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Plant Cell. Jan 2012; 24(1): 353–370.
Published online Jan 31, 2012. doi:  10.1105/tpc.111.093104
PMCID: PMC3289574

Loss of Abaxial Leaf Epicuticular Wax in Medicago truncatula irg1/palm1 Mutants Results in Reduced Spore Differentiation of Anthracnose and Nonhost Rust Pathogens[W]


To identify genes that confer nonhost resistance to biotrophic fungal pathogens, we did a forward-genetics screen using Medicago truncatula Tnt1 retrotransposon insertion lines. From this screen, we identified an inhibitor of rust germ tube differentation1 (irg1) mutant that failed to promote preinfection structure differentiation of two rust pathogens, Phakopsora pachyrhizi and Puccinia emaculata, and one anthracnose pathogen, Colletotrichum trifolii, on the abaxial leaf surface. Cytological and chemical analyses revealed that the inhibition of rust preinfection structures in irg1 mutants is due to complete loss of the abaxial epicuticular wax crystals and reduced surface hydrophobicity. The composition of waxes on abaxial leaf surface of irg1 mutants had >90% reduction of C30 primary alcohols and a preferential increase of C29 and C31 alkanes compared with the wild type. IRG1 encodes a Cys(2)His(2) zinc finger transcription factor, PALM1, which also controls dissected leaf morphology in M. truncatula. Transcriptome analysis of irg1/palm1 mutants revealed downregulation of eceriferum4, an enzyme implicated in primary alcohol biosynthesis, and MYB96, a major transcription factor that regulates wax biosynthesis. Our results demonstrate that PALM1 plays a role in regulating epicuticular wax metabolism and transport and that epicuticular wax influences spore differentiation of host and nonhost fungal pathogens.


Rusts are obligate biotrophic foliar pathogens that have evolved specialized mechanisms of invasion (Heath, 1977). To initiate rust infection, fungal urediniospores need to adhere to the leaf surface and subsequently form germ tubes. Germ tubes typically respond thigmotropically to host leaf surface features, such as stomata, forming appressoria over these openings (Hoch et al., 1987). Penetration pegs form at the appressoria-stomata interface and mature into invasive hyphae that invade mesophyll cells eventually differentiating into specialized feeding structures called haustoria (Heath, 1977; Hoch et al., 1987). Therefore, it appears that rust pathogens require specific plant surface topographical and chemical signals to trigger the formation of preinfection structures (Heath, 1977; Müller and Riederer, 2005). Traditionally, breeding for rust resistance in various crops mainly relied on host resistance mediated by gene-for-gene resistance (Ayliffe et al., 2008). However, in many cases, resistance mediated by R genes has little durability in the field due to the rapid evolution and emergence of new pathogen strains that can escape recognition by R genes and R gene–mediated downstream defenses. Alternatively, nonhost resistance (NHR) is defined as a form of resistance exhibited by an entire plant species to a particular microbial pathogen and is the most common and durable form of resistance (Heath, 2000). Therefore, identification and incorporation of traits that confer NHR to a broad range of rust fungi is an attractive and durable alternative to host resistance breeding. However, we know little about genes that regulate NHR (Mysore and Ryu, 2004).

Asian soybean rust caused by Phakopsora pachyrhizi Sydow is a major concern for soybean (Glycine max) producers in Brazil and the US (Goellner et al., 2010). Since most of the soybean cultivars and other economically important legumes are susceptible to soybean rust, there is an increasingly urgent demand for identification of durable resistance to soybean rust (van de Mortel et al., 2007). Four single resistance genes to P. pachyrhizi, Rpp1-4, have been described (Hyten et al., 2007; Garcia et al., 2008; Silva et al., 2008; Monteros et al., 2010). Interestingly, all commercial cultivars that are cultivated in the US are susceptible to rust, and no soybean varieties have been described as having broad-spectrum resistance to all isolates of P. pachyrhizi (Posada-Buitrago and Frederick, 2005). Similarly, rust disease of switchgrass (Panicum virgatum), caused by Puccinia emaculata, is a concern in Oklahoma and other parts of the US and could become an important factor once switchgrass is grown in monoculture over a long period of time (Bouton, 2007). Therefore, there is an urgent need for identification of novel genes that could be used for engineering broad-spectrum resistance to soybean rust and/or switchgrass rust isolates.

Identifying novel sources of resistance through large-scale forward or reverse genetics screens has the potential to improve crop plant resistance (Heath, 2000; Mysore and Ryu, 2004). Medicago truncatula is a rapidly emerging model plant species, especially for legumes. Several genomic tools and resources are already available for M. truncatula and include an extensive EST database, genome sequence, gene expression, protein and metabolite profiling tools, and a collection of insertion and fast neutron bombardment mutants (Young and Udvardi, 2009). For large-scale mutagenesis of the M. truncatula genome, a tobacco (Nicotiana tabacum) retrotransposon, Tnt1, has been introduced and been shown to efficiently transpose in the M. truncatula genome during tissue culture, producing insertions that are stable during seed-to-seed generations (d’Erfurth et al., 2003). We have now generated ~20,000 Tnt1 insertion lines in M. truncatula with an average of 25 insertions per line (Tadege et al., 2008). We set up a genetic screen to identify M. truncatula mutants with altered interactions with P. emaculata and/or P. pachyrhizi with an aim of identifying mutants exhibiting enhanced susceptibility to P. emaculata or P. pachyrhizi. Characterization of these susceptible mutants would lead to the identification of target genes for genetic improvement of rust resistance in switchgrass, soybeans, and other economically important crops, including wheat (Triticum aestivum) and barley (Hordeum vulgare).

The high-throughput forward genetics screen of M. truncatula Tnt1 insertion lines performed in this study unexpectedly identified an inhibitor of rust germ tube differentiation1 (irg1) mutant that failed to promote preinfection structure differentiation against nonhost rust fungi, P. emaculata and P. pachyrhizi, and also to the pathogenic anthracnose fungus Colletotrichum trifolii. Cytological, chemical, and transcriptome analyses revealed that irg1 is defective in abaxial epicuticular wax deposition and/or secretion. Flanking sequence tag sequencing of irg1 revealed that IRG1 encoded a Cys(2)His(2) zinc finger transcription factor (PALM1) that also controls dissected leaf morphology in M. truncatula (Chen et al., 2010). Our results unraveled a role for PALM1 in asymmetric epicuticular wax deposition on M. truncatula leaves, which influences fungal spore differentiation.


Tnt1 Insertional Mutant Screening in M. truncatula Identified irg1

Initial characterization of the M. truncatulaP. emaculata interaction showed that the urediniospores germinate and form germ tubes on the leaf surface but fail to recognize and form appressoria on stomata, thereby precluding successful colonization of the nonhost plant, M. truncatula (Figure 1A). These interactions were significantly different from those observed on switchgrass. On switchgrass leaves, P. emaculata spores adhered, germinated, and formed appressoria possibly after thigmotrophic signal-mediated oriented growth of the germ tubes to stomata. The penetrating infection hyphae then formed infection sites that eventually produced asexual urediniospores. Prehaustorial resistance is a very common form of NHR to parasitic rust fungi and is usually mediated by the activation of plant defense responses (Heath, 1977; Heath, 2000). Interestingly, the M. truncatula NHR response to P. emaculata was not associated with major transcriptional changes in the phenylpropanoid pathway or other pathogenesis-related (PR) genes compared with the mock-inoculated plants (see Supplemental Figure 1 online), suggesting a passive resistance mechanism. However, it is important to note that a subtle induction of Chitinase and PR-3 genes was observed at 24 h after inoculation (HAI) when compared with the wild-type plants (see Supplemental Figure 1 online). To identify mutants that compromise this particular NHR, we screened 1200 Tnt1 lines or 14,400 independent R0 or R1 plants (12 plants per each Tnt1 line) for loss of NHR to P. emaculata. Detached leaves from ~12 plants of each Tnt1 line were challenged with P. emaculata (see Methods and Supplemental Figure 2 online). Micro- and macroscopy observations of disease development were recorded at 8, 24, and 48 HAI and 5 d after inoculation to identify mutants compromised in NHR.

Figure 1.
M. truncatula irg1-1 Mutants Inhibit Preinfection Structure Differentiation by P. emaculata and P. pachyrhizi.

