driving force in the source

The evidence for at least two (Turgeon & Wimmers 1988, van Bel et al. 1992, 1994) or probably three (Turgeon & Medville 1998) may have profound consequences for the generation of turgor potential (Fig. 9). The apoplasmic mode of phloem loading (cf. Lalonde et al. in press) is highly consistent with the Münch concept; SE/CC complexes amass photo-assimilates and correspondingly high amounts of apoplasmic water (Fig. 9C). At first sight, the other modes seem to be less compatible with the original Münch model.

image

Figure 9. Speculations on the relationship between phloem loading mode and leaf water economy (drawn after Schrier 2001). A, In phloem loading occurring along a sugar gradient (cf. Fig. 8B), water to sustain phloem transport is to be collected by the mesophyll cells and the terminal sieve element/companion cell complexes. B, As the production compartment become functionally detached from the mesophyll, photosynthate accumulation by the sieve tubes will increase (cf. Fig. 8A). C, A high osmopotential of sieve tubes which are functionally and structurally detached from the mesophyll, implies that the water needed to drive mass flow is mainly taken up directly by the sieve element/companion cell complexes (cf. Fig. 8C). This short-cut of water uptake (C) as compared to species with down-hill (A) or weakly (B) up-hill sugar gradients from mesophyll to sieve elements, makes water uptake more efficient and phloem transport less vulnerable to water shortage. The size of the arrowheads are indicative of the water flux rates.

Download figure to PowerPoint

In the symplasmic mode of phloem loading, build-up of a sugar gradient may be less efficient with correspondingly lower rates of water uptake resulting in lower pressure in the phloem system (cf. van Bel 1996). Moreover, it deserves to be considered that, in these species, symplasmic connectivity between SE/CC complexes and mesophyll is intense and that much of the water entering the sieve tubes comes from the mesophyll through the plasmodesmata (Fig. 9A & B). Such a symplasmic water loading must prevail in species in which no phloem loading sensu stricto (against a concentration gradient) takes place (Turgeon & Medville 1998). A mesophyll sugar gradient as evidenced for barley (Koroleva et al. 1998) would not only drive sugars to the SE/CC complexes, but may also set up a corresponding turgor gradient through the mesophyll.

Thinking along these lines, water for mass flow propulsion is withdrawn from the whole leaf apoplast in species without distinct accumulation by minor vein phloem (Fig. 9A) (Turgeon & Medville 1998). In species with the other modes of phloem loading (Fig. 9B & C), photosynthate accumulation by SE/CC complexes in the minor veins render the sieve tubes more competitive in withdrawing water from the xylem vessels (Fig. 9B & C). The accomplished short-cut of water supply seems highly beneficial for phloem transport in times of water shortage in the mesophyll (Fig. 9C). Apoplast compartmentation achieved by water-impermeable barriers in the cell walls would radically invalidate these speculations which all rely strongly on free movement of water in the apoplast. An experimental approach to the postulate is difficult, as studies on water microflows are minefields of methodical errors (Canny 1990). Provided that the present conjectures turn out to be correct, physiological and structural detachment of the mesophyll and the conducting compartment may be interpreted as an evolutionary step forward in the battle for water engaged in the propulsion of mass flow (Schrier 2001; Fig. 9).

It is even worth considering whether pressure is involved in symplasmic sugar transport in the pre-phloem pathway. Additional pressure was invoked to explain discrepancies between calculated flux rates and the size of the post-phloem plasmodesmal corridors (see Lalonde et al. in press). A pressure-driven bulk flow through plasmodesmata from SEs to the surrounding parenchyma cells has been proposed for phloem unloading in wheat grains (Fisher & Cash-Clark 2000). Given the immense pressures needed to pressurize fluids through a glass capillary tip with a tip diameter of 100 nm (Knoblauch et al. 1999), however, it is questionable whether pressure through corridors of 2–3 nm in diameter is a viable option here, unless the hydraulic properties of plasmodesmata are radically different from those of glass microcapillaries.

Regulation of the driving force in the path

The second modification of the Münch model pertains to transport phloem. In the original, strongly physicochemical concept, sieve tubes were regarded to be hermetically sealed pipes. Actually, SE/CC complexes along the pathway in herbs (major veins, petioles, stems and major roots make up more than 99% of the phloem stretch, van Bel 1995) lose appreciable amounts of photosynthate, part of which is retrieved (Eschrich, Evert & Young 1972; Aloni, Wyse & Griffith 1986; Minchin & Thorpe 1987; Grimm et al. 1990; Schulz 1994). In Phaseolus, for instance, sieve tubes lose 6% of photosynthate per centimetre of stem, of which about two-thirds is retrieved (Minchin & Thorpe 1987). Volume flow (Eschrich et al. 1972), the more dynamic version of mass flow, reflects the dual function of transport phloem. Retention of photosynthate to nourish terminal sinks at the end of the pathway coincides with release to supply axial sinks along the pathway (van Bel 1996). The dualism in function is achieved by a rigorously regulated release/retrieval balance in the SE/CC complexes of the transport phloem. A dynamic relay mechanism as the volume flow is able to counteract potential pressure loss caused by the bottlenecks such as the sieve pores. Release and retrieval means that pressure is continuously lost and built up, sustained by countless aquaporins (Schäffner 1998; Tyerman et al. 1999) along the sieve tube path.

Release/retrieval events are controlled by carrier systems. Sucrose carrier kinetics has been measured using strips of isolated phloem (Wright & Fisher 1981; Daie 1987; van der Schoot & van Bel 1989; Grimm et al. 1990). The carriers are energized by proton-motive force (PMF) (Wright & Fisher 1981; van der Schoot & van Bel 1989). Differences in PMF in successive SE/CC complexes along the pathway could create a decreasing retrieval/release balance along the phloem path. Such a gradient would impose a decreasing turgor toward the sinks. Circumstantial evidence for a decreasing PMF from leaves to roots was reported for Phaseolus and Ricinus. In Phaseolus, a tip-to-base increase of the apoplasmic sugar was observed (Minchin & Thorpe 1984). The tip-to-base increase of sugar in the phloem apoplast is consistent with a tip-to-base decrease of pH with a corresponding drop of the sugar concentration in sieve tube sap (Vreugdenhil & Koot-Gronsveld 1989). By contrast, investigations on the membrane potential in successive internodes of Lupinus were inconclusive with regard to the existence of a PMF-gradient along the phloem pathway (van Bel & van Rijen 1994).

The strict control of release/retrieval processes seems to require a certain degree of symplasmic isolation of SE/CC complexes in transport phloem. There is a range of indications supportive of a symplasmic autonomy of SE/CC complexes in transport phloem: (a) a relative paucity of plasmodesmal connections at the interface between SE/CC complexes and phloem parenchyma cells (PPs) in stems of Phaseolus (Hayes, Offler & Patrick 1985), Lythrum, Cucurbita, Vicia and Zinnia (Kempers et al. 1998); (b) fluorochromes either injected into sieve tubes of Ricinus (van Bel & Kempers 1991), Solanum (Oparka et al. 1992), Lupinus (van Bel & van Rijen 1994), and tomato (Rhodes et al. 1996) or applied to leaves of Arabidopsis (Oparka et al. 1994, Oparka, Prior & Wright 1995) and Vicia (Knoblauch & van Bel 1998) stay contained within SE/CC complexes of transport phloem; (c) the electric conductance between adjacent PPs is at least 10 times higher than that at the interface between SE/CC complex and PP (van Bel & van Rijen 1994); and (d) the membrane potentials of SE/CCs and PPs in transport phloem often differ by more than 20 mV in the same plant (van der Schoot & van Bel 1989; van Bel & Kempers 1991; van Bel & van Rijen 1994). The distinct voltage difference points to electrical isolation of the SE/CC complexes from the surrounding tissues.

