JXB Advance Access originally published online on March 26, 2004
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Journal of Experimental Botany, Vol. 55, No. 399, pp. 1033-1044, May 1, 2004
© 2004 Oxford University Press
Regulation of Growth, Development and Whole Organism Physiology |
Tissue and cellular phosphorus storage during development of phosphorus toxicity in Hakea prostrata (Proteaceae)
Received 20 October 2003; Accepted 19 January 2004
1 School of Plant Biology, Faculty of Natural and Agricultural Sciences, the University of Western Australia, Crawley, WA 6009, Australia
2 CSIRO Division of Plant Industry, GPO Box 1600, Canberra, ACT 2601, Australia
* To whom correspondence should be addressed. Fax: +61 8 6488 1108. E-mail: hlambers{at}cyllene.uwa.edu.au
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Abstract |
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Storage of phosphorus (P) in stem tissue is important in Mediterranean Proteaceae, because proteoid root growth and P uptake is greatest during winter, whereas shoot growth occurs mostly in summer. This has prompted the present investigation of the P distribution amongst roots, stems, and leaves of Hakea prostrata R.Br. (Proteaceae) when grown in nutrient solutions at ten P-supply rates. Glasshouse experiments were carried out during both winter and summer months. For plants grown in the low-P range (0, 0.3, 1.2, 3.0, or 6.0 µmol d–1) the root [P] was > stem and leaf [P]. In contrast, leaf [P] > stem and root [P] for plants grown in the high-P range (6.0, 30, 60, 150, or 300 µmol P d–1). At the highest P-supply rates, the capacity for P storage in stems and roots appears to have been exceeded, and leaf [P] thereafter increased dramatically to approximately 10 mg P g–1 dry mass. This high leaf [P] was coincident with foliar symptoms of P toxicity which were similar to those described for many other species, including non-Proteaceae. The published values (tissue [P]) at which P toxicity occurs in a range of species are summarized. X-ray microanalysis of frozen, full-hydrated leaves revealed that the [P] in vacuoles of epidermal, palisade and bundle-sheath cells were in the mM range when plants were grown at low P-supply, even though very low leaf [P] was measured in bulk leaf samples. At higher P-supply rates, P accumulated in vacuoles of palisade cells which were associated with decreased photosynthetic rates.
Key words: Cluster roots, cryoSEM, proteoid roots, X-ray microanalysis.
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConcluding remarksReferences |
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Species of Proteaceae are more concentrated on the nutrient-impoverished soils in Western Australia than anywhere else in the world, and most of the taxa are endemic to heathlands (kwongan) and open woodlands of Mediterranean ecosystems of the south-west Botanical Province (Pate et al., 2001; Lambers et al., 2003). Two striking ecophysiological characteristics of Proteaceae are (i) a very low phosphorus (P) requirement, and (ii) a remarkable ability to mine P from poorly available soil P fractions (Specht and Groves, 1966; Jeschke and Pate, 1995; Handreck, 1997; Skene, 1998). The ability to mine P is attributed to the development of specialized ‘proteoid’ or ‘cluster’ roots (Purnell, 1960; Lamont, 1972, 2003; Dinkelaker et al., 1989) that exude large quantities of carboxylates (Grierson, 1992; Roelofs et al., 2001) and phosphatases (Grierson and Comerford, 2000). For many Proteaceae in the south-west of Western Australia, the development of proteoid roots, and hence P uptake, is restricted to wet winter months, whereas shoot growth mainly occurs during dry summer months (e.g. Banksia prionotes; Jeschke and Pate, 1995).
Since P uptake and shoot growth are separated by several months in Proteaceous species in the south-west of Australia, P acquired in winter must be stored, to be mobilized for growth in summer. The importance of storing this P in stem tissues has been demonstrated for B. prionotes (Jeschke and Pate, 1995) and B. ericifolia (Parks et al., 2000). The capacity of stem tissues to store P is limited. When P uptake exceeds the tissue storage capacity, P-toxicity symptoms develop in leaves (Parks et al., 2000).
Proteaceae and other species adapted to low-nutrient environments, typically develop foliar symptoms of phosphorus toxicity (P toxicity), even at very low rhizosphere [P] (e.g. in B. ericifolia, Ozanne and Specht, 1981; Parks et al., 2000; B. grandis, Lambers et al., 2002). The symptoms of P toxicity in Proteaceae are, generally, similar to those in other species, and include growth inhibition, early leaf senescence, and chlorotic and/or necrotic regions on leaves (e.g. B. serrata, Groves and Keraitis, 1976; B. ericifolia, Handreck, 1991; Parks et al., 2000; B. grandis, Lambers et al., 2002; Triticum aestivum, Asher and Loneragan, 1967). The physiology of P toxicity is not well understood, and symptoms are thought to result, either directly, from interference with leaf water relations at high cellular [P] (Bhatti and Loneragan, 1970), or from non-specific interactions of high cellular P with Zn or Fe (Marschner, 1995). The interaction of P with Zn and/or Fe produces symptoms in leaves that resemble micronutrient deficiencies (Robson and Pitman, 1983; Marschner, 1995; Lambers et al., 2002), and many studies that have described micronutrient deficiencies were, in fact, symptoms of P toxicity (AD Robson, personal communication).
