Rapid In Vitro Screening of Prunus Genotypes for Resistance to Armillaria Root Rot Using Roots of Young Rootstocks

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Pratima Devkota Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI 48824, USA

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Amy Iezzoni Department of Horticulture, Michigan State University, East Lansing, MI 48824, USA

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Ksenija Gasic Department of Plant and Environmental Sciences, Clemson University, Clemson, SC 29634, USA

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Gregory Reighard Department of Plant and Environmental Sciences, Clemson University, Clemson, SC 29634, USA

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Raymond Hammerschmidt Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI 48824, USA

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Abstract

Armillaria root rot (ARR), caused by Armillaria species and Desarmillaria tabescens, is a severe disease that affects stone fruit trees in the United States. One strategy to mitigate the impact of this disease is to develop ARR-resistant rootstocks. However, current techniques to screen Prunus species for resistance to ARR are time-consuming, labor-intensive, and may not fully replicate field conditions. To address these limitations, we developed a new rapid in vitro screening assay, which uses roots of 2-year-old Prunus rootstock genotypes. We screened 12 Prunus genotypes against Armillaria mellea, Armillaria solidipes, and Desarmillaria tabescens in vitro. Freshly excavated root segments were placed next to or on top of fungal cultures. After 21 days, the circumferential percentage and horizontal length of the fungal colonization and the ability of the fungus to enter through root periderm were evaluated. The root tissue surrounding the infection was also evaluated to assess any response reactions against the ARR pathogens. Our results showed that inoculated root tissues displayed signs of fungal infection, and infection and host responses varied among the Prunus genotypes. Host responses similar to those observed in the field, such as compartmentalization of infected tissue with barrier zones, necrophylactic periderm formation, and callus formation on root surfaces, were observed and were more evident in less susceptible genotypes. In conclusion, our newly developed assay, which uses freshly excavated roots from 2-year-old rootstocks, can rapidly screen Prunus genotypes for resistance to ARR.

Armillaria species cause ARR disease in more than 500 woody hosts, including stone fruit trees (Raabe 2008). In Michigan’s cherry and peach orchards, this disease is mainly caused by A. solidipes (synonym A. ostoyae) and, to a lesser extent, by A. mellea (Proffer et al. 1987). On the other hand, in the major peach-growing regions in the southern United States, D. tabescens (Scop.) R.A. Koch & Aime comb. nov. (synonyms Armillaria tabescens and Clitocybe tabescens) is primarily responsible for this disease (Koch et al. 2017; Schnabel et al. 2005). In California’s almond orchards, A. mellea is the causal ARR fungus (Baumgartner and Rizzo 2002). Armillaria spp. spread in the orchard floor through thread-like structures called rhizomorphs, or healthy root contact with fungal mycelia in infected tissues. These fungi infect, kill, and decompose tree roots and continue to thrive in the infected roots and stumps for years as saprophytes. The infected orchards suffer from yield losses, resulting in significant economic damage (Michigan Tree Fruit Commission, personal communication). If the level of Armillaria spp. infection is high, the infected orchards become unsuitable for replanting trees.

Resistant rootstocks are a time-tested approach for management of root pests and diseases in perennial crops [e.g., root-knot nematode (Saucet et al. 2016) and Phytophthora root rot (Smith et al. 2011)]. Various field trials have been conducted by several researchers to test susceptibility of different Prunus species and genotypes to different Armillaria species. Proffer et al. (1988) tested Mahaleb (cherry), Mazzard (cherry), and 17 hybrid Prunus rootstocks and results suggested that Mazzard rootstock was less susceptible to ARR than Mahaleb. Beckman and Pusey (2001) conducted a 4-year field trial in Byron, GA, and reported ‘SL0040’ (a complex plum hybrid) was less susceptible to D. tabescens compared with ‘Marianna 2624’ (plum × plum), ‘Lovell’ (peach), ‘Ishtara’ [Myrobalan plum × (Myrobalan plum × peach)], and ‘Myran’ (Myrobalan × peach). Similarly, a 10-year field trial of various ungrafted Prunus rootstocks against A. mellea in France also reported peach and peach hybrids to be highly susceptible (Guillaumin et al. 1991). However, rootstocks with plum background, Myrobalan, ‘Ishtara’, and ‘Myran’ showed tolerance to ARR, suggesting that resistance in Myrobalan is a dominant trait. Limited growth of the pathogen was observed in plums as well as defense reactions in the bark and sapwood. The variation in results between the French and the US trials could be attributed to differences in the Armillaria species tested or variations in the duration of the experiments.

Artificial inoculation assays have become fundamental tools for identifying sources of disease resistance and screening progeny from crosses. Nonetheless, extant Armillaria screening assays pose challenges, being arduous, time-intensive, and lacking reproducibility. Field inoculation methods involve a lengthy process that begins with establishing host plants in the field, which can take several years. Woody inoculum preparation alone can take 3 to 6 months. Subsequently, plants are inoculated, and the progression of the disease is assessed over several years (Beckman and Pusey 2001; Mansilla et al. 2001; Proffer et al. 1988). Achieving successful field inoculation poses challenges because of the wide variability in environmental conditions and the issue of inoculum drying. Similarly, greenhouse inoculation, which involves a comparable 3 to 6 months’ inoculum preparation period, may not always result in successful infection. Obtaining conclusive results from greenhouse assays may take several months to years of observation (Elias-Roman et al. 2019; Singh 1980).

In vitro assays using tissue-cultured plant material are available, offering the advantage of speed (Adelberg et al. 2021; Baumgartner et al. 2018). Nonetheless, when screening tissue-cultured plant material against pathogens in vitro, it may not properly emulate the host defense response and tissue infection as observed in realistic field conditions (Fenning 2019; Tripathi et al. 2008). Notably, root and shoot tissues of tissue-cultured plants in vitro may lack secondary thickening. Secondary growth in roots and stems plays a crucial role in defense mechanisms against biotic and abiotic stressors, wound healing, barrier formation, chemical defense, compartmentalization, and providing structural support (Du and Groover 2010). The periderm, composed of tissues of secondary origin such as phelloderm, cork cambium, and phellem, acts as the outermost protective barrier for roots, shielding the inner tissues from pathogens (Biggs 1992). Studying roots that have undergone secondary growth and possess periderm could enhance our understanding of host defenses. Thus, it is important to use roots with periderm in ARR screening assays.

