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Research Article
22 April 2015

Sharing a Host Plant (Wheat [Triticum aestivum]) Increases the Fitness of Fusarium graminearum and the Severity of Fusarium Head Blight but Reduces the Fitness of Grain Aphids (Sitobion avenae)


We hypothesized that interactions between fusarium head blight-causing pathogens and herbivores are likely to occur because they share wheat as a host plant. Our aim was to investigate the interactions between the grain aphid, Sitobion avenae, and Fusarium graminearum on wheat ears and the role that host volatile chemicals play in mediating interactions. Wheat ears were treated with aphids and F. graminearum inoculum, together or separately, and disease progress was monitored by visual assessment and by quantification of pathogen DNA and mycotoxins. Plants exposed to both aphids and F. graminearum inoculum showed accelerated disease progression, with a 2-fold increase in disease severity and 5-fold increase in mycotoxin accumulation over those of plants treated only with F. graminearum. Furthermore, the longer the period of aphid colonization of the host prior to inoculation with F. graminearum, the greater the amount of pathogen DNA that accumulated. Headspace samples of plant volatiles were collected for use in aphid olfactometer assays and were analyzed by gas chromatography-mass spectrometry (GC-MS) and GC-coupled electroantennography. Disease-induced plant volatiles were repellent to aphids, and 2-pentadecanone was the key semiochemical underpinning the repellent effect. We measured aphid survival and fecundity on infected wheat ears and found that both were markedly reduced on infected ears. Thus, interactions between F. graminearum and grain aphids on wheat ears benefit the pathogen at the expense of the pest. Our findings have important consequences for disease epidemiology, because we show increased spread and development of host disease, together with greater disease severity and greater accumulation of pathogen DNA and mycotoxin, when aphids are present.


Plant interactions with pathogens or insects are usually studied in isolation, even though plants are exposed to both in the field. It is likely that organisms that share the same host plant can alter its condition and thereby interact with each other. Although Fusarium pathogens and grain aphids cause substantial damage to wheat and coincide, little is known about interactions between these species.
Fusarium head blight (FHB) is a disease of small grain cereals that occurs globally and is caused by a complex of species from the genera Fusarium and Microdochium (1, 2). The most aggressive species causing FHB is Fusarium graminearum, a potent producer of the mycotoxin deoxynivalenol (DON), which is harmful to human and animal health if consumed (3). European legislation has set the acceptable limit of DON in grain marketed for human consumption at 1,250 μg kg−1 (4) in unprocessed cereals. Consequently, FHB epidemics result in large economic losses from the combined effects of reductions in yield, increased mycotoxin contamination, and reduced quality and marketability of the crop (5).
In the spring, ascospores and/or conidia of Fusarium graminearum are released from crop residues and are spread by wind or rainsplash. Wheat is most susceptible to FHB during anthesis, GS61 to GS69 (1, 6). During this time period, grain aphids are also active on wheat ears, and they may influence disease development. The English grain aphid, Sitobion avenae, is an ear-feeding species (7), and the behavior of apterous (wingless) individuals is influenced more strongly by volatile chemical emissions from wheat ears than from wheat seedlings (8). Volatile emissions from plants can be affected by abiotic or biotic stress, including that caused by pathogens, which can alter the attractiveness of the plant to visiting insects, in turn influencing the disease, where the insect may act as the vector of the pathogen (9). The interactions of phytopathogenic fungi and insect herbivores on shared host plants can have a variety of outcomes (10). It has been hypothesized that necrotrophs, such as F. graminearum (11), consistently reduce the fitness of insects feeding on the infected host, although there is evidence contradicting this claim (12, 13). It has also been shown that insect herbivory of hosts indirectly causes changes to plant chemistry, biochemistry, physiology, and growth (14) in such a way as to alter the host's capacity to withstand secondary infection. We hypothesized that F. graminearum and aphids colocalized on wheat ears could significantly influence each other's fitness by altering the condition of the shared host plant. To test this hypothesis, we designed a series of experiments investigating FHB disease progress and aphid performance on plants exposed to F. graminearum and aphids individually or in combination.
Interactions between FHB pathogens and aphid herbivores, occurring on shared plant hosts, are likely to be bidirectional. The pathogenesis of the plant and the resulting changes to induced plant host responses are likely to affect the behavior and success of insects, and insect feeding on the plant may alter the progression and severity of the disease induced by the pathogen. Pathogen-induced changes in plant secondary metabolites, toxin accumulation, and volatile chemical emissions have the potential to enhance or diminish the feeding and reproductive success of aphids on infected hosts. Aphids may facilitate weakening and injury of the host that could benefit opportunistic pathogens or prime plant defenses to be more resilient against pathogen attack (14). If there is any negative cross talk between the plant defense pathways activated by the respective organism, then prior colonization by one could facilitate the success of the other (15). Furthermore, these interactions are likely to be influenced by the changes in volatile emissions resulting from the infection of wheat ears by FHB pathogens, and the effects of these chemicals on aphid populations and behavior have not yet been explored. The impact of aphid-FHB interactions on the shared host plant needs to be better understood, since aphids and FHB pathogens occur together in real field situations, and their interactions may influence strategies for crop protection against both aphid damage and FHB disease.
The aim of this study was thus to elucidate the interactions between aphids and the pathogen F. graminearum on the ears of wheat, their shared plant host. The objectives were, first, to determine the role of aphids in FHB disease progression in terms of visually detectable symptom development as well as pathogen DNA and mycotoxin accumulation and, second, to identify the impact of pathogenesis induced by Fusarium graminearum on the volatile profile of the host plant and its effect on the survival and fecundity of grain aphids. In addition, following the first series of experiments, a third objective arose: to elucidate the importance of the timing of the interaction between aphids and F. graminearum on the host.


Plant material.

Winter wheat (Triticum aestivum cv. Gallant) was sown in compost (Levington F2+S) and was vernalized for 6 weeks at 6°C. Seed was treated with 10 g of prothioconazole and 50 g clothianidin 100 kg−1 (Redigo Deter; Bayer CropScience). Seedlings were then grown in a glasshouse in individual 5-liter pots in compost (John Innes type 2) at maximum and minimum daytime temperatures of 18°C and 15°C, respectively, and maximum and minimum nighttime temperatures of 12°C and 10°C, respectively, with a 16-h photoperiod using supplementary lighting.

Aphid rearing.