After storage of P in winter, it must be remobilized and exported to growing tissues in summer. In B. ericifolia and B. prionotes, P is stored in bark and wood tissues of the stem; according to Jeschke and Pate (1995) the ray tissues may act as the primary sites for P storage in stems of B. prionotes. In stems of jarrah (Eucalyptus marginata), which co-occurs with some of the Proteaceae in south-western Australia, Dell et al. (1987) showed that P is mostly stored in bark. Raaimakers and Lambers (1996) made a similar observation for Lecythis corrugate, a rainforest tree species from Guyana, South America. Independent of the cell type (phloem, xylem, ray cells etc.) responsible for P storage in stem tissue, and the storage form of P (organic versus inorganic P), shunting excess P to stem tissue of B. ericifolia may help regulate the [P] of the leaves (Parks et al., 2000). Slow rates of growth have been implicated as an explanation for P accumulation by Proteaceae, but this cannot fully account for the development of P toxicity at low external P concentrations. According to Shane et al. (2003) P toxicity in Proteaceae is correlated with an inability to reduce P-uptake rates at elevated, but still relatively low, external P concentration.
P analysis at the organ level gives a first impression of the total amount of P in leaves or roots, but reveals nothing of the complexity of the intact organ or P distribution within it. The measurement of [P] in vacuole and cytoplasm by 31P-NMR has shown that large quantities of P can be stored in vacuoles, buffering the [P] in the cytoplasm (Lee et al., 1990). Cryo-scanning electron microscopy (CSEM) with energy-dispersive X-ray microanalysis (EDX) has further revealed that the distribution of ions in whole tissues varies in different cell types. This is illustrated by the finding of McCully (1994) that the highest [K+] of the different root cell types was in the developing xylem elements in soybean roots; in barley, Cl– accumulates preferentially in leaf epidermal cells when plants are exposed to salt stress (Huang and Van Steveninck, 1989). The analysis of [P] at the cellular level in Proteaceae will yield data that cannot be derived from the analysis of whole tissue extracts, and which may explain the special abilities of these plants to function at remarkably low tissue [P].
Considering the importance of P toxicity in the Proteaceae, which are a major component of the very diverse flora of the south-west of Australia (Pate and Beard, 1984), the growth and development and photosynthesis rates was characterized, and P toxicity symptoms were monitored in Hakea prostrata, a prominent member of this family, grown in nutrient solution at a range of P concentrations. The response of internal [P] to increasing external P-supply rates, which ultimately led to the development of P-toxicity symptoms, was also characterized. The [P] at the whole tissue level in roots, stems, and leaves was measured using CSEM with EDX to assess the [P] in vacuoles of different leaf cell types. The influence of micronutrients on the development of P toxicity is also reported.
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Materials and methods |
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High- and low-P range studies
Plant growth and culture: Eight-month-old soil-grown Hakea prostrata R.Br. (harsh hakea) seedlings were obtained from APACE nursery (North Fremantle, Australia). The root systems were washed free of soil, and each plant was transferred to a 7.0 l black plastic pot containing 6.0 l of continuously aerated nutrient solution of the following composition (in mmol m–3, 1.0 PO43–; 400 NO3–, 200 Ca2+, 200 K+, 154 SO42–, 54 Mg2+, 20 Cl–, 2.0 Fe-EDTA, 0.24 Mn2+, 0.10 Zn2+, 0.02 Cu2+, 2.4 H3BO3, and 0.3 Mo4+ made up in deionized water (pH 5.8). Shoots were gently supported in the centre of a grey foam lid that made a light-tight seal over each pot. Plants were grown for the first 4 weeks under shade cloth in a glasshouse and root temperature was maintained at 18–20 °C by placing the pots in a root-cooling tank. The complete nutrient solution was replenished each week for the first 4 weeks of growth.
