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Keywords:

  • induced defences;
  • glucosinolates;
  • jasmonic acid;
  • salicylic acid;
  • aboveground–belowground interactions;
  • Brassica spp

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  • • 
    Interactions between shoot and root induction of glucosinolates in two crucifers, Brassica oleracea and B. nigra, were studied by applying the signalling hormones jasmonic acid (JA) and salicylic acid (SA).
  • • 
    JA application increased total shoot glucosinolate levels 1.5–3 times, but total root levels did not increase. Only root JA-application yielded a systemic response. In B. oleracea it mattered where JA was applied: root application increased aliphatic glucosinolates in the shoot, whereas shoot application increased indole glucosinolates. Plants treated with JA to both organs had profiles similar to shoot-treated plants. SA-application did not disturb the organ-specific response to JA. Increases in glucosinolate levels did not reduce plant biomass.
  • • 
    A applications reduced root glucosinolates in root-treated plants. SA root-application in B. nigra resulted in lesions on the leaves and shoot-application caused a trichome response.
  • • 
    lants thus respond specifically, depending on the organ that is induced and the hormone that is applied. We find a large potential for root-feeders to affect shoot-feeders. Glucosinolate induction in one organ is not constrained by induction in the other organ.

Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Almost all plant species that have been studied respond to phytophage attack by changing their levels of chemical compounds. In many cases, these induced responses have been found to increase plant resistance to subsequent attacks by the same or other plant enemies (Karban & Baldwin, 1997). So far, studies on induced responses have mainly focused on the aerial parts of the plant and on how these responses affect above-ground herbivores and pathogens (van Dam et al., 2003). This focus on the shoot ignores not only the impact that below-ground phytophages may have on plant survival and plant biodiversity (van der Putten et al., 2001; de Deyn et al., 2003), but also that root damage may induce defensive responses as well (Birch et al., 1990; Ludwig-Müller et al., 1997). Many induced responses are systemic throughout the plant (Baldwin et al., 1994; van Dam et al., 2001; van Dam et al., 2003), so that shoot feeders may alter food quality for root feeders, and vice versa (Bezemer et al., 2003; Wäckers & Bezemer, 2003). As plants are attacked by both root and shoot phytophages, interactions between above-ground and below-ground induced defences are likely to occur.

There are several ways in which above-ground and below-ground induced defences can interact. First, simultaneous induction in roots and shoots may result in competition for limited resources, biosynthetic capacity, or defence compounds, especially when damage alters the relative sink-strength of plant organs (van Dam & Vrieling, 1994). Alternatively, above-ground and below-ground induced responses may interact when they are elicited by phytophages that trigger different signalling pathways in the plant. It has been well documented in above-ground studies that the salicylic acid (SA) signalling pathway, involved in induced responses against pathogens (Hammerschmidt & Smith-Becker, 1999), may suppress jasmonic acid (JA)-induced responses elicited by herbivore feeding (Creelman & Mullet, 1997; Felton et al., 1999; Preston et al., 1999; Thaler et al., 1999). In several plant species, both JA- and SA-induced responses have been found to be systemic throughout the plant (van Dam et al., 2001; Rostás et al., 2003). Therefore, JA–SA interactions may also occur between root and shoot induced responses.

In this study we use two naturally occurring Brassica species to evaluate interactions between shoot and root induced responses that are elicited by JA and SA application. Although Brassicaceae contain several classes of chemical compounds that may serve as defences, such as protease inhibitors (Cipollini & Bergelson, 2000; De Leo et al., 2001), saponins (Shinoda et al., 2002) and anthocyanins (Rostás et al., 2002), the most well-known defence compounds in this plant family are the glucosinolates (GS; Halkier & Du, 1997, Fahey et al., 2001). Based on the chemical structure of their side chain, the GS can be subdivided in different classes, such as aliphatic, aromatic and indole GS (Fahey et al., 2001). GS themselves may deter generalist herbivores (Li et al., 2000), but the hydrolysis products that are formed when cell rupture brings them into contact with myrosinase, an enzyme stored in specialised plant cells (Rask et al., 2000), are generally much more potent. Depending on the chemical structure of the GS side chain and the reaction conditions (e.g. pH), combination of the enzyme with the GS results in several different noxious and toxic products, such as isothiocyanates (ITC), oxazolidine-2-thiones and nitriles (Wittstock & Gershenzon, 2002).