Recently, a new in vitro screening assay for ARR was developed, using freshly excavated and detached segments of roots from the field that had undergone secondary growth (Devkota and Hammerschmidt 2019). This technique mimics host root infection and defense under realistic field conditions. Moreover, the assay time for this method is reduced to 4 to 6 weeks; however, a primary limitation of this assay, particularly when screening numerous progenies from directed crosses, is the requirement to excavate roots from field trees, which can take a minimum of 2 to 4 years to establish, and the use of detached roots for inoculation.

Screening sources of resistant germplasm is an important first step to ensure that a breeding program uses a genetically diverse set of resistant germplasm. In this regard, conducting a rapid Armillaria screening assay using freshly excavated roots from Prunus rootstocks in their second to third year of growth could significantly expedite the screening process. Thus, the present study aims to use this in vitro screening approach to evaluate the response of 12 Prunus genotypes in the first few years to A. solidipes, A. mellea, and D. tabescens, and to determine their active and passive responses to Armillaria spp. infection. Specifically, this study seeks to characterize the active and induced responses of healthy root tissues of various Prunus genotypes to Armillaria spp. The selection of rootstock genotypes for ARR screening is based on the needs of nursery partners and Michigan State University for screening newly developed Prunus hybrids to ARR. Prunus genotypes such as Prunus cerasifera, ‘Krymsk® 86’, ‘Bright’s Hybrid® 5’, and ‘Hansen 536’ that have been screened for ARR in other studies will be screened in addition to seven other new genotypes. P. cerasifera and ‘Krymsk® 86’ have exhibited tolerance to ARR, and ‘Bright’s Hybrid® 5’ and ‘Hansen 536’ displayed higher susceptibility (Baumgartner et al. 2018; Devkota et al. 2020). Although these findings can serve as references for comparison in this study, it is important to consider that the virulence of different fungal isolates may vary across geographical regions. Therefore, the Armillaria species isolated from Michigan and the southern United States will be used in this study.

Materials and Methods

A total of 12 Prunus genotype rootstocks including nine genotypes provided by Fowler Nursery (Newcastle, CA), two Prunus genotypes provided by Michigan State University, and one P. cerasifera (Myrobalan 29c) obtained from Burnt Ridge Nursery (Onalaska, WA) were screened against single isolates of A. solidipes, A. mellea, and D. tabescens (Table 1). The Prunus rootstocks were initially propagated from softwood cuttings at the nurseries. Subsequently, they were transplanted into 1.5-gallon pots that were filled with Suremix Perlite Mix (Michigan Grower Products, Inc.TM, Galesburg, MI). This transplantation took place at the greenhouse of Michigan State University, where the temperature was maintained between 18 °C and 24 °C, and the humidity levels were kept at 65% to 75%. The rootstocks were then nurtured and grown for a duration of 1 year and 6 months.

Table 1.

Prunus genotypes and their parentage used for the in vitro inoculation assay.

Table 1.

P. cerasifera was used as a Prunus control with lower ARR susceptibility. The lineage of each Prunus genotype can be found in Table 1. Fungal isolates, A. mellea (SC.00i49) and D. tabescens (SC.GJ2.02), were isolated from peach trees with ARR symptoms from Spartanburg County, SC, and Cherokee County, SC, respectively. The A. solidipes (Warren) isolate was obtained from a tart cherry tree with ARR symptoms from Macomb County, MI. Prunus roots were excavated and cut from two to three clones of Prunus rootstocks (∼2-year-old) of those genotypes (Fig. 1A and B). Roots were gently washed with running water to remove external soil, and surface sterilized by immersing in 70% ethanol for 1 minute, washed with sterile distilled water, and blot-dried. Surface sterilized root segments were used for in vitro rapid assay (Devkota et al. 2020).

Fig. 1.
Fig. 1.

Fungal root inoculation assay. (A) Prunus species seedling growing in the greenhouse. (B) Root excavated and segmented for inoculation. (C) Fungal culture for wounded root inoculation assay. (D) Open end of root being placed in fungal culture. (E) Inoculated roots being examined. (F) Fungal culture for intact root inoculation assay. (G) Roots with intact bark being inoculated on top of fungal culture. (H) Evaluation of roots at 21 d following inoculation.

Citation: HortScience 58, 10; 10.21273/HORTSCI17196-23

Dicloran-Benomyl-Streptomycin amended Malt extract agar (DBSMEA) was used to avoid cross-contamination due to root surface microbes. DBSMEA was prepared according to Worrall (1991), and media slants were prepared in Magenta™ GA-7 Plant Culture Boxes. Four 5-mm mycelial plugs of each Armillaria spp. were placed on the top of the end of the thickest side of the slant (Fig. 1C) and culture boxes were incubated at 22 °C in the dark for 14 d. Roots of ∼5 to 10-mm diameter were cut into ∼6 cm long segments, and one end of the segment was sealed with paraffin wax. The exposed end was placed next to the culture of Armillaria spp. and D. tabescens and incubated at 22 °C in the dark (Fig. 1D). There were four samples of each Prunus spp. and the study was replicated twice. Root segments kept in culture boxes without fungal cultures were used as noninoculated controls. At 21 d after the inoculation, the periderm of the roots was peeled back to reveal any successful fungal penetration and the extent of colonization (Fig. 1E). Observations were made at 21 d, as new periderm formation occurs in 2 to 3 weeks post infection or injury (Biggs 1986; Devkota et al. 2020). Five millimeters of tissue around the inoculated side plus surrounding tissue (10 mm thick) of three roots were cut with a sharp, sterile clipper and immediately placed in formalin-acetic alcohol (FAA; formaldehyde:acetic acid: 70% ethanol, 5:5:90, v/v/v) fixative. After 48 h, FAA was decanted, and the samples were processed with ethanol:xylene series and embedded in melted paraffin wax (54 to 55 °C) overnight. A rotary microtome (model RM2245; Leica Biosystems, Deerfield, IL, USA) was used to cut sections. Four 6-µm-thick sections were prepared per root sample. The sections were mounted on positively charged glass slides and stained with 0.5% aqueous Toluidine blue to visualize fungal hyphae and host structures in the root tissue. Histochemical tests to visualize lignification and suberization of the infected tissue were used as described by Cleary et al. (2012a). The fungal infection and active host responses in the sections were visualized using a light microscope (DM2500, Leica). Active host responses such as ligno-suberized barrier zone formation, necrophylactic periderm formation around the infection point, compartmentalization of infected tissue, the activity of cambium, and callus formation on the root surface were assessed.