Twenty plants of uniform size were selected for each of a ‘high-P range’ and ‘low-P range’ experiments. Plants were grown for 10 weeks at five P (supplied as KH2PO4) supply rates (low-P range 0, 0.3, 1.2, 3.0, or 6.0 µmol P d–1) (high-P range 6.0, 30, 60, 150, or 300 µmol P d–1) in the basal nutrient solution; The volume of nutrient solution was 6.0 l, and four plants were used in each treatment. The complete nutrient solution in each pot, including the P supply (µmol P d–1), was replenished daily for the duration of the experiment (10 weeks). Plants in the high-P range were grown from October to February (spring/summer 2000/2001), and those plants grown in the low-P range from June to September (winter 2001). The average ambient temperature in the glasshouse in winter was low rising to mid 20 °C, and in summer it was low to mid 30 °C. The average light intensity was approximately 1000 µmol m–2 s–1 in summer and approximately 500 µmol m–2 s–1 in winter.
Gas-exchange measurements and plant harvests: Photosynthesis and transpiration were measured on four of the youngest fully expanded leaves for each plant using a LI-6400 portable gas exchange system (Li-Cor, Lincoln, NE, USA) at 1500 µmol quanta m–2 s–1. Plants in the high-P range and low-P range were harvested after 10 weeks of treatments, and those in the leaf P-toxicity experiment, after 6 weeks. Root systems were washed for 1 min in a 1 mmol m–3 solution of K2SO4 to remove any P from the root surface that was transferred from the nutrient solution. Each plant was separated into stems, leaves, and cluster and non-cluster roots. Samples were weighed fresh, and after drying for 8 d at 80 °C.
Total phosphorus concentrations: Dried material was ground with a mortar and pestle. Total P concentration was determined colorimetrically using a UV-VIS spectrophotometer (Shimadzu Corporation, Japan) by the malachite green method (Motomizu et al., 1983) after digestion of ground material in hot concentrated HNO3:HClO4 (3:1).
Determination of leaf micronutrient concentrations: Digests (described above) of ground material were analysed for Zn, Cu, Mn, and Fe by atomic absorption spectrometry (AAS). The micronutrient concentrations in the samples were determined by comparison with that in digests of appropriate standards from the Australasian Soil and Plant Analysis Council (ASPAC).
Development of phosphate toxicity in leaves
Plant growth and culture: H. prostrata plants (3-months-old), also obtained from APACE nursery, were washed free from soil (as above), and prepared for hydroponics (for 4 weeks as above). Eighteen plants of uniform size were selected for the P-toxicity experiments and supplied with basal nutrient solution (as above), supplemented at three P-supply rates, 0.8, 100, or 200 µmol P d–1. Nutrient solutions were replaced daily for the duration of the P treatments (6 weeks). These plants were grown during spring and summer and were used for the CSEM and EDX investigation.
Cryo-scanning electron microscopy and X-ray microanalysis: One age-class of mature leaves from plants grown at 0.8, 100, and 200 µmol P d–1 was selected. Older, mature leaves were chosen over young leaves because the youngest leaves are not a suitable indicator of nutrient status for nutrients like P that are easily retranslocated (Marschner, 1995). The leaves were frozen intact by plunging into liquid nitrogen (LN2) divided into pieces to fit into cryo-vials, shipped to Canberra in a cryo-shipper, and stored in a cryo-store. A section (
4 mm) was cut from each frozen leaf under LN2, and quickly secured in the groove of an aluminium stub containing low temperature Tissue Tek by plunging into LN2. The attached specimen was transferred under LN2 to a cryo-microtome (Reichert-Jung, Vienna) and a smooth transverse face planed with glass and diamond knives at –80 °C, transferred in LN2 to a cryo-transfer unit (Oxford Instruments, Eynsham, UK) and hence to the cryo-stage of the SEM (Jeol, 6400; Jeol Ltd., Tokyo, Japan). The planed surface was then lightly etched; the cell outlines were revealed by slowly warming to –90 °C while observing it continuously at 1 kV. The specimen was then re-cooled –170 °C, coated with 50 nm high purity aluminium, and examined at 10–15 kV. Images were captured digitally.
Microanalysis of epidermal, bundle-sheath, and palisade cells was with a Link eXL, LZ-4 detector (Oxford Instruments), using the Be-window. The accelerating voltage was 15 kV, take-off angle 33°, working distance 35 mm, and probe current set at 1.0 nA. The magnification was varied so that the 10 µm (at 1000x) scan raster covered the lumen of the cell, but did not touch the walls. Spectra were accumulated to 80 000 counts for Al. Phosphorus peaks (net counts) were expressed as percentages of the aluminium peak, and adjusted for coating thickness by dividing by the live time. Absolute [P] was determined by comparison with the appropriate standards prepared for analysis by procedures identical to those for the specimens (for further details see Huang et al., 1994; McCully et al., 2000). The X-ray microanalytical system is relatively insensitive to low concentrations of elements in frozen, fully-hydrated tissue, and the lower limit of reliable detection of elements by the X-ray microanalysis was considered about 10 mM. Therefore, data that fell between 0–10 mM were ascribed a notional value of 5.0 mM. The translucent, ropey strands that are exuded on to the surface of older leaves exhibiting toxic P symptoms were examined in the cryo-SEM on leaf pieces sputter-coated with gold or with evaporated aluminium.