Other observations, however, contradict the symplasmic autonomy of the SE/CC complexes in transport phloem: (a) the loss of 5,6-carboxyfluorescein from the sieve tubes in bean stems under sink-limiting conditions (Patrick & Offler 1996); and. (b) PCMBS- and CCCP-treatments induce a release of photosynthates along isolated petioles of Cyclamen, but not in intact plants (Grimm, Jahnke & Rothe 1997). The experiments seem to indicate collectively that SE/CC complexes are able to shift between symplasmic and apoplasmic routes by gating their plasmodesmal connections towards the PPs (see Lalonde et al., this issue).

Maintenance of the driving force by the sinks

At the sink end of the phloem system, two modes of phloem unloading may operate, either single, in series or in parallel (Lalonde in press). For most sinks, phloem unloading follows symplasmic routes. Photo-assimilates are unloaded from the SEs into arrays of sink cells connected by plasmodesmata (e.g. Warmbrodt 1985, 1986). A minor component of SE unloading will be apoplasmic, driven by large transmembrane sugar concentration differences (Patrick 1990). Rates of apoplasmic SE unloading will be relatively small given the limited plasma membrane surface areas of SEs or SE/CC complexes in the unloading zone (Patrick 1997; Schulz 1998). Yet, apoplasmic sugar may play a key role in phloem unloading. The same holds for phloem unloading modes in which the symplasmic unloading pathway is interrupted by an apoplasmic step in the post SE route. This occurs where photo-assimilate transport takes place across genomic interfaces between generations of developing seeds, mutualistic biotrophic relationships and phloem parasites (Lalonde in press). The symplasmic discontinuity imposes an apoplasmic passage of the photo-assimilates.

Irrespective of the unloading pathway, apoplasmic concentrations have an effect on local water relations and turgor. A high apoplasmic osmolarity would induce a loss of water from the SEs resulting in a local turgor drop and a steeper turgor gradient between source and sink. High apoplasmic osmolarities could also stimulate symplasmic unloading by creating a deficit of intracellular water sucking the solvent from SEs to sink cells through the symplasmic continuum (Fisher & Cash-Clark 2000). In keeping with this idea, increased unloading rates were attained in roots bathed in concentrated sugar solutions (Dick & ap Rees 1975; Schulz 1994), but the phenomenon can also be explained by widening of the plasmodesmal corridors in the unloading pathway in response to turgor relaxation (Schulz 1995). How logical they may sound for the wide sink plasmodesmata (up to 10 nm in diameter; Fisher & Cash-Clark 2000), these concepts conflict with powers in the order of 5–10 MPa required to pressurize water through 100 nm corridors (Knoblauch et al. 1999). In conclusion, the impact of extracellular osmolarity on the rate of unloading contrasts Münch's postulate that the photo-assimilate gradient to the apoplast should be as steep as possible for a rapid unloading (Münch 1930).

It should be noted that release of water in sinks is intensified when the carriers that deliver osmotic components from SEs in the sinks, are activated at low turgor values (Patrick 1997; Patrick et al. 2001). Part of the water released is used for volume growth of sink cells (e.g. Pritchard, Winch & Gould 2000). The excess water which is set free into the sink apoplast is probably re-circulated by the xylem (Pate et al. 1985; Köckenberger et al. 1997).

CONCLUSIONS AND PROSPECTS

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES

We are beginning to recognize the complexity of phloem functioning at different structural and physiological levels; the phloem is an example of evolutionary ingenuity. Viewing the stage (cf. van Bel, Ehlers & Knoblauch 2002), a number of focus points can be identified for future phloem research.

  • 1
    The evolution of SEs is still obscure. Much about SE evolution can probably be learned from the physiology of parenchyma cells adjoining SEs in mosses and ferns. Special emphasis must be laid on the increasing support to SEs either by Strasburger cells or by CCs during evolution as it may contribute to understand SE–CC interaction.
  • 2
    A vast area of SE cell biology is virtually unknown. The role of the sieve-element plastids, mitochondria, and the stacked ER needs elucidation. Further, the metabolic tasks of the multitude of soluble proteins require a thorough study as well as the question as to whether structural sieve-element proteins possess functions other than sealing sieve pores.
  • 3
    It is expected that continued research on exchange of materials and messages between SEs and CCs will reveal a number of, possibly surprising, features. The mechanism(s) of macromolecular trafficking via the PPUs is one of the focus areas to be studied in detail.
  • 4
    Exchange of macromolecules between SE and CC has profound consequences for the significance of long-distance signalling in the sieve tubes. In view of the exciting prospects, potential remote control by sugars and macromolecules deserves comprehensive research efforts.
  • 5
    Pathways and modes of phloem loading are largely uncharted. The pre-SE pathway of photo-assimilate transport in various types of phloem loading and the mode(s) of cell-to-cell transport in mesophyll are practically unknown. The modes of SE loading, in particular symplasmic phloem loading, require detailed investigations. An exact localization of the membrane proteins involved in phloem loading is fundamental to further understanding of the respective loading modes. It is also necessary to study the consequences of loading modes for leaf water management and generation of pressure flow.
  • 6
    As for transport phloem, the interplay between SE/CC complexes and adjacent phloem parenchyma deserves ample attention. In this respect, precise localization of membrane proteins would be helpful to identify the competition for solutes between phloem cells which may play a key role in photo-assimilate distribution.
  • 7
    With regard to phloem unloading, regulation of plasmodesmal gating in the unloading pathway and the corresponding water fluxes through the symplasmic bottlenecks are hot topics. Localization of membrane transport proteins, especially those on the membranes lining the apoplasmic space between genetic interfaces, requires further investigation. The importance of sugar sensing for phloem physiology and its relation to sugar metabolism in sinks and sources has to be substantiated. Furthermore, knowledge on the structural frame of sinks should be extended.
  • 8
    A point largely ignored thus far is the relationship between phloem physiology and environment. What is the impact of environmental factors on phloem transport and are the modes of phloem loading adapted to specific climatic conditions?