Strikingly, four independent Tnt1 lines showed inhibition of germ tube growth and differentiation on the abaxial leaf surface, instead of the expected phenotype of enhanced susceptibility (Figures 1A and and1B).1B). We named these mutants irg1 and found that all of the four identified independent mutants had Tnt1 insertions at different locations in the same exon of one gene. Therefore, we treated them as null mutant alleles (irg1-1, irg1-2, irg1-3, and irg1-4). Urediniospores were inoculated on the abaxial leaf surfaces (mimics natural infection on host plants), and the formation of germ tubes and the preinfection structures was visualized by the green fluorescence emitted by the wheat germ agglutinin-Alexa Fluor 488 conjugate, a fluorescent lectin that binds to N-acetyl-glucosamine in the cell walls and thus stains fungal structures (Figures 1A and and1B).1B). Unlike on the adaxial leaf surface of the wild-type and irg1-1 mutants where ~95% of the spores adhered, germinated, and formed long germ tubes, on the abaxial leaf surfaces of irg1-1 mutants, only ~60% of the spores germinated (Figures 1G and and1H).1H). By contrast, on the abaxial leaf surface of wild-type plants ~95% of the spores adhered, germinated, and differentiated into long germ tubes (Figures 1A and and1H).1H). Intriguingly, the spores that did germinate on the abaxial leaf surface failed to undergo any further differentiation on irg1 plants and did not show any further growth of the germ tubes as they did on wild-type plants (Figures 1A, 1B, and and1H).1H). Inhibition of preinfection structure formation on irg1-1 is most likely due to alterations in surface signal(s) required for the differentiation of preinfection structures of P. emaculata.

The irg1 Mutants Also Showed Inhibited Preinfection Structure Formation of P. pachyrhizi

To further test if irg1 mutants also inhibit preinfection structure formation by a broad spectrum of rust pathogens, we inoculated urediniospores of a direct-penetrating rust fungus, P. pachyrhizi, on abaxial leaf surfaces of M. truncatula wild-type R108 and irg1-1 mutants. M. truncatula is an incompatible host to P. pachyrhizi (Figures 1C to to1F).1F). Although P. pachyrhizi spores germinated, formed appressoria, and penetrated the epidermal cells causing visible necrosis, they failed to sporulate on wild-type M. truncatula when the inoculated plants were maintained in a dew chamber for 24 h with 100% humidity for spore germination and then incubated in a growth chamber with low (30 to 40%) humidity (Figure 1C). Very few necrotic lesions developed on irg1 compared with R108 plants (Figures 1C and and1D).1D). On adaxial and abaxial leaf surfaces of wild-type R108 plants and adaxial leaf surface of irg1 mutants, urediniospores adhered, germinated, and formed germ tubes, and most of the germ tubes underwent differentiation to form appressoria and directly penetrated the epidermal cells (Figures 1I to to1K).1K). However, on the abaxial leaf surface of irg1-1 mutants, the ability of the spores to adhere, germinate, and form germ tubes with appressoria was severely compromised, resulting in very low penetration of the epidermal cells (Figures 1F and and1J).1J). These results further confirmed that the abaxial leaf surface of irg1 leaves does not promote or inhibit the formation or differentiation of preinfection structures of at least two rust pathogens tested.

Inhibition of Rust Germ Tube Differentiation in irg1 Is Limited to Abaxial Leaf Surface

We noticed that the abaxial but not the adaxial leaf surfaces of irg1-1 plants were glossy in appearance when compared with the wild-type R108 plants in which neither surface is glossy (Figures 1C and and1D).1D). This suggested possible alterations in epicuticular wax loading on the abaxial leaf surface of irg1. In plant interactions with host-specific biotrophic pathogens, including Erysiphe pisi and Blumeria graminis, the components of abaxial leaf waxes were shown to specifically promote the differentiation of preinfection structures of E. pisi and B. graminis (Gniwotta et al., 2005; Hansjakob et al., 2010). To test if two different nonhost rust pathogens with different preinfection processes require surface cues for adherence or germ tube differentiation, we examined the preinfection structure formation of P. emaculata and P. pachyrhizi on the abaxial and adaxial leaf surfaces of the wild-type and irg1 plants (Figures 1G to to1J).1J). On the adaxial leaf surfaces of both wild-type and irg1 plants, almost 90% of the inoculated urediniospores of P. emaculata that germinated formed germ tubes with no appressoria on stomata (Figure 1G). Similarly, no inhibition of urediniospore germination or germ tube elongation was observed on the abaxial side of wild-type plants (Figure 1H). However, only ~60% of the spores germinated on the abaxial side of the irg1 mutants, and almost all the germinated spores failed to undergo any further differentiation (Figure 1H). These results further suggested the absence of stimulatory signals for germ tube differentiation or presence of inhibitory signals for growth of P. emaculata on the abaxial surface of irg1 plants.

Unlike P. emaculata, P. pachyrhizi is a direct-penetrating biotrophic rust fungus with a broad host range and, based on in vitro assays conducted on artificial membranes, is suggested not to require hydrophobic or chemical signals for preinfection structure formation (Koch and Hoppe, 1988; Goellner et al., 2010). Our in vivo assays conducted on the adaxial leaf surface showed no significant differences in germination, appressorium formation, or epidermal penetration by P. pachyrhizi between wild-type R108 and two independent mutant alleles of irg1 (Figures 1I). Interestingly, P. pachyrhizi showed slightly higher percentage of appressoria and penetration rate when inoculated on the abaxial leaf surface compared with adaxial leaf surface of the wild-type R108 (Figure 1J). However, on the abaxial surface of irg1-1 mutants, ~50% of the spores failed to germinate (Figure 1J). Strikingly, only ~20% of the germinated spores formed appressoria on the abaxial side of irg1-1 and irg1-2 mutant alleles, whereas ~75% of the germinated spores formed appressoria on the abaxial side of R108 (Figure 1J). Consistent with these results of reduced development of infection structures, we observed no induction of any PR and pathogen-inducible genes on the irg1-1 abaxial leaf surface inoculated with P. pachyrhizi urediniospores, suggesting that induced or constitutive defenses were not responsible for the observed phenotype (see Supplemental Figure 3A online). Furthermore, the total leaf proteins isolated from the irg1 mutants showed no significant inhibitory effects on the germination or germ tube differentiation of P. emaculata and P. pachyrhizi urediniospores on plastic (hydrophobic) surfaces, suggesting no predominant antimicrobial protein accumulation in these mutants that could account for the phenotype we observed on the abaxial leaf surface (see Supplemental Figures 3B and 3C online). These results suggested that, instead, the abaxial surface of irg1 leaves may lack the surface chemical/physical cues required for the differentiation of preinfection structure formation by P. pachyrhizi.

irg1 Showed Partial Resistance to C. trifolii but Not to Phoma medicaginis

To further test if irg1 mutants also exhibit resistance to compatible fungal pathogens, we challenged wild-type R108 and irg1-1 mutants with C. trifolii, a hemibiotrophic pathogen that forms preinfection structures (Figure 2A), and a necrotrophic pathogen, P. medicaginis, that directly penetrates without forming appressoria (Figure 2B). The percentage of C. trifolii spores that germinated and formed preinfection structures (appressoria) was significantly reduced on irg1-1 mutants, as were the consequent anthracnose symptoms developed on the abaxial leaf surface when compared with wild-type R108 (Figure 2A). The percentages of germination and appressoria formation were also slightly reduced (by ~10%) on the adaxial surface of irg1-1 and irg1-2 mutant alleles compared with the wild-type R108 (Figure 2A). However, no significant differences in symptom development or in planta fungal growth of P. medicaginis were observed upon inoculation on either abaxial or adaxial leaf surfaces of R108 and irg1-1 (Figure 2B). These results suggested that the effect of irg1 mutants may be limited to those fungi that form preinfection structures (appressoria) in response to surface signals.

Figure 2.
The irg1 Mutant Showed Partial Resistance to C. trifolii but Not to P. medicaginis.

IRG1 Encodes a Cys(2)His(2) Zinc Finger Transcription Factor and Is Allelic to PALM1

We recovered several Tnt1 flanking sequence tags from the irg1 mutant alleles, and the sequence of one of these had sequence similarity to a gene that encodes a Cys(2)His(2) zinc finger transcription factor. Chen et al. (2010) recently showed that the Cys(2)His(2) zinc finger transcription factor PALM1 controls trifoliate leaf development in M. truncatula. We decided to investigate whether there was an association between the irg1 phenotype and the PALM1 mutation. We evaluated four independent irg1 alleles identified in this study that have an insertion in PALM1 and found that irg1-1 is palm1-5 and irg1-2 is palm1-4 (see Supplemental Figure 4 online). In addition, all irg1 mutants as well as palm1 had five leaflets, unlike a typical M. truncatula leaf, which has three leaflets (see Supplemental Figure 5 online).

To further confirm if IRG1 gene is allelic to PALM1, we challenged a previously identified palm1 deletion mutant line M469 of M. truncatula ecotype A17 (palm1-1) and another independent Tnt1 line (line NF5022, palm1-6) that has an insertion in PALM1 (Chen et al., 2010) with P. pachyrhizi and P. emaculata. The formation of preinfection structures of P. pachyrhizi and P. emaculata was severely impaired on the abaxial leaf surface of all identified alleles of palm1, indicating that the loss of function of PALM1 is responsible for the irg1 phenotype (see Supplemental Figures 4 online). Since the palm1-6 Tnt1 line and the palm1-1 M469 deletion line also inhibited rust germ tube differentiation, we referred to them as irg1-5/palm1-6 and irg1-6/palm1-1, respectively. We further tested if the irg phenotype results from the loss of function of PALM1 by complementing the inhibition of germ tube differentiation phenotype of irg1-1/palm1-5 (R108 background) and irg1-6/palm1-1 (A17 background) by expressing wild-type PALM1 under the control of its native promoter in these mutants (Figure 3). The complemented lines did not show any inhibition of the rust preinfection structure formation when compared with their respective mutant lines (Figure 3). Taken together, these results showed that a gene involved in controlling leaf morphology also contributes to fungal resistance; therefore, going forward, we renamed our mutants and addressed them as irg1/palm1.