Analysis of the exudate from the leaf surface by high-pressure liquid chromatography (HPLC) and inductively coupled plasma mass spectrometry (ICPMS)
Translucent material which was exuded from the stomata of leaves showing P toxicity was solubilized in either mobile phase, 25 mM KH2PO4 (pH 2.50) (for HPLC analysis) or nitric acid (for ICPMS analysis). For HPLC analysis, organic acids were separated (Alltima C-18 [250x4.6 mm internal diameter with 5 µm packing]) and identified using Waters® HPLC (600E pump, 717 auto injector, and 996 photodiode-array detector, Milford, MA, USA). Detection was at 210 nm and spectrum matching and peak purity analysis was according to Lambers et al. (2002). The sample injection volume was 100 µl. Data acquisition and processing used Millennium© software (Waters, Milford MA, USA). Retention times of organic acid standards: tartaric, formic, malic, malonic, lactic, acetic, maleic, citric, succinic, fumaric, cis-aconitic, and trans-aconitic acid were used to identify organic acids.
For ICPMS analysis, concentrations of elements were determined using a model PE ELAN 6000 inductively coupled plasma mass spectrophotometer (ICP-MS) instrument (Perkin-Elmer, Norwalk, CT, USA), in combination with a hydride generation unit (Perkin-Elmer, B050–5540) or a flow injection unit (Perkin-Elmer, FIAS 400).
Statistics
Data were analysed with one-way analysis of variance (ANOVA) with P concentration as the main fixed factor in the model (SPSS Version 8.0, 1998). Specific differences amongst treatments were tested using LSD multiple range tests (
=0.05) following the ANOVA. To ensure normality and homogeneity of variances, data were log transformed where necessary.
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Results |
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Plant growth and cluster-root development at high and low external P supply
Plants grown at the low-P range (0, 0.3, 1.2, 3.0, and 6.0 µmol P d–1, added to a pot containing 6.0 l nutrient solution) developed slowly during the winter months. There was no significant influence of the lowest three P-supply rates (0, 0.3, and 1.2 µmol P d–1) on plant dry mass (Fig. 1A). When the external P-supply rate was increased to 3.0 µmol P d–1, the dry mass of roots, stems, and leaves increased significantly. The dry mass of stems and leaves increased further when the P-supply rate was 6.0 µmol P d–1. The cluster-root dry mass was the same for plants at all P-supply rates in the low-P range (Fig. 1A), but the proportion of cluster roots to that of total root mass decreased significantly with increasing P-supply rate (Fig. 2A). Towards the end of the experiment, the oldest leaves on plants deprived of P turned yellow and senesced, most likely due to development of P shortage.
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Plants that developed during the summer months in the high-P range (6, 30, 60, 150, and 300 µmol P d–1) accumulated far more dry mass over the 6-weeks P treatments (Fig. 1B) than those grown for 6 weeks during the winter months (Fig. 1A); even at the same rate of P supply (6.0 µmol P d–1), there was a large difference (Fig. 1B). This is probably due to the longer day-length, higher light intensity, and/or higher temperature in summer. Root dry mass increased significantly when the P-supply rate increased from 6.0 to 30 µmol P d–1; it decreased significantly for plants grown at higher external P-supply rates.
The cluster-root dry mass and the proportion of cluster roots decreased across H. prostrata plants grown at higher P-supply rates from 6 to 60 µmol P d–1 (Fig. 2A), and no cluster roots developed on plants supplied with 150 or 300 µmol P d–1 (Fig. 2B).There was no effect on stem and leaf dry mass for plants grown between 6 and 150 µmol P d–1. Plants grown at P-supply rates of 300 µmol P d–1 had a significantly decreased root and stem dry mass (Fig. 1B), and the older leaves had symptoms of P toxicity.
Net photosynthesis
Faster rates of photosynthesis of the youngest fully expanded leaves coincided with increased P-supply rates. In the low-P range, increasing the P-supply rates from 0 to 6.0 µmol P d–1 was related to ten-times faster (from 1.1 to 13 µmol CO2 m–2 s–1) net rates of photosynthesis (Fig. 3A). In the high-P range the photosynthetic rates of the youngest fully expanded leaves increased significantly when P-supply rate increased from 6.0 to 30 µmol P d–1; thereafter, rates significantly deceased at P-supply rates of 150 and 300 µmol P d–1 (Fig. 3B).