Grain aphids (Sitobion avenae) sourced from the Rothamsted Research insectary colony (Harpenden, United Kingdom) were reared on T. aestivum cv. Tybalt in a controlled-environment chamber under variable humidity at a 20°C daytime temperature with a 16-h day length.

Production of Fusarium graminearum inoculum.

Three single-spore isolates of Fusarium graminearum (isolates 212, 214, and 216; University of Nottingham) were cultured on potato dextrose agar, from which 10 plugs were transferred to carboxymethyl cellulose liquid medium (16) and were shaken at 90 rpm for 3 days at 23°C. Spore suspensions were filtered through sterile muslin, and concentrations were first calculated using a Neubauer improved hemocytometer (Marienfeld-Superior, Germany) and then adjusted to 2.5 × 105 spores ml−1 with sterile distilled water.

Production of F. graminearum-infected ears for the aphid transmission study and the study of aphid mortality and reproductive rates.

To generate an infected feedstock for aphids prior to their transfer to susceptible uninfected hosts, wheat (T. aestivum cv. Gallant) ears at GS65 (6) were spray inoculated with a spore suspension (2.5 × 105 spores ml−1) until runoff and were then bagged with polyethylene bags for 48 h. Following bag removal, plants were subjected to misting with an automated system (Access Irrigation, United Kingdom) for 1 min every hour, 10 h per day, for 7 days in a controlled-environment room at 25°C for 16 h (day) and at 18°C for 8 h (night), by which time symptoms could be observed visually. Control ears were maintained under the same glasshouse conditions.
To generate infected ears for headspace sampling, 10 single spikelets per ear (GS65) were point inoculated with 10 μl spore suspension (2.5 × 105 spores ml−1) between the lemma and the palea (layers of the spikelet) and were allowed to develop symptoms under the controlled-environment chamber conditions described above. Control plants were treated in the same way, except that sterile distilled water was used instead of the spore suspension.

Role of aphids in FHB disease caused by F. graminearum.

To test the ability of aphids to act as a vector of F. graminearum inoculum, five treatments, described in Table 1, were used. Aphids were applied to ears following transfer from their infected previous hosts and also after being artificially coated with F. graminearum spores. To test the effect of colocalization of aphids and F. graminearum on FHB disease progression in wheat ears, ears were treated simultaneously with aphids transferred from a pathogen-free previous host and with F. graminearum point inoculation. Three treatments were included as controls: negative-control ears were point inoculated with water only; positive-control ears were point inoculated with F. graminearum spores; and aphid-only control ears had no pathogen inoculum added, but aphids were added from a pathogen-free previous host. The experiment had randomized block design with six treatments and eight replicates.
TABLE 1 Treatment structure for single-ear experiment
Treatment nameAphids addedPrevious host of aphidsInoculum added
Negative controlNoneN/AaPoint inoculation with 10 μl of sterile distilled water
F. graminearum only (positive control)NoneN/APoint inoculation with 10 μl of 2.5 × 105 F. graminearum spores ml−1 pipetted between the lemma and palea
Aphid onlyTen adult wingless virginoparaeHealthy wheat earsNone
Aphid transmissionTen adult wingless virginoparaeF. graminearum-infected wheat earsOnly that occurring on aphids following feeding on the infected previous host
Spore-coated aphidsTen adult wingless virginoparaeRearing cage wheat leavesAphids shaken in spore suspension (2.5 × 105 F. graminearum spores ml−1) that had been concentrated by heating at 40°C for 2 h
F. graminearum + aphidsTen adult wingless virginoparaeRearing cage wheat leavesPoint inoculation with 10 μl of 2.5 × 105 F. graminearum spores ml−1 pipetted between the lemma and palea
N/A, not applicable.
Following treatment, ears were encased in netting for 48 h. After this time, netting and aphids (if present) were removed. Single ears of the main tillers of wheat plants at GS65 were used to assess F. graminearum disease progression with different treatments. Disease was assessed visually at five time points between 3 and 5 days apart, from 2 until 21 days after inoculation, by counting the number of spikelets with FHB symptoms (water-soaked lesions and/or bleached spikelets) (17). Disease severity was calculated as the percentage of total spikelets with symptoms per ear.

Effect of timing of aphid colonization on the progression of F. graminearum disease.

As a result of the experiment described above, we conducted further exploration into the interaction between aphids and F. graminearum when both were added to hosts. To elucidate the importance of the timing of the interaction between grain aphids and the F. graminearum inoculum on the shared host plant, aphids were added either prior to or following spray inoculation (1 × 105 spores ml−1) of all heads of single wheat plants until runoff, when the plants were bagged with polyethylene bags for 48 h. Plants were inoculated when a minimum of four ears had reached midanthesis (GS65). Eight whole-plant replicates were prepared per treatment; aphids were applied at different times in four treatments, and in one treatment, plants were kept free of aphids as a control. Aphids were added either 7 days or 3 days prior to the F. graminearum inoculum, on the same day as pathogen inoculation, or 4 days afterwards. Plants were contained within large wooden-framed netted cages, and aphids were added by placing a colonized leaf (containing a minimum of 30 adults) from the rearing cage adjacent to wheat ears; the leaf was removed after 48 h.
Plants were monitored for the aphid colonization rate every 7 days until they were considered to be thoroughly colonized (20 aphids per ear). Plants were monitored for visual signs of disease (17) every 3 to 4 days, commencing at 4 days after F. graminearum inoculation and terminating at 28 days after inoculation.

Extraction of DNA from F. graminearum and quantification by real-time PCR.

