(a) Phloem structure

In most plant species, phloem is made up of phloem fibres, phloem parenchyma, sieve cells (sieve elements) and their accompanying companion cells (Figure 5.5a). Sieve elements are ideally suited for rapid transport of substances at high rates over long distances. They are elongated and are arranged end to end in files referred to as sieve tubes (Figure 5.16b). Abutting sieve elements are interconnected through membrane-lined pores (sieve pores) with large diameters (1 to 15µm). These pores collectively form sieve plates (Figure 5.16c). The transport capacity of sieve tubes is dependent on a developmentally programmed degeneration of the sieve element protoplasm (cell contents) leaving an open, membrane-bound tube. In mature conducting sieve elements, the protoplast is limited to a functional plasma membrane enclosing a sparse cytoplasm containing low densities of plastids, mitochondria and smooth endo-plasmic reticulum distributed along the lateral walls (Figure 5.16d). These relatively empty sieve tubes provide a longitudinal network which conducts phloem sap (Figure 5.5b).

Fig 5.16ann.jpg

Figure 5.5 (a) spatial arrangement of cell types in a vascular strand from the primary stem of Phaseolus vulgaris (French bean); electron micrographs of stem phloem of Curcurbita maxima (b,c) and P. vulgaris (d) illustrating significant structural characteristics of sieve elements and companion cells. (a) Conducting cells of the phloem (sieve elements) and accompanying companion cells form groups of cells that are separated by phloem parenchyma cells. This mosaic of cells is located between the cortex and xylem and capped by phloem fibres. Scale bar = 7.3 μm. (b) A longitudinal section through two sieve elements arranged end to end to form part of a sieve tube. Companion cells can also be seen. The abutting wall (sieve plates) displays characteristic membrane-lined sieve pores (arrowheads). Cytoplasm of the sieve elements has largely degenerated leaving only endoplasmic reticulum (arrows) and a few plastids around the mature sieve element. Scale bar = 5 μm. (c) A face view of part of a sieve plate showing sieve pores (arrowheads). Scale bar = 0.5 μm. (d) Transverse section through a sieve element and its accompanying companion cell illustrating the sparse cytoplasm and low density of organelles in the sieve element contrasting with the dense ribosome-rich cytoplasm of the nucleated companion cell. Note the mitochondria and rough endoplasmic reticulum. Scale bar = 1.0 μm. c, cortex; cc, companion cell; e, epidermis; er, endoplasmic reticulum; m, mitochondrion; n, nucleus; p, pith; pf, phloem libres; pp, phloem parenchyma; se, sieve element; sp, sieve plates; vc, vascular cambium; x, xylem

Sieve elements are closely associated with one or more companion cells, forming a sieve element–companion cell (se–cc) complex (Figure 5.5d) that plays an important role in transport. These distinct cell types result from division of a common procambial mother cell. In mature se–cc complexes, relatively open sieve elements contrast with adjacent companion cells containing dense, ribosome-rich cytoplasm with a prominent nucleus and abundant mitochondria and rough endoplasmic reticulum (Figure 5.5d). High densities of extensively branched plasmodesmata in contiguous walls of sieve elements and companion cells (Figure 5.6) account for intense intercellular coupling in se–cc complexes (van Bel 1993). Thus, companion cells are considered to perform the metabolic functions surrendered by, but required for, maintenance of viable sieve elements. This functional coupling has led to the concept of se–cc complexes being responsible for phloem transport.