REFERENCES

  1. Top of page
  2. ABSTRACT
  3. INTRODUCTION
  4. ORIGIN, EVOLUTION AND PHYSIOLOGICAL FITNESS OF THE PHLOEM
  5. GENERAL FUNCTIONS OF THE PHLOEM
  6. SIEVE ELEMENT/COMPANION CELL COMPLEX, THE FUNCTIONAL UNIT OF PHLOEM CONDUITS IN ANGIOSPERMS
  7. PHYLOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  8. ONTOGENY OF THE SIEVE ELEMENT/COMPANION CELL COMPLEX
  9. STRUCTURAL EQUIPMENT OF THE SIEVE ELEMENT
  10. INTERACTION BETWEEN SIEVE ELEMENT AND COMPANION CELL
  11. LONG-DISTANCE SIGNALLING IN THE PHLOEM
  12. DIVERSITY OF SIEVE ELEMENT/COMPANION CELL COMPLEXES
  13. PLASMA-MEMBRANE-BOUND PROTEINS IN SIEVE ELEMENT/COMPANION CELL COMPLEXES
  14. DRIVING FORCES FOR MASS FLOW IN THE PHLOEM, THE COLLECTIVE POWER OF SINGLE UNITS
  15. CONCLUSIONS AND PROSPECTS
  16. ACKNOWLEDGMENTS
  17. REFERENCES
  • Aloni R. & Barnett J.R. (1996) The development of phloem anastomoses between vascular bundles and their role in xylem regeneration after wounding in Dahlia and Cucurbita. Planta 198, 595603.
  • Aloni R., Wyse R.E. & Griffith S. (1986) Sucrose transport and phloem unloading in the stem of Vicia faba: possible involvement of a sucrose carrier and osmotic regulation. Plant Physiology 81, 482486.
  • Arsanto J.P. (1986) Ca2+ binding sites and phosphatase activities in sieve element reticulum and P-protein of chickpea phloem: a cytochemical and X-ray microanalysis survey. Protoplasma 132, 160171.
  • Balachandran S., Xiang Y., Schobert C., Thompson G.A. & Lucas W.J. (1997) Phloem sap proteins from Cucurbita maxima and Ricinus communis have the capacity to traffic cell to cell through plasmodesmata. Proceedings of the National Academy of Sciences USA 94, 1415014155.
  • Barclay G.F., Oparka K.J. & Johnson R.P.C. (1977) Induced disruption of the sieve element plastids in Heracleum mantegazzianum L. Journal of Experimental Botany 28, 709717.
  • Barker L., Kühn C., Weise A., Schulz A., Gebhardt C., Hirner B., Hellmann H., Schulze W., Ward J.M. & Frommer W.B. (2000) SUT2, a putative sucrose sensor in sieve elements. Plant Cell 12, 11531164.
  • Beebe D.U. & Russin W.A. (1999) Plasmodesmata in the phloem-loading pathway. In Plasmodesmata. Structure, Function, Role in Cell Communication (eds A.J.E.Van Bel & W.J.P.Van Kesteren), pp. 261293. Springer, Berlin, Germany.
  • Behnke H.-D. (1991a) Distribution and evolution of forms and types of sieve-element plastids in the dicotyledons.ALISO 3, 167182.
  • Behnke H.-D. (1991b) Non dispersive protein bodies in sieve elements: a survey and review of their origin, distribution and taxonomic significance.International Association of Wood Anatomists Bulletin 12, 143175.
  • Behnke H.-D. & Sjolund R.D. (1990) Sieve Elements. Comparative Structure, Induction and Development. Springer, Berlin, Germany.
  • Van Bel A.J.E. (1993) The transport phloem. Specifics of its functioning. Progress in Botany 54, 134150.
  • Van Bel A.J.E. (1995) The low-profile directors of carbon and nitrogen economy in plants: parenchyma cells associated with translocation channels. In Plant Stems. Physiology and Functional Morphology. (ed. B.L.Gartner), pp. 205222. Academic Press, San Diego, CA, USA.
  • Van Bel A.J.E. (1996) Interaction between sieve element and companion cell and the consequences for photoassimilate distribution. Two structural hardware frames with associated software packages in dicotyledons? Journal of Experimental Botany 47, 11291140.
  • Van Bel A.J.E. (1999) Evolution, polymorphology and multifunctionality of the phloem system. Perspectives in Plant Ecology, Evolution and Systematics 2, 163184.
  • Van Bel A.J.E. & Ehlers K. (2000) Symplasmic organization of the transport phloem and implications for photosynthate transfer to the cambium. In Cell and Molecular Biology of Wood Formation (eds R.Savidge, J.R.Barnett & R.Napier), pp. 8599. Bios, Oxford, UK.
  • Van Bel A.J.E. & Kempers R. (1991) Symplastic isolation of the sieve element-companion cell complex in the phloem of Ricinus communis and Salix alba. Planta 183, 6976.
  • Van Bel A.J.E. & Kempers R. (1997) The pore/plasmodesm unit; key element in the interplay between sieve element and companion cell. Progress in Botany 58, 278291.
  • Van Bel A.J.E. & Knoblauch M. (2000) Sieve element and companion cell: the story of the comatose patient and the hyperactive nurse. Australian Journal of Plant Physiology 27, 477487.
  • Van Bel A.J.E. & Van Rijen H.V.M. (1994) Microelectrode-recorded development of the symplasmic autonomy of the sieve element/companion cell complex in the stem phloem of Lupinus luteus L. Planta 192, 165175.
  • Van Bel A.J.E., Ammerlaan A. & Van Dijk A.A. (1994) A three-step screening procedure to identify the mode of phloem loading in intact leaves. Evidence for symplasmic and apoplasmic phloem loading associated with the type of companion cell. Planta 192, 3139.
  • Van Bel A.J.E., Ehlers K. & Knoblauch M. (2002) Sieve elements caught in the act. Trends in Plant Science 7, 126132.
  • Van Bel A.J.E., Gamalei Y.V., Ammerlaan A. & Bik L.P.M. (1992) Dissimilar phloem loading in leaves with symplastic and apoplastic minor vein configurations. Planta 186, 518525.
  • Blackman L.M., Boevink P., Santa Cruz S., Palukaitis P. & Oparka K.J. (1998) The movement protein of cucumber mosaic virus traffics into sieve elements in minor veins of Nicotiana clevelandii. Plant Cell 10, 527537.
  • Bostwick D.E., Dannenhoffer J.M., Skaggs M.I., Lister R.M., Larkins B.A. & Thompson G.A. (1992) Pumpkin phloem lectin genes are specifically expressed in companion cells. The Plant Cell 4, 15391548.
  • Botha C.E.J. & Cross R.H.M. (2000) Towards reconciliation of structure with function in plasmodesmata – who is the gatekeeper? Micron 21, 713721.
  • Botha C.E.J. & Cross R.H.M. (2001) Regulation within the supracellular highway – plasmodesma are the key. South African Journal of Botany 67, 19.
  • Bouché-Pillon S., Fleurat-Lessard P., Fromont J.-C., Serrano R. & Bonnemain J.-L. (1994) Immunolocalisation of the plasma membrane H+-ATPase in minor veins of Vicia faba in relation to phloem loading. Plant Physiology 105, 691697.
  • Büttner M., Truernit E., Baier K., Scholz-Starke J., Sontheim M., Lauterbach C., Huss V.A.R. & Sauer N. (2000) AtSTP3, a green leaf-specific, low-affinity monosaccharide-H+ symporter of Arabidopsis thaliana. Plant, Cell and Environment 23, 175184.
  • Canny M.J. (1990) What becomes of the transpiration stream? New Phytologist 114, 341368.
  • Chiou T.J. & Bush D.R. (1998) Sucrose is a signal molecule in assimilate partitioning. Proceedings of the National Academy of Sciences USA 95, 47844788.
  • Christeller J.T., Farley P.C., Ramsay R.J. & Laing W.A. (1998) Purification, characterization and cloning of an aspartic protease inhibitor from squash phloem exudate. European Journal of Biochemistry 254, 160167.