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For plants grown in the high-P range the decreased photosynthetic rates at higher P-supply rate also coincided with the higher leaf [P] (Fig. 4). The photosynthetic rates were highest when leaf [P] was 0.9–1.6 mg g–1 leaf DM (Fig. 4), similar to that determined in bulked leaf samples from plants grown at the same P supply rate (Fig. 5B).
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Differences in P concentrations amongst leaves, roots, and stems
In the low-P range the root, stem, and leaf [P] approximately tripled when the P-supply rate was increased from 0 to 6.0 µmol P d–1. Root [P] significantly increased at each of the three lowest P-supply rates (0, 0.3, and 1.2 µmol P d–1). Even at these low P-supply rates the stem [P] increased before leaf [P]. When the P-supply rate was 3.0 or 6.0 µmol P d–1, root, stem, and leaf [P] significantly increased (Fig. 5A).
In the high-P range the root, stem, and leaf [P] increased to different degrees with increasing P-supply rate (Fig. 5B). Stem [P] saturated when the P-supply rate was 60 µmol P d–1. Root [P] also saturated, but not until the rate of P supply was 150 µmol P d–1. When the P-supply rate was 300 µmol P d–1, after both root and stem [P] were saturated, leaf [P] increased dramatically (Fig. 5B). On plants grown at the two highest P-supply rates (150 and 300 µmol P d–1), leaves that senesced after developing symptoms of P toxicity had a leaf [P] of 10±0.44 mg P g–1 dry mass.
For comparison of the hydroponically grown plants with plants in their natural environment, living (mature) and senesced (litter) leaves of H. prostrata plants growing in native bushland at Kings Park and Botanic Garden, Perth, were examined. The mature leaf [P] of native plants was similar (0.44±0.04 mg P g–1 DM) to that of the hydroponically grown plants supplied with a 6.0 µmol P d–1 both in winter (low-P range; 0.38±0.023 mg P g–1 DM) and in summer (high-P range; 0.36±0.048 mg P g–1 DM).
Leaf Cu, Fe, Mn, and Zn concentrations
The leaf [Cu] and [Fe] did not change significantly with increasing P-supply rate for plants grown in the low-P range, whereas the [Mn] and [Zn] decreased significantly when the P-supply rate was increased from 0 to 6.0 µmol P d–1 (Table 1). In the high-P range the [Cu], [Fe], [Mn], and [Zn] did not vary significantly amongst P supplies from 6.0–300 µmol P d–1 (Table 1). Leaves that had senesced after developing symptoms of P toxicity had elevated [Mn] and, to some extent, [Fe] and [Zn]. By contrast, the [Fe] and [Mn] in leaves senesced naturally from old age in native bushland plants were increased relative to those of the mature (living) leaves (Table 1).
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Microscopy and in situ analysis of leaves with symptoms of P toxicity
Development and composition of leaf exudates, and leaf structure: A separate batch of H. prostrata plants (i.e. at 0.8, 100, and 200 µmol P d–1) was cultivated, see Materials and methods to provide the additional material for the investigation of leaf P toxicity by CSEM. All of the symptoms of P toxicity subsequently described for plants grown at 100 and 200 µmol P d–1 were also observed in the earlier, high-P range experiments when plants were grown at 150 and 300 µmol P d–1.
Plants grown at low P supply (i.e. 0.8 µmol P d–1) looked healthy and had dark-green leaves while those grown at 100 or 200 µmol P d–1 looked healthy initially, but after 3 weeks the oldest leaves developed symptoms of P toxicity. All the plants supplied with 100 or 200 µmol P d–1 developed olive-green patches on one or two of their oldest leaves (Fig. 6A) just as observed in earlier high-P range experiments (i.e plants grown at 150 and 300 µmol P d–1). The olive green patches on these leaves subsequently turned a uniform light ‘chalky-green’ colour, dried out, and senesced. At least three leaves senesced on each of these plants during the experiment. On some of these leaves, gel-like, ropey, strands of translucent material covered portions of the lamina (Fig. 6B). These strands appeared to emerge through the stomata over the olive-green patches (Figs 6A, B, 7A, B), like toothpaste squeezed from a tube. They eventually spread over large patches of the leaf surface with some strands longer than 10 mm (Fig. 7B). The strands somewhat resembled fungal hyphae, but did not stain with Toluidine blue at pH 4.4 (O’Brien and McCully, 1981).