Necrophylactic periderm (np) formation was rated according to a 0 to 2 scale: 0 = no np formation and successful infection, 1 = formation of np that partially blocked fungal infection, and 2 = formation of strong necrophylactic periderm that completely blocked fungal infection. Barrier zone formation with or without lignified and/or ligno-suberized barrier zone was rated using a 0 to 1 scale where 0 = no barrier zone formation and 1 = barrier zone formation. The activity of the cambium was rated accordingly in a 0 to 1 scale: 0 = cambium was dead and inactive and 1 = cambium was still active and forming a callus. Similarly, the formation of callus on the root surface was also rated according to a scale of 0 to 2: 0 = no callus formation, 1 = formation of callus initials, formation, 2 = formation of distinct callus masses and initiation of new root formation.

To evaluate the ability of the intact periderm of different Prunus genotypes to resist fungal invasion surface sterilized, roots of ∼5 to 10 mm diameter were cut into 3-cm segments and sealed securely at both ends with paraffin wax. Three root segments from each Prunus genotype were placed on top of DBSMEA agar (100 mm × 15 mm) alongside Armillaria cultures and rhizomorphs (Fig. 1F and G). The study was replicated twice. Root segments placed in petri plates with media and without fungal cultures were used as noninoculated controls. At 21 d, the periderm of the roots was carefully peeled back to reveal successful infection and the extent of colonization by fungi. Furthermore, root segments were sectioned to visualize the fungal infection and host defense responses (Fig. 1H).

Based on various parameters related to host infection and response, a five-scale ARR susceptibility index was developed, categorizing genotypes as resistant, less susceptible, moderately susceptible, susceptible, and highly susceptible. The major variable used to determine relative susceptibility was the circumferential percentage of fungal infection (Shigo and Tippett 1981). Genotypes were considered resistant when no fungal infection was observed. Genotypes were considered less susceptible when the circumferential infection was limited to <30% accompanied by a low level of fungal spread and infection with robust host responses. Genotypes were considered moderately susceptible to ARR when the circumferential infection ranged from 30% to 60%. Genotypes were considered susceptible when the circumferential infection ranged from 60% to 80%. Genotypes with a circumferential infection exceeding 80% were classified as highly susceptible.

The general linear model was used to analyze the data (PROC GLM, SAS 9.4). Assumptions of normality and homogeneity of variance were tested for all the response variables. Tukey’s honestly significant differences were calculated to determine the statistical differences of response variables among the Prunus genotypes at α = 0.05.

Results

Fresh root segment inoculation allowed successful inoculation and assessment of pathogen development and host response at 21 d post-inoculation. White mycelial fans of Armillaria spp. and D. tabescens were observed underneath the root periderm, spreading both longitudinally and circumferentially. The spread of fungal hyphae was followed by the development of tissue wetness and necrosis. Prunus genotype × fungal interactions were significant [F(22, 251) = 14.14, P ≤ 0.0001], suggesting the length of the spread of each fungus varied among Prunus genotypes. The longitudinal length of fungal growth differed among all genotypes [F(11, 251) = 115.03, P ≤ 0.00001; Table 2]. Fungal growth length varied among the fungus [F(2, 251) = 50.55, P ≤ 0.00001], with A. mellea having the highest growth. A shorter length of fungal growth was observed in P. cerasifera, ‘BB 106®’, ‘Krymsk® 86’, and ‘Controller™ 9’, whereas there was a significantly greater length of fungal growth in ‘Corette® 5’ and ‘Benzie’. Growth of A. mellea was significantly higher than that of A. solidipes and D. tabescens in genotypes ‘Hansen 536’, ‘FN1-23’, and ‘Corette® 5’.

Table 2.

Least square means of the length of fungal penetration (mm) underneath the root periderm of Prunus genotypes.

Table 2.

In addition to the longitudinal spread, fungal hyphae spread around the root circumference underneath the periderm. Tissue necrosis was present in advance of the presence of fungal hyphae. Host response reactions were deterring the fungal spread in a circumferential fashion. Prunus genotype × fungal interactions were significant [F(22, 251) = 9.41, P ≤ 0.0001]. The circumferential percentage of fungal spread varied significantly among the Prunus genotypes [F(11, 251) = 219.53, P ≤ 0.0001; Table 3]. P. cerasifera and ‘Krymsk® 86’ had the lowest circumferential percentage of infection, and fungal hyphae encircled the full circumference of ‘Corette® 5’ and ‘Benzie’ periderms. Other Prunus genotypes had 30% to 60% of their root circumference covered by fungus. Three fungi differed in their ability to spread circumferentially [F(2, 251) = 40.66, P ≤ 0.0001]. A. mellea had a high ability to spread circumferentially compared with A. solidipes and D. tabescens. The circumferential spread of A. mellea was higher in ‘Hansen 536’ and ‘Bright’s Hybrid® 5’ compared with D. tabescens.

Table 3.

Least square means of circumferential percentage of fungal infection in the root of Prunus genotypes.

Table 3.

The fungal penetration varied among the intact root periderm of different Prunus genotypes [F(11, 186) = 10.46, P ≤ 0.0001; Fig. 2]; however, the three fungi did not differ in their ability to penetrate [F(2, 186) = 0.86, P = 0.42412]. There was no genotype × fungal interaction [F(22, 186) = 1.10, P = 0.34589]. All three fungal species were least successful in penetrating the intact periderm of P. cerasifera and ‘Krymsk® 86’ and most successful in penetrating the intact periderm of ‘Corette® 5’, ‘Benzie’, and ‘FN1-23’.

Fig. 2.
Fig. 2.

Frequency of success of fungal penetration through intact bark of various Prunus genotypes.

Citation: HortScience 58, 10; 10.21273/HORTSCI17196-23

None of the tested genotypes showed resistance to ARR. Overall, P. cerasifera and ‘Krymsk® 86’ were the less susceptible genotypes. ‘Controller™ 9’, ‘BB 106®’, and ‘Krymsk® 7’ were moderately susceptible. ‘Controller™ 6’, ‘Hansen 536’, ‘Bright’s Hybrid® 5’, and ‘FN1-23’ were susceptible genotypes. ‘Corette® 5’ and its open-pollinated offspring ‘Benzie’ were highly susceptible genotypes. The success of fungal penetration of various genotypes correlated with the rating but was not considered a significant variable because the ability of the fungus to spread in root circumference largely determines ARR susceptibility. Some genotypes showed slight differences in susceptibility among the three fungi (Table 3). For example, a genotype highly susceptible to A. mellea may be moderately susceptible to A. solidipes.

