AOBPreview originally published online on May 30, 2006
Annals of Botany 2006 98(2):309-315; doi:10.1093/aob/mcl109
Effects of Phosphorus and Nitrogen on Nodulation are Seen Already at the Stage of Early Cortical Cell Divisions in Alnus incana
1 Department of Agricultural Research for Northern Sweden, Crop Science Section, Swedish University of Agricultural Sciences, Box 4097, S-904 03 Umeå, Sweden and 2 Research Programme on Biological Interactions, Department of Science and Technology, Universidad Nacional de Quilmes, R. Sáenz Peña 180, B1876 BXD Bernal, Argentina
* For correspondence. Present address: Department of Plant Physiology, Umeå Plant Science Centre (UPSC), Umeå University, S-901 87 Umeå, Sweden. E-mail Francesco.Gentili{at}plantphys.umu.se
Received: 28 November 2005 Returned for revision: 3 March 2006 Accepted: 4 April 2006 Published electronically: 30 May 2006
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ABSTRACT |
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• Background and Aims The present work aimed to study early stages of nodulation in a chronological sequence and to study phosphorus and nitrogen effects on early stages of nodulation in Alnus incana infected by Frankia. A method was developed to quantify early nodulation stages in intact root systems in the root hair-infected actinorhizal plant A. incana. Plant tissue responses were followed every 2 d until 14 d after inoculation. Cortical cell divisions were already seen 2 d after inoculation with Frankia. Cortical cell division areas, prenodules, nodule primordia and emerging nodules were quantified as host responses to infection.
• Methods Seedlings were grown in pouches and received different levels of phosphorus and nitrogen. Four levels of phosphorus (from 0·03 to 1 mM P) and two levels of nitrogen (0·71 and 6·45 mM N) were used to study P and N effects on these early stages of nodule development.
• Key Results P at a medium concentration (0·1 mM) stimulated cell divisions in the cortex and a number of prenodules, nodule primordia and emerging nodules as compared with higher or lower P levels. A high N level inhibited early cell divisions in the cortex, and this was particularly evident when the length of cell division areas and presence of the nodulation stages were related to root length.
• Conclusions Extended cortical cell division areas were found that have not been previously shown in A. incana. The results show that effects of P and N are already expressed at the stage when the first cortical cell divisions are induced by Frankia.
Key words: Alnus incana, cortical cell division area, Frankia, nitrogen, nodulation, phosphorus
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INTRODUCTION |
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Alnus is an actinorhizal plant, infected by the N2-fixing actinomycete Frankia via root hairs (Berry et al., 1986
; Berry and Sunell, 1990). In the host cells, Frankia is surrounded by an invaginated cell membrane. In Alnus, the sequence of events leading to root nodules begins with root hair deformation and penetration of root hairs by Frankia. Between the Frankia cell wall and the plant cell membrane there is an accumulation of host-derived wall material continuous with the plant cell wall at the site of Frankia penetration (Lalonde and Knowles, 1975; Van Dijk and Merkus, 1976; Huss-Danell, 1990; Berg, 1999). Cell divisions in the root cortex result in a so-called prenodule and, finally, cell divisions in the pericycle are the origin of a nodule primordium that will grow and develop into the true nodule (Callaham and Torrey, 1977; Lalonde, 1977; Torrey and Callaham, 1979; Berry et al., 1986; Newcomb and Wood, 1987; Berry and Sunell, 1990; Akkermans and Hirsch, 1997). Knowledge of this sequence of events is mainly based on detailed studies of transverse sections at various positions on the root. There is, however, a succession of developmental stages from the root tip to older parts of the root.
At the moment of inoculation, the susceptibility of root tissues to infection and nodule development is confined to a region of the root just behind the root tip, from the location of young emerging root hairs to the location of mature root hairs. As a consequence, a chronological succession of nodule development is seen from the proximity of the root tip (root hair deformation and cell divisions) towards the older parts of the root (nodule primordia and emerging nodules). The earliest nodules and the highest number of nodules develop in the middle part of the susceptible region as would be expected because of the phenomenon of autoregulation of nodulation described for legumes (Caetano Anollés and Gresshoff, 1991) and some actinorhizal plants (Valverde and Wall, 1999).