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Only Ca was detected in the strands by X-ray microanalysis, but trace amounts of P as well as Ca was detected by ICPMS (data not shown). The strands were partly soluble in water and alcohol, and HPLC analysis indicated that no organic acids were present with retention times similar to those of the standards.
Cryo-SEM of the transverse faces of cryo-planed leaves clearly revealed that the leaf palisade mesophyll is arranged bilaterally with two or three layers on each side of the leaf. Spongy mesophyll was absent. The amount of airspace was surprizing large for a xeromorphic plant and in leaves of plants grown with 0.8 µmol P d–1, there were abundant internal air spaces around the palisade mesophyll (Fig. 8A).
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A similar arrangement of cells was observed in some leaves of plants grown with 100 and 200 µmol P d–1, but, in other leaves of these plants, areas of palisade mesophyll cells were swollen so that no internal airspaces remained (Fig. 8B). These regions probably coincided with the olive green patches observed on the surface of fresh leaves, but this was not determined definitively.
P concentration in leaf cells determined by X-ray microanalysis: Cellular [P] was measured in fully expanded, mature leaves. For plants grown at the highest P-supply rates (i.e. 100 and 200 µmol P d–1) the leaves had not yet developed visible P-toxicity symptoms and looked ‘healthy’ (i.e. one age-class younger than those that had developed visible P toxicity symptoms) (Fig. 9).
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The response of leaf cell [P] to P-supply rate (0.8, 100, and 200 µmol P d–1) varied amongst epidermal, bundle-sheath, and palisade cells. The palisade cell [P] increased significantly from 5.0±2.0 mM at a P-supply rate of 0.8 µmol P d–1 to 136±8.0 mM at a P-supply rate of 100 µmol P d–1 (c. 42x) (Fig. 9). The [P] increased further (c. 1.7x) to 233±22.6 mM when the P-supply rate was 200 µmol P d–1 (Fig. 9). Bundle-sheath cell [P] increased (c. 5x) from 5.0 mM to 29 mM at P-supply rates from 0.8–200 µmol P d–1, but was not significant. The epidermal cell [P] tended to increase when the P-supply rate was increased from 0.8 to 100 µmol P d–1 and was significantly increased (c. 12x) when the P-supply rate was 200 µmol P d–1 (Fig. 9).
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Discussion |
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Is P toxicity in H. prostrata different from that in other species?
The P-toxicity symptoms in H. prostrata are clearly similar to those of other species. The ‘wet-like’ olive-green patches that were observed in the initial phase of P-toxicity symptoms are similar to the ‘translucent’ patches described by Warren and Benzian (1959) for Lupinus luteus (yellow lupin). The chlorotic and necrotic leaf symptoms in H. prostrata show many similarities to those reported for other Proteaceae, as referenced in the Introduction, and for a range of crop species (Table 2). Exceptionally in H. prostrata was the extrusion of the ropey strands of material from the stomata which, as far as the authors are aware, has not been reported elsewhere in the literature. The significance and composition of this translucent material is not known, but its formation is probably associated with the death of leaf cells from P toxicity.
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A comparison of the leaf [P] at which P toxicity occurred in H. prostrata (approximately 10 mg g–1 DM) with that of other species, showed that this concentration agrees closely with the leaf [P] (or shoot [P]) (range 0.9– 47 mg P g–1 DM) associated with development of P-toxicity symptoms in many other Proteaceae (e.g. Hakea gibbosa, Grundon, 1972; Banksia species, e.g. B. serrata, Groves and Karaitis, 1976; B. robur, B. aemula, and B. oblongifolia, Grundon, 1972; B. ericifoloia; Handreck, 1991) (Table 2). In fact, these leaf P concentrations for H. prostrata are similar to values reported for many crop plants (e.g. wheat, Bhatti and Loneragan, 1970; tomato, Jones, 1998) (Table 2).
It was found, using the present data in comparison with those in the literature, that the plant [P] associated with P toxicity most certainly depends upon the tissues analysed; lower concentrations for bulked samples of shoots and leaves, and very high [P] for necrotic lesions (Table 2). The present results on H. prostrata and published measurements (Table 2) demonstrate that leaves and shoot tissue in Proteaceae are not more sensitive to [P] than those of other species. Some of the species in Table 2, including H. prostrata, are, however, at the lower end of the range of leaf [P] that is related to the development of P-toxicity symptoms (Table 2). The second essential component of the authors’ argument is to consider the validity of the comparison of the [P] in mesic leaves of crop species with those in scleromorphic leaves of Proteaceae (Grose, 1989). They are not readily comparable on a dry mass basis, because P is ‘diluted’ by the increasing proportion of sclerenchyma in leaves from species in Mediterranean regions of Australia (Specht and Rundel, 1990; Foulds, 1993). However, the data can also be compared on a fresh mass basis. The dry matter content for leaves of H. prostrata was 30%, assuming a value of 15% for crop species in general. This estimation of leaf (or shoot) [P] at which P toxicity occurs for H. prostrata (and the Proteaceae in Table 2) is 0.27–14.1 mg P g–1 FM (not including necrotic regions), and for crop species in general it is 0.51–4.2 mg P g–1 FM. This shows that enhanced tissue sensitivity to [P] in H. prostrata cannot explain their greater susceptibility to P-toxicity symptoms at relatively lower external [P].