Nonspecific host defense mechanisms were triggered in most of the inoculated root segments (Figs. 35). Response reactions were also noted in the wounded noninoculated controls; however, these reactions were observed at minimal levels. On the other hand, no response reactions were detected in the intact root inoculated controls. The essential feature of responses of root tissues to Armillaria infection includes forming ligno-suberized barrier zones from the cells previously present, followed by the development of phellogen cells internally and differentiation of or the initiation of formation of necrophylactic periderm around the infected tissue. There was a high incidence of barrier zone formation around the infected tissue in the less susceptible genotypes, such as P. cerasifera and ‘BB 106®’ (Fig. 3D–F). Those barrier zones were made up of lignified cells and or ligno-suberized cells. Necrophylactic periderm in less susceptible genotypes like P. cerasifera was nearly always complete in most cases, and breaching was less frequently observed. In highly susceptible genotypes such as ‘Corette® 5’ and ‘Benzie’, fungi were successfully established in the roots as the host defenses were either absent or low (Fig. 4). Cambial tissues were fully or partly healthy, with actively producing calli in the least susceptible and moderately susceptible genotypes, respectively. Cambial tissue was either dead or inactive in highly susceptible genotypes, ‘Corette® 5’ and ‘Benzie’. Furthermore, another nonspecific induced defense was callus formation on the root surface as well in some genotypes. On the root surface of less susceptible genotypes, a high level of callus formation was observed. The callus was initiating to form new roots in some cases. Overall anatomical and morphological changes associated with fungal treatments in all Prunus genotypes are summarized in Table 4.

Fig. 3.
Fig. 3.

(A) Longitudinal section of Prunus cerasifera inoculated with Armillaria mellea showing barrier formation. (B) Walling off the infected region with the barrier zone and necrophylactic periderm formation in P. cerasifera. (C) Root of intact ‘Krymsk® 86’ root segment with no sign of fungal invasion without any visible host defense reaction. (D) Formation of barrier zone around the infection point in Prunus genotype ‘BB 106®’. (E) Lignified cells in the barrier zone. (F) Suberized cells in the barrier zone. Bars = 500 µm. White arrows show a fungal infection, and the blue arrows show defense responses.

Citation: HortScience 58, 10; 10.21273/HORTSCI17196-23

Fig. 4.
Fig. 4.

Cross and longitudinal sections of various Prunus species infected with Armillaria spp. (A) Desarmillaria tabescens successfully penetrating in ‘Corette® 5’ (Bars = 200 µm). (B) Armillaria solidipes infecting ‘Corette® 5’ without any visible host defense. (C) Longitudinal section of ‘Controller™ 6’ successfully infected with Armillaria mellea. (D) ‘Controller™ 6’ responding to A. mellea infection. (E) Absence of any barrier zone in phloroglucinol-HCl stained section of ‘Controller™ 6’. (F) A. mellea successfully penetrating ‘Benzie’. Bars = 500 µm. White arrows show fungal infection, and blue arrows indicate defense responses.

Citation: HortScience 58, 10; 10.21273/HORTSCI17196-23

Fig. 5.
Fig. 5.

(A–C) Transverse section of Prunus cerasifera intact root inoculated with Armillaria solidipes. (A) No fungal penetration through intact periderm (Bar = 500 µm). (B) Formation of barrier around fungal infection point (Bar = 500 µm). (C) Cambium is active in the inoculated root (Bar = 200 µm). (D and E) ‘Krymsk® 7’ root with intact periderm inoculated with A. solidipes (Bars = 200 µm). (D) Initiation of formation of necrophylactic periderm. (E) Fungal penetration underneath the bark and barrier being formed to stop the spread. (F) ‘Benzie’ inoculated with A. mellea without distinct host defense response (Bar = 500 µm). White arrows indicate point where root periderm touched fungal inoculum, and blue arrows indicate host defense responses.

Citation: HortScience 58, 10; 10.21273/HORTSCI17196-23

Table 4.

Overall Prunus host response inoculated with Armillaria species and Desarmillaria tabescens.

Table 4.

Discussion

In this study, we successfully infected freshly detached woody roots of Prunus genotypes with Armillaria spp. and D. tabescens growing in solid medium. Inoculation resulted in fungal infection and active host responses that mimic field conditions. Fungal hyphae spread longitudinally and circumferentially underneath the bark in the roots. The extent of fungal spread and infection varied among the roots of Prunus genotypes indicating variation in ARR susceptibility. Although this is the first study to test the ARR susceptibility of most of these genotypes, susceptibility findings are consistent for those previously tested in other studies. Past field trials and screening studies have also shown that P. cerasifera (aka Myrobalan plum) and ‘Krymsk® 86’ were less susceptible to A. mellea and D. tabescens (Baumgartner et al. 2018; Devkota et al. 2020; Guillaumin et al. 1991), which is consistent with our finding. Our observation that ‘Hansen 536’ is susceptible to A. mellea is consistent with the findings of Baumgartner et al. (2018).

This study provides insight into disease resistance in Prunus spp., as the induced response following the fungal infection in the root periderm aids in understanding ARR resistance. Nonspecific anatomical responses, including barrier zone formation, necrophylactic periderm formation to compartmentalize infected tissue, the activity of cambium, and callus tissue formation in bark surface parallel those described elsewhere (Aslam and Magel 2018; Cleary et al. 2012b; Mullick 1977; Robinson et al. 2004; Solla et al. 2002). Wounding and fungal infection triggered necrophylactic periderm formation as one of the evident host response reactions. Breaching of this new barrier was only rarely observed in less susceptible genotypes. P. cerasifera made a thinner layer of necrophylactic periderm but was less frequently breached by the invading pathogen. It is likely that the presence of a high level of antifungal compounds in the bark and/or the characteristics and constituents of different components of the root cell wall may have played a role. The presence of antifungal compounds in the periderm may give more time for the roots to form anatomical antifungal barriers. Several studies suggest that a complex range of phenolic compounds in periderm has antifungal activity (Stange et al. 2001; Treutter 2006). In the case of the susceptible genotypes, ‘Corette® 5’ and ‘Benzie’, any barrier zones or new periderms initiating to form were easily breached by inoculated fungus. In these genotypes, Armillaria spp. advanced to the deeper bark tissues and killed the cambium.

Our assay to screen Prunus genotypes to ARR aligns with the popular concept of compartmentalization of the decay of trees. Shigo and Tippett (1981) studied root systems of several trees from different species that appeared from suppressed and dying to dominant and healthy because of Armillaria spp. infection and concluded that host compartmentalization of decay was the strategy to survive the fungal infection. Compartmentalization may occur in the xylem and bark. Effective compartmentalization is thought to ensure the survival of the tree as it tolerates fungal infection. Significant loss or death of trees occurs when the trees cannot effectively compartmentalize the infected tissue. Tree roots respond to wall off the spread of the fungus as long as some cambium remains alive (Shigo and Tippett 1981). If the living circumference of the root collar decreases, then any additional infection may lead to tree death. In the present assay, the measured circumferential spread of fungal infection in the periderm provided an idea about the ability of each genotype to respond rapidly to ARR-causing pathogens.