In N2-fixing plants, legumes and actinorhizal plants, phosphorus is often the most limiting nutrient for growth (Vance, 2001). P deficiency can impair nodulation and N2 fixation (Tang et al., 2001; Valverde et al., 2002). In plants relying on N2 fixation, P stimulated nodulation and N2 fixation more than it stimulated plant growth (Israel, 1987; Leidi and Rodriguez-Navarro, 2000; Hellsten and Huss-Danell, 2001; Gentili and Huss-Danell, 2002, 2003; Valverde et al., 2002). On the other hand, stimulation of N2 fixation by P has also been ascribed to a general growth effect in legumes (Robson et al., 1981; Jakobsen, 1985) and in an actinorhizal symbiosis (Reddell et al., 1997).
The fact that N inhibits nodulation in both actinorhizal plants and legumes is well known (e.g. Bond and Mackintosh, 1975; Streeter, 1988; Arnone et al., 1994; Huss-Danell, 1997; Hellsten and Huss-Danell, 2001; Gentili and Huss-Danell, 2002, 2003). To some extent stimulation of nodulation by P can counteract the inhibition by N (Wall et al., 2000; Gentili and Huss-Danell, 2002). However, little is known about the effects of P, N and their interaction on early stages of plant responses following inoculation by Frankia. The present work aimed to study the early stages of nodulation in a chronological sequence and to study P and N effects on early stages of nodulation in Alnus incana infected by Frankia. To accomplish this, we developed a light microscopy technique for intact A. incana root systems to analyse early cell and tissue responses at different times after inoculation.
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MATERIALS AND METHODS |
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Plant material and growth conditions
Seeds from a clone of A. incana (L.) Moench (Huss-Danell, 1991
) were shaken in water with a few drops of detergent, rinsed, and then incubated for 1 h in hydrogen peroxide (30 %, v/v) with occasional shaking, and finally rinsed in sterile water. The seeds were sown in Magenta jars (Sigma, St Louis, MO, USA) containing glass beads moistened with modified Evans solution (Huss-Danell, 1978) diluted to one-tenth of full strength and with 0·71 mM N added as ammonium nitrate. This solution is hereafter referred to as NP solution. Seedlings were transferred to growth pouches (Mega International, Minneapolis, MN, USA) 3 weeks after sowing. All material and all solutions used were sterilized before use. Each pouch contained four seedlings and each treatment comprised five growth pouches. The bottom of the pouches was cut open and the five pouches in each treatment were placed together in a dark plastic bag to protect roots from light. Nutrient solution was added to the plastic bags such that the paper towel in the growth pouch could adsorb solution by capillarity. The solution level was kept at 2 cm from the bottom of the pouches. Fresh solution was added twice a week to adjust the solution level, and the solution was renewed once a week.
Nutrient treatments
Treatments with the five different nutrient solutions (Table 1) began 1·5 weeks after seedlings were transferred to pouches. The paper towel in each pouch was rinsed with new solution and the pouches were then placed in new dark plastic bags. The rinsing minimized carry-over of old solution into new solution. The pH of all solutions was adjusted to 6·8 and all solutions were then passed through a 0·22 µm filter (Nalgene Co., Rochester, NY, USA). The electrical conductivity of the solutions in the bags showed no sign of salt accumulation. Plants were grown in a greenhouse in Umeå (63°45'N), Sweden, under a diurnal regime of 17 h photoperiods, with supplementary light provided by Philips HPI/T 400 W lamps at approx. 25 °C, and 7 h dark periods at approx. 15 °C. Relative humidity was approx. 40 %.
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Inoculation
Seedlings were inoculated with the effective Frankia strain ArI3 6·5 weeks after germination, i.e. 2 weeks after the start of treatments with different nutrient solutions. This time sequence was used to ensure that seedlings had adapted to the growth conditions in the pouches. The seedlings were uniform in size and approx. 2 cm high.
Frankia inoculum was grown on P+N medium (Hafeez et al., 1984) for 1 month. Each seedling received approx. 1 mg (wet weight) of bacteria suspended in NP solution. The suspension was pipetted onto the entire root system of each seedling.