We applied JA and SA solutions to roots and shoots of Brassica oleracea and B. nigra plants, which are two naturally occurring species from Western Europe. Although both species belong to the same genus, they are quite different in morphology, life-history and chemistry. B. oleracea is a spring perennial that has smooth, waxy leaves. The shoots of B. oleracea contain several different glucosinolates, among which the aliphatic gluconapin (3-butenyl GS) and the indole GS glucobrassicin are the most prominent (Mithen et al., 1987). B. nigra, on the other hand, is a summer annual, which has a rough appearance because its leaves are covered with trichomes (Traw & Dawson, 2002a). Next to the main GS sinigrin (2-propenyl GS) the plant species produces minor quantities of indole GS such as glucobrassicin (Traw & Dawson, 2002b,a). Moreover, when both plant species are grown under similar conditions in the glasshouse, the total GS levels in shoots of B. nigra are 3.5 times higher than those in shoots of B. oleracea (Harvey et al., 2003).

Based on previous studies in cultivated Brassica species, we expect that both JA and SA application will elicit a GS response and that these responses are systemic (Ludwig-Müller et al., 1997; Bartlet et al., 1999). Individual classes of GS, however, may respond differently to induction treatments (Mikkelsen et al., 2003). Indole GS, for example, consistently increased after application of JA or methylJA to shoots of the oilseed rapes Brassica napus and B. rapa, the mustard B. juncea or the Chinese cabbage B. campestris cv. pekinensis (Bodnaryk, 1994; Doughty et al., 1995; Ludwig-Müller et al., 1997; Bartlet et al., 1999). The levels of aromatic GS increased as well only in B. campestris, whereas in the other species the levels of aliphatic and aromatic GS levels remained unchanged after JA treatment (Doughty et al., 1995; Ludwig-Müller et al., 1997; Bartlet et al., 1999). By contrast, SA application to the roots increased all classes of GS in the shoots of B. napus, but aromatic GS levels increased more than indole or aliphatic GS levels (Kiddle et al., 1994). Because the structure of GS is closely related to their efficacy as defences against different phytophages (Brown & Morra, 1997; Potter et al., 1999), differential induction of GS indicates that plants are able to specifically tailor their response to the enemy that is feeding. Interactions between simultaneous induction events in roots and shoots may interfere with the optimisation of the response, and eventually affect the amount of damage to the plant.

By applying JA and SA solutions to roots and shoots in all possible combinations, we specifically addressed the question whether application of JA or SA to one organ elicits a systemic response in the other organ, and whether these responses interact. By analyzing the different classes of GS, we were able to study whether JA and SA applications trigger differential induction of specific classes of GS. In addition to levels of total and individual classes of GS, we also analyzed changes in biomass, which may indicate potential costs involved in the production of inducible compounds.

Materials and Methods

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Plant rearing

Seeds of B. nigra (L.) Koch and B. oleracea L. were mass collected in natural populations near Heteren, The Netherlands, in 2000 and stored in the dry and in the dark at 10°C. Seeds were germinated on glass beads in water in 10 × 10 cm plastic containers with a clear plastic lid. The containers were placed in a germination cabinet at 24°C and with a 16 h:8 h L:D photoperiod. Twelve days later, the seedlings were transferred to 1.3-l pots, containing 1500 g fine river sand. Before the seedlings were planted, three batches of sand were weighed, dried for 12 h at 110°C and weighed again to determine the initial water content of the sand. After transfering the seedlings, the pots were supplied with nutrient solution so that the total water content in the pots amounted to 15% of the sand dry mass. The sand in the pots was covered with aluminium foil to reduce evaporation. The pots with the seedlings were placed in a glasshouse that was kept at 21°C during the day and 16°C at night, under ambient light conditions that were supplied with sodium lamps to maintain the minimum PAR at 225 µmoles m−2 s−1 for at least 16 h d−1.

Every 2–3 d, five randomly chosen pots were weighed to determine the volume of solution needed to maintain the water content in the pots at 15%. Previous studies had shown that B. nigra and, to a lesser extent, B. oleracea suffer from phosphorus deficiency when grown on plain sand and 0.5 Hoagland solution. Therefore, we doubled (B. oleracea) or tripled (B. nigra) the concentration of KH2PO4 in the 0.5 Hoagland solution. Thus, over the 34 d this experiment lasted, the B. nigra plants each received 755 ml 0.5 Hoagland (1.1 mmol P) and B. oleracea plants received 790 ml 0.5 Hoagland (0.79 mmol P) per pot.