For Armillaria spp. to survive, it must infect when the host defenses are weakest and spread as far as possible before recognition and compartmentalization by the host. We observed the lowest level of host defense in Prunus genotypes, such as ‘Corette® 5’ and ‘Benzie’, with a high level of fungal spread. For the host tree to survive, it must recognize the fungus and restrict it to a small volume of the bark and wood as rapidly as possible. To restrict infection, the host must generate new root tissues. In the roots of susceptible genotypes, the fungus begins to spread into the wood beneath the bark and the cambium. The woody tissue beneath the cambium is mostly secondary xylem, which is already dead and has limited biochemical mechanisms for defense. If the bark and cambium cannot form effective barriers, the fungus can easily invade the secondary xylem.

Formation of a high level of callus in the wounded root in the less and moderately susceptible genotypes suggests that the callus may have contributed to reducing the severity of the disease. On the other hand, the absence of proper callus formation in susceptible Prunus genotypes and their failure to achieve wound closure indicates that fungi can effectively establish themselves in the roots of these species under field conditions. Moreover, there was a notable abundance of callus formation on the root surface of less and moderately susceptible genotypes. Interestingly, in some cases, this callus initiated the growth of new roots. This phenomenon of new root formation could potentially aid these genotypes in compensating for any root death caused by the disease, as it allows them to produce fresh and healthy roots.

In this study, we observed less variability in the root samples within the same genotype because the genotypes were propagated from softwood cuttings. Softwood cuttings are a common method of horticultural propagation to produce plants that are genetically identical to the parent plant. These cuttings are taken from actively growing plants and are rooted in a growing medium, providing necessary nutrients and water for strong and healthy roots. Consequently, softwood cuttings typically produce more mature roots, with less variability compared with seed propagation or tissue culture. Although tissue culture has the advantage of rapidly producing a substantial number of genetically identical plants, it may lead to underdeveloped root systems due to the artificial growth conditions. Using roots developed from softwood or hardwood cuttings may provide reliable results during rapid in vitro screening. In conclusion, our study revealed varying degrees of susceptibility to Armillaria spp. infection among the roots of 12 Prunus genotypes, highlighting the usefulness and efficiency of this in vitro inoculation technique in screening additional genotypes for resistance to ARR.

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  • Biggs AR. 1992 Anatomical and physiological responses of bark tissues to mechanical injury, p 13–26. In: Blanchette RA, Biggs AR (eds). Defense mechanisms of woody plants against fungi. Springer Series in Wood Science, Springer Berlin, Heidelberg.

  • Cleary MR, Van Der Kamp BJ, Morrison DJ. 2012a. Effects of wounding and fungal infection with Armillaria ostoyae in three conifer species. II. Host response to the pathogen. For Pathol. 42(2):109123. https://doi.org/10.1111/j.1439-0329.2011.00727.x.

    • Search Google Scholar
    • Export Citation
  • Cleary MR, van der Kamp BJ, Morrison DJ. 2012b. Effects of wounding and fungal infection with Armillaria ostoyae in three conifer species. I. Host response to abiotic wounding in non‐infected roots. For Pathol. 42(2):100108. https://doi.org/10.1111/j.1439-0329.2011.00726.x.

    • Search Google Scholar
    • Export Citation
  • Devkota P, Hammerschmidt R. 2019. A rapid and holistic approach to screen susceptibility of Prunus species to Armillaria root rot. For Pathol. 49(5):e12547. https://doi.org/10.1111/efp.12547.

    • Search Google Scholar
    • Export Citation
  • Devkota P, Iezzoni A, Gasic K, Reighard G, Hammerschmidt R. 2020. Evaluation of the susceptibility of Prunus rootstock genotypes to Armillaria and Desarmillaria species. Eur J Plant Pathol. 158(1):177193. https://doi.org/10.1007/s10658-020-02065-y.

    • Search Google Scholar
    • Export Citation
  • Du J, Groover A. 2010. Transcriptional regulation of secondary growth and wood formation. J Integr Plant Biol. 52(1):1727. https://onlinelibrary.wiley.com/doi/10.1111/j.1744-7909.2010.00901.x.

    • Search Google Scholar
    • Export Citation
  • Elias-Roman RD, Calderon-Zavala G, Guzman-Mendoza R, Vallejo-Perez MR, Klopfenstein NB, Mora-Aguilera JA. 2019. ‘Mondragon’: A clonal plum rootstock to enhance management of Armillaria root disease in peach orchards of Mexico. Crop Prot. 121:8995. https://doi.org/10.1016/j.cropro.2019.03.011.

    • Search Google Scholar
    • Export Citation
  • Fenning TM. 2019. The use of tissue culture and in-vitro approaches for the study of tree diseases. Plant Cell Tissue Organ Cult. 136(3):415430. https://doi.org/10.1007/s11240-018-01531-0.

    • Search Google Scholar
    • Export Citation
  • Guillaumin JJ, Pierson J, Grassely C. 1991. The susceptibility to Armillaria mellea of different Prunus species used as stone fruit rootstocks. Scientia Hortic. 46(1-2):4354. https://doi.org/10.1016/0304-4238(91)90091-C.

    • Search Google Scholar
    • Export Citation
  • Koch RA, Wilson AW, Séné O, Henkel TW, Aime MC. 2017. Resolved phylogeny and biogeography of the root pathogen Armillaria and its gasteroid relative, Guyanagaster. BMC Evol Biol. 17:133. https://doi.org/10.1186/s12862-017-0877-3.

    • Search Google Scholar
    • Export Citation
  • Mansilla JP, Aguín O, Sainz MJ. 2001. A fast method for production of Armillaria inoculum. Mycol. 93(3):612615. https://doi.org/10.1080/00275514.2001.12063191.

    • Search Google Scholar
    • Export Citation
  • Mullick DB. 1977. The non-specific nature of defense in bark and wood during wounding, insect and pathogen attack, p 395–441. In: Loewus FA, Runeckles VC (eds). The structure, biosynthesis, and degradation of wood. Springer, Boston, MA. https://doi.org/10.1007/978-1-4615-8873-3_10.

  • Proffer TJ, Jones AL, Perry RL. 1988. Testing of cherry rootstocks for resistance to infection by species of Armillaria. Plant Dis. 72(6):488490. https://doi.org/10.1094/PD-72-0488.