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Figure 5.6 (a) Electron micrograph and (b) diagrammatic interpretation of a secondary plasmodesma interconnecting a mature sieve element and its companion cell in a tobacco leaf. Note the characteristic branching of the plasmodesma within the wall of a companion cell. Scale bar = 0.2 μm. c, callose; other symbols as for Figure 5.16 (Based on Ding et al. 1993; reproduced with permission of Blackwell Science)

(b) Visualising the translocation stream

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Figure 5.7 Microautoradiographs of (a) transverse and (b) longitudinal sections of Phaseolus vulgaris stem tissue illustrating localisation of ¹⁴C-labelled photosynthate in sieve tubes. These sections are obtained by snap freezing plant tissue and removing frozen water by sublimation (e.g. freeze-drying or freeze substitution). ¹⁴C-labelled compounds do not move during preparation. Tissues are embedded in absolute dryness and thin sections are cut, mounted dry on microscope slides and overlain with a thin film of photographic emulsion. Silver grains are visible in the emulsion where ¹⁴C, an ideal radioisotope for these experiments, irradiates the film. Abbreviations: se, sieve element; pp, phloem parenchyma; vb, vascular bundle; other symbols as for Figure 5.6. Scale bar in (a) = 20μm; in (b) = 10 μm

Transport of radioactively labelled substances through phloem has been demonstated using microautoradiography (Figure 5.7), providing irrefutable evidence that sieve elements are conduits for transport of phloem sap. Experimentally, a pulse of ¹⁴CO₂ is fixed photosynthetically and ¹⁴C-labelled sugars are given time to reach the stem, which is then excised and processed for microautoradiography. As ¹⁴C first moves through the stem, most of the isotope is confined to the transport pathway and very little has had time to move laterally into storage pools. High densities of ¹⁴C-labelled sugars are found in sieve elements (Figure 5.7), demonstrating that these cells constitute a transport pathway.

(c) Phloem sealing mechanisms

Herbivory or environmental factors causing physical damage could pose a threat to transport through sieve tubes and has undoubtedly imposed strong selection pressure for the evolution of an efficient and rapid sealing mechanism for damaged sieve tubes. Since sieve tube contents are under a high turgor pressure (P), severing would cause phloem contents to surge from the cut site, incurring excessive assimilate loss in the absence of a sealing mechanism. For dicotyledonous species, an abundant phloem-specific protein (P-protein) provides an almost instantaneous seal. P-protein is swept into sieve pores where it becomes entrapped, thus sealing off the damaged sieve tubes. Production of callose (β-1,3 glucan) in response to wounding or high-temperature stress is another strategy to seal off damaged sieve tubes. Callose also seals off sieve pores during overwintering in deciduous plants. Callose is deposited between the plasma membrane and cell wall, eventually blocking sieve pores. Whether deposited in response to damage or overwintering, callose can be degraded by β-1,3 glucanase, allowing sieve tubes to regain transport capacity.

Since phloem translocation is confined to sieve elements embedded within a tissue matrix, it is difficult to obtain uncontaminated samples of translocated sap. The least equivocal approach has been to take advantage of the high P of sieve tube contents. Puncturing or severing sieve tubes should cause exudation of phloem sap provided a sealing mechanism is not activated.

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Figure 5.8 Aphids can be used to collect phloem sap. Top photograph: a feeding aphid with its stylet embedded in a sieve tube (see insert); scl, sclerenchyma; st, stylet; x, xylem; p, phloem. Note the drop of ‘honeydew’ being excreted from the aphid’s body. Plates (a) to (e) show a sequence of stylet cutting with an RF microcautery unit at about 3-5 s intervals (a to d) followed by a two-minute interval (d to e) which allowed exudate to accumulate. The stylet has just been cut in (b); droplets of hemolymph (aphid origin) are visible in (b) and (c); once the aphid moves to one side the first exudate appears (d), and within minutes a droplet (e) is available for microanalysis. Scale bars: top = 1 mm, bottom = 1.5 mm (Courtesy D. Fischer)

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Figure 5.9. Exudation of phloem contents from lupins. A, following incision of the vasculature at the stylar tip and of the ventral suture of fruits of Lupinus angustifolius. B, following incision of the vasculature of a stem of L. angustifolius. C, following incisions to the vasculature of pre and post anthesis stage flowers on the inflorescence of L. angustifolius. D. exudation at the abscission zone following abscission of two flowers 5 minutes earlier on the raceme of L. mutabilis. Photographs courtesy Craig. Atkins.