Treatment and harvest of the plants

Twenty-seven days after transplantation of the seedlings, 50 plants per species of equal size and stature were selected and randomly distributed over the 10 treatment groups. At that time, the B. nigra plants had an average of 11 leaves, whereas B. oleracea had on average eight leaves. Each plant received both a root and a shoot treatment in a full reciprocal design of JA/SA/water treatments (Table 1). Shoots were treated with 500 µl solution on two fully expanded leaves. The solution was gently rubbed onto the leaves with a latex-gloved finger. The roots were treated by injecting 25 ml of the proper solution in the sand next to the root crown, using a plastic syringe supplied with a pipette tip that was inserted in the soil. The original pH (4.7) of the JA solution that was applied to the roots was lowered with HCl to 3.0 to control for acid effects. The pH, 3.1, of the acidic solution (HCl in 0.1% Triton X-100) used for the ‘water’ treatments was chosen so that it was between the pH of the JA solution (3.5 and 3.0) and SA solution (2.7). The jasmonic acid (JA) treatments consisted of 2.4 µmoles (500 µg) of JA (±– jasmonic acid, Sigma, St Louis, IL, USA) to either roots or shoots, and the salicylic acid (SA; sodium salicylate, Baker, Deventer, The Netherlands) treatments were either 5.4 µmoles (725 µg) to the shoots or 180 µmoles (25 mg) to the roots. Both solutions were in 0.1% Triton X-100 to facilitate application to the leaf surface and absorption through the cuticle (Bodnaryk, 1994; Ludwig-Müller et al., 1997). These amounts were chosen because previous studies using other species of Brassicaceae had shown that they effectively induced glucosinolate levels in Brassica species at 7 d after treatment (Bodnaryk, 1994; Kiddle et al., 1994; Ludwig-Müller et al., 1997). Despite the low pH of the solutions, we did not observe any necrosis nor other direct phytotoxic effects on the leaves on which the hormones were applied.

Table 1.  Treatment combinations with jasmonic acid (JA) and salicylic acid (SA) solutions in 0.1% Triton in water
Shoots[DOWNWARDS ARROW]Roots[RIGHTWARDS ARROW]Water pH = 3.1500 µg JA pH = 3.025 mg SA pH = 2.7
  1. For all treatment groups n= 5.

Water Water/waterWater/JAWater/SA
pH = 3.1
500 µg JA JA/waterJA/JAJA/SA
pH = 3.5
725 µg SA SA/waterSA/JASA/SA
pH = 2.7

Additionally, we treated an extra group of plants with the same volumes of plain water. This treatment group was used to test whether dry masses and total GS levels were affected by the addition of an acidic solution with 0.1% Triton alone. In neither species did we find significant differences in dry masses or GS levels between the plain water and acid water groups (t-tests). Therefore we used the acid water group as the control group (water/water treatment in Table 1), so that we are sure to measure real effects of the JA and SA addition and not the effect of adding acidic water alone.

Seven days after treatment, the plants were harvested. The sand was removed from the roots by flushing with ample water and the roots were dried with paper tissue. Roots and shoots were separated, weighed to determine fresh masses and placed in separate paper bags to be flash frozen in liquid nitrogen. Plant parts were stored at −80°C until they were freeze-dried to constant weight. After the dry masses had been determined, the plant parts were ground to powder in a coffee mill and a 100-mg aliquot was weighed in a 15-ml tube for glucosinolate extraction.

Glucosinolates

GS were extracted with boiling 70% methanol solution, desulphatased with arylsulphatase (Sigma, St. Louis, IL, USA) on a DEAE-Sephadex A 25 column (EC, 1990) and separated on a reversed phase C-18 column on HPLC with an acetonitril-water gradient as described in Graser et al. (2000). Detection was performed with a single wavelength detector set to 226 nm. Sinigrin (sinigrin monohydrate, ACROS, New Jersey, USA) was used as an external standard. We used the correction factors for detection at 226 nm from Buchner (1987) to calculate the concentrations of the different types of GS in both plant species. Samples were analyzed by ESI-MS using a Hewlett-Packard (Avondale, PA, USA) HP 1100 HPLC coupled to a Micromass Quattro II (Waters, Micromass, Manchester, UK) tandem quadrupole mass spectrometer (geometry quadrupole-hexapole-quadrupole) equipped with an electrospray (ESI) source. The capillary and cone voltages in ESI mode were 3.3 kV and 18 V, respectively. Nitrogen was used for nebulization (15 l h−1) and as drying gas (250 l h−1 250°C). Source and capillary were heated at 80°C and 250°C, respectively. The mass spectrometer was operated in conventional scanning mode using the first quadrupole. Positive-ion full-scan mass spectra were recorded over the range from m/z 50–650 in a scan time of 1.5 s. Fixed precursor ion (MS/MS) spectra (a daughter ion scan) were recorded by setting the first quadrupole to transmit the parent ion of interest and scanning the product ions obtained after collision of parent ions in the hexapole gas cell using the second quadrupole analyzer. Argon was used for collision-induced dissociations (CID) at c. 1.5 × 10−3 mbar and the collision energy was varied from 16 to 25 eV for fragmentation. Separation of compounds was achieved on a reverse phase column (5 µm C18 phase, 100 × 2.1 mm i.d., Supelco) equipped with a precolumn (Supelco). Solvent system and gradient program were used as indicated above. The flow was maintained at 0.4 ml min-1 at a column temperature of 30°C. The UV detector was set at 226 nm. The compounds were identified based on their measured masses. The ions having masses which indicated presumptive desulphated glucosinolates were subjected to MS/MS analysis and daughter spectra were measured. Where the loss of m/z 162 was observed, the compound structure was declared to be a glucosinolate.