    • Search Google Scholar
    • Export Citation
  • Proffer TJ, Jones AL, Ehret GR. 1987. Biological species of Armillaria isolated from sour cherry orchards in Michigan. Phytopathology. 77(6):941943. https://doi.org/10.1094/Phyto-77-941.

    • Search Google Scholar
    • Export Citation
  • Raabe RD. 2008. Plants resistant or susceptible to Armillaria mellea, the oak root fungus. University of California, Berkeley. https://alamedabackyardgrowers.org/wp-content/uploads/2019/01/List-of-Oak-Root-Fungus-Resistant-or-Susceptible-Trees-Plants.pdf. [accessed 28 Jul 2023].

  • Robinson RM, Morrison DJ, Jensen GD. 2004. Necrophylactic periderm formation in the roots of western larch and Douglas‐fir trees infected with Armillaria ostoyae. II. The response to the pathogen. For Pathol. 34(2):119129. https://doi.org/10.1111/j.1439-0329.2004.00354.x.

    • Search Google Scholar
    • Export Citation
  • Saucet SB, Van Ghelder C, Abad P, Duval H, Esmenjaud D. 2016. Resistance to root‐knot nematodes Meloidogyne spp. in woody plants. New Phytol. 211(1):4156. https://doi.org/10.1111/nph.13933.

    • Search Google Scholar
    • Export Citation
  • Schnabel G, Ash JS, Bryson PK. 2005. Identification and characterization of Armillaria tabescens from the southeastern United States. Mycol Res. 109(11):12081222. https://doi.org/10.1017/S0953756205003916.

    • Search Google Scholar
    • Export Citation
  • Shigo AL, Tippett JT. 1981. Compartmentalization of decayed wood associated with Armillaria mellea in several tree species. Broomall, PA, US Department of Agriculture, Forest Service, Northeastern Forest Experiment Station. 20:488. https://doi.org/10.2737/NE-RP-488.

  • Singh P. 1980. Armillaria root rot: Artificial inoculation and development of the disease in greenhouse. Eur J Plant Pathol. 10(7):420–431. https://doi.org/10.1111/j.1439-0329.1980.tb00059.x.

  • Smith LA, Dann EK, Pegg KG, Whiley AW, Giblin FR, Doogan V, Kopittke R. 2011. Field assessment of avocado rootstock selections for resistance to Phytophthora root rot. Australas Plant Pathol. 40:3947. https://doi.org/10.1007/s13313-010-0011-0.

    • Search Google Scholar
    • Export Citation
  • Solla A, Tomlinson F, Woodward S. 2002. Penetration of Picea sitchensis root bark by Armillaria mellea, Armillaria ostoyae, and Heterobasidion annosum. For Pathol. 32(1):5570. https://doi.org/10.1046/j.1439-0329.2002.00265.x.

    • Search Google Scholar
    • Export Citation
  • Stange RR Jr , Midland SL, Holmes GJ, Sims JJ, Mayer RT. 2001. Constituents from the periderm and outer cortex of Ipomoea batatas with antifungal activity against Rhizopus stolonifer. Postharvest Biol Technol. 23(2):8592. https://doi.org/10.1016/S0925-5214(01)00105-3.

    • Search Google Scholar
    • Export Citation
  • Treutter D. 2006. Significance of flavonoids in plant resistance: A review. Environ Chem Lett. 4(3):147157. https://doi.org/10.1055/s-2005-873009.

    • Search Google Scholar
    • Export Citation
  • Tripathi L, Odipio J, Tripathi JN, Tusiime G. 2008. A rapid technique for screening banana cultivars for resistance to Xanthomonas wilt. Eur J Plant Pathol. 121:919. https://doi.org/10.1007/s10658-007-9235-4.

    • Search Google Scholar
    • Export Citation
  • Worrall JJ. 1991. Media for selective isolation of hymenomycetes. Mycol. 83:296302. https://doi.org/10.1080/00275514.1991.12026013.

  • Fig. 1.

    Fungal root inoculation assay. (A) Prunus species seedling growing in the greenhouse. (B) Root excavated and segmented for inoculation. (C) Fungal culture for wounded root inoculation assay. (D) Open end of root being placed in fungal culture. (E) Inoculated roots being examined. (F) Fungal culture for intact root inoculation assay. (G) Roots with intact bark being inoculated on top of fungal culture. (H) Evaluation of roots at 21 d following inoculation.

  • Fig. 2.

    Frequency of success of fungal penetration through intact bark of various Prunus genotypes.

  • Fig. 3.

    (A) Longitudinal section of Prunus cerasifera inoculated with Armillaria mellea showing barrier formation. (B) Walling off the infected region with the barrier zone and necrophylactic periderm formation in P. cerasifera. (C) Root of intact ‘Krymsk® 86’ root segment with no sign of fungal invasion without any visible host defense reaction. (D) Formation of barrier zone around the infection point in Prunus genotype ‘BB 106®’. (E) Lignified cells in the barrier zone. (F) Suberized cells in the barrier zone. Bars = 500 µm. White arrows show a fungal infection, and the blue arrows show defense responses.

  • Fig. 4.

    Cross and longitudinal sections of various Prunus species infected with Armillaria spp. (A) Desarmillaria tabescens successfully penetrating in ‘Corette® 5’ (Bars = 200 µm). (B) Armillaria solidipes infecting ‘Corette® 5’ without any visible host defense. (C) Longitudinal section of ‘Controller™ 6’ successfully infected with Armillaria mellea. (D) ‘Controller™ 6’ responding to A. mellea infection. (E) Absence of any barrier zone in phloroglucinol-HCl stained section of ‘Controller™ 6’. (F) A. mellea successfully penetrating ‘Benzie’. Bars = 500 µm. White arrows show fungal infection, and blue arrows indicate defense responses.

  • Fig. 5.

    (A–C) Transverse section of Prunus cerasifera intact root inoculated with Armillaria solidipes. (A) No fungal penetration through intact periderm (Bar = 500 µm). (B) Formation of barrier around fungal infection point (Bar = 500 µm). (C) Cambium is active in the inoculated root (Bar = 200 µm). (D and E) ‘Krymsk® 7’ root with intact periderm inoculated with A. solidipes (Bars = 200 µm). (D) Initiation of formation of necrophylactic periderm. (E) Fungal penetration underneath the bark and barrier being formed to stop the spread. (F) ‘Benzie’ inoculated with A. mellea without distinct host defense response (Bar = 500 µm). White arrows indicate point where root periderm touched fungal inoculum, and blue arrows indicate host defense responses.

