The immunocytochemical localization of the plasma membrane H+-ATPase in epidermal cells of tomato roots was studied using a monoclonal antibody raised against purified maize P-type H+-ATPase. Plants subjected to iron starvation exhibited increased proton extrusion that was confined to the root elongation zones. Immunogold labelling of the H+-ATPase on the plasma membrane was considerably higher in rhizodermal cells within zones with intense proton extrusion than in non-acidifying areas of the roots. Transfer cells were formed in rhizodermal cells of Fe-deficient plants. Quantitative determination of immunolabelling revealed that the density of PM H+-ATPase in transfer cells was about twice that of ordinary epidermal cells. In transfer cells, H+-ATPase was most abundant on the plasma membrane lining the labyrinthine invaginations of the peripheral cell wall. While the number of immunologically detectable ATPase molecules in transfer cells was not spatially correlated with proton extrusion activity, the frequency of transfer cells was considerably higher in acidifying root areas relative to non-active segments. Split-root experiments indicated that both the steady-state level of plasma membrane H+-ATPase and proton extrusion activity are systemically regulated, indicating inter-organ regulation of rhizosphere acidification. Exogenous application of the auxin analog 2,4-dichlorophenoxyacetic acid and the ethylene precursor 1-aminocyclopropane-1-carboxlic acid caused the formation of transfer cells at a frequency similar to that observed in Fe-deficient roots. However, the number of proton pumps was not affected by the hormone treatment, suggesting that both responses are regulated independently. It is concluded that transfer cells in the rhizodermis may be important but not crucial for rhizosphere acidification.
Root epidermal cells may develop root hairs, and they may undergo highly regulated structural and functional differentiation processes which are controlled by multiple gene loci and are affected by the environment (Schiefelbein 2000; Cavell & Grierson 2000). Shortage in iron supply, for example, causes an increase in root hair length and number, providing for a more efficient uptake of nutrients. The development of rhizodermal transfer cells, bearing wall ingrowths located primarily along their outer tangential walls, has been observed in several so-called strategy I species in response to suboptimal Fe availability, as part of a suite of adaptive mechanisms (Römheld & Marschner 1986). The strategy I syndrome includes the induction of a plasma membrane-bound Fe(III) chelate reductase encoded by the FRO2 gene, an enhanced expression of a Fe(II) transporter mediating the uptake of ferrous iron from the soil, and an increased activity of a plasma membrane-bound H+-ATPase providing a reduction of the rhizosphere pH that may enhance the mobilization of sparingly soluble iron species (Eide et al. 1996; Robinson et al. 1999; Dell'Orto et al. 2000; Guerinot 2000; Eckhardt, Mas Marques & Buckhout 2001; see Schmidt 1999 for a review).
Clear evidence for an involvement of transfer cells in the physiological responses to iron shortage has not yet been presented. Root zones of high Fe(III)-reducing capacity did not correlate with transfer cell frequency, thus questioning an involvement of transfer cells in the reduction of Fe chelates (Schmidt & Bartels 1996). Transfer cells in other tissues such as developing seeds, cotelydons, and leaf minor veins have shown to be enriched in H+-ATPase (Bouché-Pillon et al. 1994a, b; Tegeder et al. 1999; Farley, Patrick & Offler 2000). Enhanced H+-ATPase expression can be assumed for rhizodermal transfer cells induced by salt stress, since NaCl regulation of plasma membrane H+-ATPase gene expression has been reported for several plant species (Niu et al. 1993; Zhang et al. 1999; Hasegawa et al. 2000). No data on the number of plasma membrane ATPase molecules are available for transfer cells induced by iron stress. The role of rhizodermal transfer cells in the physiological responses to Fe deficiency is thus still open.
In this study we examined whether transfer cell formation is coupled to enhanced acidification of the rhizosphere, and demonstrated, using monoclonal antibodies, that the steady-state level of H+-ATPase was affected by the iron nutritional status and differed between cell types in the root epidermis in Fe-deficient plants. A further objective of this work was to elucidate the involvement of hormones in the induction of plasma membrane H+-ATPase activity.