Individual wheat ears (n = 8) were harvested at 30 and 35 days after inoculation for the aphid transmission study and the aphid timing study, respectively. Whole ears were freeze-dried and were milled intact in a centrifugal mill (ZM 200; Retsch GmbH, Germany). Flour samples (0.5 g) were weighed into 15-ml tubes, and DNA was extracted as described previously (18) in cetyltrimethylammonium bromide (CTAB) buffer. Volumes of CTAB and potassium acetate were adjusted to 3.75 ml and 1.25 ml, respectively, to account for the small sample mass, and the duration of incubation of flour and CTAB at 65°C was 2 h. The DNA concentration was first calculated by examination of absorbances at the wavelengths of 260, 280, 328, and 360 nm by a Cary 50 Probe UV-visible spectrophotometer (Varian, CA, USA), then computed with Simple Reads software, and adjusted to 20 ng μl−1. The DNA was stored at −20°C.
A diagnostic PCR assay was carried out to confirm the presence of fungal DNA by using primers ITS4 and ITS5 (ITS4, 5′-TCCTCCGCTTATTGATATGC-3′; ITS5, 5′-GGAAGTAAAAGTCGTAACAAGG-3′) and a program described previously (19). The resulting samples were run on a 1% gel and were visualized under UV light (Gel Doc 2000 system; Bio-Rad, USA). Then two technical repeats of a real-time PCR assay were performed in a total volume of 13 μl comprising 2.5 μl template DNA, SYBR green 2× master mix (Bio-Rad, USA), and 50 M F. graminearum-specific primers (Fg16NF, 5′-ACAGATGACAAGATTCAGGCACA-3′; Fg16NR, 5′-TTCTTTGACATCTGTTCAACCCA-3′) (20) with a template product size of 280 bp. The cycling protocol, which has been described previously (21), comprised a 1.5-min initial denaturation step at 95°C, followed by 35 cycles of 30 s at 94°C, 45 s at 64°C, and 45 s at 72°C, with a final elongation for 5 min at 72°C. Target DNA from plant extracts was quantified using six DNA standards in 10-fold dilutions, originally extracted from a pure culture of F. graminearum isolate 212 (University of Nottingham). Linear regression was used to calculate the quantity of target DNA. Quantities of pathogen DNA are expressed as the amount of target DNA (in picograms) per total DNA (in nanograms).

Analysis of mycotoxin content.

The flour remaining after DNA extraction was used in mycotoxin extractions for deoxynivalenol (DON). No flour remained for any of the replicates of ears treated with spore-coated aphids; thus, mycotoxin quantification could not be performed for this treatment.
For calibration, flour samples (1 g) were spiked with a 13C-labeled DON internal standard (Biopure; Romer Labs, Runcorn, United Kingdom) and a 13C-labeled zearalenone internal standard (Biopure; Romer Labs) to produce a final concentration of 20 ppb each. Extraction was performed in acetonitrile (84%) for 1 h on a rotary shaker; then acetonitrile (100%) was added; the mixture was shaken by hand to mix and was filtered through filter paper (Whatman no. 1); and the filtrate was passed through a cleanup column (MycoSep 226; Romer Labs). The solvent of 5 ml filtrate was evaporated off to dryness with nitrogen gas; the filtrate was redissolved in 500 μl methanol (100%); the solvent was evaporated off once again; and then the filtrate was redissolved in 100 μl methanol (10%) for use in liquid chromatography (LC)-mass spectrometry (MS).
Aliquots of the samples (10 μl) were chromatographed using a Luna C18(2) high-performance LC (HPLC) column (length, 250 mm; inside diameter [i.d.], 3 mm; particle size, 5 μm; Phenomenex, Macclesfield, United Kingdom) fitted to an Agilent 1100 HPLC system (Stockport, United Kingdom). The compounds were eluted (flow rate, 0.5 ml · min−1) using a linear gradient from 10% methanol (Sigma-Aldrich) to 100% methanol over 15 min. This was first held at 100% for 7 min, then decreased to 0% over 0.2 min, and finally held at 0% for 4.8 min.
Mass spectrometry was performed on a Micromass Platform LCZ mass spectrometer (Micromass, Manchester, United Kingdom), controlled by MassLynx software (version 3.2). The mass spectrometer was equipped with an electrospray ionization (ESI) source operated in negative-ion mode. The source temperature was 75°C; the desolvation temperature, 450°C; the gas flow rate, 646 liters · h−1; the capillary voltage, 3.5 kV; the cone voltage, 21 V. DON and 13C-labeled DON were detected at m/z 331 and 346 in selected ion mode, with a dwell time of 0.1 s.
Calibration was achieved using a series of standards that contained 1,000 ppb of each standard and 20 ppb of each internal standard. The peak area for DON in samples was compared with that of standards after compensating for variation in the peak area of the internal standard. Mycotoxin extraction was unsuccessful for the aphid timing study, and insufficient flour remained to repeat the extraction process.

Assessment of aphid mortality and population growth.

Aphids that were transferred to aphid-only controls or to transmission treatment ears in the single-ear experiment were monitored to measure mortality and population growth on healthy versus diseased ears. Aphids were fed and contained in netting on control and symptomatic ears for 48 h (n = 8), after which time the netting was opened, and the ear and netting were inspected for living and dead aphids. The numbers of living and dead aphids were recorded prior to the transfer of surviving insects to new, pathogen-free hosts. Mortality was calculated as the number of dead aphids as a percentage of the starting population. After transfer to the new host ears, aphids were once again entrapped in netting, and after 48 h of aphid feeding, the numbers of living and dead aphids were recorded again. Aphid population growth following transfer to the new host was calculated as the total number of dead and live aphids found on the ears after a 48-h feeding period, expressed as a percentage of the starting population.

Volatile collection.

Fourteen days after inoculation, 4 symptomatic ears per plant were placed in partially sealed glass vessels and volatiles collected using headspace sampling methods described previously (22). Control ears were mock-inoculated with sterile distilled water instead of a spore suspension. Charcoal-filtered air was pumped in through the base at 600 ml min−1 and was drawn out through a PoraPak filter in the top of the jar at 400 ml min−1 for 48 h. Volatile samples were eluted from filters with 0.5 ml dichloromethane (DCM). Samples were collected simultaneously from four plants: two pathogen-free control plants and two plants point inoculated with F. graminearum.

Olfactometer bioassays.

Behavioral assays were carried out using a Perspex four-arm olfactometer (23) lit from above by diffuse, uniform lighting and maintained at 23°C. The bottom of the apparatus was lined with filter paper (Whatman no. 1), and air was drawn through the four arms toward the center at 350 ml min−1. Single winged S. avenae virginoparae (females reproducing by parthenogenesis) (n = 10) were introduced into the central chamber, and the time spent and the number of entries into each arm were recorded using specialist software (Olfa; Francesco Nazzi, Udine, Italy) over a 16-min period. The apparatus was rotated a quarter of a turn every 2 min to eliminate any directional bias. A headspace sample (10 μl) was applied to a filter paper strip, and the solvent was allowed to evaporate for 30 s. The filter paper was then placed at the end of the treated side arm. The three control arms were similarly treated with 10 μl of the solvent alone on filter paper.

Chemical analysis and identification.