Figure 3.
Expression of the Cys(2)His(2) Zinc Finger Transcription Factor PALM1 in the irg1 Mutant Background Restores the Mutant Phenotype.

The irg1/palm1 Mutant Affects Abaxial Leaf Epicuticular Wax

Surface chemical and physical signals are known to affect adhesion, germination, and differentiation of preinfection structures in pathogenic fungi (Podila et al., 1993; Kolattukudy et al., 1995; Uppalapati and Fujita, 2000). Therefore, based on our pathogen assays and our observation that the abaxial leaf surface of irg1/palm1 is glossy, we speculated that irg1 mutants may have defects in cuticle structure or epicuticular wax deposition. Consistent with our hypothesis, scanning electron microscopy analyses of air-dried leaf samples showed that irg1 leaves, unlike the wild-type leaves, completely lacked epicuticular wax crystals on the abaxial leaf surface but did have wax crystals on the adaxial surface (Figure 4). The leaves from the complemented line (irg1-1:PALM1) showed restored wax crystals on the abaxial surface as seen on wild-type leaves.

Figure 4.
Loss of Function of IRG1/PALM1 Results in Loss of Epicuticular Wax Crystal Deposition on the Abaxial Leaf Surface of M. truncatula.

Epicuticular Waxes of irg1/palm1 Mutant Leaves Have Less Alcohols and More Alkanes

To further understand the nature of compositional changes in wax (aldehydes and primary alcohols; see Supplemental Figure 6 online) in irg1/palm1, we extracted the epicuticular waxes from R108, irg1-1/palm1-5, and irg1-2/palm1-4 lines. The total epicuticular waxes isolated from intact leaves were ~1.71-fold higher per leaf area in R108 compared with irg1 mutants. The amount of total acids and alcohols in irg1-1/palm1-5 leaves were 3.2- and 2.3-fold lower, respectively, than the R108 leaves (Figure 5A). However, the total alkanes were 2.2-fold higher in irg1-1/palm1-5 than the R108 leaves (Figure 5A). A similar trend in total acids, alcohols, and alkanes was observed for irg1-2/palm1-4 allele (Figure 5A).

Figure 5.
Loss of Function of IRG1/PALM1 Results in Dramatically Reduced Alcohol Content on the Abaxial Leaf Surface of M. truncatula.

Since our scanning electron microscopy pictures showed a lack of epicuticular wax crystals on the abaxial surface and the inhibition of rust spore differentiation was observed only on the abaxial surfaces, we hypothesized that the abaxial surface of irg1 may either lack a particular wax constituent that is required for the promotion of germ tube differentiation or may accumulate an inhibitory factor. To test this hypothesis, we isolated the epicuticular waxes separately from the abaxial and adaxial leaf surfaces. The amount of total alcohols, the predominant constituent of M. truncatula leaf waxes, and their composition were similar on the adaxial leaf surfaces of wild-type R108 and irg1/palm1 mutants (Figure 5B; see Supplemental Figures 7A and 7B online). However, on the abaxial surface, significant changes were observed in the amount and composition of alcohols and alkanes between R108 and irg1 mutants (Figure 5C; see Supplemental Figure 7 online). The total alcohols in the abaxial waxes were ~17-fold lower in irg1/palm1 mutant alleles than R108 (Figure 5C). Dramatic reductions of C28 and C30 alcohols in irg1 mutants were the major contributors for ~17-fold reduction in primary alcohols (see Supplemental Figures 7A and 7B online). The total alkanes in the abaxial waxes were approximately threefold higher in irg1 mutants than R108 (Figure 5C). C29 and C31 alkanes were found in higher amounts on both abaxial and adaxial surfaces of irg1/palm1 mutant alleles when compared with R108 leaves (Figure 5C; see Supplemental Figures 7C and 7D online). It is important to note that the total alkanes extracted from intact leaves using hexane were higher and did not exactly reflect the total of alkanes from abaxial and adaxial waxes isolated using gum arabic (Figure 5). It is possible that gum arabic absorbs alkanes poorly or most of the epicuticular waxes that are stripped with gum arabic are predominantly alcohols with some alkanes. It is also possible that the extracts from the whole leaf using hexane might also extract some of the intercuticular waxes (alkanes). Nevertheless, these results clearly demonstrate that loss-of-function mutation of IRG1/PALM1 leads to dramatic alterations in the amount and chemical composition of waxes.

Epicuticular Waxes/Hydrophobicity Promotes Germination and Appressorium Formation by P. pachyrhizi and P. emaculata

Our cytological analyses demonstrated that irg1/palm1 mutant alleles were defective in formation of epicuticular wax crystals and accumulation of alcohols on abaxial surface of leaves (Figures 4 and and5).5). We therefore hypothesized that the components of the epicuticular waxes/hydrophobic surface are required for P. pachyrhizi and P. emaculata to form preinfection structures. To test this hypothesis, we first quantified the differences in surface hydrophobicity by measuring the contact angle at the interface of a drop of liquid (water) with the leaf surface (sessile drop technique; Curvers et al., 2010). A higher contact angle (>90°) is indicative of poor wetting or a hydrophobic surface. No significant differences in the contact angle were observed between adaxial leaf surfaces of the wild-type and irg1-1 or irg1-2 mutants, which exhibited an average contact angle of 140° (Figure 6). However, very distinct differences in contact angles were observed between the abaxial leaf surfaces of the wild-type and irg1-1or irg1-2 mutants (Figure 6). The abaxial leaf surface of wild-type plants exhibited an average contact angle of 138°, whereas the mutant alleles showed a dramatic decrease in contact angle (average of 92°), which is indicative of a hydrophilic surface. Although the abaxial surface of the mutants contained dramatically reduced levels of alcohols, these results suggested that the lack of primary alcohols and epicuticular wax crystals increased the hydrophilicity of irg1/palm1 mutant leaves, and the three-dimensional surface morphologies of epicuticular waxes and their polymerization patterns may play an important role in the hydrophobicity of M. truncatula leaves.

Figure 6.
Loss of Function of IRG1/PALM1 Results in Reduced Contact Angle and Hydrophobicity of Water Droplets on the Abaxial Leaf Surface of M. truncatula.

To further test the role of M. truncatula epicuticular waxes and/or hydrophobicity contributed by the epicuticular wax in stimulating the differentiation of fungal structures, we made quantitative analyses of fungal development (spore germination, germ tube elongation, and appressorium differentiation) on hydrophilic (glass) surfaces, which were uncoated or coated with epicuticular waxes isolated from soybean, switchgrass, wild-type M. truncatula, and irg1 mutants. We also studied the effect of C30 alcohol, the predominant alcohol that was absent on the abaxial leaf surfaces of irg1/palm1 mutants, by quantifying fungal development on slides coated with C30 alcohol. In addition, we also studied fungal development on host leaf surfaces where the waxes were manually removed.

On hydrophilic uncoated glass surfaces mock coated with hexane, germination and differentiation of P. pachyrhizi urediniospores were severely impaired, wherein only 13.70% (±5.13) of the urediniospores germinated and <5% of the total spotted spores on the glass slides formed appressoria (Figure 7A). The glass slides coated with total waxes isolated from the adaxial leaf surface of the wild-type R108 and irg1/palm1 mutants (irg1-1 and irg1-2) showed a significant increase in percent germination (53.53 to 70.7%) and appressorium formation (19.15 to 23.59%) compared with the percentage of germination on the mock-coated slides (Figure 7A). No significant differences in their ability to promote spore germination and appressorium formation were observed between the adaxial waxes isolated from wild-type R108 and irg1/palm1 mutants (irg1-1 and irg1-2) (Figure 7A). However, consistent with our observations on the abaxial leaf surfaces, the total waxes isolated from the abaxial leaf surface of irg1-1/palm1-5 or irg1-2/palm1-4 promoted only ~30% germination of the P. pachyrhizi urediniospores compared with the ~70% germination induced by the abaxial waxes of R108 (Figure 7B). Although the total abaxial waxes from irg1 alleles promoted ~30% germination and germ tube growth, they failed to promote the formation of appressoria, and the percentage of total spores that formed appressoria was only ~5 to 8%, comparable to the mock-coated slide glass (Figure 7B). These results implicated that the physical (hydrophobicity) or chemical (primary alcohols) cues imparted by the adaxial or abaxial epicuticular waxes extracted into hexane from the wild-type plants and adaxial surfaces of irg1 mutant alleles can promote P. pachyrhizi spore germination and appressorium formation. These results further suggested the requirement of specific plant signals for fungal development on leaf surfaces.