  • Citovsky V. & Zambryski P. (2000) Systemic transport of RNA in plants. Trends in Plant Science 5, 5254.
  • Clark A.M., Jacobsen K.R., Dannenhoffer J.M., Skaggs M.I. & Thompson G.A. (1997) Molecular characterization of a phloem-specific gene encoding for the filament protein, phloem protein 1 (PP1) from Cucurbita maxima. Plant Journal 12, 4961.
  • Cronshaw J. & Sabnis D.D. (1990) Phloem proteins. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 257283. Springer, Berlin, Germany.
  • Daie J. (1987) Sucrose uptake in the isolated phloem of celery is a single saturable system. Planta 171, 474482.
  • Dannenhoffer J.M., Schulz A., Skaggs M.I., Bostwick D.E. & Thompson G.A. (1997) Expression of the phloem lectin is developmentally linked to vascular differentiation in cucurbits. Planta 201, 405414.
  • Dannenhoffer J.M., Suhr S.C. & Thompson G.A. (2001) Phloem-specific expression of the pumpkin fruit trypsin inhibitor. Planta 212, 155162.
  • Davis J.D. & Evert R.F. (1970) Seasonal development of phloem in woody vines. Botanical Gazette 131, 128138.
  • Deeken R., Sanders C., Ache P. & Hedrich R. (2000) Development and light-dependent regulation of a phloem-localised K+ channel of Arabidopsis thaliana. Plant Journal 23, 285290.
  • DeWitt N.D. & Sussman M.R. (1995) Immunocytological localisation of an epitope-tagged plasma membrane proton pump (H+-ATPase) in phloem companion cells. Plant Cell 7, 20532067.
  • DeWitt N.D., Harper J.F. & Sussman M.R. (1991) Evidence for a plasma membrane proton pump in phloem cells of higher plants. Plant Journal 1, 121128.
  • DeWitt N.D., Hong B., Sussman M.R. & Harper J.F. (1996) Targeting of two Arabidopsis H+-ATPase isoforms to the plasma membrane. Plant Physiology 112, 833844.
  • Dick P.S. & Ap Rees T. (1975) The pathway of sugar transport in roots of Pisum sativum. Journal of Experimental Botany 26, 305314.
  • Ding B., Haudenshield J.S., Willmitzer L. & Lucas W.J. (1993) Correlation between arrested plasmodesmatal development and the onset of accelerated leaf senescence in yeast invertase transgenic tobacco plants. Plant Journal 4, 179189.
  • Ehlers K., Binding H. & Kollmann R. (1999) The formation of symplasmic domains by plugging of plasmodesmata: a general event in plant morphogenesis? Protoplasma 209, 181192.
  • Ehlers K., Knoblauch M. & Van Bel A.J.E. (2000) Ultrastructural features of well-preserved and injured sieve elements: minute clamps keep the phloem transport conduits free for mass flow. Protoplasma 214, 8092.
  • Eleftheriou E. (1990) Monocotyledons. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 139159. Springer, Berlin, Germany.
  • Esau K. (1969) The Phloem. Encyclopedia of Plant Anatomy, Vol. 5. Bornträger, Berlin, Germany.
  • Eschrich W., Currier H.B., Yamaguchi S. & McNairn R.B. (1965) Der Einfluss verstärkter Callosebildung auf den Stofftransport in Siebröhren. Planta 65, 4964.
  • Eschrich W., Evert R.F. & Young J.H. (1972) Solution flow in tubular semipermable membranes. Planta 107, 279300.
  • Evert R.F. (1990a) Seedless vascular plants. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 3562. Springer, Berlin, Germany.
  • Evert R.F. (1990b) Dicotyledons. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 103137. Springer, Berlin, Germany.
  • Evert R.F. & Russin W.A. (1993) Structurally phloem unloading in the maize leaf cannot be symplastic. American Journal of Botany 80, 13101317.
  • Evert R.F., Tucker C.M., Davis J.D. & Deshpande B.P. (1969) Light microscope investigation of sieve element ontogeny and structure in Ulmus americana. American Journal of Botany 56, 9991017.
  • Fagard M. & Vaucheret M. (2000) (Trans) gene silencing in plants: how many mechanisms? Annual Review of Plant Physiology and Plant Molecular Biology 51, 167194.
  • Fischer W.-N., André B., Rentsch D., Krolkiewicz S., Tegeder M., Breitkreuz K. & Frommer W.B. (1998) Amino acid transport in plants. Trends in Plant Science 3, 188195.
  • Fisher D.B. & Cash-Clark C.E. (2000) Gradients in water potential and turgor pressure along the translocation pathway during grain filling in normally watered and water-stressed wheat plants. Plant Physiology 123, 139147.
  • Fisher D.B., Wu K. & Ku M.S.B. (1992) Turnover of soluble proteins in the wheat sieve tube. Plant Physiology 100, 14331441.
  • Gamalei Y.V. (1989) Structure and function of leaf minor veins in trees and herbs. Trees 3, 96110.
  • Gamalei Y.V. (1990) Leaf Phloem. Nauka, Leningrad, Russia. (in Russian)
  • Gamalei Y.V. (1991) Phloem loading and its development related to plant evolution from trees to herbs. Trees 5, 5064.
  • Geigenberger P., Lerchl J., Stitt M. & Sonnewald U. (1996) Phloem-specific expression of pyrophosphatase inhibits long-distance transport of carbohydrates and amino acids in tobacco plants. Plant, Cell and Environment 19, 4355.
  • Goeschl J.D. & Magnuson C.E. (1986) Physiological implications of the Münch–Horwitz theory of phloem loading: effect of loading rates. Plant, Cell and Environment 9, 95102.
  • Goggin F.L., Medville R. & Turgeon R. (2001) Phloem loading in the tulip tree. Mechanisms and evolutionary implications. Plant Physiology 124, 891899.
  • Golecki B., Schulz A., Carstens-Behrens U. & Kollmann R. (1998) Evidence for graft transmission of structural phloem proteins or their precursors in heterografts of Cucurbitaceae. Planta 206, 630640.
  • Golecki B., Schulz A. & Thompson G.A. (1999) Translocation of structural P proteins in the phloem. Plant Cell 11, 127140.
  • Gottwald J.R., Krysan P.J., Young J.C., Evert R.F. & Sussman M.R. (2000) Genetic evidence for the in planta role of phloem-specific plasma membrane sucrose transporters. Proceedings of the National Academy of Sciences USA 97, 1397913984.
  • Graham L.E. (1993) The Origin of Land Plants. Wiley, New York, USA.
  • Grimm E., Bernhardt G., Rothe K. & Jacob F. (1990) Mechanisms of sucrose retrieval along the phloem path – a kinetic approach. Planta 182, 480485.
  • Grimm E., Jahnke S. & Rothe K. (1997) Photoassimilate translocation in the petiole of Cyclamen and Primula is independent of lateral retrieval. Journal of Experimental Botany 48, 10871094.
  • Haebel S. & Kehr J. (2001) Matrix-assisted desorption/ionization time of flight mass spectrometry peptide mass fingerprints and post source decay: a tool for the identification and analysis of phloem proteins from Cucurbita maxima Duch. separated by two-dimensional polyacrylamide gel electrophoresis. Planta 213, 586593.
  • Haritatos E., Ayre B.G. & Turgeon R. (2000) Identification of phloem involved in assimilate loading in leaves by the activity of the galactinol synthese promoter. Plant Physiology 123, 929937.
  • Hartmann T. (1999) Chemical ecology of pyrrolizidine alkaloids. Planta 207, 483495.