For some plant species, sieve-pore sealing develops slowly, or can be experimentally down-regulated by massage or repeated excisions (Milburn and Kallarackal 1989) or slowed by puncturing the vasculature while it is snap frozen in liquid N₂ (Pate et al 1984). Carefully placed incisions that do not disturb the underlying xylem, which in any case is more likely to be under tension, permit collection of relatively pure phloem exudate through the severed sieve tubes. Nevertheless, contamination with the contents of cells other than sieve tubes damaged at the site of incision is inevitable.  For the major solutes of phloem such as sugars or amino acids that are present in high concentrations this problem is minimal but for less abundant molecules like hormones or other signals, particularly proteins or nucleic acids, conclusions about the origin and functions of these must be made with caution. The ‘natural hemophiliacs’ of the plant world are few and include a number of cucurbits, some brassicas, castor bean, species of the genus Yucca and some species of lupin (Lupinus albus, L. angustifolius, L. mutabilis and L. cosentinii). The excision technique has been expanded to plant species that do not readily exude, by chemically inhibiting the sealing mechanism. Callose production is blocked when wounded surfaces are exposed to the chelating agent ethylenediaminetetraacetic acid (EDTA) by complexing with calcium, a cofactor for callose synthase. Immersing whole, excised organs in EDTA solution, which is essential to inhibit blockage, risks contaminating sap with solutes lost from the apoplast as well as non-conducting cells. This is not an ideal technique.

Enlisting sap-sucking aphids or leaf hoppers to sample sap has been more successful. Aphids can guide a long syringe-like mouthpart (a stylet) into conducting sieve elements (Figure 5.8). Pressure normally forces sieve-tube sap through the stylet into the aphid’s gut where it becomes food or is excreted as ‘honeydew’. By detaching the aphid from its mouthpart pure phloem contents can be collected from the cut end of the implanted stylet. Detaching the aphid body can be achieved by surgery following rapid anesthesia in high CO₂ or by severing the stylet using a laser.  While stylectomy has been successful with a number of monocotyledons (rice, wheat and barley) the technique has proved more difficult to use with dicotyledons, yielding at best a few microlitres of phloem contents. On the other hand collection of milliliter volumes of exudate from one of the natural hemophiliacs is possible permitting extensive analysis of solutes and macromolecules. In the case of lupins, exudation occurs readily at many sites on the plant so that solutes translocated from source tissues as well as entering sinks can be collected and analysed (Figure 5.9).

(a) Chemical analysis of phloem exudate

Chemical analyses of phloem exudate collected from a wide range of plant species have led to a number of generalisations (e.g. Milburn and Baker 1989) about the contents of sieve tubes. Phloem exudate is a concentrated solution (10–12% dry matter), generating an osmotic pressure (Π) of 1.2 to 1.8MPa. pH is characteristically alkaline (pH 8.0 to 8.5). The principal organic solutes are non-reducing sugars (sucrose), amides (glutamine and asparagine), amino acids (glutamate and aspartate) and organic acids (malate). Of these solutes, non-reducing sugars generally occur in the highest concentrations (300–900 mM). Nitrogen is transported through the phloem as amides and amino acids; nitrate is absent and ammonium only occurs in trace amounts. Calcium, sulphur and iron are scarce in phloem exudate while other inorganic nutrients are present, particularly potassium which is commonly in the range of 60–120 mM. Physiological concentrations of auxins, gibberellins, cytokinins and abscisic acid have been detected in phloem exudate along with nucleotide phosphates. The principal macromolecule group is protein but low levels of peptides and nucleic acids are also present.  While in cucurbits the protein in exudate is comprised largely of P-protein, a diverse array of proteins, many of them enzymes, have also been detected.

(b) Significance of the chemical forms translocated 

Phloem sap provides most inorganic and all organic substrates necessary to support plant growth. Non-transpiring tissues are particularly dependent on resources delivered in the phloem (Section 5.1). That translocated sugars represent the major chemical fraction of the phloem sap is consistent with the bulk of plant dry matter (90%) being composed of carbon, hydrogen and oxygen. Carbon transport is further augmented by transport of nitrogen in organic forms.