Headspace samples were analyzed by gas chromatography (GC)-MS using 4 μl per sample. An HP-1 capillary GC column (length, 50 m; i.d., 0.32 mm) fitted with a cold on-column injector was directly coupled to a mass spectrometer (VG AutoSpec; Fisons Instruments, Manchester, United Kingdom). Ionization was carried out by electron impact at 70 eV and 250°C. The oven temperature was maintained at 30°C for 5 min and was then programmed at 5°C · min−1 to 250°C. Tentative GC-MS identifications were confirmed by peak enhancement with authentic samples on both the polar and nonpolar GC columns (24).


Electroantennogram (EAG) recordings were made from aphid antennae using Ag-AgCl glass microelectrodes filled with Ringer solution (7.55 g · liter−1 sodium chloride, 0.64 g · liter−1 potassium chloride, 0.22 g · liter−1 calcium chloride, 1.73 g · liter−1 magnesium chloride, 0.86 g · liter−1 sodium bicarbonate, 0.61 g · liter−1 sodium orthophosphate). The head of a winged S. avenae virginopara was separated from the body with a scalpel, and the tips of the antennae were removed to ensure good contact with the electrode. The head was placed in the tip of the indifferent electrode, and the tips of the antennae were positioned in the end of the recording electrode. The coupled GC-electrophysiology system, in which the effluent from the GC column is simultaneously directed to the antennal preparation and the GC detector, has been described previously (25). The volatiles were separated on an HP-1 column (length, 50 m; i.d., 0.32 mm) fitted into an HP6890 GC system equipped with a cold on-column injector and a flame ionization detector (FID). The oven temperature was first maintained at 40°C for 2 min and was then programmed at 5°C · min−1 to 100°C and then at 10°C · min−1 to 250°C. The carrier gas was hydrogen (flow rate, 42 cm · s−1). The outputs from the EAG amplifier and the FID were monitored simultaneously and were analyzed with a software package (Syntech, the Netherlands).

Preparation of synthetic components.

Synthetic versions of six electrophysiologically active chemicals were acquired (suppliers are listed in Table 2), and 1-mg · ml−1 solutions were prepared in hexane. Blends of synthetic chemicals were prepared using the same ratio and concentration as those in natural samples of Fusarium graminearum-induced volatile organic compounds (FgVOCs) from wheat. A two-component blend, comprising 2-pentadecanone and 2-heptanone in hexane, and a six-component blend, comprising all six chemicals listed in Table 2, also in hexane, were made.
TABLE 2 Electrophysiologically active FgVOC components used in olfactometer bioassays
Chemical nameAbundance in FgVOCs (ng μl−1)Supplier
Phenyl acetic acid0.027Aldrich
2-Pentadecanone128.8Alfa Aesar

Statistical analysis.

GenStat, version 15.1 (VSN International, United Kingdom), was used for all data analyses. Disease severity data were angularly transformed prior to analysis of disease progress over time by repeated-measurements analysis of variance (ANOVA). DNA and mycotoxin data were log10 transformed prior to ANOVA to normalize the residuals. Aphid mortality and population growth were analyzed using one-way two-sample t tests.
The mean times spent in treated and control arms and the numbers of entries into treated and control arms were compared using a paired t test. Data from olfactometer bioassays of replicate FgVOC samples were subjected to the Bonferroni correction for multiple t tests. The time spent in arms of the comparative olfactometer assays was analyzed by general ANOVA. Treatment means generated by individual and repeated-measurements ANOVA were considered significantly different at a P value of ≤0.05.


Role of aphids in FHB disease progression and accumulation of Fusarium graminearum DNA and DON in wheat ears.

The first differences in disease severity between treatments were observed 6 days after inoculation, when ears treated with both aphids and Fusarium graminearum inoculum showed significantly more symptoms than those subjected to other treatments (P < 0.001) (Fig. 1). For the duration of the experiment, ears thus treated developed the greatest disease severity, and by 28 days after inoculation, 83.1% mean disease severity was observed. Ears point inoculated with F. graminearum developed significantly more-severe disease at 18 days after inoculation than negative-control ears, spore-coated aphid-treated ears, or aphid transmission ears, with a final disease severity of 41.6% by 21 days after inoculation. Aphid transmission ears and ears treated with spore-coated aphids developed mean disease severities of 15.7% and 14.3%, respectively, not significantly different from that for negative controls.
FIG 1 Disease progress curve for aphid transmission experiment analyzed by repeated-measurements ANOVA, in which treatment, time, and treatment plotted against time were all significant (n = 8; P < 0.001; 5% LSD = 12.14; Greenhouse-Geisser ε = 0.3932). Fg, Fusarium graminearum. Treatments are described in Table 1.
Ears treated with aphids and F. graminearum inoculum had 2- and 5-fold greater quantities of pathogen DNA and DON, respectively, than pathogen-only controls (Fig. 2). A significantly greater quantity of pathogen DNA was found in ears treated with both the pathogen and aphids than in ears treated with the pathogen only, and both treatments yielded significantly greater pathogen DNA quantities than negative controls. DON accumulated more in ears treated with both the pathogen and aphids than in ears subjected to all other treatments. However, DON accumulation in pathogen-only controls was not significantly different from that in negative controls. All other treatments, except for spore-coated aphids, for which DON was not quantified, yielded pathogen DNA and DON quantities that were not significantly different from those for negative controls. The means for quantities of pathogen DNA and DON across treatments ranged from 0.001 to 48 pg · ng−1 and from 180 to 36,000 ppb, respectively. Both pathogen DNA and DON were detected in negative controls and aphid-only controls at low levels, presumed to be due to cross-contamination aided by the circulation of air within the controlled-environment chamber. Furthermore, regression analysis of log10-transformed DON concentration data on log10-transformed DNA concentration data showed that pathogen DNA and DON levels were significantly positively related (P > 0.001; R2 = 0.48); DON concentrations were increased in samples with increased pathogen DNA concentrations.
FIG 2 Mean log-transformed concentrations (± standard errors of the means) of F. graminearum DNA (expressed as picograms per nanogram of total DNA) (a) and DON (expressed in parts per billion) (b) in treated ears. Bars marked with the same lowercase letter are not significantly different at 5% LSD. The 5% LSD was 0.34 (a) or 0.91 (b). The effect of treatment was significant for both variables (n = 8; P < 0.001).

Effect of the timing of aphid colonization on FHB disease progression and F. graminearum DNA.