MATERIALS AND METHODS
Plant materials and growth conditions
Lycopersicon esculentum Mill. seeds were germinated in perlite moistened with medium of the following composition (mm): KNO3 (3), MgSO4 (0·5), CaCl2 (1·5), K2SO4 (1·5), NaH2PO4 (0·5); (µm): H3BO3 (25), MnSO4 (1), ZnSO4 (0·5) (NH4)6Mo7O24 (0·05), CuSO4 (0·3), and 40 µm Fe-ethylenediaminetetraacetic acid (FeEDTA). Fourteen days after emergence, seedling were transferred to 11 L tanks (40 plants/tank) containing continuously aerated nutrient solution and grown in a greenhouse for an additional 10 d under the following conditions: 40 to 55% relative humidity, 16 h, 26 °C/8 h, 22 °C day/night regime, photon flux density of approximately 150 µmol m−2 s−1 supported by Philips IP23 lamps (Eindhoven, The Netherlands). The seedlings were then transplanted into either complete solution (+Fe plants) or iron-free medium (–Fe plants) for 5 d. The nutrient solution was replaced every 7 d, and the pH was adjusted to 6·0 with KOH. Hormone-treated plants were grown for 2 d in medium supplemented with 0·1 µm 2,4-dichlorophenoxyacetic acid (2,4-D) or 1 µm 1-aminocyclopropane-1-carboxlic acid (ACC).
For the split-root experiments, plants were grown hydroponically as described above in the presence of 40 µm FeEDTA. After 6 d, plants were grown with their root systems divided into two approximately equal parts, using containers with two separate compartments (5·5 L each) holding nutrient solution with either 40 µm FeEDTA or no Fe added.
Visualization of proton extrusion
Spatial localization of acidification capacity was determined by embedding the roots of plants in an agar (0·7%) medium containing 0·5 mm CaSO4 and the pH indicator Bromcresol Purple (0·005%) for 20 min. The pH of the medium was adjusted to pH 6·0. The roots had been grown for 5 d in either full nutrient solution, Fe-free medium, or in medium supplemented with 10 µm ACC or 0·1 µm 2,4-D for 2 d before the pH assay was carried out. For electron microscopic analysis, stained root segments were cut off the gel with a razor blade and used for the determination of transfer cell frequency after washing in 0·5 mm CaSO4.
Analysis of root hair patterns was performed by light microscopy in dark field. Photomicrographs were recorded on negative film (Agfa 100, Leverkusen, Germany).
Standard electron microscopy
Roots were cut into approximately 1 cm long segments and subsequently washed in 0·5 mm CaSO4. The segments were fixed overnight in 0·1 m potassium phosphate buffer, pH 7·4, containing 0·5% (w/v) glutaraldehyde and 1·5% (w/v) paraformaldehyde. After being rinsed three times in 0·1 m potassium phosphate buffer, pH 7·4, the tissue was dehydrated through a graded ethanol series of 20, 40 and 50%, and post-fixed in 0·3% (w/v) osmium tetroxide for 8 h in 50% ethanol at −20 °C. Root segments were washed again in 50% ethanol (three times for 10 min) and treated in a solution of 0·3% (w/v) uranyl acetate in 50% ethanol overnight at 4 °C. The material was washed again twice in 50% ethanol (10 min each). The samples were then dehydrated in an ethanol series of 75%, 90% and twice at 100% for 30 min each, infiltrated with London Resin White (London Resin Co. Ltd. London, UK), and polymerized at 55 °C for 24 h in vacuo. Ultrathin sections were cut with a Reichert (Vienna, Austria) Ultracut E microtome and stained with uranyl acetate and lead citrate. Sections used for electron microscopy were examined in a Zeiss (Jena, Germany) EM 902 A electron microscope.
For the cytochemical localization of the ATPase a similar protocol was used with the following changes. The segments were fixed without glutaraldehyde and subsequently washed and dehydrated at 0 °C. The tissue was post-fixed in 0·05% (w/v) osmium tetroxide for 2 h in 50% ethanol at −20 °C. Infiltration with London Resin White was carried out at 0 °C. The samples were polymerized at 4 °C under UV light (365 nm) for 60 h.
On-section immunogold staining was carried out on formvar-coated nickel grids at 20 °C. The sections were etched by 0·5 mm HCl and 0·56 m NaIO4, and washed three times for 3 min. Non-specific sites were blocked for 30 min with 1·0% (w/v) bovine serum albumin (BSA) and 0·2% acetylated BSA in TBS buffer (100 mm Tris-HCl, 0·9% (w/v) NaCl, pH 7), and then for 5 min in the same solution. Monoclonal antibodies 46E5B11D5 against maize plasma membrane H+-ATPase (Villalba, Lützelschwab & Serrano 1991; Palmgren & Christensen 1994; Baur et al. 1996; Jahn et al. 1998) were diluted 1 : 50 with blocking solution and incubated overnight at room temperature. After washing in blocking solution three times for 5 min each, grids were incubated for 2 h in goat antimouse IgG, 1 : 50 dilution, labelled with 15 nm gold particles. The grids were washed in blocking solution three times for 5 min each and three times for 5 min each in double-distilled H2O. The sections were observed with a Zeiss EM 902 A electron microscope. For estimating plasma membrane ATPase density, gold particles of an average of about 30 epidermal cells from 12 root segments per growth type (from eight plants in average) were analysed. Quantification of gold particles was made per 10 µm of plasma membrane contour.