Carbohydrate is translocated as non-reducing sugars in which the metabolically reactive aldehyde or ketone group is reduced to an alcohol (mannitol, sorbitol) or combined with a similar group from another sugar to form an oligosaccharide. Apart from sucrose, transported oligosaccharides belong to the raffinose series. In this series, sucrose is bound with increasing numbers of galactose residues to form raffinose, stachyose and verbascose respectively. However, sucrose is the most common sugar species transported. In a small number of plant families, other sugar species predominate. For example, the sugar alcohol sorbitol is the principal transport sugar in the Rosaceae (e.g. apple) and stachyose predominates in the Cucurbitaceae (e.g. pumpkin and squash). Exclusive transport of non-reducing sugars probably reflects packaging of carbohydrate in a chemical form which protects it from being metabolised. Metabolism of these transported sugars requires their conversion to an aldehyde or ketone by enzymes which are thought to be absent from sieve-tube contents.

Plant physiologists have long regarded the two long distance translocation streams of xylem and phloem as having functions additional to the distribution of nutrients and assimilates. Specifically, each serves as a means of communication between the source and sink organs such that systemic signals are thought to transmit molecular responses to endogenous and environmental cues. Furthermore, evidence is accumulating that some of these signals regulate gene expression as a consequence of their translocation (see below). 

(c) Macromolecule composition of phloem

Proteomic and transcriptomic analyses have demonstrated a widely diverse composition of proteins, peptides and nucleic acids, including mRNA and small RNAs, in phloem exudates. While the origin of each individual protein or nucleic acid remains to be verified the limited compositional data available from stylectomy confirms that indeed each group of macromolecules is present in phloem. In cucurbit phloem exudate some 1110 different proteins have been detected along with a large number of mRNAs and similar data have been obtained for exudates from other species (Brassica napus, Ricinus communis and Lupinus albus). Compositional data for phloem proteins of these species show a common complement that includes phloem-specific P proteins together with proteins involved in sugar metabolism and transport, protein turnover and transport, detoxification of reactive oxygen species, as well as proteins that provide defence against insect herbivores and pathogens (Figure 10).  Some undoubtedly play a role in maintenance of the SE system while others, such as the Flowering Locus T (FT) protein associated with the flowering response (‘florigen’), appear to be systemic ‘signals’ (Rodriguez-Medina et al 2011) and there may be many more. Because sieve tubes are enucleate and lack ribosomes (5.2.2 a), proteins in the translocation stream are not formed in situ but are transported from sites of synthesis in the companion cells.

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Figure 5.10. Two dimensional polyacrylamide gel electrophoresis separation of proteins in phloem exudate from Lupinus albus.  The gel was developed in the first dimension by isoelectric focusing with a linear pH gradient of 3-10 followed by separation due to differences in molecular mass.  The positions of mass standards are shown on the right hand side of the gel.  After staining with Coomassie Blue to locate the spots they were excised for digestion with trypsin. The peptides were then analysed by partial sequence determination using MS/MS and identified using database searches. (Courtesy of Craig Atkins)

Functional analysis of the cDNA identified in transcriptome studies of phloem exudates revealed transcripts involved in a wide range of processes that include metabolism, plant responses to stresses, transport, DNA/RNA binding and protein turnover.  The presence of transcripts in phloem exudate supports the idea of an RNA-based signalling network that is thought to function in control of processes associated with plant growth and development (Lough and Lucas, 2006).  However, the functional role of transcripts in the contents of sieve tubes as well as their actual translocation is yet to be determined.

Small RNA molecules (18-25 nt) have been identified in phloem exudate collected from rape, white lupin, pumpkin, castor bean and Yucca filamentosa as well as in aphid stylet exudate collected from apple stems. The population includes both microRNAs (miRNA) and small interfering RNAs (siRNA) a large number of which target mRNA of transcription factors that themselves regulate genes expressions. miRNAs are also involved in mediating environmental responses, including responses to salinity, drought, nutrient limitations, as well as hormone interactions. Their small size and powerful functions in targeting mRNAs to regulate expression suggest that those in phloem exudate are likely to be systemic signals.