Although there were no significant differences in disease development as assessed visually between treatments with different timings of aphid colonization at 5% LSD (least significant difference), plants with a 7-day duration of aphid colonization (T − 7) manifested the highest disease severity (19.13%), followed by plants treated with aphids 3 days before inoculation (T − 3) (14.43%) (Fig. 3). Plants treated with aphids either simultaneously with pathogen inoculation (T0) or 4 days after pathogen inoculation (T + 4) and aphid-free plants manifested similar severities as assessed visually (13.51%, 13.22%, and 11.0%, respectively).
FIG 3 Disease progress curve for F. graminearum-inoculated ears with aphids applied at different times, analyzed by repeated-measurements ANOVA (n = 8; P = 0.53; 5% LSD = 6.20; Greenhouse-Geisser ε = 0.1845).
The timing of aphid colonization in relation to FHB infection significantly influenced the quantity of pathogen DNA in flour samples (P = 0.003). Treatment of plants with the inoculum in the absence of aphids produced significantly lower levels of pathogen DNA than all treatments where aphids were present (Fig. 4). The amount of pathogen DNA increased with the duration of aphid colonization; thus, the highest levels were found in plants treated with aphids 7 days prior to the addition of the pathogen inoculum, which contained 7-fold more than aphid-free controls. Significant differences in pathogen DNA quantity were also observed between aphid-free controls and plants treated with aphids 3 days prior to or simultaneously with pathogen inoculation; mean quantities for the treatments ranged from 2.9 to 22.0 pg · ng of total DNA−1.
FIG 4 Mean log-transformed concentrations of F. graminearum DNA (± standard errors of the means) in F. graminearum-inoculated plants treated with aphids at different times. The effect of treatment was significant (n = 8; P = 0.003; 5% LSD = 0.55). Bars marked with the same lowercase letter are not significantly different at 5% LSD.

Aphid mortality and reproductive rate.

After a 48-h feeding period, aphids fed on ears heavily infected with F. graminearum had a mean mortality rate of 59.4%, significantly greater than the mortality rate of 36.5% for aphids fed on control ears (P = 0.011) (Fig. 5a). Surviving aphids from F. graminearum-infected ears transferred to pathogen-free wheat ears exhibited a depressed reproductive rate (P = 0.042), producing, on average, 1.43 offspring per aphid, in contrast to 2.0 offspring produced per aphid when the previous host ear was not infected (Fig. 5b).
FIG 5 (a) Aphid population mortality (± standard errors of the means) after 48 h of feeding on mock-inoculated or F. graminearum-inoculated ears (P = 0.011). (b) Aphid population growth (as a proportion of the starting population) after surviving aphids from the experiment for which results are shown in panel a were transferred to pathogen-free wheat ears (P = 0.042). Asterisks indicate significant differences.

Olfactometer bioassays with natural samples.

Headspace samples from healthy wheat ears did not alter aphid behavior, and there were no significant differences in the amount of time spent in olfactometer arms treated with volatiles from healthy wheat and that in arms treated with pure solvent (data not shown). However, volatiles from FHB-exposed plants (FgVOCs) elicited an aversive response from aphids, which consistently spent significantly less time in areas treated with headspace samples from F. graminearum-infected ears than in areas treated with pure solvent (P = 0.03) (Fig. 6).
FIG 6 Olfactometer bioassay results showing mean time spent in olfactometer arms treated with VOCs from healthy wheat, pathogen-induced plant VOCs, individual chemical components of the natural sample (FgVOCs), or blends of chemicals according to natural abundances compared to time spent in control arms (± standard errors of the means). 2c, 2-component; 6c, 6-component. Asterisks indicate significant differences from controls (*, P < 0.05; **, P < 0.01; ***, P < 0.001).

Analysis of FgVOCs by GC-MS and GC-EAG.

Gas chromatograms of FgVOCs showed an increase in 2-pentadecanone levels over those in healthy-ear samples and the presence of several other compounds not found in healthy-ear samples. GC-coupled electroantennography (GC-EAG) of FgVOCs identified several compounds that were detectable by grain aphids, including 2-pentadecanone. Six of these were used in further olfactometer assays and are listed in Table 2.

Olfactometer assays with synthetic chemicals.

Of the six individual synthetic components of the FgVOCs that were used in olfactometer bioassays, 2-pentadecanone and 2-heptanone were strongly and weakly repellent, respectively (P, 0.013 and 0.091, respectively); (−)-α-gurjunene was weakly attractive (P = 0.058); and phenyl acetic acid, (−)-α-cedrene, and 2-tridecanone had no statistically significant influence on aphid behavior (Fig. 6).
A two-component blend of 2-pentadecanone and 2-heptanone was significantly repellent to aphids in olfactometer bioassays (P < 0.001) (Fig. 6), as was a six-component blend of all the individually tested chemicals (P = 0.004). In choice test olfactometer bioassays, designed to compare aphid responses to the natural FgVOC sample with those to synthetic blends, there was a preference for the 2-component blend over the natural sample (P < 0.001) (Fig. 7a). However, in bioassays using the six-component blend, there was no significant difference between the time spent in areas treated with FgVOCs and the time spent in areas treated with the blend, but both of these treatments yielded means significantly different from those for the control arms (P = 0.004) (Fig. 7b), indicating that both treatments were repellent.
FIG 7 Choice test olfactometer bioassay comparing a natural sample (FgVOC) to 2-component (a) and 6-component (b) blends of synthetic chemicals. (Bars marked with the same lowercase letter are not significantly different at 5% LSD.) The 5% LSD was 0.94 (a) or 1.35 (b). The treatment effect was significant in both cases, with P values of <0.001 (a) and 0.004 (b).