Iron deficiency alters patterning of epidermal root cells
Consistent with previous findings, the growth of plants in iron-free nutrient solution caused the formation of extra root hairs and induced the development of transfer cells in the root epidermis. The ultrastructure of rhizodermal cells is shown in Fig. 1. Ingrowth depositions in the rhizodermis were also induced by application of the auxin analog 2,4-D or the ethylene precursor ACC. As in other species, tomato rhizodermal transfer cells exhibited an extensive labyrinth of secondary wall material, numerous mitochondria, Golgi vesicles, and intense endoplasmic reticulum. The wall invaginations were polarized to outer periclinal walls. No differences in cell structure were observed between transfer cells induced by iron deficiency and those formed in plants grown in the presence of hormones. In the roots of Fe-sufficient plants, secondary wall ingrowths were occasionally observed. In these cases the transfer cells were restricted to small zones of rhizosphere acidification (see below). No transfer cells were observed in non-active zones of Fe-sufficient roots.
Proton extrusion is correlated with the distribution of plasma membrane ATPase
To determine whether the expression of H+-ATPase correlates spatially with proton efflux, we embedded plants in an agar medium containing the pH-sensitive dye Bromcresol Purple. Visualization of proton extrusion from plants grown with or without iron is shown in Fig. 2A and B, respectively. A slight staining of the dye was observed along the whole root system of Fe-sufficient plants. In roots of Fe-deficient plants, acidification spots were clearly more pronounced and restricted to distinct zones located in the elongation zones behind the tips (Fig. 2B). Figure 2C and D will be discussed below. In general, acidification spots were located above the elongation zone of the roots (Fig. 3). The zone of metaxylem differentiation, in which the transfer cells are mainly located, only partly overlaps the area of active acidification.
To investigate the subcellular localization of H+-ATPase in root epidermal cells, ultrathin sections from root elongation zones were immunolabelled with the monoclonal antibody 46E5B11F6 against the N-terminal region of plasma membrane ATPase from maize roots (Villalba et al. 1991; Baur et al. 1996; Jahn et al. 1998). This antibody was shown to immunoreact with PM-H+-ATPase from roots of both monocots and dicots. For example, the antibody cross-reacts with different H+-ATPase isoforms of Arabidopsis (AHA1, AH2 and AHA3; Palmgren & Christensen 1994). In addition, the antibody has been positively tested with H+-ATPase from roots of squash (W. Michalke, unpublished) and tomato (Schikora & Schmidt 2002). Immunolabelling was concentrated at the plasma membrane of the cells (Fig. 4A & B). Only very low non-specific labelling was observed in controls in which the first antibody was omitted (Fig. 4C). As expected, considerable differences in H+-ATPase density were obvious between cross-sections of iron-deficient roots through zones that are actively extruding protons and non-active segments from iron-sufficient roots (P = 0·01). Statistically significant differences were also apparent when acidifying sections from –Fe plants were compared with those showing no net proton efflux (Table 1). In acidifying segments H+-ATPase density was increased by 35% when compared with non-active zones. No significant variation in H+-ATPase density was observed between cross-sections from non-acidifying areas of –Fe roots and those of +Fe roots (Table 1).