An important question that relates to the significance of macromolecules in the contents of sieve tubes is proof that they are translocated and that translocation is essential for their function at a sink. A diversity of studies that have exploited cucurbit root stocks and grafted scions has provided clear evidence that P proteins among others are graft transmissible. In a series of elegant experiments Aoki et al. (2005) labelled and injected two isolated pumpkin phloem proteins (CmPP16‑1 and CmPP16-2) into the vasculature of intact rice plants through severed leaf hopper stylets and showed their translocation as well as some evidence for specificity in protein translocation.  The Flowering Locus T (FT) protein formed in leaves mediates the flowering transition of shoot apical meristems and the evidence that it is translocated is compelling. The long distance movement of RNA molecules was first demonstrated for plant viruses and there is now good evidence for phloem translocation of a number of transcripts (Lough and Lucas 2006). A recent compilation identified 13 miRNAs involved in plant responses to drought/salt stress (Covarrubias and Reyes 2010).  Eight of these were identified in lupin phloem exudate (Rodriguez-Medina et al. 2011) and, importantly, six were also recovered from PCR amplification of apple stylet exudate (Varkonyi-Gasic et al 2010).  There is thus a possibility that the responses to drought and salinity are mediated through miRNAs translocated from sites where the stress is sensed to sites where a response is initiated. 

The most convincing case for a translocated miRNA in phloem regulating gene expression relates to Pi homeostasis. While both local and systemic signals are involved, miR399 is phloem mobile and acts directly in roots to down regulate the expression of PHO2 (a ubiquitin conjugating enzyme) that results in greater expression of Pi transporters to increase Pi uptake under conditions of deficiency. Systemic signaling has also been implicated in homeostasis of other nutrients, including N, S and Cu with, in each case, miRNAs involved. 

Phloem flux can be estimated in a number of ways. The simplest is to determine dry weight gain of a discrete organ connected to the remainder of a plant by a clearly definable axis of known phloem cross-sectional area. Developing fruits or tubers meet these criteria. Sequential harvests from a population of growing fruit or tubers provide measures of the organ’s net gain of dry matter imported through the phloem. Net gains or losses of dry matter resulting from respiration or photosynthesis are incorporated into calculations to give gross gain in dry matter by the organ. Flux of dry matter through the phloem (specific mass transfer — SMT; Canny 1973) can then be computed on a phloem or preferably on a sieve-tube lumen cross-sectional area basis. Area estimates can be obtained from histological sections of the pedicel or stolon that connects a test organ to its parent plant. Expressed on a phloem cross-sectional area basis, SMT estimates are normally in the range of 2.8–11.1 g m^–2 phloem s^–1 (Canny 1973). Flux on the basis of sieve-tube lumen cross-sectional area is preferable but relies on identification of sieve tubes and the assumption that they are equally functional as transport conduits. Sieve tubes account for some 20% of phloem cross-sectional area, suggesting fluxes are about five-fold higher through a sieve-tube lumen.

Speed of phloem translocation can be determined from simultaneous measurements of SMT and phloem sap concentrations as shown in Equation 5.1 below:

\[\mathrm{Speed} (m \cdot s^{-1}) = \mathrm{SMT}(g \cdot m^{-2} \cdot s^{-1})/ \mathrm{concentration} (g \cdot m^{-3}) \tag{5.1} \]

For a sucrose concentration of 600 mM (or 2.16 x 10⁵ g m^-3) and the highest SMT values shown above, Equation 5.1 estimates that phloem sap can move at speeds of up to 56 × 10^–5 m s^–1 or 200 cm h^–1. These estimates have been verified by following the movement of radioisotopes introduced into the phloem translocation stream.

These estimates of transport rates and speeds tacitly assume that phloem sap moves through sieve tubes by mass flow (water and dissolved substances travel at the same speed). Independent estimates of transport rate, concentration of phloem sap and translocation speed lend support to, but do not verify, the assumption that movement occurs as a mass flow.