  • Adelberg J, Naylor-Adelberg J, Miller S, Gasic K, Schnabel S, Bryson P, Saski C, Parris S, Reighard G. 2021. In vitro co-culture system for Prunus spp. and Armillaria mellea in phenolic foam rooting matric. In Vitro Cell Dev Biol Plant. 57:387397. https://doi.org/10.1007/s11627-020-10136-2.

    • Search Google Scholar
    • Export Citation
  • Aslam AJ, Magel EA. 2018. Influence of drought and season on compartmentalization of black locust (Robinia pseudoacacia L.) inoculated with Armillaria mellea. Eur J Plant Pathol. 152:2131. https://doi.org/10.1007/s10658-018-1444-5.

    • Search Google Scholar
    • Export Citation
  • Baumgartner K, Fujiyoshi P, Ledbetter C, Duncan R, Kluepfel DA. 2018. Screening almond rootstocks for sources of resistance to Armillaria root disease. HortScience. 53(1):48. https://doi.org/10.21273/HORTSCI12038-17.

    • Search Google Scholar
    • Export Citation
  • Baumgartner K, Rizzo DM. 2002. Spread of Armillaria root disease in a California vineyard. Am J Enol Viticult. 53(3):197203. https://doi.org/10.5344/ajev.2002.53.3.197.

    • Search Google Scholar
    • Export Citation
  • Beckman TG, Pusey PL. 2001. Field testing peach rootstocks for resistance to Armillaria root rot. HortScience. 36(1):101103. https://doi.org/10.21273/HORTSCI.36.1.101.

    • Search Google Scholar
    • Export Citation
  • Biggs AR. 1986. Wound age and infection of peach bark by Cytospora leucostoma. Can J Bot. 64(10):23192321.

  • Biggs AR. 1992 Anatomical and physiological responses of bark tissues to mechanical injury, p 13–26. In: Blanchette RA, Biggs AR (eds). Defense mechanisms of woody plants against fungi. Springer Series in Wood Science, Springer Berlin, Heidelberg.

  • Cleary MR, Van Der Kamp BJ, Morrison DJ. 2012a. Effects of wounding and fungal infection with Armillaria ostoyae in three conifer species. II. Host response to the pathogen. For Pathol. 42(2):109123. https://doi.org/10.1111/j.1439-0329.2011.00727.x.

    • Search Google Scholar
    • Export Citation
  • Cleary MR, van der Kamp BJ, Morrison DJ. 2012b. Effects of wounding and fungal infection with Armillaria ostoyae in three conifer species. I. Host response to abiotic wounding in non‐infected roots. For Pathol. 42(2):100108. https://doi.org/10.1111/j.1439-0329.2011.00726.x.

    • Search Google Scholar
    • Export Citation
  • Devkota P, Hammerschmidt R. 2019. A rapid and holistic approach to screen susceptibility of Prunus species to Armillaria root rot. For Pathol. 49(5):e12547. https://doi.org/10.1111/efp.12547.

    • Search Google Scholar
    • Export Citation
  • Devkota P, Iezzoni A, Gasic K, Reighard G, Hammerschmidt R. 2020. Evaluation of the susceptibility of Prunus rootstock genotypes to Armillaria and Desarmillaria species. Eur J Plant Pathol. 158(1):177193. https://doi.org/10.1007/s10658-020-02065-y.

    • Search Google Scholar
    • Export Citation
  • Du J, Groover A. 2010. Transcriptional regulation of secondary growth and wood formation. J Integr Plant Biol. 52(1):1727. https://onlinelibrary.wiley.com/doi/10.1111/j.1744-7909.2010.00901.x.

    • Search Google Scholar
    • Export Citation
  • Elias-Roman RD, Calderon-Zavala G, Guzman-Mendoza R, Vallejo-Perez MR, Klopfenstein NB, Mora-Aguilera JA. 2019. ‘Mondragon’: A clonal plum rootstock to enhance management of Armillaria root disease in peach orchards of Mexico. Crop Prot. 121:8995. https://doi.org/10.1016/j.cropro.2019.03.011.

    • Search Google Scholar
    • Export Citation
  • Fenning TM. 2019. The use of tissue culture and in-vitro approaches for the study of tree diseases. Plant Cell Tissue Organ Cult. 136(3):415430. https://doi.org/10.1007/s11240-018-01531-0.

    • Search Google Scholar
    • Export Citation
  • Guillaumin JJ, Pierson J, Grassely C. 1991. The susceptibility to Armillaria mellea of different Prunus species used as stone fruit rootstocks. Scientia Hortic. 46(1-2):4354. https://doi.org/10.1016/0304-4238(91)90091-C.

    • Search Google Scholar
    • Export Citation
  • Koch RA, Wilson AW, Séné O, Henkel TW, Aime MC. 2017. Resolved phylogeny and biogeography of the root pathogen Armillaria and its gasteroid relative, Guyanagaster. BMC Evol Biol. 17:133. https://doi.org/10.1186/s12862-017-0877-3.

    • Search Google Scholar
    • Export Citation
  • Mansilla JP, Aguín O, Sainz MJ. 2001. A fast method for production of Armillaria inoculum. Mycol. 93(3):612615. https://doi.org/10.1080/00275514.2001.12063191.

    • Search Google Scholar
    • Export Citation
  • Mullick DB. 1977. The non-specific nature of defense in bark and wood during wounding, insect and pathogen attack, p 395–441. In: Loewus FA, Runeckles VC (eds). The structure, biosynthesis, and degradation of wood. Springer, Boston, MA. https://doi.org/10.1007/978-1-4615-8873-3_10.

  • Proffer TJ, Jones AL, Perry RL. 1988. Testing of cherry rootstocks for resistance to infection by species of Armillaria. Plant Dis. 72(6):488490. https://doi.org/10.1094/PD-72-0488.

    • Search Google Scholar
    • Export Citation
  • Proffer TJ, Jones AL, Ehret GR. 1987. Biological species of Armillaria isolated from sour cherry orchards in Michigan. Phytopathology. 77(6):941943. https://doi.org/10.1094/Phyto-77-941.

    • Search Google Scholar
    • Export Citation
  • Raabe RD. 2008. Plants resistant or susceptible to Armillaria mellea, the oak root fungus. University of California, Berkeley. https://alamedabackyardgrowers.org/wp-content/uploads/2019/01/List-of-Oak-Root-Fungus-Resistant-or-Susceptible-Trees-Plants.pdf. [accessed 28 Jul 2023].

  • Robinson RM, Morrison DJ, Jensen GD. 2004. Necrophylactic periderm formation in the roots of western larch and Douglas‐fir trees infected with Armillaria ostoyae. II. The response to the pathogen. For Pathol. 34(2):119129. https://doi.org/10.1111/j.1439-0329.2004.00354.x.