Table 1. Effects of iron status and hormones on the distribution of H+-ATPase in epidermal cells of tomato roots. The number of gold particles is reported per 10 µm of plasma membrane (mean ± SE, n = 20)
Outer tangential walls
Other cell walls
Average of all walls
Outer tangential wall
Other cell walls
Average of all walls
+Fe, non-acidifying segments
6·2 ± 0·3
4·3 ± 0·3
5·0 ± 0·3
No transfer cells
No transfer cells
No transfer cells
+Fe, acidifying segments
9·4 ± 0·6
5·7 ± 0·5
7·0 ± 0·6
23·0 ± 2·0
10·0 ± 0·7
15·8 ± 1·3
–Fe, acidifying segments
11·1 ± 0·8
6·4 ± 0·5
8·1 ± 0·7
23·1 ± 3·4
9·7 ± 1·2
15·7 ± 2·3
–Fe, non-acidifying segments
5·7 ± 0·4
6·5 ± 0·6
6·2 ± 0·5
16·6 ± 2·2
15·3 ± 1·8
15·9 ± 2·0
6·7 ± 1·3
5·3 ± 1·2
5·8 ± 1·2
8·8 ± 1·6
9·0 ± 1·0
9·0 ± 1·3
6·2 ± 0·6
5·0 ± 0·6
5·5 ± 0·6
7·9 ± 0·7
7·3 ± 0·7
7·6 ± 0·7
Split-roots +Fe half
7·8 ± 0·6
6·3 ± 0·5
6·9 ± 0·5
11·1 ± 0·7
6·7 ± 0·7
8·7 ± 0·7
Split-roots –Fe half
7·4 ± 0·5
3·5 ± 0·3
4·9 ± 0·4
8·6 ± 0·9
5·9 ± 0·4
7·1 ± 0·7
Transfer cells exhibit higher H+-ATPase density
Transfer cells were formed in both acidifying and non-acidifying zones of Fe-deficient roots. However, in active spots their frequency was significantly higher than in non-acidifying areas (23·8 and 3·2%, respectively). In addition, few acidifying spots bearing transfer cells were observed in Fe-sufficient roots; no wall ingrowth formation occurred in non-acidifying segments of this growth type. To further investigate the role of transfer cells in the rhizodermis, their plasma membrane-associated gold particles were examined. The immunogold label clearly followed the contours of the plasma membrane and was not associated with the cell wall (Fig. 4B). The density of immunologically detectable H+-ATPase in cells with wall ingrowths was about two-fold higher than in ordinary rhizodermal cells when calculated on a unit surface basis (Table 1). No divergence in immunolabelling was observed between transfer cells in acidifying and non-acidifying segments. In cells with wall ingrowths, the distribution of label was clearly more polarized to the outer periclinal walls of the plasma membrane lining. In acidifying areas about 70% of labelling was confined to walls facing the root–soil interface. In transfer cells and normal rhizodermal cells of non-active zones a uniform labelling of immunologically detectable H+-ATPase around the cell perimeter was observed (Table 1).
Auxin and ethylene induce cell wall ingrowths but do not affect ATPase expression
To elucidate whether the expression of H+-ATPase is affected by auxin and ethylene, the plants were grown in the presence of either ACC or 2,4-D for 2 d. This treatment has previously been shown to induce transfer cells in tomato roots (Schmidt et al. 2000a). Roots of hormone-treated plants exhibited an acidification pattern similar to that of Fe-sufficient plants (data not shown, see Fig. 2A). This behaviour was mirrored in the density of plasma membrane H+-ATPases in root epidermal cells, which was within the range of non-acidifying roots of untreated plants (Table 1). In hormone-treated plants transfer cells occurred with a frequency similar to that of roots from Fe-deficient plants (data not shown). No differences were apparent between plants treated with ACC or 2,4-D (Table 1). The level of antibody binding was clearly less intense than in transfer cells of Fe-deficient plants, indicating that expression of plasma membrane H+-ATPase and induction of transfer cell formation are regulated separately. In addition, transfer cells induced by hormone application exhibited no asymmetry in immunolabelling on the plasma membrane in the ingrowth area relative to other walls (Table 1; Fig. 4D).
Plasma membrane H+-ATPase in iron-deficient roots is systemically regulated
To prove whether the level of the H+-ATPase is regulated by the presence of iron in the root or whether expression of the enzyme is systemically controlled, tomato plants were subjected to a split-root treatment. During 6 d of growth, one-half of the root system received 40 µm Fe, while the other half root was fed with iron-free nutrient medium. Visualization of acidification capacity of the split-root plants revealed significant dissimilarities between two root halves (Fig. 2). Contrary to what we expected, the split roots cultivated in iron-containing nutrient medium exhibited a proton extrusion pattern characteristic of undivided iron-deficient plants (Fig. 2C), whereas –Fe root halves showed a staining similar to that observed in +Fe control roots. A similar behaviour was described for the regulation of the Fe(III) chelate reductase activity, the other major response of strategy I-species to iron deficiency (Schmidt, Boomgaarden & Ahrens 1996). The density of plasma membrane H+-ATPase in ordinary rhizodermal cells corresponded to the acidification pattern (Table 1). While the frequency of immunologically detectable H+-ATPase in the +Fe root half was typical of acidifying segments, the abundance of gold particles in the –Fe split roots matched that observed in Fe-sufficient epidermal cells. A significantly higher (P = 0·05) labelling of +Fe split roots was only observed when the membranes lining the inner tangential and anticlinal walls were analysed. Transfer cells were induced in both root halves; the frequency was, however, more than four-fold higher in the –Fe root half (Schikora & Schmidt 2001), indicating that a high frequency of transfer cells is not imperative for rhizosphere acidification. The distribution of immunologically detectable H+-ATPase in transfer cells from the split-root plants followed a pattern that was different from ordinary rhizodermal cells. Only a non-significant trend towards higher density was noted in transfer cells of the +Fe root half, the absolute value of which was in the range of the non-active transfer cells induced by hormones rather than showing the high level of immunolabel characteristic of the plasma membrane of transfer cells in untreated roots (Table 1). A survey of the changes in response to the various treatments is given in Table 2.