A simple and direct test for mass flow is to determine experimentally whether water and dissolved substances move at the same speed. This test should be relatively easy to apply using radioactively labelled molecules. Unfortunately, in practice it turns out that different molecular species are not loaded into the sieve tubes at the same rates and the plasma membranes lining the sieve tubes are not equally permeable to each substance. Thus, the analysis is complicated by the necessity to use model-based corrections for rates of loading into and losses from the sieve tubes. Nevertheless, the speed estimates obtained from such experiments are found to be similar for dissimilar molecules, supporting the proposition that mass flow accounts for most transport through sieve tubes.

Phloem translocation is generally believed to be driven by pressure. Münch (1930) proposed that a passive mass flow of phloem sap through sieve tubes was driven by the osmotically generated pressure gradient between source and sink regions (Figure 5.11). At source regions, the principal osmotica of phloem sap are actively or passively loaded into sieve tubes from companion cells or mesophyll cells (see 5.3.2), thereby driving water towards the lower water potentials within sieve tubes. As water enters, P rises. Unloading of solutes from sieve tubes at sink regions reverses water potentials; water flows out of sieve tubes and P falls relative to that of sieve tubes in source regions.

The pressure-flow hypothesis can be modelled using the relationship that rate of mass flow (F_f) of a substance is given by the product of speed (S) of solution flow, path cross-sectional area (A) and its concentration (C). That is:

 \[F_f = S \cdot A \cdot C \tag{5.2}\]

 Speed (m s^–1) has the same units as volume flux (J_v — m³ m^–2 s^–1) of solution passing through a transport conduit. Poisseuille’s Law describes the volume flux (J_v) of a solution of a known viscosity (h) driven by a pressure difference (DP) applied over the length (l) of pathway of radius (r) as:

\[J_v = \pi r^4 \Delta P/8 \eta l \tag{5.3} \]

Fig_p_5.20.png

Figure 5.11 Scheme describing the pressure flow hypothesis of phloem transport (Based on Münch 1930)

The term πr⁴/8ηl in Equation 5.3 provides an estimate of hydraulic conductivity (L_p) of the sieve-tube conduit which is set by the radius of the sieve pores. Raised to the fourth power, small changes in the sieve-pore radius will exert profound effects on the hydraulic conductivity of the sieve tubes (Section 5.2). The viscosity of sieve-tube sap is determined by the chemical species (particularly sugars) and their concentrations in the phloem sap.

Key features of the pressure-flow hypothesis are encapsulated in Equation 5.3. The central question is whether a pressure gradient exists in sieve tubes with the expected direction and of sufficient magnitude to support observed rates of sap flow. Indirect estimates of P in sieve tubes made through determination of intra- and extracellular P support the pressure-flow hypothesis. Direct measurements of sieve-tube P are technically challenging because of the inaccessibility of these small, highly turgid cells. They are, for instance, too small for pressure-probe measurements. However, manometric pressure measurements obtained using severed aphid stylets agree with indirect estimates (Wright and Fisher 1980). Experimental manipulation of the pressure gradient between the source and sink also results in alterations in phloem translocation rates consistent with the pressure-flow model.

Whether the pressure gradient is sufficiently steep is a more vexing question. The pressure gradient required to drive phloem translocation at observed rates is determined by the transport resistance of the phloem path, according to Ohm’s Law. Dimensions of the sieve pores set a limiting radius for volume flux of transported sap (Equation 5.3) and hence transport resistance. If the sieve pores were open and unoccluded by P-protein, a number of studies have demon-strated that the measured pressure gradients are sufficient to support the observed rates of flow. However, the in situ radii of sieve pores remain unknown.