  • Hayashi H., Fukuda A., Suzui N. & Fujimaki S. (2000) Proteins in the sieve tube-companion cell complexes: their detection, localization and possible functions. Australian Journal of Plant Physiology 27, 489496.
  • Hayes P.M., Offler C.E. & Patrick J.W. (1985) Cellular structures, plasma membrane surface areas and plasmodesmatal frequencies of the stem of Phaseolus vulgaris L. in relation to radial photosynthate transfer. Annals of Botany 56, 125138.
  • Hellmann H., Barker L., Funck D. & Frommer W.B. (2000) The regulation of assimilate allocation and transport. Australian Journal of Plant Physiology 27, 583594.
  • Hsu F.C. & Kleier D.A. (1996) Phloem mobility of xenobiotics VIII A short review. Journal of Experimental Botany 47, 12651271.
  • Imlau A., Truernit E. & Sauer N. (1999) Cell-to-cell and long-distance trafficking of the green fluorescent protein in the phloem and symplastic unloading of the protein into the sinks. Plant Cell 11, 309322.
  • Iqbal M. (1995) Ultrastructural differentiation of sieve elements. In The Cambial Derivatives (ed. M.Iqbal), pp. 241270. Bornträger, Berlin, Germany.
  • Ishiwatari Y., Fujiwara T., McFarland K.C., Nemoto K., Hayashi H., Chino M. & Lucas W.J. (1998) Rice phloem thioredoxin h has the capacity to mediate its own cell-to-cell transport through plasmodesmata. Planta 205, 1222.
  • Jacinto T., McGurl B., Franceschi V., Delano-Freier J. & Ryan C.A. (1997) Tomato prosystemin promoter confers wound-inducible, vascular bundle-specific expression of the β-glucoronidase gene in transgenic tomato plants. Planta 203, 406412.
  • Jang J.-C., Leon P., Zhou L. & Sheen J. (1997) Hexokinase as a sugar sensor in higher plants. Plant Cell 9, 519.
  • Johnson R.P.C., Freundlich A. & Barclay G.F. (1976) Transcellular strands in sieve tubes: what are they? Journal of Experimental Botany 101, 11271136.
  • Jorgensen R.A., Atkinson R.G., Forster R.L.S. & Lucas W.J. (1998) An RNA-based information superhighway in plants. Science 279, 14861487.
  • Kehr J., Haebel S., Blechschmidt-Schneider S., Willmitzer L., Steup M. & Fisahn J. (1999) Analysis of phloem protein patterns from different organs of Cucurbita maxima Duch by matrix-assisted laser desorption/ioniosation time of flight mass spectroscopy combined with sodium dodecyl sulfate polyacrylamide gel electrophoresis. Planta 207, 612619.
  • Kempers R., Ammerlaan A. & Van Bel A.J.E. (1998) Symplasmic constriction and ultrastructural features of the sieve element/companion cell complex in the transport phloem of apoplasmically and symplasmically phloem-loading species. Plant Physiology 116, 271278.
  • Kempers R. & Van Bel A.J.E. (1997) Symplasmic connections between sieve element and companion cell in the stem phloem of Vicia faba L. have a size exclusion limit of at least 10 kDa. Planta 201, 195201.
  • Kim M., Canio W., Kessler S. & Sinha S. (2001) Developmental changes due to long-distance movement of a homeobox fusion transcript in tomato. Science 293, 287289.
  • Kim J.-Y., Mahé A., Brangeon J. & Prioul J.-L. (2000) A vascular maize invertase, Ivr2, is induced by water stress. Organ/tissue specificity and diurnal modulation of expression. Plant Physiology 124, 7184.
  • Knoblauch M. & Van Bel A.J.E. (1998) Sieve tubes in action. Plant Cell 10, 3550.
  • Knoblauch M., Hibberd J.M., Gray J.C. & Van Bel A.J.E. (1999) A galinstan expansion femtosyringe for microinjection of eukaryotic organelles and prokaryotes. Nature Biotechnology 17, 906909.
  • Knoblauch M., Peters W.S., Ehlers K. & Van Bel A.J.E. (2001) Reversible calcium-regulated stopcocks in legume sieve tubes. Plant Cell 13, 12211230.
  • Koch K.E. & Nolte K.D. (1995) Sugar-modelled expression of genes for sucrose metabolism and their relationship to transport pathways. In Carbon Partitioning and Source–Sink Interactions in Plants (eds M.A.Madore & W.J.Lucas), pp. 145155. American Society of Plant Physiologists, Rockville, MD, USA.
  • Köckenberger W. (2001) Nuclear magnetic resonance micro-imaging in the investigation of plant cell metabolism. Journal of Experimental Botany 52, 641652.
  • Köckenberger W., Pope W.J., Xia Y., Jeffrey K.R., Komor E. & Callaghan P.T. (1997) A non-invasive measurement of phloem and xylem water flow in castor bean seedlings by nuclear magnetic resonance microimaging. Planta 201, 5363.
  • Kollmann R. (1973) Cytologie des Phloems. In Grundlagen der Cytologie (eds H.Ruska & P.Sitte), pp. 479505. Gustav Fischer, Jena, Germany.
  • Koroleva O.A., Farrar J.F., Tomos A.D. & Pollock C.J. (1998) Carbohydrates in individual cells of epidermis, mesophyll, and bundle sheath in barley leaves with changed export or photosynthetic rate. Plant Physiology 118, 15251532.
  • Kragler F., Monzer J., Xoconostle-Cazares B. & Lucas W.J. (2000) Peptide antagonists of the plasmodesmal macromolecular trafficking pathway. EMBO Journal 19, 28562868.
  • Kühn C., Franceschi V.R., Schulz A., Lemoine R. & Frommer W.B. (1997) Macromolecular trafficking indicated by localization and turnover of sucrose transporters in enucleate sieve elements. Science 275, 12981300.
  • Kühn C., Quick W.P., Schulz A., Riesmeier J.W., Sonnewald U. & Frommer W.B. (1996) Companion cell-specific inhibition of the potato sucrose transporter SUT1. Plant, Cell and Environment 19, 11151123.
  • Lacombe B., Pilot G., Michard E., Gaymard F., Sentenac H. & Thibaud J.-B. (2000) A shaker-like K+ channel with weak rectification is expressed in both source and sink phloem tissues of Arabidopsis. Plant Cell 12, 837851.
  • Lalonde S., Boles E., Hellmann H., Barker L., Patrick J.W., Frommer W.B. & Ward J.M. (1999) The dual function of sugar carriers: transport and sugar sensing. Plant Cell 11, 707726.
  • Langhans M., Ratajczak R., Lützelschwab M., Michalke W., Wächter R., Fischer-Schliebs E. & Ullrich C.E. (2001) Immunolocalization of plasma membrane H+-ATPase and tonoplast-type pyrophosphatase of the sieve element-companion cell complex in the stem of Ricinus communis L. Planta 213, 1119.
  • Lawton D.M. (1978a) P-protein crystals do not disperse in uninjured sieve elements in roots of runner bean (Phaseolus multiflorus) fixed with glutaraldehyde. Annals of Botany 42, 353361.
  • Lawton D.M. (1978b) Ultrastructural comparison of the tailed and tailless P-proteins respectively of runner bean (Phaseolus vulgaris) and garden pea (Pisum sativum) with tilting stage electron microscopy. Protoplasma 97, 111.
  • Lehmann J. (1979) Nachweis von ATP und ATPase in den Siebröhren von Cucurbita pepo. Zeitschrift für Pflanzenphysiologie 94, 331338.
  • Leineweber K., Schulz A. & Thompson G.A. (2000) Dynamic transitions in the translocated phloem filament protein. Australian Journal of Plant Physiology 27, 733741.