Wheat ears treated with a combination of aphids and Fusarium graminearum inoculum showed greatly accelerated disease progression, increased disease severity, and mycotoxin accumulation relative to those of plants treated only with the F. graminearum inoculum. However, aphid fitness was markedly reduced on diseased ears. Thus, in this pathosystem, colocalization of organisms has significant effects on fitness, and the effects are unequal, with the fitness of one organism rising substantially but that of the other declining sharply. It appears that the host colonization strategy of the fungal pathogen Fusarium graminearum is detrimental to the aphids but that host colonization by aphids may act to suppress plant defenses against the pathogen, so that the accumulation of pathogen DNA in aphid-colonized hosts increases in a dose-dependent manner. The sequence of arrival of attacking organisms on a plant may influence their performance, due to differences in the activation of induced defense pathways (15).
Sitobion avenae did not transmit the pathogen to new hosts; it only increased the development and within-ear spread of disease that was already present, perhaps because aphid-feeding damage makes it easier for the pathogen to colonize the plant. The interaction has large benefits for the pathogen overall, since aphid feeding increased the susceptibility of the host to the pathogen, so that the host reached advanced stages of disease earlier in its life cycle. Our data showed a larger area under the disease progress curve when aphids were present than when they were absent, indicating that the presence of aphids facilitated faster colonization by F. graminearum. This could provide more sources of inoculum early enough in the growing season to overlap the flowering period of neighboring plants or later-emerging ears and thus facilitate further spread of the pathogen. In this case, the influence of insects on pathogenesis is synergistic and positive, meaning that aphids are highly relevant for disease epidemiology. This will impact disease management protocols but, to our knowledge, has not been reported previously except as anecdotal field observations.
These results also have implications for the management of mycotoxin accumulation, since we observed a 5-fold increase in DON accumulation in wheat ears when aphids and F. graminearum occurred together over that in hosts treated with F. graminearum alone. When treatment with F. graminearum alone was compared with treatment with F. graminearum plus aphids, there was a greater difference in DON concentration than in visually detectable symptoms of disease. The presence of aphids resulted in the accumulation of increased levels of DON by infected plants. Crucially, pathogen inoculation with added aphid stress resulted in a greater mycotoxin output per unit of pathogen DNA than pathogen inoculation alone. DON is considered important for virulence during plant pathogenesis (26). Conversely, host resistance has been correlated with reduced mycotoxin contamination (27) via mechanisms including resistance to initial infection and pathogen movement throughout host tissues, as well as by the detoxification of mycotoxins within the ear (28, 29). Aphids appear to suppress plant defenses against F. graminearum, because when aphid and pathogen stresses occur together, or when aphids have established themselves prior to pathogen infection, the host plant may be less able to inhibit pathogen spread or to detoxify DON that is produced in ears.
In corroboration of these findings, previous studies of FHB in maize found that interaction with insects resulted in increased mycotoxin accumulation. When transgenic Bacillus thuringiensis (Bt) maize plants and nontransgenic controls were manually infested with European corn borer larvae (ECB), not only did the Bt maize have less ECB feeding than wild-type plants, but it also had fewer symptoms of fusarium ear rot and a ≤10-fold-lower concentration of fumonisin mycotoxins (30). Spray application of pyrethroid pesticides to reduce ECB infestation in Italian maize fields reduced fumonisin levels by 75% on average (31). It should be noted that in this case, ECB larvae both damage the host and act as a vector to deliver Fusarium conidia to corn ears, where infection is initiated (32). In contrast, in our study, the aphids are not involved as a vector, yet a rise in mycotoxin accumulation in insect-affected wheat was achieved through other mechanisms. In wheat field trials, the application of insecticide to reduce grain aphid populations at booting and/or heading was shown to reduce the incidence and severity of FHB by 20% and 31%, respectively (33). In terms of direct evidence that aphid involvement increases the success of pathogens on wheat, it has been shown previously that aphid feeding increases the frequency of lesions caused by Microdochium nivale on wheat leaves (34). Furthermore, wheat leaves fed on by Rhopalosiphum padi aphids for a 7-day period that was terminated 7 days prior to the infection of wheat ears with either Fusarium culmorum or F. graminearum showed no significant difference in disease severity but significantly increased ergosterol and DON contents in grain over those in aphid-free infected plants (35). This work shows that aphid feeding does not need to be at the same site as pathogen inoculation for increased host susceptibility to FHB disease.
It appears that the risk posed to wheat is greater when aphid colonies are present on ears prior to infection than when both stressors occur simultaneously, and aphid feeding shortly before or after inoculum arrival increases host susceptibility to the pathogen. This information is required to understand the impact of aphid involvement in FHB disease epidemiology and to inform those designing management strategies of the importance of pest management at different epidemiological stages. The mechanism by which aphid feeding increases host susceptibility to this fungal pathogen as yet remains unknown. Honeydew deposits on plant organs following aphid feeding may promote successful colonization by FHB-causing pathogens. It is also plausible that effectors within aphid saliva can act to downregulate the host defenses in such a way as to benefit the pathogen as well as the herbivore (36, 37). Since it has been shown that aphid feeding on wheat leaves can lead to increased mycotoxin content in grain upon infection with Fusarium spp. (35), it appears that molecular mechanisms above and beyond the possible influence of honeydew play a significant role in the increase in host susceptibility to FHB brought about by coincidental aphid stress. The elucidation of these mechanisms is necessary for full understanding of the fundamental cause of the effects observed.
Olfactometer bioassays showed that aphids were significantly repelled by FgVOC headspace samples. In contrast, olfactometer bioassays using volatile samples from healthy ears showed no effect on aphid behavior. Thus, pathogen-induced plant volatiles can affect the behavior of herbivorous insects so as to discourage infestation. This could have evolved as a means for aphids to avoid poor-quality hosts. In this case, the aggressive pathogen F. graminearum turns aphid-hospitable wheat ears into strongly repellent sites through the production of deterrent semiochemicals. 2-Pentadecanone was produced in F. graminearum-infected ear samples in greater amounts than in healthy ear samples and was repellent to aphids by itself. However, the pathogen-induced volatiles were more potently repellent as a mixture than as individual compounds. The effect of 2-pentadecanone on aphid behavior was not as strong as the effect observed with the full blend of electrophysiologically active FgVOCs. The six-component blend exhibited repellence comparable to that of natural samples, so it can be concluded that this is an adequate model for the main compounds responsible. 2-Pentadecanone is a novel compound that is repellent to aphids, and this effect should be explored for application within integrated pest management. The timing within the plant and pathogen life cycles at which healthy host volatile emissions begin to resemble FgVOCs remains unknown. Since the hosts were expressing visually detectable symptoms at the time of VOC collection, we do not yet have evidence that disease-induced volatiles can be used for early detection of F. graminearum infection. While the in-field consequences of aphid repellence as a result of FHB symptom induction have not been explored yet, it is plausible that relocation of repelled aphids to hosts with fewer FHB symptoms could increase aphid stress on these plants and perpetuate the promotion of increased FHB severity and mycotoxin accumulation across whole-field systems.
Fusarium pathogens have been shown previously to cause changes in the volatile chemical profiles of their hosts, for example, young wheat plants (38), stored wheat grain (39), and maize plants (40). Inoculation of maize leaves with a composite inoculum comprising Fusarium avenaceum, F. culmorum, F. graminearum, and Fusarium oxysporum induced volatile emissions that were not present in controls. The concentration of these chemicals increased over time, and particular components [(Z)-3-hexenal, (Z)-3-hexenyl acetate, β-caryophyllene, and linalool] were attractive to cereal leaf beetles (40). Another study sampled volatiles prior to ear emergence from wheat plants that were inoculated with a single-species inoculum of F. avenaceum, F. culmorum, or F. graminearum through the soil, damaged by cereal leaf beetle herbivory, or damaged mechanically (38). Of the chemicals that were not found in controls, (Z)-3-hexenyl acetate was the most abundant in F. graminearum-infected plants. The abundances of induced volatiles differed between Fusarium species used to inoculate the host. Moreover, volatiles induced by the fungi were distinct in both quantity and quality from those induced by either herbivory or mechanical damage.
The cause of the observed increase in aphid mortality and the decreased reproductive rate on diseased plants is unknown. Changes to the free amino acid composition of leaves has been suggested as a mechanism behind both the increased fitness of aphids on birch leaves infected with Marssonina betulae (41) and the decreased fitness of Aphis fabae aphids feeding on bean plants infected with the necrotroph Botrytis cinerea (14). As FHB symptoms develop on the host ears, green tissue becomes bleached due to fungal blockages in plant vasculature (41) from which aphids would otherwise feed; therefore, the removal of feeding sites could contribute to increased aphid mortality. However, the decreased rate of reproduction following aphid relocation to healthy hosts implies that contact with toxins produced in the infected host may have a lasting deleterious effect on the aphid. It is of interest for putative chemical pest control to identify the chemicals responsible for this effect, although they may be the same mycotoxins that are deleterious to human health.
The repellent effect observed in olfactometer bioassays implies that Sitobion avenae would be an incompatible vector for F. graminearum, and this is further demonstrated by the failure of aphids fed on symptomatic ears to produce disease in subsequent hosts. This work strongly suggests that Fusarium graminearum is not transmissible by grain aphids. Instead, as summarized in Fig. 8, colonization by aphids prior to FHB symptom induction improves the spread and development of F. graminearum. However, as the disease progresses, the host is transformed into a hostile and toxic environment for the aphid colonies. Aphid dispersal as a result of the creation of an inhospitable host environment can be viewed as a behavioral adaptation by the herbivore to escape a harmful environment. There may also be a further benefit to the pathogen if relocating aphids spread throughout a crop, enhancing susceptibility to the pathogen in less-affected hosts. Overall, interactions between F. graminearum and grain aphids on wheat ears benefit the pathogen at the expense of the herbivore and result in increased damage to the host plant from the disease.
FIG 8 Postanthesis (GS65) pest-pathogen interactions (between grain aphids and Fusarium graminearum) on wheat ears.