Figure 7.
Effect of Epicuticular Wax on Development of Urediniospores of P. pachyrhizi and P. emaculata.

To further test the specificity of the waxes in promoting P. pachyrhizi spore germination and differentiation, we evaluated the effects of total waxes isolated from the abaxial and adaxial leaf surfaces of soybean, the host for P. pachyrhizi, and switchgrass, a monocot nonhost of P. pachyrhizi (Figures 7A and and7B).7B). Although the adaxial and abaxial waxes from soybean promoted germination comparable to waxes from M. truncatula R108, they promoted more appressorium formation (~40%) compared with the total waxes from adaxial or abaxial surfaces of M. truncatula R108 (Figures 7A and and7B).7B). Interestingly, the total waxes from the adaxial or abaxial surfaces of switchgrass promoted spore germination that is comparable to R108 or soybean (Figures 7A and and7B).7B). However, the total waxes from switchgrass leaves failed to induce a high percentage of appressorium formation; specifically, waxes from the abaxial leaf surfaces of switchgrass promoted a very low percentage of appressorium formation (~9%), comparable to the abaxial leaf surface waxes of irg1 mutants (Figures 7A and and7B).7B). These results further suggested that waxes (or hydrophobicity) in general promote spore germination but that appressorium formation requires more specific signals, and the constituents of the waxes may affect these processes.

Chemical analyses showed that the abaxial surfaces of the irg1 alleles failed to accumulate primary alcohols and form abaxial wax crystals. Therefore, we tested if the C30 primary alcohol, the main constituent of M. truncatula waxes, affects the differentiation of spores. Hydrophilic glass slides coated with C30 alcohols at 5 μg/cm2 promoted a 3.5-fold increase in percentage of germination and a 5.4-fold increase in appressorium formation by the germinating spores of P. pachyrhizi (Figure 8A). The C30 alcohol concentration of 5 μg/cm2 is similar to the concentration present on M. truncatula leaf surfaces, and stimulation of appressorium formation was observed even at 0.5 μg/cm2. C30 alcohols also promoted a significant increase in percentage of germination of P. emaculata spores at high concentration (5 μg/cm2; Figure 8B). Unlike P. pachyrhizi spores, on hydrophilic (uncoated glass) surfaces, the urediniospores of P. emaculata germinated to a higher level (~20 to 30%) and formed germ tubes that continued to grow without forming appressoria (Figure 8C). In addition, P. emaculata spores also failed to form appressoria on glass slides coated with waxes or primary alcohols isolated from leaf surfaces (data not shown). Although we were unable to test all the different chain length alcohols and other alkanes or aldehydes, our results clearly suggested that the primary alcohols present in M. truncatula leaf surfaces provide the chemical and physical cues for promotion of infection structure formation by P. pachyrhizi and P. emaculata.

Figure 8.
Effect of Primary Alcohol and Epicuticular Wax on Development of Urediniospores of P. pachyrhizi and P. emaculata.

To further confirm the requirement for hydrophobic waxes and to corroborate the results obtained with the irg1/palm1 mutants (Figure 1) and glass slides (Figures 7, 8A, and 8B), we followed the development of P. pachyrhizi urediniospores on the abaxial surface of the native host plant, soybean (Wax+), and on the abaxial surface of soybean that was gently rubbed with a buffer solution containing celite and bentonite to remove most if not all the epicuticular wax layer (Wax). On Wax+ abaxial surfaces, 70% of P. pachyrhizi urediniospores formed appressoria (Figure 8D), whereas on Wax surfaces, ~45% of the inoculated spores formed appressoria and penetrated the epidermal layer (Figure 8D). Consistent with these spore differentiation defects, a significant reduction in Asian soybean rust infection was observed in detached soybean Wax leaf surfaces (Figure 8E). It is important to note that celite and bentonite remove epicuticular wax crystals but may also damage the cuticle for a short time (24 h) and subsequent cuticle-mediated signals are shown not to alter the local responses to pathogen or the outcome of susceptibility (Xia et al., 2009). However, we could not completely rule out any such small contributions of signals in the reduced pathogenicity on surfaces where the waxes were physically removed. Nevertheless, taken together with the experiments done on glass slides, these results further confirmed that surface waxes are critical or function as stimulatory signals for P. pachyrhizi spore germination and differentiation of appressoria.

We further tested the requirement for surface waxes (hydrophobicity) for prepenetration development (i.e., germination, germ tube elongation, and appressorium differentiation) of P. emaculata urediniospores on Wax+ and Wax abaxial surfaces of the host plant switchgrass. A 35 to 40% reduction in appressoria (appressoria formation on stomata) was observed on Wax abaxial surfaces (Figure 8F). On Wax+ switchgrass, the germinated spores formed appressoria over the stomatal openings (Figure 8F, top panel). On the Wax surfaces, although the germinated spores oriented to recognize the stomata, a significant number of them failed to form appressoria on the stomata (Figure 8F, bottom panel). These results along with data about germination on slides coated with primary alcohols suggested that P. emaculata spores require waxy surface signals for appressorium formation and for enhanced germination but not for initial germ tube growth.

Transcript Profiling Identifies a Role for IRG1/PALM1 in Regulating Expression of Genes Involved in Long-Chain Fatty Acid Biosynthesis and Transport

One of the findings of our study is that the loss-of-function mutation of a transcription factor involved in leaf morphogenesis impacts epicuticular wax loading in M. truncatula, which in turn affects germination and differentiation of fungal spores (Figures 4 and and5).5). To understand this phenomenon at a molecular level, we compared the transcript profiles of wild-type R108 and three independent irg1/palm1 homozygous null mutant lines (irg1-1, irg1-2, and irg1-5) using Affymetrix GeneChip Medicago Genome Array (Figure 9A). It is important to note that these lines have Tnt1 insertions in different locations of the same IRG1/PALM1 exon (see Supplemental Figure 4 online) and also have multiple insertions in other independent locations in the genome. Therefore, all three mutants tested had both common and unique differential gene expression patterns compared with R108 (Figure 9A, Table 1; see Supplemental Data Set 1 online). We used a set of 400 upregulated and 48 downregulated genes that were commonly altered in all three mutant alleles compared with R108 to identify the major pathways targeted by IRG1/PALM1 (Figure 9A, Table 1; see Supplemental Data Set 1 online).

Figure 9.
Overview of Global Transcript Changes in irg1/palm1 Mutants.
Table 1.
A List of Selected Wax or Lipid Biosynthesis-Related Genes Differentially Regulated in irg1/palm1 Mutant Alleles (irg1-1/palm1-5, irg1-2/palm1-4, and irg1-5/palm1-6) Compared with the Wild Type

The majority of the most significantly regulated genes (twofold change, P value < 8.15954E-07), a major portion are predicted to function in wax/lipid biosynthesis (Table 1; see Supplemental Data Set 1 online). In irg1/palm1, genes involved in cuticular wax accumulation, including ECERIFERUM2 (CER2), which encodes a nuclear-localized protein (Xia et al., 1997), CER1, which encodes a putative decarbonylase known to promote long-chain alkane biosynthesis (Aarts et al., 1995), and several genes encoding lipid transfer proteins (LTPs) and ABC transporters were upregulated (more than twofold), whereas most of the putative wax biosynthetic genes represented on the microarray chip, including those encoding ECERIFERUM, CER4-2, CER2-2, CER5 (Pighin et al., 2004), CER8 (Lü et al., 2009), CER6, and KCS1, were downregulated (Table 1; see Supplemental Data Set 1 online). To further understand the effects of PALM1 mutation on wax biosynthesis, we studied the expression of M. truncatula orthologs of Arabidopsis thaliana genes implicated in wax biosynthesis using real-time quantitative RT-PCR (qRT-PCR; Table 1, Figure 9B; see Supplemental Table 1 online). Consistent with the microarray results, a significant upregulation of CER2-1 (sevenfold) and CER1 (fivefold) was observed in irg1/palm1 (Figure 9B; see Supplemental Table 1 online). Several other genes implicated in wax biosynthesis, including CER4, which encodes an alcohol-forming fatty acyl-CoA reductase, CER6-3, CER8, β-keto acyl-CoA reductase (KCR1 and KCR2), and wax synthase (WSD1) were downregulated. Approximately fourfold downregulation was observed for CER4-2 and CER6-4 genes, while others were moderately downregulated (1.5-fold to twofold) in irg1/palm1 compared with wild-type R108 (Figure 9B; see Supplemental Table 1 online). Interestingly, a homolog of Arabidopsis MYB96, a key transcription factor involved in regulation of wax biosynthesis (Seo et al., 2011), was strongly downregulated in irg1/palm1 mutant alleles (Figure 9B; see Supplemental Table 1 online). Furthermore, the transcriptome data also identified several P450 genes and PR genes, including chitinases and β-1-3-glucanases that were preferentially upregulated in irg1/palm1 mutants (see Supplemental Data Set 1 online). In summary, the transcript profiles of the three independent irg1/palm1 alleles showed alterations in expression of several genes involved in wax biosynthesis and transport. Functional analysis of the transporters (ABC/LTPs) and overexpression of CER4, CER2, and MYB96 in M. truncatula will help us to better understand the mechanism of wax biosynthesis and asymmetric epicuticular wax loading in irg1 mutants.