To facilitate evaluation of the data, the GS were grouped according to the chemical structure of their functional side chain (Fahey et al., 2001). The structure of the functional group is indicative for the biosynthetic origin of the GS as well as their effect on herbivores and pathogens (Halkier & Du, 1997; Wittstock & Halkier, 2002). In B. nigra we discriminated three groups of GS side chains: aliphatic (sinigrin), indole (glucobrassicin, 4-OH glucobrassicin, 4-methoxy glucobrassicin and neoglucobrassicin), and aromatic GS (gluconasturtiin). In B. oleracea we found five groups: aliphatic (glucobrassicanapin and gluconapin), 2-OH alcohols (progoitrin and gluconapolifeirin), sulphur containing (glucoraphanin, glucoalyssin and glucoerucin), indoles (glucobrassicin, 4-OH glucobrassicin, 4-methoxy glucobrassicin and neoglucobrassicin), and aromatic GS (gluconasturtiin). The total concentration (µmoles g−1 d. wt) of GS was calculated by summation of all GS that were detected in either plant species.

Statistical analysis

The effect of induction treatments on dry mass and total as well as specific GS levels in roots and shoots were analyzed simultaneously using manova, with treatment group as a fixed effect in the model. Using the planned contrast option we analyzed six protected orthogonal contrasts between: (1) control and all JA treatments (JA (to shoot)/water (to root), water/JA, JA/JA); (2) control and all SA treatments (SA/water, water/SA, SA/SA); (3) JA/water and JA/SA (interaction JA and SA); (4) water/JA and SA/JA (idem); (5) JA/water vs water/JA (local or systemic response to JA?); (6) JA/JA vs JA/water and water/JA (synergistic effects?). Normality of the data was checked by posthoc analysis of the residuals using the Kolmogorov-Smirnov test of normality. The alpha values used in individual anovas and protected contrast analyses generated after the overall manova analysis were corrected for multiple comparisons by the sequential Bonferroni correction, using the number of variables that were compared simultaneously in one analysis as the highest multiplication factor for P-values (Holm, 1979). All statistical analysis were performed with STATISTICA 6.0 Software (Statsoft inc., Tusla, OK, USA) using sigma restricted Type VI sum of squares.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Brassica oleracea

B. oleracea plants responded significantly to treatments with JA and SA (manova, F112, 173 = 1.90, P < 0.001), but this did not result in significant differences in root or shoot dry mass between treatments (Fig. 1 top panel; anovas, P > 0.13). Total shoot and root GS levels, however, significantly changed after treatments with hormone solution (Fig. 1 bottom panel; anova shoot GS, F8,36 = 5.04, P < 0.01, root GS F8,36 = 3.78, P < 0.05). Not all types of GS responded equally to the treatments (Table 2). In the shoots only indole GS (F8,36 = 17.09, P < 0.001) and aliphatic GS (F8,36 = 3.93, P < 0.05) levels changed due to hormone application, whereas in the roots the levels of indole GS (F8,36 = 4.21, P < 0.05), sulphur GS (F8,36 = 4.38, P < 0.01) and 2OH-alcohol GS (F8,36 = 4.59, P < 0.01) were affected.

image

Figure 1. Shoot (gray bars) and root (open bars) dry masses (top panel, g + SE) and total glucosinolate levels (bottom panel, µmoles g−1 dry mass + SE) of Brassica oleracea plants treated with different combinations of jasmonic acid (JA) and salicylic acid (SA). The treatment is indicated in the bar belonging to each organ. If no treatment is indicated, the organ was treated with acidic water (pH = 3.1) to control for pH effects of JA and SA applications.

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Table 2.  Levels of glucosinolates (µmoles g−1 dry mass, (SE)) in shoots and roots of Brassica oleracea treated with jasmonic acid (JA) and salicylic acid (SA) to roots and shoots. The glucosinolates are grouped according to the chemical structure of their functional side chain
Shoot/root treatmentSHOOT Aliphatic2OH-alcohSulphurIndoleAromaticROOT Alipahtic2OH-alcohSulphurIndoleAromatic
  1. Aliphatic; glucobrassicanapin and gluconapin: 2-OH alcohols; progoitrin and gluconapoleiferin: sulphur containing; glucoraphanin, glucoalyssin and glucoerucin: indole; glucobrassicin, 4-OH glucobrassicin, 4-methoxy glucobrassicin and neoglucobrassicin: aromatic; gluconasturtiin.