This work was supported by a Lawes Trust Ph.D. studentship. Rothamsted Research receives grant aid support from the Biotechnology and Biological Sciences Research Council (BBSRC).


Parry DW, Jenkinson P, McLeod L. 1995. Fusarium ear blight (scab) in small-grain cereals—a review. Plant Pathol 44:207–238.
Glynn NC, Hare MC, Parry DW, Edwards SG. 2005. Phylogenetic analysis of EF-1α gene sequences from isolates of Microdochium nivale leads to elevation of varieties majus and nivale to species status. Mycol Res 109:872–880.
Desjardins AE. 2006. Fusarium mycotoxins: chemistry, genetics, and biology. APS Press, St. Paul, MN.
. 20 December 2006. Commission Regulation (EC) No 1881/2006 of 19 December 2006 setting maximum levels of certain contaminants in foodstuffs. Off J Eur Union L 364/5:5–24.
Nganje WE, Bangsund DA, Leistritz FL, Wilson WW, Tiapo NM. 2004. Regional economic impacts of Fusarium head blight in wheat and barley. Appl Econ Perspect Pol 26:332–347.
Zadoks JC, Chang TT, Konzak CF. 1974. Decimal code for growth stages of cereals. Weed Res 14:415–421.
Wratten SD. 1975. Nature of effects of aphids Sitobion avenae and Metopolophium dirhodum on growth of wheat. Ann Appl Biol 79:27–34.
De Zutter N, Audenaert K, Haesaert G, Smagghe G. 2012. Preference of cereal aphids for different varieties of winter wheat. Arthropod Plant Interact 6:345–350.
Mayer CJ, Vilcinskas A, Gross J. 2008. Phytopathogen lures its insect vector by altering host plant odor. J Chem Ecol 34:1045–1049.
Stout MJ, Thaler JS, Thomma BPHJ. 2006. Plant-mediated interactions between pathogenic microorganisms and herbivorous arthropods. Annu Rev Entomol 51:663–689.
Goswami RS, Kistler HC. 2004. Heading for disaster: Fusarium graminearum on cereal crops. Mol Plant Pathol 5:515–525.
Al-Naemi F, Hatcher PE. 2013. Contrasting effects of necrotrophic and biotrophic plant pathogens on the aphid Aphis fabae. Entomol Exp Appl 148:234–245.
Zebitz CPW, Kehlenheck H. 1991. Performance of Aphis fabae on chocolate spot disease-infected faba bean plants. Phytoparasitica 19:113–119.
Ohgushi T. 2005. Indirect interaction webs: herbivore-induced effects through trait change in plants. Annu Rev Ecol Evol Syst 36:81–105.
Bruce TJA, Pickett JA. 2007. Plant defence signalling induced by biotic attacks. Curr Opin Plant Biol 10:387–392.
Tuite J. 1969. Plant pathological methods. Fungi and bacteria. Burgess Publishing Company, Minneapolis, MN.
Bai G. 1996. Variation in Fusarium graminearum and cultivar resistance to wheat scab. Plant Dis 80:975–979.
Edwards SG, Pirgozliev SR, Hare MC, Jenkinson P. 2001. Quantification of trichothecene-producing Fusarium species in harvested grain by competitive PCR to determine efficacies of fungicides against fusarium head blight of winter wheat. Appl Environ Microbiol 67:1575–1580.
White TJ, Bruns T, Lee S, Taylor J. 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics, p 315–322. In Innis MA, Gelfand DH, Sninsky JJ, White TJ (ed), PCR protocols: a guide to methods and applications. Academic Press, San Diego, CA.
Nicholson P, Simpson DR, Weston G, Rezanoor HN, Lees AK, Parry DW, Joyce D. 1998. Detection and quantification of Fusarium culmorum and Fusarium graminearum in cereals using PCR assays. Physiol Mol Plant Pathol 53:17–37.
Brandfass C, Karlovsky P. 2008. Upscaled CTAB-based DNA extraction and real-time PCR assays for Fusarium culmorum and F. graminearum DNA in plant material with reduced sampling error. Int J Mol Sci 9:2306–2321.
Birkett MA, Bruce TJA, Martin JL, Smart LE, Oakley J, Wadhams LJ. 2004. Responses of female orange wheat blossom midge, Sitodiplosis mosellana, to wheat panicle volatiles. J Chem Ecol 30:1319–1328.
Petersson J. 1970. An aphid sex attractant. Part 1. Biological Studies. Entomol Scand 1:63–73.
Pickett J. 1990. Gas chromatography-mass spectrometry in insect pheromone identification: three extreme case histories, p 299–309. In McCaffery AR, Wilson ID (ed), Chromatography and isolation of insect hormones and pheromones. Plenum Press, New York, NY.
Wadhams LJ. 1990. The use of coupled gas chromatography electrophysiological techniques in the identification of insect pheromones, p 289–298. In McCaffery AR, Wilson ID (ed), Chromatography and isolation of insect hormones and pheromones. Plenum Press, New York, NY.
Maier FJ, Miedaner T, Hadeler B, Felk A, Salomon S, Lemmens M, Kassner H, Schäfer W. 2006. Involvement of trichothecenes in fusarioses of wheat, barley and maize evaluated by gene disruption of the trichodiene synthase (Tri5) gene in three field isolates of different chemotype and virulence. Mol Plant Pathol 7:449–461.