Development of P toxicity in H. prostrata at low external [P] is characteristic of many Proteaceae, and raises the question whether plants with cluster roots are more susceptible to P toxicity. It has been reported that other root adaptations that enhance nutrient uptake are associated with the development of higher internal [P] at relatively low external [P]. For example, plants with mycorrhizas accumulate higher internal [P] at lower external P supply (Mosse, 1973), and in studies with soybean plants, Foote and Howell (1964) showed that the rootstocks from a ‘P-sensitive’ line were more important than the shoots for determining P toxicity. Whether or not cluster roots increase the susceptibility of Proteaceae to P toxicity requires further investigation, but measurements of net P-uptake rates in H. prostrata grown at a range of P in nutrient solution revealed an inability to down-regulate net P-uptake rates at elevated external or internal [P] that may lead to the development of P toxicity (Shane et al., 2003).
P storage in root and stem tissue may prevent leaf P toxicity
The ability to store P is an important trait for Proteaceae (e.g. Banksia prionotes; Jeschke and Pate, 1995) in their natural habitats, because P uptake and shoot growth are temporally separated. The storage of [P] in stems of H. prostrata grown at increasing P-supply rate is similar to that in B. prionotes (Jeschke and Pate, 1995) and B. ericifolia (Parks et al., 2000). The fact that roots of H. prostrata were also an important storage organ for P suggests that P storage is a characteristic of woody tissues in H. prostrata, and perhaps most Proteaceae. However, further investigation to localize the cell type responsible for P storage is needed.
The development of P-toxicity symptoms in H. prostrata highlights the second major function of P storage; that is, to shunt excess P to root and stem tissues, rather than to leaf tissue, thereby maintaining a buffer against large increases in leaf [P]. In most natural habitats the capacity of stem and root tissues for P storage would exceed the P influx. By contrast, in hydroponics or fertilized soil cultures the availability of P can be drastically enhanced, and exceed the capacity of stem and root tissues to store it. Then P accumulates in leaves, leading to symptoms of P toxicity.
Implications of directing P to palisade cells
This is the first demonstration of high [P] in leaf cells of a Proteaceous species when bulked leaf [P] on a dry mass basis was very low. For H. prostrata, cellular [P] in the intact leaf was not dramatically lower than that reported for crops grown under low-P conditions (Treeby et al., 1987; Marschner, 1995). Millimolar [P] were measured in vacuoles of epidermal, bundle-sheath, and palisade leaf cells when bulked leaf [P] was as low as 0.23–0.33 mg P g–1 DM.
The reliability of the present microanalysis estimates for vacuolar [P] in different leaf cell types can be verified by estimating it from the data on bulk leaf [P], since the cellular [P] can be roughly calculated from those data. The leaves of H. prostrata (grown under the present conditions) contained approximately 70% water; therefore, based on the measured leaf [P] (on plants grown with 0.8 µmol P d–1), they contained, on average, a total of 0.476 mg P. Assuming that all this P is in solution, and that the distribution of P across leaf cells (all types) is homogeneous (which it is not) the final concentration of P would be 3.7 mM. For plants grown at P-supply rates of 100 and 200 µmol P d–1 the final P concentration would be around 92 mM. The measurements of [P] determined here by X-ray microanalysis in leaf cells of fully hydrated frozen leaves were clearly in this range. As outlined in the Materials and methods, only one age class of leaves was measured, but it is expected, based on data from bulk leaf [P], that [P] would be even higher in vacuoles of cells from leaves with visible symptoms of P toxicity.