Overall, the pressure-flow hypothesis accounts for many observed features of phloem translocation, including distribution of resources. While conclusive evidence supporting this hypothesis is still sought, less attention is now focused on this issue with a growing appreciation that the phloem pathway has spare transport capacity. Evidence from Kallarackal and Milburn (1984), for example, showed that the specific mass transfer (SMT – see preceding section) to an intact fruit of castor bean could be doubled on removal of competing fruits. Moreover, if P of sieve elements at the sink end of the phloem path was reduced to zero, by severing the pedicel and allowing exudation, SMT rose to an incredible 305 g m^–2 sieve-tube area s^–1! In another experiment, when half the conducting tissue was removed from the peduncle of sorghum or wheat plants, grain growth rate was not impaired (Wardlaw 1990). Together, these observations imply that phloem has excess carrying capacity in both dicotyledons and monocotyledons. Particularly in monocotyledonous plants, a strong selection pressure for spare transport capacity must exist because there is no vascular cambial activity to replace damaged sieve elements.

Loading of sugars, potassium and accompanying anions into sieve tubes at sources determines solute concentrations in phloem sap (Table 5.1). The osmotic pressure (Π) of these solutes influences P generated in sieve tubes. Thus, source output determines the total amount of assimilate available for phloem transport as well as the pressure head driving transport along the phloem path to recipient sinks. Withdrawal of assimilates from sieve tubes at the sink end of the phloem path, by the combined activities of phloem unloading and metabolism/compartmentation (Table 5.1), determines Π of phloem sap. Other sink-located membrane transport processes influence Π around sieve tubes. The difference between intra- and extracellular Π of sieve tubes is a characteristic property of each sink and determines P in sink sieve tubes.

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The pressure difference between source and sink ends of the phloem pathway drives sap flow (Equation 5.3) and hence phloem translocation rate (Equation 5.2) from source to sink. The source and sink processes governing the pressure dif-ference (Table 5.1) are metabolically dependent, thus rendering phloem translocation rates susceptible to cellular and environmental influences. The pressure-flow hypothesis predicts that the phloem path contribution to longitudinal transport is determined by the structural properties of sieve tubes (Table 5.1). Variables of particular importance are cross-sectional area (A) of the path (determined by numbers of sieve pores in a sieve plate and sieve-tube numbers) and radius of these pores (sets r in Equation 5.3). These quantities appear in Equations 5.2 and 5.3. Thus, the individual properties of each sink and those of the phloem path connecting that sink to its source will determine the potential rate of assimilate import to the sink (Figure 5.12).

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Figure 5.12. Scheme describing photoassimilate flow from a source leaf linked to two competing sinks, Sink 1 and Sink 2. Assimilate flows through alternative phloem paths (Path 1 and Path 2) each with its own conductance (K_path) and pressure difference (P) between source and sink. Hence Path 1 is distinguished by K_path1 and P_sink1 and Path 2 by K_path2and P_sink2

The transport rate (R) of assimilate along each phloem path, linking a source with each respective sink, can be predicted from the pressure-flow hypothesis (see Equations 5.2 and 5.3) as:

\[ R = K_{path} (P_{source} - P_{sink}) C \tag{5.4}\]

where path conductance (K_path) is the product of path hydraulic conductivity (L_p) and cross-sectional area (A). Hence, the relative flows of assimilates between hypothetical sinks (sink 1 and sink 2) shown in Figure 5.12 may be expressed by the following ratio:

\[ \frac{K_{path1} (P_{source} - P_{sink1}) C} {K_{path2} (P_{source} - P_{sink2}) C} \tag {5.5} \]

Partitioning of assimilates between two competing sinks is thus a function of path conductance and P at the sink end of the phloem path (Equation 5.5). Since phloem has spare capacity, any differences in the conductance of the inter-connecting paths (Figure 5.12) would exert little influence on the rate of phloem transport to the competing sinks. Assimilate partitioning between competing sinks would then be determined by the relative capacity of each sink to depress sieve-tube P at the sink end of the respective phloem path. Even when differences in path conductance are experi-mentally imposed, phloem transport rates are sustained by adjustments to the pressure differences between the source and sink ends of the phloem path (Wardlaw 1990).

These conclusions have led to a shift in focus from phloem transport to phloem loading and unloading, which are instrumental in determining the amount of assimilate translocated and its partitioning between competing sinks, respectively.