In this study, we identified and characterized an epicuticular wax mutant (irg1) of M. truncatula in a gene encoding a Cys(2)His(2)zinc finger transcription factor and showed that a loss-of-function mutation of IRG1 causes major changes in epicuticular wax content and composition on abaxial leaf surfaces. Most intriguingly, these changes in the irg1 mutant impart resistance to certain biotrophic fungal pathogens by inhibiting the differentiation of preinfection structures. Our results showed that irg1/palm1 mutants were completely devoid of wax crystals on the abaxial leaf surface and suggested a possible role for signals at the cuticle interface, especially the components of the leaf wax in promotion or inhibition of rust preinfection structure differentiation on the abaxial leaf surface of the irg1 mutants (Figures 1 and and2).2). Consistent with our results, two other previous studies have shown normal development of fungal preinfection structures on the adaxial surface and short germ tubes with few appressoria on abaxial leaf surfaces during host-specific pathogenic interactions involving wild-type pea (Pisum sativum)–E. pisi and wild-type ryegrass (Lolium spp)–Erysiphe graminis interactions (Carver et al., 1990; Gniwotta et al., 2005). Variations in wax crystal composition between adaxial and abaxial leaf surfaces were implicated in pea–E. pisi interactions (Gniwotta et al., 2005). However, the genes or mechanism(s) responsible for this variation in wax crystals between adaxial and abaxial surfaces were not identified. Furthermore, our results also provided evidence for requirement of waxes at contact surfaces for appressorium differentiation by P. pachyrhizi and further strengthened the earlier hypothesis that surface composition and hydrophobicity play important roles in appressorium formation and penetration by direct penetrating fungi (Lee and Dean, 1994).

Interestingly, hydrophobicity was not very critical for germination or germ tube elongation or preinfection structure development by P. emaculata urediniospores. Although primary alcohols enhanced the percentage germination (Figure 8B), P. emaculata urediniospores germinated quite efficiently (20 to 25%) and formed long germ tubes on the hydrophilic (glass) surface (Figure 8C) and on host leaf surfaces where epicuticular waxes were removed (Figures 8F, bottom panel). However, on the abaxial surface of irg1 mutants, although the spores germinated, they failed to elongate and undergo any further morphogenesis. These results suggested a possible role for unknown chemical signals on the abaxial leaf surface of irg1 in inhibiting P. emaculata germ tube elongation. Plants with defective cuticle structure and hydrophobicity (Bessire et al., 2007; Chassot et al., 2007, 2008; Curvers et al., 2010) have strong resistance to Botrytis cinerea, possibly through increased release of antimicrobial compounds from more permeable epidermal cells. Very-long-chain aldehydes have been shown to promote preinfection structure formation of B. graminis (Hansjakob et al., 2010) and Puccinia graminis f. sp tritici (Reisige et al., 2006). Chemical analyses of irg1 mutants showed increased accumulation of alkanes in comparison to wild-type R108 (Figure 5; see Supplemental Figure 7 online), indicating a possible role in inhibiting the growth of P. emaculata germ tubes. It is possible that the effects of other inhibitory signals may be more conspicuous in the absence of alcohols that show some stimulatory effects on overall percentage of germination. We tried to isolate any possible surface antimicrobials (proteins) from leaves using the methods described for tomato (Solanum lycopersicum) fruit surfaces (Yeats et al., 2010) but failed to obtain sufficient protein concentration to conduct spore germination assays. However, our results using total leaf proteins isolated from irg1/palm1 mutant did not show any inhibitory activity.

The asymmetric distribution of leaf epicuticular waxes to the abaxial side in the irg1/palm1 mutants is intriguing. Complete absence of wax crystals on abaxial and adaxial leaf surfaces of wild-type Arabidopsis (Jenks et al., 1995) and absence of wax crystals on abaxial but not adaxial surfaces of wild-type Lolium perenne (Ringelmann et al., 2009) have been reported. Several studies have reported complex changes in wax composition in epicuticular wax mutants, including cer1-cer6, resulting from increased flux of precursors into other metabolic pathways in Arabidopsis stems (Aarts et al., 1995; Jenks et al., 1995; Rowland et al., 2006; Kunst and Samuels, 2009). Our chemical analyses further demonstrated that loss of function of a Cys(2)His(2) zinc finger transcription factor (PALM1) also results in changes in the distribution of different classes of waxes among adaxial and abaxial surfaces. These complex changes in composition and distribution of waxes in irg1 can be attributed to the increased flux of metabolites into the decarbonylation pathway (see Supplemental Figure 6 online).

Our transcriptome analysis revealed that IRG1/PALM1 regulates several target genes involved in lipid metabolism and transport. Based on the complete absence of wax crystals phenotype on the abaxial side and 50% reduction in primary alcohols, the major wax component, we expected a dramatic downregulation of target genes involved in wax biosynthesis. However, only homologs of CER4 and CER6 showed significant downregulation (less than or equal to twofold) in irg1 plants compared with the wild type (Figure 9B, Table 1). Interestingly, homologs of Arabidopsis CER2 were upregulated in irg1 plants (Table 1, Figure 9B). CER6/CUT1 encodes a putative KCS that is potentially involved in elongation of fatty acyl-CoAs longer than C22, and deletion of CER6/CUT1 results in 93 to 94% reduction of total wax loading and almost inactive decarbonylation pathway in Arabidopsis (Millar et al., 1999; Fiebig et al., 2000). CER4 encodes fatty acyl-CoA reductase, which is responsible for primary alcohol formation in Arabidopsis (Rowland et al., 2006). Consistent with our chemical analyses, the transcriptome analysis provided the genetic evidence that the downregulation of a CER4 was responsible for the reduced alcohols on the abaxial surface of irg1/palm1 mutant leaves. Furthermore, CER1, a major enzyme that promotes long-chain alkane biosynthesis (Aarts et al., 1995; Bourdenx et al., 2011), and homologs of CER1, including gl1/cer3/wax2 (Rowland et al., 2007; Mao et al., 2012), were upregulated in the microarray analyses. Taken together with the chemical analyses, our results also further suggested that the homologs of Arabidopsis CER4 and CER1 function to promote alcohol and alkane biosynthesis, respectively, in M. truncatula. Based on these findings and our results, it is tempting to speculate that downregulation of CER4 and concomitant upregulation of CER1 result in a reduced accumulation of primary alcohols and increased flux of precursors into the decarbonylation pathway, resulting in the accumulation of alkanes in irg1 mutants of M. truncatula (Table 1, Figure 5; see Supplemental Figure 6 online). Therefore, the irg1/palm1 mutant may be helpful in elucidating the biochemical function of CER1 in C30 alcohol biosynthesis in M. truncatula and other crop legumes. Primary alcohols appear to be the predominant form of very-long-chain fatty acids in the epicuticular waxes of fabaceae, including M. truncatula (Zhang et al., 2005, 2007) and pea (Gniwotta et al., 2005). Furthermore, our results also showed that the plate-type wax morphologies in M. truncatula mainly contained primary alcohols and were required for three-dimensional structure formation of surface waxes and surface hydrophobicity. Our results also suggested that the physical (hydrophobicity) and chemical surface cues promote spore germination and germ tube elongation of several biotrophic fungi. In addition, the physical and chemical surface cues also promoted appressorium formation by P. pachyrhizi (Figures 7 and and8).8). Altered wax composition impacts the initial events of pathogenesis and spore differentiation during compatible plant–fungal interactions (Kolattukudy et al., 1995; Gniwotta et al., 2005; Zabka et al., 2008; Hansjakob et al., 2010, 2011). Due to the absence of the long-chain aldehydes from the leaf cuticular wax, the glossy11 mutant of maize (Zea mays) doesn’t support appressorium formation and subsequent prepenetration by B. graminis and thus is resistant to this fungus (Hansjakob et al., 2010, 2011). Future studies involving overexpression of CER1 and Myb96 in wild-type and irg1/palm1 mutants may shed new light on the role of different classes of very-long-chain fatty acids and their quantities in formation of epicuticular wax structures and their contributions to hydrophobicity/fungal differentiation in M. truncatula. The abaxial leaf surface of irg1/palm1 mutants may also provide a natural leaf surface with altered wax content and composition that would allow us to test new hypotheses for the role of epicuticular waxes in fungal spore germination and germ tube differentiation.