    • Search Google Scholar
    • Export Citation
  • Saucet SB, Van Ghelder C, Abad P, Duval H, Esmenjaud D. 2016. Resistance to root‐knot nematodes Meloidogyne spp. in woody plants. New Phytol. 211(1):4156. https://doi.org/10.1111/nph.13933.

    • Search Google Scholar
    • Export Citation
  • Schnabel G, Ash JS, Bryson PK. 2005. Identification and characterization of Armillaria tabescens from the southeastern United States. Mycol Res. 109(11):12081222. https://doi.org/10.1017/S0953756205003916.

    • Search Google Scholar
    • Export Citation
  • Shigo AL, Tippett JT. 1981. Compartmentalization of decayed wood associated with Armillaria mellea in several tree species. Broomall, PA, US Department of Agriculture, Forest Service, Northeastern Forest Experiment Station. 20:488. https://doi.org/10.2737/NE-RP-488.

  • Singh P. 1980. Armillaria root rot: Artificial inoculation and development of the disease in greenhouse. Eur J Plant Pathol. 10(7):420–431. https://doi.org/10.1111/j.1439-0329.1980.tb00059.x.

  • Smith LA, Dann EK, Pegg KG, Whiley AW, Giblin FR, Doogan V, Kopittke R. 2011. Field assessment of avocado rootstock selections for resistance to Phytophthora root rot. Australas Plant Pathol. 40:3947. https://doi.org/10.1007/s13313-010-0011-0.

    • Search Google Scholar
    • Export Citation
  • Solla A, Tomlinson F, Woodward S. 2002. Penetration of Picea sitchensis root bark by Armillaria mellea, Armillaria ostoyae, and Heterobasidion annosum. For Pathol. 32(1):5570. https://doi.org/10.1046/j.1439-0329.2002.00265.x.

    • Search Google Scholar
    • Export Citation
  • Stange RR Jr , Midland SL, Holmes GJ, Sims JJ, Mayer RT. 2001. Constituents from the periderm and outer cortex of Ipomoea batatas with antifungal activity against Rhizopus stolonifer. Postharvest Biol Technol. 23(2):8592. https://doi.org/10.1016/S0925-5214(01)00105-3.

    • Search Google Scholar
    • Export Citation
  • Treutter D. 2006. Significance of flavonoids in plant resistance: A review. Environ Chem Lett. 4(3):147157. https://doi.org/10.1055/s-2005-873009.

    • Search Google Scholar
    • Export Citation
  • Tripathi L, Odipio J, Tripathi JN, Tusiime G. 2008. A rapid technique for screening banana cultivars for resistance to Xanthomonas wilt. Eur J Plant Pathol. 121:919. https://doi.org/10.1007/s10658-007-9235-4.

    • Search Google Scholar
    • Export Citation
  • Worrall JJ. 1991. Media for selective isolation of hymenomycetes. Mycol. 83:296302. https://doi.org/10.1080/00275514.1991.12026013.

Pratima Devkota Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI 48824, USA

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Amy Iezzoni Department of Horticulture, Michigan State University, East Lansing, MI 48824, USA

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Ksenija Gasic Department of Plant and Environmental Sciences, Clemson University, Clemson, SC 29634, USA

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Gregory Reighard Department of Plant and Environmental Sciences, Clemson University, Clemson, SC 29634, USA

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Raymond Hammerschmidt Department of Plant, Soil and Microbial Sciences, Michigan State University, East Lansing, MI 48824, USA

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Contributor Notes

This work was funded by US Department of Agriculture (USDA) Agricultural Marketing Service–Specialty Crop Multi-State Program grant 2015‐10.170, USDA-National Institute of Food and Agriculture–Specialty Crop Research Initiative grant 2020-51181-32142, Michigan State University Project GREEEN, and Michigan State University AgBioResearch. We thank Fowler Nursery for providing Prunus genotypes.

P.D. is the corresponding author. E-mail: devkotap@msu.edu.

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  • Fig. 1.

    Fungal root inoculation assay. (A) Prunus species seedling growing in the greenhouse. (B) Root excavated and segmented for inoculation. (C) Fungal culture for wounded root inoculation assay. (D) Open end of root being placed in fungal culture. (E) Inoculated roots being examined. (F) Fungal culture for intact root inoculation assay. (G) Roots with intact bark being inoculated on top of fungal culture. (H) Evaluation of roots at 21 d following inoculation.

  • Fig. 2.

    Frequency of success of fungal penetration through intact bark of various Prunus genotypes.

  • Fig. 3.

    (A) Longitudinal section of Prunus cerasifera inoculated with Armillaria mellea showing barrier formation. (B) Walling off the infected region with the barrier zone and necrophylactic periderm formation in P. cerasifera. (C) Root of intact ‘Krymsk® 86’ root segment with no sign of fungal invasion without any visible host defense reaction. (D) Formation of barrier zone around the infection point in Prunus genotype ‘BB 106®’. (E) Lignified cells in the barrier zone. (F) Suberized cells in the barrier zone. Bars = 500 µm. White arrows show a fungal infection, and the blue arrows show defense responses.

  • Fig. 4.

    Cross and longitudinal sections of various Prunus species infected with Armillaria spp. (A) Desarmillaria tabescens successfully penetrating in ‘Corette® 5’ (Bars = 200 µm). (B) Armillaria solidipes infecting ‘Corette® 5’ without any visible host defense. (C) Longitudinal section of ‘Controller™ 6’ successfully infected with Armillaria mellea. (D) ‘Controller™ 6’ responding to A. mellea infection. (E) Absence of any barrier zone in phloroglucinol-HCl stained section of ‘Controller™ 6’. (F) A. mellea successfully penetrating ‘Benzie’. Bars = 500 µm. White arrows show fungal infection, and blue arrows indicate defense responses.

  • Fig. 5.

    (A–C) Transverse section of Prunus cerasifera intact root inoculated with Armillaria solidipes. (A) No fungal penetration through intact periderm (Bar = 500 µm). (B) Formation of barrier around fungal infection point (Bar = 500 µm). (C) Cambium is active in the inoculated root (Bar = 200 µm). (D and E) ‘Krymsk® 7’ root with intact periderm inoculated with A. solidipes (Bars = 200 µm). (D) Initiation of formation of necrophylactic periderm. (E) Fungal penetration underneath the bark and barrier being formed to stop the spread. (F) ‘Benzie’ inoculated with A. mellea without distinct host defense response (Bar = 500 µm). White arrows indicate point where root periderm touched fungal inoculum, and blue arrows indicate host defense responses.

 

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