Water/water1.55 (0.46)1.92 (0.51)0.56 (0.08)0.45 (0.07)0.28 (0.05)0.46 (0.14)2.25 (0.38)2.81 (0.46)2.59 (0.35)6.36 (1.35)
JA/water1.83 (0.39)3.72 (0.68)0.82 (0.14)4.48 (0.58)0.21 (0.03)0.64 (0.12)4.69 (0.88)2.66 (0.34)3.76 (0.51)5.01 (0.79)
Water/JA4.13 (0.46)4.83 (0.96)0.89 (0.19)1.10 (0.14)0.34 (0.06)1.04 (0.29)4.11 (0.48)2.03 (0.23)2.70 (0.44)5.06 (0.79)
JA/JA1.41 (0.27)2.72 (0.75)0.83 (0.07)9.72 (2.0)0.20 (0.04)0.21 (0.05)2.54 (0.17)1.35 (0.14)3.59 (0.30)4.19 (0.58)
SA/water1.87 (0.53)2.10 (0.38)0.46 (0.06)0.43 (0.12)0.26 (0.07)0.64 (0.13)2.62 (0.34)2.03 (0.23)2.13 (0.37)5.20 (0.82)
Water/SA2.45 (0.56)2.56 (0.58)0.67 (0.15)0.64 (0.17)0.26 (0.06)0.60 (0.11)1.73 (0.19)1.35 (0.14)1.70 (0.20)3.53 (0.51)
SA/SA2.78 (0.34)2.60 (0.32)0.57 (0.03)0.57 (0.14)0.24 (0.03)0.49 (0.07)1.49 (0.21)0.73 (0.17)1.06 (0.17)1.91 (0.35)
JA/SA1.67 (0.53)2.61 (0.46)0.87 (0.31)3.66 (0.83)0.18 (0.04)0.50 (0.06)2.18 (0.12)1.97 (0.23)2.61 (0.77)3.84 (0.81)
SA/JA3.18 (0.45)3.69 (0.37)0.80 (0.16)1.21 (0.27)0.27 (0.04)0.63 (0.18)3.26 (0.79)1.94 (0.25)2.54 (0.50)4.12 (1.07)

Plants treated with JA to either roots, shoots, or both roots and shoots, overall had 2–3 times higher shoot GS levels than control plants treated with acid water (Fig. 1, bottom panel, bars labelled ‘JA treatment’; Protected contrasts (PC), t = 4.47, P < 0.001). Total root GS levels of these plants were unaffected by JA treatment. The strongest shoot response was shown by the indole GS levels, which increased 3- to 20-fold compared with the control plants (Table 2; PC t = 5.40, P < 0.001). The 2OH-alcohol GS levels also increased significantly in response to JA (PC, t= 2.65, P < 0.05).

Total shoot GS levels of both JA/water and water/JA treated B. oleracea plants equally increased compared with controls, despite possible differences in uptake efficiency between both organs, indicating that root treatment with JA induces a systemic increase in the shoot (Fig. 1, bottom panel). Interestingly, it mattered for the individual classes of GS to which organ JA was applied. Contrast analysis showed a significant difference in indole and aliphatic GS levels between these treatment groups (PC JA/water vs water/JA, t= 3.32, P < 0.05, and t = 3.74, P < 0.01, respectively). If the JA solution was applied to the shoot, the indole GS levels increased significantly in the shoots, whereas plants treated with the same amount of JA to their roots induced mainly aliphatic GS in their shoots (Table 2, lines 2 and 3). A similar pattern was found in shoots of the JA/SA and SA/JA treatment groups, indicating that this organ-specific response to JA was not disrupted by SA application. Simultaneous application of JA to roots and shoots (JA/JA treatment) resulted in significantly higher total shoot GS levels than in plants treated with JA to either roots or shoots (Fig. 1 top; PC, t = 2.16, P < 0.05). This increase was mainly caused by a strong increase in indole GS levels in the shoots (PC, t = 7.64, P < 0.001). Aliphatic and 2OH-alcohol GS levels of JA/JA plants, on the other hand, were lower, indicating that the induced GS pattern of JA/JA treated plants was most similar to that of plants treated with JA to their shoots only (Table 2, lines 2–4).

SA treatment significantly affected total GS root levels (PC, SA treated plants vs controls, t = 2.57, P < 0.05), whereas shoot levels remained similar to those of the control group (Fig. 1, bottom panel, bars labelled ‘SA treatment’). Root application of SA strongly reduced root GS levels compared with those of control plants, specifically those of aromatic GS (Table 2, PC t = 2.90, P < 0.01). SA application to the roots did not reduce the local shoot response to JA in JA/SA treated plants, nor did SA application to the shoots interfere with the systemically induced shoot response of plants treated with JA to their roots (PC JA/SA vs JA/water and PC SA/JA vs water/JA, P > 0.1). Similar to the plants treated with SA only, the roots of JA/SA plants had lower total GS levels than those of JA/water plants (PC t = 2.22, P < 0.05). JA application to the shoot thus did not relieve the effect of SA application to the roots.