  • Lemoine R., Kühn C., Thiele N., Delrot S. & Frommer W.B. (1996) Antisense inhibition of the sucrose transporter in potato: effects on amount and activity. Plant, Cell and Environment 19, 11241131.
  • Lerchl J., Geigenberger P., Stitt M. & Sonnewald U. (1995) Impaired photoassimilate partitioning caused by phloem-specific removal of pyrophosphate can be complemented by a phloem-specific cytosolic yeast-derived invertase in transgenic plants. Plant Cell 7, 259270.
  • Lichtner F. (2000) Phloem mobility of crop protection products. Australian Journal of Plant Physiology 27, 609614.
  • Linstead P., Dolan L. & Roberts K. (1993) Roots: wild type structure. Arabidopsis. An Atlas of Morphology and Development. (ed. J.Bowman), pp. 102103. Springer, Berlin, Germany.
  • Lohaus G., Winter H., Riens B. & Heldt H.W. (1995) Further studies of the phloem loading process in leaves of barley and spinach. The comparison of metabolite concentrations in the apoplastic compartment with those in the cytosolic compartment and in the sieve tubes. Botanica Acta 108, 270275.
  • Lucas W.J. (1999) Plasmodesmata and the cell-to-cell transport of proteins and nucleoprotein complexes. Journal of Experimental Botany 50, 979987.
  • Lucas W.J., Ding B. & Van Der Schoot C. (1993) Plasmodesmata and the supracellular nature of plants. New Phytologist 125, 435476.
  • Lucas W.J., Yoo B.C. & Kragler F. (2001) RNA as a long-distance information macromolecule in plants. Nature Reviews/Molecular Cell Biology 2, 849857.
  • Magnuson C.E., Goeschl J.D. & Fares Y. (1986) Experimental tests of the Münch–Horwitz theory of phloem transport: effect of loading rates. Plant, Cell and Environment 9, 103109.
  • Marten I., Hoth S., Deeken R., Ketchum K.A., Hoshi T. & Hedrich R. (1999) AKT3, a phloem-localised K+ channel is blocked by protons. Proceedings of the National Academy of Science USA 96, 75817586.
  • Martin T., Frommer W.B., Salanabout M. & Willmitzer L. (1993) Expression of an Arabidopsis sucrose synthase gene indicates a role in metabolization of sucrose both during phloem loading and in sink organs. Plant Journal 4, 367377.
  • McNairn R.B. & Currier H.B. (1968) Translocation blockage by sieve plate callose. Planta 82, 369380.
  • Minchin P.E.H. & Thorpe M.R. (1984) Apoplastic phloem unloading in the stem of bean. Journal of Experimental Botany 35, 538550.
  • Minchin P.E.H. & Thorpe M.R. (1987) Measurement of unloading and reloading of photo-assimilate within the stem of bean. Journal of Experimental Botany 38, 211220.
  • Moriau L., Michelet B., Bogaerts P., Lambert L. & Michel A. (1999) Expression analysis of two gene subfamilies encoding the plasma membrane H+-ATPase in Nicotiana plumbaginifolia reveals the major transport functions of this enzyme. Plant Journal 19, 3141.
  • Münch E. (1930) Die Stoffbewegungen in der Pflanze. Gustav Fischer, Jena, Germany.
  • Murray C. & Christeller J.T. (1995) Purification of a trypsin inhibitor (PFT1) from pumpkin fruit phloem exudate and isolation of putative trypsin and chymotrypsin inhibitor cDNA clones. Biological Chemistry 276, 281287.
  • Nakamura S., Hayashi H., Mori S. & Chino M. (1993) Protein phosphorylation in the sieve tubes of rice plants. Plant and Cell Physiology 34, 927933.
  • Narvaez-Vasquez J., Orozco-Cardenas M.L., Franceschi V.R. & Ryan C.A. (1995) Autoradiographic and biochemical evidence for the systemic translocation of systemin in tomato plants. Planta 195, 593600.
  • Nelson R.S. & Van Bel A.J.E. (1998) The mystery of virus trafficking into, through and out of the vascular bundles. Progress in Botany 59, 476533.
  • Niklas K.J. (1997) The Evolutionary Biology of Plants. University of Chicago Press, Chicago, London.
  • Noiraud N., Delrot S. & Lemoine R. (2000) The sucrose transporter of celery. Identification and expression during salt stress. Plant Physiology 122, 14471455.
  • Noiraud N., Maurousset L. & Lemoine R. (2001) Identification of a mannitol transporter, AgMaT1, in celery phloem. Plant Cell 13, 695705.
  • Nolte K.D. & Koch K.E. (1993) Companion-cell specific localization of sucrose synthase in zones of phloem loading and unloading. Plant Physiology 101, 899905.
  • Öczan S., Dover J. & Johnston M. (1998) Glucose sensing and signaling by two glucose receptors in the yeast Saccharomyces cerevisiae. EMBO Journal 17, 25662673.
  • Offler C.E. & Patrick J.W. (1984) Cellular structures, plasma membrane surface areas and plasmodesmatal frequencies of seed coats of Phaseolus vulgaris L. in relation to photosynthate transfer. Australian Journal of Plant Physiology 11, 7990.
  • Oparka K.J. & Gates P. (1981) Transport of assimilates in the developing caryopsis of rice (Oryza sativa L.). Ultrastructure of the pericarp vascular bundle and its connections with the aleurone layer. Planta 151, 561573.
  • Oparka K.J. & Santa Cruz S. (2000) The great escape: phloem transport and unloading of macromolecules. Annual Review of Plant Physiology and Plant Molecular Biology 51, 323347.
  • Oparka K.J. & Turgeon R. (1999) Sieve elements and companion cells – traffic control centers of the phloem. Plant Cell 11, 739750.
  • Oparka K.J., Duckett C.M., Prior D.A.M. & Fisher D.B. (1994) Real-time imaging of phloem unloading in the root tip of Arabidopsis. Plant Journal 6, 759766.
  • Oparka K.J., Prior D.A.M. & Wright K.M. (1995) Symplastic communication between primary and developing lateral roots of Arabidopsis thaliana. Journal of Experimental Botany 46, 187197.
  • Oparka K.J., Viola R., Wright K.M. & Prior D.A.M. (1992) Sugar transport and metabolism in the potato tuber. In Carbon Partitioning Within and Between Organs (eds C.J.Pollock, J.F.Farrar & A.J.Gordon), pp. 91114. Bios, Oxford, UK.
  • Palauqui J.C., Elmayan T., Dorlhac de Borne F., Crété P., Charles C. & Vaucheret H. (1996) Frequencies, timing, and spatial patterns of co-suppression of nitrate reductase and nitrite reductase in transgenic tobacco plants. Plant Physiology 112, 14471456.
  • Palauqui J.C., Elmayan T., Pollien J.-M. & Vaucheret H. (1997) Systemic acquired silencing: transgene-specific post-translational silencing is transmitted by grafting from silenced to non-silenced scions. EMBO Journal 16, 47384745.
  • Palevitz B.A. & Newcomb E.H. (1971) The ultrastructure and development of tubular and crystalline P protein in the sieve elements of certain papilionaceous legumes. Protoplasma 72, 399427.
  • Pate J.S., Peoples M.B., Van Bel A.J.E., Kuo J. & Atkins C.A. (1985) Diurnal water balance of the cowpea fruit. Plant Physiology 77, 148156.
  • Patrick J.W. (1990) Sieve element unloading: cellular, mechanism and control. Physiologia Plantarum 78, 298308.
  • Patrick J.W. (1997) Phloem unloading. Sieve element unloading and post-sieve element transport. Annual Review of Plant Physiology and Plant Molecular Biology 28, 165190.