In addition to CER1, CER2 and two genes encoding LTP-like proteins were upregulated in irg1 mutants when compared with wild-type R108 (Table 1). Based on the EST information, we could not confirm if the LTP genes encoded glycosylphosphatidylinositol-anchored LTPs. However, one of the LTPs (Affy ID, Mtr.13293.1.S1_at) showed high similarity to nonspecific LTPs (ns-LTPs). Some LTPs have been shown to be secreted and accumulate extracellularly where they can play a role in diverse functions, including cuticular wax transport and defense against pathogens (Segura et al., 1993; Kader, 1996; Kunst and Samuels, 2003). Loss of function or reduced expression of a glycosylphosphatidylinositol-anchored LTP results in reduced alkane accumulation at the plant surface and plays a role in lipid export (Debono et al., 2009; Lee et al., 2009). Therefore, it is tempting to speculate that upregulation of the LTP (Mtr.13293.1.S1_at) could be one of the reasons for increased alkane accumulation in irg1 mutants.

Unlike the contributions of the constituents and morphologies of the epicuticular waxes, the role of cuticle and cuticular lipids has been well studied in plant–pathogen interactions. Studies done with cuticle mutants, including gpat4/gpat8 (Li et al., 2007), lacs2 and att1 (Xiao et al., 2004; Tang et al., 2007; Lee et al., 2009), and gl1 (Xia et al., 2010) have shown that these mutants are more susceptible to pathogens due to a range of alterations, including stomatal or substomatal spaces or cuticle-derived active signaling. By contrast, enhanced resistance of att1 and lacs2 mutants to B. cinerea was reported either via enhanced perception of the fungal elicitors because of the permeable surfaces of these mutants leading to the accumulation of antimicrobials or enhanced upregulation of defense-related genes (Bessire et al., 2007). Increased susceptibility to the biotrophic pathogen Erysiphe cichoracearum and resistance to necrotrophic fungal pathogens B. cinerea and Alternaria brassicicola were demonstrated in an Arabidopsis rst1 mutant. Interestingly, rst1 was shown to be a cuticular wax mutant with 59.1% reduction in waxes (and wax crystal deposition) on stem but 43% increase of waxes in the leaves (Chen et al., 2005). RST1 was shown to influence plant defense responses by altering the interactions with jasmonic acid– and salicylic acid–mediated pathways (Mang et al., 2009). However in our study, irg1/palm1 showed cuticular wax defects but did not show any alteration in the expression of genes involved in SA pathway or other phytoalexin-mediated pathways in mock- or pathogen-inoculated leaves compared with the wild type, suggesting a predominant role of altered abaxial leaf surface properties (hydrophobicity or wax constituents) in the promotion of biotrophic fungal differentiation.

In conclusion, we provide evidence for an increased disease resistance phenotype of irg1/palm1 plants possibly due to altered abaxial leaf surface signals that inhibit differentiation of fungal preinfection structure in M. truncatula. Although both developmental and environmental cues affect wax biosynthesis, only a very few transcription factors that regulate wax biosynthesis have been isolated (Samuels et al., 2008; Kunst and Samuels, 2009; Seo et al., 2011). Overexpression of the transcription factor WXP1 causes increased accumulation of acyl-reduction pathway products in M. sativa leaves (Zhang et al., 2005). Recently, a homeodomain-Leu zipper IV family transcription factor was shown to regulate genes involved in cuticle biosynthesis (Javelle et al., 2010). Furthermore, overexpression of wax inducer/SHINE family in Arabidopsis and APETALA2 (AP2)/ethylene-responsive element binding protein–type transcription factors, WXP1 and WXP2, in Medicago positively regulate wax biosynthesis (Aharoni et al., 2004; Broun et al., 2004; Zhang et al., 2005, 2007). Interestingly, Cys(2)His(2) zinc finger transcription factor (IRG1/PALM1) contains an ERF-associated amphiphilic repression domain at the C-terminal region and is conserved in the class II ERF transcriptional repressors of the AP2/ERF domain proteins (Chen et al., 2010). How a zinc transcription factor with an ERF transcriptional repressor domain regulates wax biosynthesis, and asymmetric distribution of epicuticular wax is a subject for future research. The irg1/palm1 mutant identified in this study confers altered leaf morphology and wax deposition on only one side of the leaf surface. However, it is important to note that the epicuticular waxes are also shown to regulate nonstomatal water loss and are important in protecting plants against water loss (Riederer and Schreiber, 2001). Our preliminary results suggested that irg1 mutants were not more susceptible to water loss than the wild type, but the effect of loss of waxes on the abaxial side of the leaf on drought susceptibility warrants further systematic study. Further characterization of irg1/palm1 mutants may help to improve our understanding of asymmetric epicuticular wax loading on leaf surfaces and may provide us with approaches to specifically engineer adaxial or abaxial leaf surface waxes to improve fungal resistance both by regulating fungal differentiation and by improving the penetration ability of pesticides in agricultural spray applications.


Plant Materials

Seeds of Medicago truncatula cv Jemalong A17, R108, and Tnt1 insertion lines of M. truncatula mutant collection, NF0227 (irg1-1/palm1-5), NF1271 (irg1-2/palm1-4), NF1432 (irg1-3), NF4045 (irg1-4), and NF5022 (irg1-5/palm1-6) in R108 background, and M469 (irg1-6/palm1-1) in A17 background from the deletion mutant collection of M. truncatula were scarified for 8 min using concentrated sulfuric acid, washed thrice with distilled water, and germinated on moist filter papers. Two days after germination in darkness at 24°C, 12 seedlings from each Tnt1 line were transferred to soil (one seedling per cell in 6 × 12 celled trays). Following 3 weeks incubation in the greenhouse, the plants were transferred to growth chambers located in a USDA–Animal and Plant Health Inspection Service–approved BSL2+ facility to conduct soybean rust or switchgrass (Panicum virgatum) rust inoculation assays.

Screening of M. truncatula Tnt1 Insertion Population

To identify Tnt1 mutants with altered resistance to switchgrass rust, we set up a forward genetic screen using a detached leaf assay. An isolate of the switchgrass rust causative agent Puccinia emaculata collected from Oklahoma (PE-OK1) was maintained on a susceptible lowland switchgrass (P. virgatum cv Summer). Fresh urediniospores of PE-OK1 were collected using a gelatin capsule spore collector designed by the Cereal Disease Laboratory, St. Paul, MN, and suspended in distilled water with 0.001% Tween 20. The abaxial side of the detached leaves were spray inoculated with 105 spores/mL (0.001% Tween 20) using an artist airbrush (Paasche Airbrush) set at 2 p.s.i. with a portable air pump (Gast Manufacturing) for uniform spore deposition. The inoculated leaves were maintained on moist filter papers and incubated overnight in dark and then maintained at 24°C with 16-h-light/8-h-dark cycle. It is important to note that we used an R0 or R1 segregating Tnt1 population for the forward genetic screen, and one detached leaf from each of 12 plants representing one Tnt1 line was spray inoculated with P. emaculata spores as described above. Tnt1 population is shown to contain multiple copies of the Tnt1 tag in each line, and most of the phenotypes are suggested to segregate in the R1 progeny (Tadege et al., 2008). Therefore, we used a sufficiently large segregating R1 population (12 plants per each line) with a possibility of recovering at least one Tnt1 line with homozygous insertion in a given gene that also confers altered NHR response to P. emaculata.

Asian Soybean Rust Maintenance and Inoculation Procedures

An isolate of Asian soybean rust pathogen, Phakopsora pachyrhizi, from Illinois was maintained on the susceptible soybean cultivar (Glycine max cv Williams) grown in a growth chamber at 22°C/19°C with a 12-h-light/12-h-dark cycle (1000 μmol·m–2·s–1). P. pachyrhizi urediniospores collected and prepared as described above for switchgrass rust were used to inoculate detached leaves or whole plants grown in 72-cell trays. The inoculated plants were maintained in a dew chamber for 24 h with 100% humidity maintained at 19°C with a 0-h-light/24-h-dark cycle. The plants were then transferred to a growth chamber (22°C/19°C with a 12-h-light/12-h-dark cycle) and incubated further to allow symptom development.

Light, Confocal, and Scanning Electron Microscopy

Initial interactions of P. pachyrhizi or P. emaculata with M. truncatula were recorded by direct observations of inoculated leaves using an Olympus stereomicroscope (SZX19) or compound microscopes (BX 41) equipped with fluorescence attachment. For fluorescence microscopy, fungal mycelia were stained with wheat germ agglutinin (WGA), coupled to the green fluorescent dye Alexa Fluor 488 (WGA-Alexa Fluor 488; Invitrogen) as described previously (Uppalapati et al., 2009). Inoculated leaves were stained with 10 μg/mL WGA-Alexa Fluor 488 by a brief vacuum infiltration in PBS followed by a 20-min incubation at room temperature. For microscopy observations, after washing with PBS, whole leaves or sections of the leaf were placed on a glass slide and mounted using a cover glass with Dow Corning high vacuum grease for microscopy. Fluorescence microscopy to document the infection process was done using an Olympus epifluorescence microscope (BX 41) or a Leica TCS SP2 AOBS confocal laser scanning microscope (Leica Microsystems) equipped with ×20 (numerical aperture of 0.70) and ×63 (numerical aperture of 1.2, water immersion) objectives using appropriate laser and excitation filter settings (WGA-Alexa Fluor 488 to 488 nm). Chloroplast autofluorescence was captured by exciting with the 647-nm line of the argon-krypton laser and emission detected at 680 nm. A series of optical sections (z series) were acquired by scanning multiple sections, and the z-series projections were done with the software provided with the Leica TCS SP2 AOBS confocal laser scanning microscope.