Brassica nigra

JA and SA treatments overall induced a significant response in B. nigra plants (manovaF64,163 = 2.27 P < 0.001). Root dry mass was significantly different between treatment groups (anova root dry mass F8,34 = 3.79, P < 0.05), whereas shoot dry masses were not affected by either JA or SA application (Fig. 2, top panel). Both total shoot and total root GS levels responded significantly to the different treatments (Fig. 2, bottom panel; anova shoot GS F8,34 = 10.29, P < 0.001; root GS, F8,34 = 3.28, P < 0.05). The changes in total GS levels were to a large extent caused by changes in the major GS, sinigrin (Table 3, aliphatic GS column), in both shoots (anova, F8,34 = 10.47, P < 0.001) and roots (F8,34 = 3.00, P = 0.066 after Bonferroni correction).

image

Figure 2. Shoot (gray bars) and root (open bars) dry masses (top panel, g + SE) and total glucosinolate levels (bottom panel, µmoles g−1 dry mass + SE) of Brassica nigra plants treated with different combinations of jasmonic acid (JA) and salicylic acid (SA). The treatment is indicated in the bar belonging to each organ. If no treatment is indicated, the organ was treated with acidic water (pH = 3.1) to control for pH effects of JA and SA applications.

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Table 3.  Levels of glucosinolates (µmoles g−1 dry mass, (SE)) in shoots and roots of Brassica nigra treated with jasmonic acid (JA) and salicylic acid (SA) to roots and shoots. The glucosinolates are grouped according to the chemical structure of their functional side chain
Shoot/root treatmentSHOOT AliphaticIndoleAromaticROOT AlipahticIndoleAromatic
  1. Aliphatic; sinigrin: indole; glucobrassicin, 4-OH glucobrassicin, 4-methoxy glucobrassicin and neoglucobrassicin: aromatic; gluconasturtiin.

Water/water21.01 (1.88)0.09 (0.03)0.11 (0.02) 8.87 (1.45)0.83 (0.15)5.96 (0.44)
JA/water29.29 (1.04)0.08 (0.01)0.22 (0.05) 9.60 (0.58)1.09 (0.21)8.12 (0.83)
Water/JA30.19 (1.78)0.08 (0.02)0.15 (0.03) 7.99 (1.26)0.76 (0.12)8.37 (1.78)
JA/JA35.02 (3.68)0.07 (0.02)0.24 (0.05)11.12 (2.00)1.32 (0.28)8.86 (0.78)
SA/water22.76 (1.32)0.15 (0.03)0.15 (0.01) 9.79 (1.41)0.98 (0.11)7.67 (0.62)
Water/SA18.62 (0.87)0.16 (0.02)0.17 (0.03) 7.69 (1.53)1.40 (0.21)6.94 (0.82)
SA/SA17.28 (0.96)0.19 (0.04)0.20 (0.04) 4.74 (0.50)0.82 (0.04)7.62 (0.97)
JA/SA27.26 (2.05)0.22 (0.09)0.16 (0.03) 4.73 (0.46)0.86 (0.16)6.40 (1.09)
SA/JA31.61 (1.84)0.09 (0.02)0.27 (0.05) 9.68 (1.39)1.22 (0.20)9.75 (0.85)

Plants treated with JA only had significantly higher levels of total shoot GS levels than the control group (Fig. 2, bars labelled ‘JA treatment’; PC, t = 4.68, P < 0.001). This 1.5-fold increase in total GS was caused by increases in both levels of the aliphatic GS (PC t = 4.68, P < 0.001) and the aromatic GS (PC, t = 2.11, P < 0.05). Although total root GS levels were not significantly increased by JA application, aromatic GS levels were significantly higher in the roots of JA treated plants (PC t = 2.32, P < 0.05). Shoot or root application of JA resulted in similar patterns of GS induction, which indicates that the shoot response to JA application is fully systemic and independent of differences in uptake efficiency of JA by shoots or roots (Table 3; PC, all P > 0.2). Simultaneous application of JA to roots and shoots resulted in significantly higher shoot GS levels than application to either one of the organs alone (PC JA/water and water/JA vs JA/JA, t = 2.19 P < 0.05). Again this was mainly caused by a significant increase in aliphatic GS levels (PC, t = 2.18 P < 0.05). JA application did not significantly affect root or shoot dry mass (Fig. 2 top panel).