  • Patrick J.W. & Offler C.E. (1996) Post-sieve element transport of photoassimilates in sink regions. Journal of Experimental Botany 47, 11651177.
  • Patrick J.W., Zhang W., Tyerman S.D., Offler C.E. & Walker N.A. (2001) Role of membrane transport in phloem translocation of assimilates and water. Australian Journal of Plant Physiology 28, 695707.
  • Pfluger J. & Zambryski P. (2001) Cell growth: the power of symplastic isolation. Current Biology 11, 436439.
  • Phillipar K., Fuchs I., Lüthen H., et al. (1999) Auxin-induced K+ channel expression represents an essential step in coleoptile growth and gravitropism. Proceedings of the National Academy of Science USA 96, 1218612191.
  • Pritchard J., Winch S. & Gould N. (2000) Phloem water relations and root growth. Australian Journal of Plant Physiology 27, 539548.
  • Prüfer D., Schmitz J., Tacke E., Kull B. & Rohde W. (1997) In vivo expression of full-length cDNA copy of potato leafroll virus (PLRV) in protoplasts and transgenic plants. Molecular and General Genetics 253, 609614.
  • Radford J., Vesk M. & Overall R.L. (1998) Callose deposition at plasmodesmata. Protoplasma 201, 3037.
  • Raven J.A. (1991) Long-term functioning of enucleate sieve elements: possible mechanisms of damage avoidance and damage repair. Plant, Cell and Environment 14, 139146.
  • Read S.M. & Northcote D.H. (1983) Chemical and immunological similarities between the phloem proteins of three genera of Cucurbitaceae. Planta 158, 119127.
  • Rhodes J., Thain J.F. & Wildon D.C. (1996) The pathway for systemic electrical signal transduction in the wounded tomato plant. Planta 200, 5057.
  • Riesmeier J.W., Hirner B. & Frommer W.B. (1993) Potato sucrose transporter expression in minor veins indicates a role in phloem loading. Plant Cell 5, 15911598.
  • Riesmeier J.W., Willmitzer L. & Frommer W.B. (1994) Evidence for an essential role of the sucrose transporter in phloem loading and assimilate partitioning. EMBO Journal 13, 18.
  • Robidoux J., Sandborn E.B., Fensom D.S. & Cameron M.L. (1973) Plasmatic filaments and particles in mature sieve elements of Heracleum sphondylium under the electron microscope. Journal of Experimental Botany 79, 349359.
  • Robinson D.G., Sieber H., Kammerloher W. & Schäffner A.R. (1996) PIP1 aquaporins are concentrated in plasmalemmasomes of Arabidopsis thaliana mesophyll. Plant Physiology 111, 645649.
  • Rokitta M., Peuke A.D., Zimmermann U. & Haase A. (1999) Dynamic studies of phloem and xylem flow in fully differentiated plants using fast NMR micro-imaging. Protoplasma 209, 126131.
  • Rook F., Weisbeek P.J. & Smeekens S.C.M. (1998) The light-controlled Arabidopsis bZIP transcription factor factor gene ATB2 encodes a protein with an unusually long leucine zipper domain. Plant Molecular Biology 37, 171178.
  • Ruan Y.-L., Llewellyn D.J. & Furbank R.T. (2001) The control of single-celled cotton fiber elongation by developmentally reversible gating of plasmodesmata and coordinated expression of sucrose and K+ transporters and expansin. Plant Cell 13, 4760.
  • Ruiz-Medrano R. & Xoconostle-Cazares B. & Lucas W.J. (1999) Phloem long-distance transport of CmNACP-1 mRNA: implications for supracellular regulation in plants. Development 126, 44054419.
  • Ruiz-Medrano R., Xoconostle-Cazares B. & Lucas W.J. (2001) The phloem as a conduit for inter-organ communication. Current Opinion in Plant Biology 4, 202209.
  • Ryals J.A., Neuenschwander U.H., Willits M.G., Molina A., Steiner H.-Y. & Hunt M.D. (1996) Systemic acquired resistance. Plant Cell 8, 18091819.
  • Sakuth T., Schobert C., Pecsvaradi A., Eichholz A., Komor E. & Orlich G. (1993) Specific proteins in the sieve-tube exudate of Ricinus communis L. seedlings: separation, characterization and in-vivo labelling. Planta 191, 207213.
  • Salmon S., Lemoine R., Jamaï A. & Bouché-Pillon S. (1995) Study of sucrose and mannitol transport in plasma-membrane vesicles from phloem and non-phloem tissue of celery (Apium graveolens L.) petioles. Planta 197, 7684.
  • Santa Cruz S. (1999) Perspective: phloem transport of viruses and macromolecules – what goes in must come out. Trends in Microbiology 7, 237241.
  • Sasaki T., Chino M., Hayashi H. & Fujiwara T. (1998) Detection of several mRNA species in rice phloem sap. Plant Cell Physiology 39, 895897.
  • Schäffner A.R. (1998) Aquaporin function, structure, and expression: are there more surprises to surface in water relations? Planta 204, 131139.
  • Scheirer D.C. (1990) Mosses. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 1933. Springer, Berlin, Germany.
  • Schmitz K. (1990) Algae. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 118. Springer, Berlin, Germany.
  • Schobert C., Baker L., Szederkenyi J., Großmann P., Komor E., Hayashi H., Chino M. & Lucas W.J. (1998) Identification of immunologically related proteins in sieve-tube exudate collected from monocotyledonous and dicotyledonous plants. Planta 206, 245252.
  • Schobert C., Großmann P., Gottschalk M., Komor E., Pecsvaradi A. & Zur Nieden U. (1995) Sieve-tube exudate from Ricinus communis L. seedlings contains ubiquitin and chaperones. Planta 196, 205210.
  • Van Der Schoot C. & Van Bel A.J.E. (1989) Glass microelectrode measurements of sieve tube membrane potentials in internodes and petioles of tomato (Solanum lycopersicum L.). Protoplasma 149, 144154.
  • Schrier A.A. (2001) Der Einfluß der Temperatur auf das Funktionieren, die Evolution und die Verbreitung apoplasmatischer und symplasmatischer Phloembeladung. Thesis. University of Gießen, Gießen, Germany.
  • Schulz A. (1990) Conifers. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 6388. Springer, Berlin, Germany.
  • Schulz A. (1994) Phloem transport and differential unloading in pea seedlings after source and sink manipulations. Planta 192, 239248.
  • Schulz A. (1995) Plasmodesmal widening accompanies the short-term increase in symplasmic phloem unloading in root tips under osmotic stress. Protoplasma 118, 2237.
  • Schulz A. (1998) Phloem. Structure related to function. Progress in Botany 59, 429475.
  • Schulz A., Kühn C., Riesmeier J.W. & Frommer W.B. (1998) Ultrastructural effects in potato leaves due to anti-sense inhibition of the sucrose transporter indicate an apoplasmic mode of phloem loading. Planta 206, 533543.
  • Scott P.C., Miller L.W., Webster B.D. & Leopold A.C. (1967) Structural changes during bean leaf abcission. American Journal of Botany 54, 730734.
  • Sjolund R.D. (1997) The phloem sieve element. A river flows through it. Plant Cell 9, 11371146.
  • Sjolund R.D. & Shih C.Y. (1983) Freeze-fracture analysis of phloem structure in plant tissue cultures. I. The sieve element reticulum. Journal of Ultrastructural Research 82, 111121.
  • Smeekens S. (2000) Sugar-induced signal transduction in plants. Annual Review of Plant Physiology and Plant Molecular Biology 51, 4981.