Sample preparation and microscopy
Plants were harvested at 2, 4, 6, 8, 10 and 14 DAI (days after inoculation) by placing the entire pouch at –20 °C. Pouches were later thawed at room temperature and the root systems were carefully released from the paper towel by using a spatula. Intact root systems were bleached with sodium hypochlorite [approx. 5 % (v/v)] for 20 min and then rinsed in deionized water. The roots were placed in a gas-tight vial under vacuum for 30 min to get rid of any gas bubbles in the root tissues. Such cleared root systems could then be kept in Petri dishes with deionized water at 8 °C for up to 4 weeks. Pieces from a few of the frozen root systems were thawed and then fixed in 2·5 % (v/v) glutaraldehyde and 4 % paraformaldehyde in phosphate buffer pH 7·2 for 3·7 h at room temperature and rinsed in phosphate buffer. Samples were post-fixed in 2 % (w/v) OsO4 in phosphate buffer for 1 h at room temperature and then rinsed with phosphate buffer. After dehydration in ethanol (from 20 % up to 100 %) and acetone, samples were embedded in Epon resin and acetone (1 : 1 v/v) overnight and then in only Epon resin for 24 h at 60 °C. Semi-thin sections (1–2 µm) were cut and stained with toluidine blue.
Intact root systems and root sections were observed under an Olympus BX 60 light microscope. Photographs were taken with an Olympus camera mounted on the microscope using a Kodak epy 64 T Ektachrome film.
Definitions
The following definitions were used in the present work:
- RHD (root hair deformation), deformation of root hairs after inoculation with Frankia. Deformed root hairs appeared shorter than non-deformed root hairs.
- CCDA (cortical cell division area), area with cell divisions in the cortex, seen after inoculation with Frankia, but not in non-inoculated plants.
- Prenodule, a limited proliferation of the cortex underneath the infected root hair in response to the invasion of Frankia (Callaham and Torrey, 1977) resulting in a visible bump on the root surface (Berry and Sunell, 1990).
- Nodule primordium, produced by cell divisions in the pericycle, and as it grows it will be seen as a bump on the root surface. A nodule primordium develops close to a prenodule.
- Emerging nodule, a nodule that has erupted through the root epidermis.
- Lateral root primordium, produced by cell divisions in the pericycle but without any CCDA. A lateral root primordium has a narrower base than a nodule primordium.
Measurements
The length of CCDA was measured, with a calibrated scaled ocular, as the length of a continuous area where all cells in the outermost cortical cell layer had divided. Sometimes a CCDA included one or more prenodules, nodule primordia or emerging nodules. Root length was measured in a root scanner (Delta-T Devices Ltd, Cambridge, UK). The scans were calibrated against a dark blue sewing thread of known length. Prior to scanning, root systems were stained with methylene blue (0·1 % aqueous solution) for 30 s to facilitate the detection of thin roots.
Statistical analyses
One-way analysis of variance (ANOVA) and t-test were used to identify statistically significant differences among treatments at P < 0·05 significance level. Minitab software release 13·1 was used (Minitab Inc., State College, PA, USA).
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RESULTS |
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Time course of nodule development
Non-inoculated seedlings did not show any RHD or any cell divisions in the cortex (Fig. 1A). In inoculated seedlings, a developmental sequence was seen in cleared roots.
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At 2 DAI, a few anticlinal cell divisions were observed in the outermost cortical cell layer in the cleared roots. The CCDA was located in a region where root hairs were slightly deformed. With the technique we used to examine intact roots, the cell divisions were clearly seen under the microscope at this stage of nodule development but are difficult to visualize in photographs at this stage with only one focal plane (Fig. 1B).
At 4 DAI, there was extensive RHD (cf. Callaham et al., 1979). The CCDA comprised up to 12 dividing cells in the outermost cortical cell layer and up to five dividing cells in the next cell layer of the cortex (Fig. 1C).
At 6 DAI, dividing cells were seen in the three outermost cortical cell layers. The CCDA was longest in the outermost layers and gradually became shorter in the deeper cortical layers. On the other hand, cells in the outermost layer remained small while cells in the second and especially in the third cortical cell layers appeared to be enlarged in size (Fig. 1D). Altogether the area of dividing cells started to produce a bump in that region of the root, a prenodule (Fig. 1D), where strongly deformed root hairs were also seen.