Mesterhazy A. 2002. Role of deoxynivalenol in aggressiveness of Fusarium graminearum and F. culmorum and in resistance to Fusarium head blight. Eur J Plant Pathol 108:675–684.
Bai GH, Desjardins AE, Plattner RD. 2002. Deoxynivalenol-nonproducing Fusarium graminearum causes initial infection, but does not cause disease spread in wheat spikelets. Mycopathologia 153:91–98.
Boutigny AL, Richard-Forget F, Barreau C. 2008. Natural mechanisms for cereal resistance to the accumulation of Fusarium trichothecenes. Eur J Plant Pathol 121:411–423.
Munkvold GP, Hellmich RL, Rice LG. 1999. Comparison of fumonisin concentrations in kernels of transgenic Bt maize hybrids and nontransgenic hybrids. Plant Dis 83:130–138.
Saladini MA, Blandino M, Reyneri A, Alma A. 2008. Impact of insecticide treatments on Ostrinia nubilalis (Hübner) (Lepidoptera: Crambidae) and their influence on the mycotoxin contamination of maize kernels. Pest Manag Sci 64:1170–1178.
Sobek EA, Munkvold GP. 1999. European corn borer (Lepidoptera: Pyralidae) larvae as vectors of Fusarium moniliforme, causing kernel rot and symptomless infection of maize kernels. J Econ Entomol 92:503–509.
Bagga PS. 2008. Fusarium head blight (FHB) of wheat: role of host resistance, wheat aphids, insecticide and strobilurin fungicide in disease control in Punjab, India. Cereal Res Commun 36(Suppl 6):667–670.
Diehl T, Fehrmann H. 1989. Wheat fusarioses—influence of infection date, tissue injury and aphids on leaf and ear attack. Z Pflanzenkrankh Pflanzensch 96:393–407.
Liu Y, Buchenauer H. 2005. Interactions between Barley yellow dwarf virus and Fusarium spp. affecting development of Fusarium head blight of wheat. Eur J Plant Pathol 113:283–295.
Anathakrishnan R, Sinha DK, Murugan M, Zhu KY, Chen MS, Zhu YC, Smith CM. 2014. Comparative gut transcriptome analysis reveals differences between virulent and avirulent Russian wheat aphids, Diuraphis noxia. Arthropod Plant Interact 8:79–88.
Bos JI, Prince D, Pitino M, Maffei ME, Win J, Hogenhout SA. 2010. A functional genomics approach identifies candidate effectors from the aphid species Myzus persicae (green peach aphid). PLoS Genet 6:e1001216.
Piesik D, Lemnczyk G, Skoczek A, Lamparski R, Bocianowski J, Kotwica K, Delaney KJ. 2011. Fusarium infection in maize: volatile induction of infected and neighboring uninfected plants has the potential to attract a pest cereal leaf beetle, Oulema melanopus. J Plant Physiol 168:1534–1542.
Presicce DS, Forleo A, Taurino AM, Zuppa M, Siciliano P, Laddomada B, Logrieco A, Visconti A. 2006. Response evaluation of an E-nose towards contaminated wheat by Fusarium poae fungi. Sensors Actuat B Chem 118:433–438.
Piesik D, Wenda-Piesik A, Lamparski R, Tabaka P, Ligor T, Buszewski B. 2010. Effects of mechanical injury and insect feeding on volatiles emitted by wheat plants. Entomol Fenn 21:117–128.
Guenther JC, Trail F. 2005. The development and differentiation of Gibberella zeae (anamorph: Fusarium graminearum) during colonization of wheat. Mycologia 97:229–237.

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Published In

cover image Applied and Environmental Microbiology
Applied and Environmental Microbiology
Volume 81Number 1015 May 2015
Pages: 3492 - 3501
Editor: H. Goodrich-Blair
PubMed: 25769834


Received: 23 January 2015
Accepted: 8 March 2015
Published online: 22 April 2015


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Jassy Drakulic
University of Nottingham, School of Biosciences, Division of Plant and Crop Sciences, Sutton Bonington, United Kingdom
Rothamsted Research, Harpenden, Hertfordshire, United Kingdom
John Caulfield
Rothamsted Research, Harpenden, Hertfordshire, United Kingdom
Christine Woodcock
Rothamsted Research, Harpenden, Hertfordshire, United Kingdom
Stephen P. T. Jones
University of Nottingham, School of Biosciences, Division of Plant and Crop Sciences, Sutton Bonington, United Kingdom
Robert Linforth
University of Nottingham, School of Biosciences, Division of Plant and Crop Sciences, Sutton Bonington, United Kingdom
Toby J. A. Bruce
University of Nottingham, School of Biosciences, Division of Plant and Crop Sciences, Sutton Bonington, United Kingdom
Rothamsted Research, Harpenden, Hertfordshire, United Kingdom
Rumiana V. Ray
University of Nottingham, School of Biosciences, Division of Plant and Crop Sciences, Sutton Bonington, United Kingdom


H. Goodrich-Blair


Address correspondence to Rumiana V. Ray, [email protected].

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