  • Smith J.A.C. & Milburn J.A. (1980a) Osmoregulation and the control of phloem-sap composition in Ricinus communis L. Planta 148, 2834.
  • Smith J.A.C. & Milburn J.A. (1980b) Phloem turgor and the regulation of sucrose loading in Ricinus communis L. Planta 148, 4248.
  • Sokolova M., Prüfer D., Tacke E. & Rohde W. (1997) The potato leaf roll virus 17K movement protein is phosphorylated by a membrane-associated protein kinase from potato with biochemical features of proten kinase C. FEBS Letters 400, 201205.
  • Soltis D.E., Soltis P.S., Chase M.W., et al. (2000) Angiosperm phylogeny inferred from 18S rDNA, rbcL, and atpB sequences. Botanical Journal of the Linnean Society 133, 381461.
  • Stadler R. & Sauer N. (1996) The Arabidopsis thaliana AtSUC2 gene is specifically expressed in companion cells. Botanica Acta 109, 299306.
  • Stadler R., Brandner H., Schulz A., Gahrtz M. & Sauer N. (1995) Phloem loading by the PmSUC2 sucrose carrier from Plantago major occurs in companion cells. Plant Cell 7, 15451554.
  • Staehelin L.A. (1997) The plant ER: a dynamic organelle composed of a large number of discrete functional domains. Plant Journal 11, 11511165.
  • Taylor E.L. (1990) Phloem evolution: an appraisal based on the fossil record. In Sieve Elements. Comparative Structure, Induction and Development (eds H.-D.Behnke & R.D.Sjolund), pp. 285298. Springer, Berlin, Germany.
  • Thaine R. (1962) A translocation hypothesis based on the structure of plant cytoplasm. Journal of Experimental Botany 13, 152160.
  • Thaine R. (1969) Movement of sugars through plants by cytoplasmic pumping. Nature 222, 873875.
  • Thompson G.A. & Schulz A. (1999) Long-distance transport of macromolecules. Trends in Plant Science 4, 354360.
  • Thorsch J. & Esau K. (1981a) Changes in the endoplasmic reticulum during differentiation of a sieve element in Gossypium hirsutum. Journal of Ultrastructural Research 74, 183194.
  • Thorsch J. & Esau K. (1981b) Nuclear degeneration and the association of endoplasmic reticulum with the nuclear envelope and microtubules in maturing sieve elements of Gossypium hirsutum. Journal of Ultrastructural Research 74, 195204.
  • Truernit E. & Sauer N. (1995) The promoter of the Arabidopsis thaliana SUC2 sucrose-H+ symporter gene directs expression of β-glucuronidase to the phloem: evidence for phloem loading by SUC2. Planta 196, 564570.
  • Truernit E., Schmid J., Epple P., Illig J. & Sauer N. (1996) The sink-specific and stress-regulated Arabidopsis STP4 gene: enhanced expression of a gene encoding a monosaccharide transporter by wounding, elicitors, and pathogen challenge. Plant Cell 8, 21692182.
  • Turgeon R. (1991) Symplastic phloem loading and the sink-source transition in leaves: A model. In Recent Advances in Phloem Transport and Assimilate Compartmentation (eds J.-L.Bonnemain, S.Delrot, W.J.Lucas & J.Dainty), pp. 1822. Ouest Editions, Nantes, France.
  • Turgeon R. (1996) Phloem loading and plasmodesmata. Trends in Plant Science 1, 418423.
  • Turgeon R. & Hepler P.K. (1989) Symplastic continuity between mesophyll and companion cells in the minor veins of Cucurbita pepo leaves. Planta 179, 2431.
  • Turgeon R. & Medville R. (1998) The absence of phloem loading in willow leaves. Proceedings of the National Academy of Sciences USA 95, 1205512060.
  • Turgeon R. & Wimmers L.E. (1988) Different patterns of vein loading of exogenous [14C]-sucrose in leaves of Pisum sativum and Coleus blumei. Plant Physiology 87, 179182.
  • Turgeon R., Medville R. & Nixon K.C. (2001) The evolution of minor vein phloem and phloem loading. American Journal of Botany 88, 13311339.
  • Tyerman S.D., Bohnert H.J., Maurel C., Steudle E. & Smith J.A.C. (1999) Plant aquaporins: their molecular biology, biophysics and significance for plant water relations. Journal of Experimental Botany 50, 10551071.
  • Voinnet O., Vain P., Angell S. & Baulcombe D.C. (1998) Systemic spread of sequence-specific transgene RNA degradation in plants is initiated by localized intoduction of ectopic promoterless DNA. Cell 95, 177187.
  • Volk G. & Franceschi V.R. (2000) Localization of a calcium-channel-like protein sieve element plasma membrane. Australian Journal of Plant Physiology 27, 779786.
  • Vreugdenhil D. & Koot-Gronsveld E.A.M. (1989) Measurements of pH, sucrose and potassium ions in the phloem of castor bean (Ricinus communis) plants. Physiologia Plantarum 77, 385388.
  • Wang Y.Q. & Kollmann R. (1996) Vascular differentiation in the graft union of in vitro grafts with different compatibility. Structural and functional aspects. Journal of Plant Physiology 147, 521533.
  • Warmbrodt R.D. (1985) Studies on the root of Hordeum vulgare L. – Ultrastructure of the seminal root with special reference to the phloem. American Journal of Botany 72, 414432.
  • Warmbrodt R.D. (1986) Structural aspects of the primary tissues of the Cucurbita pepo L. root with special reference to the phloem. New Phytologist 102, 175192.
  • Webster D.H. & Currier H.B. (1965) Callose: lateral movement of assimilates from phloem. Science 150, 16101611.
  • Weise A., Barker L., Kühn C., Lalonde S., Buschmann H., Frommer W.B. & Ward J.M. (2000) A new subfamily of sucrose transporters, SUT4, with low affinity/high capacity localized in enucleate sieve elements of plants. Plant Cell 12, 13451355.
  • Wergin W.P., Palevitz B.A. & Newcomb E.H. (1975) Structure and development of P-protein in phloem parenchyma and companion cells of legumes. Tissue and Cell 7, 227242.
  • Wright J.P. & Fisher D.B. (1981) Measurement of the sieve tube membrane potential. Plant Physiology 67, 845848.
  • Xoconostle-Cazares B., Xiang Y., Ruiz-Medrano R., Wang H.-L., Monzer J., Yoo B.-C., McFarland K.C., Franceschi V.R. & Lucas W.J. (1999) Plant paralog to viral movement protein potentiates transport of mRNA into the phloem. Science 283, 9498.
  • Zahur M.S. (1959) Comparative Study of the Secondary Phloem of 423 Species of Woody Dicotyledons Belonging to 85 Families. Cornell University Agricultural Station Memoir 358.
  • Zhao R., Dielen V., Kinet J.-M. & Boutry M. (2000) Cosuppression of a plasma membrane H+-ATPase isoform impairs sucrose translocation, stomatal opening, plant growth, and male fertility. Plant Cell 12, 535546.
  • Ziegler H. (1975) Nature of transported substances I. In Phloem Transport. Encyclopedia of Plant Physiology (eds M.H.Zimmermann & J.A.Milburn), pp. 59100. Springer, Berlin, Germany.
  • Zimmermann M.H. & Ziegler H. (1975) List of sugars and sugar alcohols in sieve-tube exudates. In Encyclopedia of Plant Physiology (eds M.H.Zimmermann & J.A. Milburn), pp. 480503. Springer, Berlin, Germany.

Received 25 October 2001;received inrevised form 3 April 2002;accepted for publication 18 April 2002