There is no doubt that H. prostrata and other Proteaceae are able to function well at very low leaf [P] (in H. prostrata 0.33 mg P g–1 DM) compared with other species (approximately 1 mg P g–1 DM in crop plants) (Marschner, 1995; Lambers et al., 1998), but there are no physiological explanations for this. Even if leaf [P] values are expressed on a fresh mass basis for plants at the lowest P-supply rates, they are still very much lower than those reported as being adequate for growth of crops (Epstein, 1972; Foulds, 1993; Marschner, 1995). At the highest P-supply rates of 100 and 200 µmol P d–1, P preferentially accumulated in leaf palisade cells over that of epidermal and bundle sheath cells, which ultimately leads to P becoming toxic. By contrast, studies that measured [P] in leaf cells of Vicia faba (Outlaw et al., 1984) and Lupinus luteus (Treeby et al., 1987) grown at a range of P supplies, showed that epidermal cells accumulated greater vacuole [P] than mesophyll cells. If preferential accumulation in palisade cells also occurred at much lower external P concentrations, then it could explain the maintenance of reasonable photosynthetic rates at very low leaf [P], and the significantly increased rates of photosynthesis in H. prostrata even when bulk leaf [P] rises only slightly. If the P arriving in leaf tissue via the xylem (and possibly recirculated in the phloem) were preferentially directed to where it was most needed, as evident from the increased [P] in palisade cells, it indicates that the very low P requirement of the Proteaceae may, in part, be related to a high capacity for optimizing distribution at the leaf level. The usefulness of heterogeneous elemental ion distribution is illustrated by other studies that have shown that sequestration of Cl– in epidermal cells maintains low mesophyll [Cl–] in barley plants grown under salt stress (Huang and Van Steveninck, 1989). In H. prostrata further evidence is provided supporting this notion as observed by the lack of response of bundle-sheath cell [P] at higher P-supply rate. This is interesting from both an anatomical and a physiological point of view. The P transported from the xylem or phloem sap to the palisade cells must pass through or between bundle sheath cells that surround the vascular tissues, on the way to the palisade cells. This indicates bundle sheath cells removed very little P.
Leaf [Zn], [Mn], [Fe], and [Cu], are independent of the progression of leaf P toxicity
As shown in Table 1, the differences in micronutrient concentrations ([Fe], [Zn], [Mn], or [Cu]) in leaves of H. prostrata grown at the various P-supply rates do not explain the differences in growth or the development of different proportions of cluster roots. The differences in leaf [Fe] probably depend on transpiration rate, with little redistribution due to its low phloem mobility (Marschner, 1995; Lambers et al., 1998). This would also account for higher [Fe] in the senesced leaves of the native bushland plants, and the slightly elevated [Fe] in mature leaves that senesced prematurely, after developing symptoms of P toxicity. The present hydroponically grown H. prostrata plants had similar micronutrient status ([Fe], [Zn], [Mn], and [Cu]), even when grown at different P-supply rates, to that of leaves in native-bush plants.
P toxicity developed in H. prostrata leaves when no significant change occurred in leaf [Fe] or in that of the other micronutrients in response to P-supply rate. Similar findings have been reported for the ‘phosphate-accumulator’ mutant of Arabidopis thaliana where leaf P toxicity corresponded with leaf [P] of 14.5 mg P g–1 DM, but micronutrient status was unchanged (Delhaize and Randall, 1995). Even though the concentration of the micronutrients is similar, high leaf P concentrations such as those in Table 2, would decrease the solubility of Zn (Cakmak and Marschner, 1987), and might even precipitate Fe as Fe-phosphates, thus causing P-induced micronutrient deficiencies. Furthermore, it is expected that 80% of the P would be present largely as orthophosphate salts in necrotic regions of leaves with P toxicity (Table 2), having a negative impact on the leaf water relations (Bhatti and Loneragan, 1970).
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Concluding remarks |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionConcluding remarksReferences |
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Roots and stems of H. prostrata have a large capacity to store P, to be used, in their natural habitat, during the summer when shoot growth predominates. Storage of phosphorus is necessary, both because of the seasonality of the cluster-root development and P uptake (winter) and shoot growth (summer), and to maintain low leaf [P]. When the capacity of stem and root tissue to store P is exceeded, leaf [P] rises dramatically, leading to development of P-toxicity symptoms. In H. prostrata, and perhaps many other Proteaceae, leaf P toxicity, through accumulation of excess P in leaves, is not due to enhanced tissue sensitivity. The distribution of [P] amongst different cell types in leaves is not homogeneous. Accumulation of P by palisade cells may be advantageous or typical of plants adapted to function at very low [P], but may lead to toxic [P] in palisade cells at elevated external P supply.
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Acknowledgements |
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We thank Cheng Huang, Research School of Biological Sciences, Australian National University, Canberra, and Celia Millar and Rosemary White, CSIRO Microscopy Centre, Canberra for help with the cryo-analysis. Martin de Vos and Sytze de Roock helped grow the plants, and were supported by Utrecht University. MW Shane was the recipient of an International Postgraduate Research Scholarship and a University of Western Australia Postgraduate Award. We also thank Greg Cawthray (HPLC) and Michael Smirk (ICPMS and AAS) for their assistance.
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