Table 2. Summary of the changes in transfer cell number, density of immunologically detectable H+-ATPase, and acidification activity in response to the various treatments
The intensity of the responses is indicated by + and – symbols using the following order: – < + < ++. Values in parentheses indicate a density of transfer cells < 6%. Transfer cell number is indicated by ++ when more than 20% of the rhizodermal cells were differentiated into transfer cells.
Split-roots +Fe half
Split-roots –Fe half
Transfer cells are generally regarded as a means to amplify the surface area in zones of high solute flux. In the root epidermis, cells with wall protuberances were suggested to denote the site of the physiological responses to Fe-deficiency stress, providing the structural basis for the transmembrane flux of electrons and protons (Landsberg 1996). In the present study, we have demonstrated that the number of immunologically detectable H+-ATPase in rhizodermal transfer cells of tomato roots was considerably increased relative to ordinary epidermal cells. The frequency of gold particles on a given length of plasma membrane of wall ingrowth depositions in tomato roots is comparable with that reported for transfer cells in other tissues (Bouché-Pillon et al. 1994a, 1994b). Although the number of transfer cells appeared to be controlled by the local iron concentration, that is, repressed by iron in the nutrient solution, proton extrusion capacity was clearly increased in the +Fe half of the split roots, negating an absolute requirement of transfer cells for an effective acidification. This assumption is supported by several lines of evidence. In the Fe-hyperaccumulating pea mutant dgl, enhanced proton extrusion is constitutively expressed, whereas the formation of transfer cells is repressed by adequate iron levels in the nutrient medium, arguing against a requirement of wall ingrowths for lowering rhizosphere pH (Grusak & Pezeshgi 1996; Schikora & Schmidt 2001). Moreover, an enhanced efflux of protons in Fe-deficient roots has been observed in roots that are not capable of developing plasmalemma infoldings in the rhizodermis, such as Arabidopsis thaliana (Yi & Guerinot 1996; Schmidt, Tittel & Schikora 2000b). Vice versa, some species that do form transfer cells in response to Fe deficiency are not capable of acidifying the medium (Schmidt & Bartels 1996). Further evidence against a function of transfer cells in rhizosphere acidification comes from the split-root experiments. Finally, no differences in labelling density were observed between transfer cells in acidifying and non-acidifying areas of Fe-deficient roots. On the other hand, some data seem to prove that transfer cells are actively involved in rhizosphere acidification. First, the frequency of transfer cells in Fe-deficient roots was considerably higher in acidifying areas relative to non-active zones. Thus, the high proton extrusion activity of transfer cells might be masked by cells that do not contribute to the acidification of the medium. Secondly, acidifying zones in Fe-sufficient roots revealed a high number of transfer cells (14·8%), whereas virtually no transfer cells were found in non-active areas (data not shown). Considering the data at hand, it is reasonable to assume that transfer cells are equipped with a high number of H+-ATPase sites contributing to rhizosphere acidification without being essential for the reaction.
A possible role that is consistent with the occurrence and structure of transfer cells might be seen in the transport of Fe across the plasma membrane. Iron uptake in plants is mediated by a protein encoded by a member of the ZIP gene family, designated IRT1 (Eide et al. 1996; Korshunova et al. 1999; Guerinot 2000; Eckhardt et al. 2001). In pea roots, RIT1, a homologue of IRT1, is repressed by external iron in both the wild type and in the brz mutant which constitutively displays up-regulated reductase and H+-ATPase activity (Grusak & Pezeshgi 1996; Waters, Blevins & Eide 2000). Hence, RIT1 appears to be co-regulated with transfer cell formation (Schikora & Schmidt 2001). It is thus reasonable to assume a role of transfer cells in the mediation of iron uptake. However, the multitude of iron-regulated transporters in plant roots (Fox & Guerinot 1998; Curie et al. 2000, 2001) offers the possibility that different transporters are expressed in different cell types and that RIT1 is not or only partially responsible for Fe uptake in brz roots.