Scanning electron microscopy analysis of the air-dried leaves was performed as described previously (Zhang et al., 2005). Briefly, leaves from the top two internodes were harvested and air-dried at room temperature in a Petri dish, since the conventional scanning electron microscopy sample preparation would wash the surface waxes away. Air-dried leaves were mounted on stubs and coated with ~20 nm of 60/40 Gold-Palladium particles using a Hummer VI sputtering system (Anatech). Coated surfaces were viewed using a JEOL JSM-840A scanning electron microscope at 15 kV.

Surface Hydrophobicity Measurements

Leaves were fixed to glass slides with double-sided tape and 10-μL droplets of distilled water were dropped using a micropipette. The photographs of the droplets were taken, and the contact angle (θC) was measured using the angle tool incorporated in the image J version 1.44p software. A total of 12 independent measurements per abaxial or adaxial leaf surfaces were performed for wild-type and two mutant alleles. Two water droplets per leaf surface were evaluated on three plants grown in independent pots from three different experiments. The average of all the 12 replicates was used for contact angle measurements.

Preinfection Structure Formation and Penetration Assays

Approximately 100 spores of P. emaculata or P. pachyrhizi in 10-μL aliquots were placed on the adaxial or abaxial surface of the detached leaves from 4-week-old M. truncatula wild-type R108 or irg1 mutant plants and incubated in dark overnight and then transferred to a growth chamber (22°C/19°C with 12-h-light/12-h-dark cycle). To capture the early stages of preinfection structure formation, 24 HAI, the inoculated leaves were washed two to three times in a Petri dish with PBS to remove the free-floating spores and were stained by floating in a PBS solution supplemented with 0.05% Tween 20 and 10 μg/mL WGA-Alexa Fluor 488 to visualize the fungal germ tubes and appressoria. Urediniospore-forming germ tubes >60 μm in length were counted as long germ tubes, whereas germ tubes ≤50 μm in length were counted as short germ tubes. The number of P. pachyrhizi and P. emaculata spores that germinated and formed germ tubes >10 μm in length were evaluated at 24 HAI and were counted as the percentage of germination. The subsequent developments were followed 72 HAI, and the germinated tubes forming differentiated appressoria were counted as appressoria, and the differentiated germ tubes without appressoria that grew on the surface were also counted from 20 random fields on three independent leaves. On M. truncatula, the urediniospores of P. emaculata germinated and formed long germ tubes but failed to form appressoria (on the stomata) and penetrate. Therefore, only the percentage of germination and the differentiated germ tubes without appressoria were evaluated for P. emaculata on M. truncatula. The number of dead autofluorescing epidermal cells resulting from direct penetration of P. pachyrhizi was counted 72 HAI from 20 random fields per each inoculated site and is used to calculate the percentage of penetration. It is important to note that the percentage of germination reflects the total percentage of urediniospores that germinated and is a representation of differentiated germ tubes without appressoria + differentiated appressoria + percentage of penetration.

Fungal Differentiation on Artificial Surfaces and Leaf Surfaces Manipulated to Remove the Surface Wax Coating

The abaxial and adaxial leaf surface epicuticular waxes from M. truncatula wild-type and irg1/palm1 mutants, soybean, or switchgrass were isolated using a polymer film of gum arabic as described (Gniwotta et al., 2005). The polymer films peeled from two to three leaflets were pooled and extracted with hexane, evaporated, and resuspended by sonication in fresh hexane. The hexane solution of waxes from wild-type R108 or irg1 mutants was adjusted to a final concentration of 0.05 mL/cm2 leaf area. Uncoated, frosted glass slides (25 × 75 mm; VWR International) were coated (three times) with a hexane solution to cover the whole surface. Hexane was allowed to completely evaporate between each application as described (Podila et al., 1993). The C30 alcohol 1-triacontanol (Sigma-Aldrich) was dissolved in hexane to a final concentration of 7 × 10−3 mol/L (~5 μg/cm2) and 7 × 10−4 mol/L (~0.5 μg/cm2) and was applied to the glass as described above for the leaf waxes. The approximate coverage per cm2 was estimated based on the assumptions made by Hansjakob et al. (2010). Uncoated glass slides were coated with same amount of hexane, the solvent used for wax preparations, and were referred as mock coated/control. To measure the effect of surface hydrophobicity or surfaces waxes on appressorium formation, a spore suspension of P. pachyrhizi was applied on the mock control and glass slides coated with leaf waxes or C30 alcohol.

For removal of epicuticular waxes, detached leaf surfaces of switchgrass and soybean were gently rubbed with a cotton swab saturated with a solution containing bentonite (0.02% [w/v]; Sigma-Aldrich) and celite (1% [w/v]; Sigma-Aldrich) as described (Xia et al., 2009). The urediniospores of P. emaculata and P. pachyrhizi were inoculated on the respective host leaf surfaces by spotting 10 μL aliquots of 104 spores/mL in distillated water with 0.001% Tween 20 on the native and surfaces manipulated to remove wax. The percentage of preinfection structures was evaluated as described above in the preinfection structure formation and penetration assays section.

Infection Assays with Colletotrichum trifolii

C. trifolii race 1 was maintained on potato dextrose agar media (Becton Dickson and Company). Conidia from 10- to 14-d-old cultures were harvested, washed in water, and resuspended in sterile distilled water. Leaves from 4-week-old wild-type R108 and irg1 mutants were harvested and spot inoculated with 10 μL of suspension or spray inoculated with a suspension containing 1 × 106 spores/mL in 0.005% Tween 20. The fungal structures were stained with lactophenol trypan blue, and the percentage of spores forming different preinfection structures (bottom panel) was evaluated, 72 HAI, by counting 20 random fields. The data represent the mean of three independent experiments.

Complementation and Stable Transformation

The irg1-6/palm1-6 complementation lines were previously described (Chen et al., 2010). In this study, we developed stable transgenic lines of irg1-1 for complementation studies using pCAMBIA3330 carrying a genomic fragment, including 2.718-kb 5′-flanking sequence, 0.756-kb open reading frame, and 1.028-kb 3′-downstream sequence of PALM1 as described (Chen et al., 2010).

Gas Chromatography–Mass Spectrometry Analysis of Cuticular Wax Content

Total leaf cuticular wax extraction and analysis was conducted as described previously (Zhang et al., 2005). Leaf samples were collected from one leaflet of the top two expanded trifoliates/pentafoliates excised from the major stems of well-watered wild-type R108 and irg1 mutant alleles. The two leaflets from the same plant were combined as one leaf sample to represent one biological replicate. Each sample was added to 10 mL of gas chromatography–mass spectrometry (GC-MS)–grade hexane (Sigma-Aldrich). Tissues were agitated for 2 min, and the solvent was decanted into new glass tubes. The same amount of hexane was used to rinse the tissues and tubes for 10 s and was pooled to the sample tube. Hexane was evaporated to ~1 mL under a nitrogen stream, transferred to 2-mL autosampler vials, and then evaporated completely. The abaxial and adaxial leaf waxes were extracted as described (Gniwotta et al., 2005). The polymer films from two leaflets were pooled to represent one biological replicate, extracted with hexane, and evaporated to 1 mL as described above for the whole leaves. Six replicates for the whole plant leaf extracts represent six biological replicates from plants grown in different pots from the same experiment. The abaxial or adaxial leaf waxes represent the average of four replicates biological replicates from plants grown in different pots from the same experiment. These experiments were repeated three independent times, and data from a representative experiment are presented.

The dried extracts were derivatized using N-methyl-N-trimethylsilyl trifluoroacetamide (+1% trimethylchlorosilane; Pierce Biotechnology) and were run on an Agilent 7890A gas chromatograph using a splitless injection as described (Zhang et al., 2005). Quantification was done using the area of the ions with mass-to-charge ratios of M-15, 117, 57, and 218 for fatty alcohols, fatty acids, alkanes, and sterols respectively. The amount of total wax composition and each cuticular wax constituent was expressed per unit of leaf area. Leaf areas were determined using a leaf area meter (LI-3000; Li-Cor). All values represent averages of four biological replicates ± sd.