At 8 DAI, nodule primordia emerging from the pericycle were observed, and the vascular tissue developing in the nodule primordia could also be seen (Fig. 1E).
At 10 DAI, nodule primordia had developed further and were extending over a long region in the cortex and producing a broad bump on the root surface (Fig. 1F).
By 14 DAI, seedlings had a shoot length of 3–4 cm. Nodules had emerged through the root surface. From this point on, all stages from deformed root hairs to emerged nodules were present in the same root.
All cell division areas and new tissues described above were easily distinguished from a lateral root primordium (Fig. 1G). The lateral root developed from the pericycle and grew through the root cortex where no cell divisions and only regularly sized cells were found. The shape of lateral root primordia was narrower and more pointed than that of nodule primordia (cf. Fig. 1G and F).
Longitudinal sections at 8 DAI verified the patterns observed in intact roots. Early cell division areas were distinguished as long rows of dividing cells. When several longitudinal sections were mounted together, it was possible to get a picture of a part of the root including different stages of tissue responses from single rows of dividing cortical cells to long and wide areas of cell division, where nodule primordia could be found in the centre of the cell division areas (data not shown).
N and P effects on nodulation stages
The different structures described above were recorded at 14 DAI in cleared roots of plants grown under different N and P regimes (Table 1).
However, prenodules and nodule primordia could be distinguished with certainty only when the structures were in parallel to the focal plan. Prenodules, nodule primordia and emerging nodules were therefore counted together in each plant. Increasing P from low (NlP) to medium (NP) concentration caused an increase, although not statistically significant, in the total length of CCDA per plant (Fig. 2). A further increase in P from medium (NP) to a moderately high level (NmhP), a high level (NhP) or a moderately high P level combined with a high N level (hNmhP) resulted in a decrease in cell division area per plant (Fig. 2). A more pronounced effect of P was observed on the number of prenodules, nodule primordia and emerging nodules. The number recorded per plant in NP treatment was significantly higher than in all other treatments (Fig. 3). However, when the number of prenodules, nodule primordia and emerging nodules per plant was expressed in relation to the length of the CCDA, there were no significant differences among P concentrations, but high N combined with moderately high P gave significantly higher numbers (Fig. 4).
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A low P concentration stimulated plants to produce longer roots as compared with plants receiving P at medium concentration (cf. NlP and NP in Fig. 5). Plants receiving moderately high or high P had intermediate root length. High N concentration stimulated root growth and resulted in the longest root systems (Fig. 5).
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As root length varied among treatments, it was appropriate to relate length of CCDA as well as number of prenodules, nodule primordia and emerging nodules to root length (Figs 6 and 7). In both cases, a medium level of P gave values almost twice as high as all other P levels. A high N level decreased the values approx. 3-fold.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSLITERATURE CITED |
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The analysis of A. incana root cell responses to Frankia inoculation revealed details of early steps in the actinorhizal interaction that have not been previously described. The novelty is our description of a CCDA comprising one, two or three cell layers (Fig. 1B–D). A similar phenomenon is observed in indeterminate legume nodule development (Diaz et al., 2000
) and in some cases in determinate nodulation in legumes (van Spronsen et al., 2001). The CCDA preceded prenodule formation in A. incana. It is reasonable to think that the CCDA has the ability to develop into nodules in the case of damage or excision of existing nodules, and that the new nodules will appear in the same area of the root as where the excised nodules were located. This is in agreement with autoregulation studies (Wall and Huss-Danell, 1997) where the excision of developed nodules in A. incana resulted in formation of new nodules in the same area of the root. In a previous work dealing with Casuarina glauca, it has been shown that the prenodule cells are infected by Frankia (Laplaze et al., 2000). Furthermore, prenodule cells showed the same characteristics as the nodule cells, indicating that the prenodule respresents a primitive symbiotic organ (Laplaze et al., 2000). In our work, the presence of Frankia inside cells has been clearly seen 10 DAI in primordia cell sections (data not shown) but not in the area of dividing cells which constitute the first stages of the prenodule structure. However, one may speculate that already at 6 DAI prenodule cells will be infected and that infection will increase with time. In the work dealing with C. glauca (Laplaze et al., 2000), prenodules and nodules were harvested 2–4 weeks after inoculation. In contrast, in the present work, A. incana showed formation of prenodules much earlier already at 6 DAI when no primordium was seen (Fig. 1C and D). Concerning Frankia infection of prenodules cells, it seems of great relevance to consider when the prenodule is sampled, at the beginning of its formation (only cortical cell divisions) or later, when the nodule primordium is present. The yellow-brown colour of the prenodule region observed around a developing primordium (Fig. 1E) suggests the presence of oxidized compounds that would be an indication of infected cells in that area.