The split-root experiments demonstrated that up-regulation of acidification, probably brought about by increased H+-ATPase activity, was evident in the Fe-sufficient root half when the other portion of the roots was grown in Fe-free medium. A similar pattern has been observed with respect to ferric reduction activity (Schmidt et al. 1996; Schikora & Schmidt 2001). Although the basis for this behaviour cannot be clarified at the molecular level, the data are indicative of an inter-organ regulation of acidification and, thus, of H+-ATPase activity, possibly involving signals originating from the shoot. Phloem-mobile molecules involved in the regulation of iron homeostasis have not been identified until now, but there is some evidence suggesting that iron itself could be the signal that interacts with regulatory pathways (Schmidt et al. 1996). Iron-transporting proteins (ITPs), exhibiting high similarity to members of the LEA (late embryogenesis abundant) protein family, have recently been identified from the phloem of castor bean seedlings (Krüger, Hell & Stephan 2001). One of these proteins, designated ITP2, shows high Fe-binding capacity and represents a likely candidate for an iron carrier in the phloem. Shoot control of root responses involved in nutrient uptake is not unique to Fe nutrition and has recently been demonstrated for phosphate and sulphate uptake at the level of gene expression (Burleigh & Harrison 1999; Lappartient et al. 1999).
Information regarding the mechanisms that underlie induction of rhizodermal transfer cells is scarce. A Myb-like gene that can transactivate the promotors of transfer cell-specific genes has recently been isolated from maize endosperm (Gómez et al. 2002). In roots, application of the plant hormones auxin and ethylene mimics an iron deficiency-like phenotype, that is it induces the formation of extra root hairs and transfer cells that are structurally similar to those induced by iron deficiency (Schmidt & Bartels 1996; Schmidt & Schikora 2001). In the present investigation, the frequency of transfer cells induced by exogenously applied hormones was in the same range as that in Fe-deficient plants (data not shown), without affecting physiological reactions such as proton extrusion or ferric reduction (this study; Schmidt & Schikora 2001). The density of H+-ATPase in hormone-induced transfer cells was clearly lower than in those induced by Fe deficiency, suggesting uncoupling of wall ingrowth formation and H+-ATPase expression. Thus, the induction of plasmalemma infoldings and ATPase molecules appears to be regulated separately under some physiological conditions. The existence of several PM ATPase subfamilies offers the possibility that other isoforms, not detected by the antibody used in the present investigation, are more abundant after hormone treatment. The lack of proton extrusion activity of hormone-treated plants, however, argues against this assumption.
Whether all physiological responses to Fe deficiency stress are subject to a similar control is still under debate. Physiological uncoupling was inferred from temporal and spatial separation of rhizosphere acidification and increased Fe(III) reductase activity in Fe-deficient pea and Trifolium roots (Grusak, Welch & Kochian 1989; Grusak & Pezeshgi 1996; Cohen, Norvell & Kochian 1997; Wei, Loeppert & Ocumpaugh 1997). Both responses appear, however, to be systemically regulated. A plausible scenario implies that a phloem-translocatable compound, which mediates long-distance communication between shoot and root, can be differentially translated in root cells to induce either rhizosphere acidification or Fe(III) chelate reductase activity.
In conclusion, the experiments show that increased H+-ATPase density in epidermal cells of Fe-deficient roots spatially correlates with increased proton extrusion. Similar to the reduction of Fe(III) chelates, the level of H+-ATPase is systemically regulated, indicating inter-organ control of these processes involved in iron acquisition. Transfer cells induced by Fe-deficiency stress appeared to exhibit a higher steady-state level of H+-ATPase enzyme than ordinary epidermal cells. However, neither the immunolabelling nor the frequency of transfer cells resembled in vivo acidification patterns in all cases. These data suggest that transfer cells contribute to but are not required for rhizosphere acidification. Application of hormones induced the formation of wall ingrowths but not H+-ATPase activity, implying that both reactions are controlled by separate signalling pathways.
This work was supported by the Deutsche Forschungsgemeinschaft (DFG).
Received25 May 2002; received in revised form 26 August 2002; accepted for publication 26 August 2002