Total Protein Isolation and Germination Inhibition Assay

Total protein was extracted from wild-type R108, irg1-1, and irg1-2 M. truncatula leaves. One gram of leaf tissue was homogenized in buffer (1:2 [w/v]) containing 50 mM Tris-Cl, pH 7.4, 100 mM NaCl, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride, 10 mM MgCl2, 1 mM EDTA, and complete miniprotease inhibitor cocktail according to the manufacturer’s instructions (Roche Diagnostics). Homogenates were centrifuged at 12,000g for 5 min at 4°C to remove debris, and protein concentrations in the homogenates were determined by the Bradford assay (Bio-Rad). One milliliter of the urediniospores (2 × 106 spores/mL) of P. emaculata and P. pachyrhizi were mixed with an equal volume of total protein isolated from M. truncatula wild type or irg1 mutants at a final concentration of 20 mg of protein/mL, then the urediniospores were incubated in 6-well cell plastic culture plates (BD Falcon) for 12 h under dark condition. After incubation, the numbers of P. emaculata and P. pachyrhizi spores that germinated and germ tubes with and without differentiated appressoria were counted from five random fields. The average is used to calculate the percentage of germination and spores with and without an appressorium.

Statistical Analysis

Significant differences (P ≤ 0.05) within the data sets were tested by one-way analysis of variance followed Duncan’s multiple range test using SAS 9.2 (SAS Institute). We fitted a general linear model to our factorial data to estimate the significance of both main effects and interactions for the cuticular wax content analyses performed using GC-MS (Figure 5). Similarly, the statistical significances in urediniospore differentiation were determined using analysis of variance followed by Duncan’s multiple range test (Figures 2 and and7).7). Student’s paired t test was used to analyze the significant differences between wild-type and mutant alleles or mock and wax treatments (Figures 1, ,6,6, and and88).

Microarray Analyses

To represent different alleles of the irg1 mutant, two independent sampling of three alleles, irg1-1, irg1-2, and irg1-5, totaling six biological replicates, were used in the microarray analyses in comparison with three biological replicates of wild-type plants. All plants were planted in the greenhouse in separate pots in a single experiment. Total RNA was purified from leaf tissues of 4-week-old M. truncatula R108 and irg1 mutants using TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. Three plants were used for each biological replicate. Total RNA was extracted from individual plants, and following isolation, was pooled to represent one biological replicate. The integrity of the RNA was checked on an Agilent Bioanalyzer 2100, and 10 μg of total RNA was used as a template for amplification. Probe labeling, chip hybridization, and scanning were performed according to the manufacturer’s instructions (Affymetrix). All the raw data were imported into Robust Multichip Average software and normalized as described (Irizarry et al., 2003). The presence/absence call for each probe set was obtained from dCHIP (Fiebig et al., 2000), and differentially expressed genes between sample pairs were selected using associative analyses, as described (Dozmorov and Centola, 2003). Genes that showed significant differences in transcript levels (twofold or greater and P value < 8.15954E-07) between the wild-type and mutant plants were selected for further analyses. Type I family-wise error rate was reduced using the Bonferroni-corrected P value threshold of 0.05/N, where N represents the number of genes present on the chip, which is 61,278 for M. truncatula. In this method, the background noise presented among replicates and technical noise during microarray experiments was measured by the residual presented among a group of genes whose residuals are homoscedastic. Genes whose residuals between the compared sample pairs that are significantly higher than the measured background noise level were considered to be differentially expressed. Due to the large gene number in the background measurement, this method enabled Bonferroni correction without being overly stringent. The data and complete MIAME information are deposited at ArrayExpress (http://www.ebi.ac.uk/microarray-as/ae/) under accession number E-MEXP-3190.

Real-Time Quantitative RT-PCR and RNA Gel Blot Analyses

Real-time qRT-PCR was performed as described (Uppalapati et al., 2009) using the gene-specific primers (see Supplemental Table 2 online). Total RNA was treated with Turbo DNase (Ambion) to eliminate genomic DNA, and 5 μg of DNase-treated RNA was reverse transcribed using Superscript III reverse transcriptase (Invitrogen) with oligo(dT)20 primers. The cDNA (1:10) was then used for qRT-PCR. The qRT-PCR was performed using Power SYBR Green PCR master mix (Applied Biosystems) in an optical 384-well plate with an ABI Prism 7900 HT sequence detection system (Applied Biosystems). Melt-curve analysis was performed to monitor primer-dimer formation and to check amplification of gene-specific products. The average threshold cycles (CT) values calculated from triplicate biological samples were used to determine the fold expression relative to the controls. Primers specific for ubiquitin were used to normalize small differences in template amounts. For RNA gel blot analysis, 10 μg of total RNA was denatured, fractionated by electrophoresis on a 1% agarose gel, and then blotted onto nylon membranes (Amersham Biosciences). Digoxigenin-labeled DNA probes were made with DIG PCR labeling mix (Boehringer Mannheim) using gene-specific forward and reverse primer sets described earlier (Uppalapati et al., 2009). Hybridization was performed at 65°C overnight in a solution containing 50 mM Church buffer (pH 7.2; Church and Gilbert 1984), 5× SSC (1× SSC is 0.15 M NaCl and 0.015 M sodium citrate), 0.1% laurylsarcosine, 7% SDS (w/v), and 2% blocking solution (Boehringer Mannheim). Further washes were done at 65°C in a solution containing 0.5× SSC and 0.1% SDS. Hybridized mRNAs were detected with anti-DIG antibody conjugated with alkaline phosphatase (Boehringer Mannheim) and its chemiluminescent substrate, CDP-Star (Boehringer Mannheim).

Accession Numbers

Sequence data from this article for M. truncatula genes can be found in the Dana Farber Cancer Institute Medicago gene index, J. Craig Venter Institute–Medicago Genome Resource, or GenBank/EMBL databases under the following TC numbers or locus number: CER1-1 (TC130292), CER2 (TC115187), CER3 (TC125004), CER4-1 (TC122585), CER4-2 (TC160800), CER6-1 (TC116151), CER6-2 (TC125487), CER6-3 (TC121408), CER6-4 (TC113573), CER8-1 (TC140235), CER10 (TC125186), PAS2 (GE348322), KCR1 (TC113588), KCR2 (TC145787), WSD1 (TC145247), FATB (TC148248), MYB30 (TC142663), MYB96 (Medtr4g135250/Medtr8g112130), IRG1/PALM1 (GI:298201167), and LTP (TC97568).

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure 1. Expression Profiles for Selected Genes in the Phenylpropanoid Pathway (PAL, CHS, CHR, CHI, IFS, and IFR) and Pathogenesis-Related Genes (PR3 and PR10) in the Wild Type (R108) at 0, 8, 24, and 48 h after Inoculation with P. emaculata Urediniospores.
  • Supplemental Figure 2. A Schematic Showing the Sequence of Events for Conducting the Forward Genetic Screens to Identify M. truncatula Genes Involved in Nonhost Resistance against P. emaculata.
  • Supplemental Figure 3. Expression Profiles for Selected Defense-Related Genes and Antimicrobial Activity of Protein Extracts on Fungal Differentiation.
  • Supplemental Figure 4. Rust Germ Tube Differentiation on Multiple Mutant Alleles of irg1.
  • Supplemental Figure 5. Leaf Phenotype of 4-Week-Old Wild-Type R108 and irg1-1 Plants of M. truncatula.
  • Supplemental Figure 6. A Simplified Wax Biosynthesis Pathway and Some CER Genes Implicated in Wax Biosynthesis in Arabidopsis (Modified from Kunst and Samuels, 2003; Samuels et al., 2008).
  • Supplemental Figure 7. Composition of Alcohols and Alkanes in Wild-Type and irg1 Mutant Alleles of Medicago truncatula.
  • Supplemental Table 1. Fold Changes in Expression of Corresponding M. truncatula Orthologs of Arabidopsis Wax Biosynthesis-Related Genes Using qRT-PCR and Microarray Analysis.
  • Supplemental Table 2. List of Genes and Corresponding Primers Used for qRT-PCR.
  • Supplemental Data Set 1. A List of Differentially Regulated Genes (Minimum Twofold Up- or Downregulated) in Three Different Alleles of irg1 (irg1-1/palm1-4, irg1-2 /palm1-5, and irg1-5/palm1-6).


We thank Markus Frank for helpful discussions, Yongfeng Zhang for assistance with mutant screening, and Kelly Craven, Clemencia Rojas, and Jackie Kelley for critical reading of the manuscript. We also thank Azhaguvel Perumal and Muhammet Sakiroglu for guidance with statistical analyses and Janie Gallaway and Colleen Elles for plant maintenance in the greenhouse. This work was supported through a grant to K.S.M. and S.R.U. from BASF Plant Sciences, Germany, and National Science Foundation--Experimental Program to Stimulate Competitive Research (EPS-0814361) to K.S.M. The insertion mutant lines of M. truncatula were created through research support, in part, from the National Science Foundation (DBI-0703285).


S.R.U. identified the mutant, designed research, analyzed data, and wrote the article. Y.I. designed research, performed research, and analyzed data. K.S.M. designed research and wrote the article. S.R.U. and Y.I. performed the bulk of the experiments, and V.D. and S.M. helped them with genetic screens. J.N. helped with scanning electron microscopy. M.B. performed wax analysis using GC-MS. Y.T. analyzed microarray data. J.C., M.T., P.R., R.C., and H.S. contributed reagents and provided seed stocks for Tnt1 mutant populations or PALM1 complemented lines.


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