The clearing of A. incana roots at 14 DAI allowed us to observe and quantify different stages of nodulation from RHD to emerging nodules in the same root, and to relate those stages to root length. This also allowed an analysis of P and N effects on early steps of nodulation. Even though N inhibition of nodulation has been observed in a range of actinorhizal plants (Arnone et al., 1994; Valverde, 2000; Wall et al., 2000; Gentili and Huss-Danell, 2002, 2003), an N effect has not been demonstrated on early stages of the nodulation process other than RHD (Prin and Rougier, 1987).
We observed that the number of prenodules, nodule primordia and emerging nodules per CCDA was independent of the level of P (Fig. 4). However, in the case of hNmhP, the number of prenodules, nodule primordia and emerging nodules per CCDA was increased (Fig. 4). This suggests a general positive interaction of N and P that is also seen in root growth (Fig. 5). Furthermore, we found that high N combined with moderately high P clearly inhibited early cell divisions (Fig. 2) but stimulated root growth (Fig. 5), and this inhibitory effect on CCDA was significant when related to root length (Fig. 6). The subsequent stages, recorded as the number of prenodules, nodule primordia and emerging nodules, were affected in the same way as CCDA when expressed per root length (Fig. 7). The inhibition already expressed at the CCDA level is still evident in the following steps of development. Taken together, our data suggest that the limiting step regulated by N would be at early cortical cell divisions, but not after that.
In this work, we found that P supplied at medium concentration, compared with lower or higher concentrations, clearly stimulated all the stages that are described here as steps in nodule formation, i.e. CCDA, prenodules, nodule primordia and emerging nodules (Figs 2, 3, 6 and 7). There is hardly any information on P effects on early stages in nodulation, except that Quispel (1958) noted stimulation when P was added during the first week after inoculation. The stimulatory effect of P at medium concentration (NP) was more pronounced when the nodulation stages were related to root length (cf. Fig. 2, and Figs 6, 3 and 7), indicating that the effect of P was specific for the nodule developmental process and not indirect via root growth. This confirms previous works where stimulation of nodulation by P was reported for A. incana (Gentili and Huss-Danell, 2003) and Hippophaë rhamnoides (Gentili and Huss-Danell, 2002) when analysed 4–10 weeks after inoculation.
The observed P and N effects already occurred at early stages in nodulation, before N2 fixation was substantial for plant growth, i.e. when Frankia is mainly a parasite for the plant. We cannot judge from this study how large a proportion of nodule primordia and emerging nodules will develop into mature nodules and how a large proportion will remain as arrested stages (Wall and Huss Danell, 1997; Valverde and Wall, 1999) in the roots, but we conclude from our results that in A. incana the regulation of the final number of nodules starts very early, already at the first cortical cell divisions.
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ACKNOWLEDGEMENTS |
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The Frankia strain ArI3 was a kind gift from Dr Bjørn Solheim. We appreciate the stimulating discussions with Dr Alison Berry and Dr Birgitta Bergman on symbiotic systems in general and actinorhizal symbioses in particular. We are grateful to Ann-Sofi Hahlin, Susanne Lindwall and Joaquín Wall for technical assistance. F.G. was supported by the EU (Grant no. FAIR-BM-972009) and the Kempe Foundation. L.G.W. is a research member of CONICET. K.H.D. received financial support from the Swedish Research Council, The Swedish Research Council for Environment, Agricultural Sciences and Spatial Planning, and The Swedish Foundation for International Cooperation in Research and Higher Education.
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