SA treatment alone did not significantly affect any of the GS levels, but reduced root dry mass significantly when compared with controls, especially if the SA was applied to the roots (Fig. 2 top panel, bars labelled ‘SA treatment’; PC, t = 3.78, P < 0.01). The same effect was observed in plants that were treated with SA to their roots and JA to their shoots. They also had significantly lower root dry masses than plants treated with JA to their shoots only (Fig. 2; PC JA/SA vs JA/water, t = 2.20, P < 0.05). Moreover, the JA/SA plants had lower total root GS than JA/water plants (Fig. 2 bottom: PC t = 2.62, P < 0.05) because of a decrease in aliphatic GS levels (Table 3; PC 2.53, P < 0.05). Nevertheless, simultaneous application of SA to plants treated with JA to the other organ did not reduce the total GS response in the shoot significantly compared with plants treated with JA only. Remarkably, JA/SA plants had significantly higher levels of indole GS in their shoots than JA/water plants (Table 3; PC t = 2.41, P < 0.05). A similar response of the indole GS, though not statistically significant, was found in plants treated with SA only, which suggest that SA treatment specifically induces indole GS production (Table 3, line 5 through 7).

By contrast to what may be concluded form the GS response, B. nigra plants did show an induced, systemic response to SA. One day after treatment, all 15 plants treated with SA to their roots, showed necrotic lesions at the leaf margins of the leaves that were present at the time of hormone application. Younger leaves that were formed later in the experiment, did not show any signs of necrosis. Similarly, the 15 plants treated with SA to their leaves, all responded by changes in trichome morphology, which was visible also within 1 d of SA application. Both treated and younger, untreated, leaves had enlarged trichome bases, which were surrounded by a dark, purple ring. Because the experiment was designed to study chemical responses and morphological responses were not anticipated at the beginning of the experiment, we had no extra plants available to quantify the changes in trichome morphology in more detail.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

This study shows that root induction events, as mimicked by the application of JA, have a significant systemic impact on total shoot GS levels in two wild Brassica species. This indicates that root phytophages, which trigger the JA-signalling pathway, may significantly alter food quality for their above-ground feeding counterparts by changing the levels of defence compounds in the shoot. This may not only affect the performance of shoot phytophages on induced plants, but also determine how much damage will be done to the shoot (Bezemer et al., 2003). JA application to the shoot mainly increased local GS levels. In B. nigra, however, the average level of aromatic GS (in this case gluconasturtiin) in roots of JA-treated plants significantly increased to more than 8 µmoles g−1 dry mass. Potter et al. (1999) found that this is a critical threshold level for gluconasturtiin in B. napus roots to significantly reduce susceptibility to the generalist nematode Pratylenchus penetrans. The increase in aromatic GS was the strongest in plants that were treated with JA to their roots, but was also clearly present in shoot-treated plants. Thus even if total GS levels in roots are not significantly affected, shoot feeders that induce the JA signalling pathway in B. nigra may alter host-plant suitability for specific groups of root feeders, such as nematodes. As in B. oleracea, B. nigra plants treated with JA to both roots and shoots had higher total GS levels in their shoots than plants treated with a single application of JA only. Although the effect was not completely additive, it indicates that in both plant species the total GS response could have been even stronger than observed in this experiment.

We found that shoot and root treatments with similar quantities of JA result in similar increases of total GS, but significantly different GS profiles in B. oleracea shoots. Such a shift in GS profiles may be of great ecological importance, as the structure of a GS is closely related to its potency as defence to phytophages. Moreover, because application of jasmonates also may also increase the levels of the hydrolytic enzyme myrosinase in the leaves (Taipalensuu et al., 1997), the JA-induced differences in GS levels probably translate directly into different profiles of the biologically active hydrolysis products upon damage. The aliphatic GS that were induced after JA application to the roots, yield isothiocyanates, which are generally recognized as potent antimicrobial and antifungal compounds as well as deterrents of a wide range of generalist and specialist herbivores (Brown & Morra, 1997; Fahey et al., 2001; Lambrix et al., 2001; Agrawal & Kurashige, 2003). By contrast, the indole GS, which were induced after shoot application, produce isothiocyanates that are unstable and decompose spontaneously (Fahey et al., 2001). Because of this instability, indole GS are far less effective deterrents or toxins, and their presence may even attract certain herbivores to the plant (Chew, 1988; Moyes et al., 2000). These effects may also extend to third and higher trophic levels, because the isothiocynates of aliphatic GS are partly volatile and thus may be used as host-finding cues by predators and parasitoids that are specialised on hosts that feed on GS-containing plants (Brown & Morra, 1997; Bradburne & Mithen, 2000). Plants that were treated with JA to both organs simultaneously showed an induced GS profile that was similar to plants treated with JA to their shoots. This indicates that the shoot-induced response has priority over the root-specific response. The specific GS response of B. oleracea to root and shoot JA application was not altered by SA application to the other organ, as has been observed in above-ground induced responses in several other plant species (Karban & Baldwin, 1997). This does not completely preclude interaction between JA and SA signalling pathways: it was shown that exogenous application of SA reduces the expression of myrosinase transcripts (Taipalensuu et al., 1997). Myrosinase activity, however, was not measured in this study and moreover it is not known which level of myrosinase activity suffices to hydrolyze all GS at a wound site. Therefore, the outcome of these potential interactions can only be analysed at the phenotypic level, that is by measuring the volatile profile of damaged plants that were pretreated with JA and SA. In B. nigra plants, we did not find a significant difference in shoot GLS levels or patterns between the different JA treatments.

Unlike in cultivated Brassica species (Kiddle et al., 1994; Ludwig-Müller et al., 1997), SA applications to B. nigra and B. oleracea did not increase overall GS levels in shoots or roots. In both plant species, total root GS levels even decreased in plants treated with SA to the roots, especially in B. oleracea. This is consistent with the GS levels found in SA-over-expressing A. thaliana mutants, which also had lower GS levels than wild-type plants (Mikkelsen et al., 2003). A decrease in root GS synthesis would not necessarily decrease shoot levels as well, because above-ground and below-ground GS levels and profiles are regulated independently (Sang et al., 1984; Potter et al., 1999).

Additionally, we observed a combined morphological-chemical response to SA leaf application in B. nigra plants. The dark purple ring around the trichomes that were formed after treatment most likely was a result of accumulation of anthocyanins in cells surrounding the trichome bases. It is known from studies on other, cultivated, Brassica species, that herbivory as well as fungal infection of the leaves may increase shoot anthocyanin levels (Rostás et al., 2002). Trichome responses in B. nigra also occur after leaf damage by lepidopteran larvae, which increases trichome densities on younger leaves (Traw & Dawson, 2002a). This indicates that trichome responses are an intrinsic part of the induced response in this plant species.

Root application of SA to B. nigra, on the other hand, induced the formation of lesions on the leaves, which is known as the hypersensitive response, a common plant response to pathogen attack (Hammerschmidt & Nicholson, 1999). Because we did not observe any direct phytotoxic effects when SA was applied directly to the shoot, this response most likely is part of the systemic acquired resistance response, which is controlled by the SA signaling pathway (Pieterse et al., 2002). As the lesion formation is a systemic response, it shows that the lack of response in shoot GS levels after SA application is not caused by a lack of SA uptake or transport from roots to shoots.

Interestingly, in neither of the plant species was SA application found to depress the GS response to JA application, as has been reported in other plants species (Creelman & Mullet, 1997; Felton et al., 1999; Preston et al., 1999; Thaler et al., 1999). Similarly, simultaneous application of JA to roots and shoots did not decrease the total GS response in the shoot, probably because root and shoot biosynthesis are regulated independently. This indicates that the GS response in these plant species is not constrained when different phytophages attack the plant at the roots and at the shoots simultaneously. Brassicaceae, however, are known to produce many other inducible compounds that have defensive properties, and that are induced in response to different types of phytophages. Possibly, there are constraints on the simultaneous induction of different classes of compounds. The eventual effects of such interactions on naturally occurring root and shoot phytophages, as well as on the plant's reproductive output, remain to be assessed.

In our experiment we found no indications that increased GS levels following JA applications reduced plant biomass or growth within 1 wk after induction. This indicates that the enhanced production of these compounds per se is not costly to the plant in this short time period. However, we can not rule out that there may be other, ecological, costs to induction of GS and possibly other inducible compounds in these plants. These costs may only emerge under circumstances that are ecologically more realistic, for example when plants that are induced are in competition for nutrients and light (van Dam & Baldwin, 2001). Especially in B. nigra, which is a rapidly growing annual species found in dense stands of up to 150 individuals per square meter (J. Harvey, pers. obs.), such cost may be of crucial importance.

Our results have shown that there is a great potential for induced responses to mediate interactions between above-ground and below-ground phytophages feeding on wild Brassica species. This potential only increases if one realizes that next to GS and trichomes, there are several other defensive compounds found in these species that are inducible by herbivores and pathogens, for example protease inhibitors or volatile compounds that attract natural enemies (Geervliet et al., 1997; De Leo et al., 2001). We expect that in natural environments, where plants are constantly under attack by a wide range of root and shoot phytophages, such interactions may be of great ecological importance to plants, phytophages and their natural enemies alike.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

The authors thank H. Duyts and R. Wagenaar for practical assistance with the harvest of the plants. M. Reichelt kindly provided several reference compounds for glucosinolate analysis. We thank W. van der Putten and T.M. Bezemer for critically reading an earlier version of the manuscript and A. Biere for statistical advice. This research was funded by VIDI grant no. 864-02-001 of the Netherlands Organisation for Scientific Research (NWO) to NM van Dam. A. Svatoš was supported by the Max Planck Society. Publication 3234 NIOO-KNAW Netherlands Institute of Ecology.

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  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
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