408 – Baumgartner and Rizzo

Relative Resistance of Grapevine Rootstocks to Root Disease

Kendra Baumgartner1* and David M. Rizzo2

Abstract: Grapevine rootstocks were screened for resistance to infection by , the pathogen that causes Armillaria root disease. The first objective was to determine which factors hasten colonization of grape- vines by A. mellea in order to develop the most rapid inoculation technique. Results showed that wounding the root collar bark and vascular cambium did not significantly increase infection rate (p = 1.0), that young vines were infected significantly faster than older vines (p = 0.03), and that fine roots (≤5 mm diam) were not susceptible to infection. Based on these findings, the inoculation technique was modified and used to inoculate dormant rootings of eight rootstocks in the greenhouse. After two years, the root collars were examined for the presence of myce- lial fans of A. mellea and infection was confirmed by culture. The rootstock Freedom had the lowest frequency of infection (7%) and was the most resistant rootstock (p = 0.0016). Rootstocks O39-16, 5C, Riparia Gloire, and 3309C had the highest frequencies of infection with 63%, 73%, 79%, and 85%, respectively. Rootstocks St. George, Ramsey, and 110R had intermediate frequencies of infection. Results suggest that Freedom and, to a lesser extent, St. George, Ramsey, and 110R will be useful components of an integrated management strategy for Armillaria root disease. Resistant rootstocks may decrease the rate of colonization of grapevines in A. mellea-infested vineyards, thereby reducing yield losses from Armillaria root disease. Key words: Armillaria mellea, disease resistance, screening technique, Vitis vinifera

Armillaria root disease of grapevine (Vitis vinifera L.) is cronutrients, delayed ripening of fruit, and eventual death caused by Armillaria mellea (Vahl:Fr.) P. Kumm., a fungal of infected vines (Baumgartner and Warnock 2006). Foliar pathogen that also infects the roots of many forest symptoms include stunted shoots, dwarfed leaves, wilting, in California (Baumgartner and Rizzo 2001b). The pathogen and premature defoliation. survives as a saprobe on woody roots after the host dies Armillaria root disease affects vineyards in all major (Redfern and Filip 1991). Its mycelium decomposes cellu- grapegrowing regions of California, including the Central lose and other cell wall components for nutrients as it Valley, the North Coast, and the Central Coast (Baum- grows, decaying the root wood. Where forests are con- gartner and Rizzo 2001a). Because of the broad host range verted to vineyards, mycelium can survive in partially de- of A. mellea, which includes many species of fruit and cayed roots that remain buried in the soil and can in- nut trees (Raabe 1962), planting a different perennial crop fect grapevines for up to several years after planting in place of diseased vines is unlikely to eliminate the (Baumgartner and Rizzo 2002). Infection occurs when vine Armillaria root disease problem. The common name of A. roots come in direct contact with residual tree roots and mellea, “oak root ,” misleads many growers to be- are subsequently colonized by A. mellea mycelium. Armil- lieve that establishing vineyards on sites without oaks laria root disease causes significantly decreased yield and minimizes the risk of Armillaria root disease. Avoidance of growth (Baumgartner 2004), lower root absorption of ma- infested forest sites is extremely difficult because A. mel- lea infects many different tree species in addition to oaks and aboveground symptoms are rare (Baumgartner and Rizzo 2001b). Furthermore, the mycelial fans and mush- 1Research plant pathologist, USDA–Agricultural Research Service, and 2Pro- rooms of other Armillaria species, none of which cause fessor, Department of Plant Pathology, University of California, Davis, Armillaria root disease on agronomic hosts, can be mis- CA 95616. taken for A. mellea (Baumgartner and Rizzo 2001b). *Corresponding author [email: [email protected]; tel: 530 754- 7461; fax: 530 754-7195] Pesticides are not the best means of controlling Acknowledgments: Research was supported by the USDA–Viticulture Con- Armillaria root disease. Methyl bromide can kill A. mellea sortium and the USDA–Agricultural Research Service. Grapevines were in residual roots (Munnecke et al. 1981), but it does not generously donated by Sunridge Nurseries Inc., Bakersfield, CA. penetrate the soil deeply enough to reach all infected The authors thank J. Warren, B. Craughwell, and A. Euliss for greenhouse roots (Bliss 1951). Vines planted in fumigated soil are typi- and laboratory assistance and S. Krebs (Napa Valley College, CA), P. Cous- cally healthy for several years, but they eventually be- ins (USDA-ARS, Geneva, NY), and G. Schnabel (Clemson University, SC) for their comments on this manuscript. come infected when their roots contact inoculum that es- Manuscript submitted April 2006; revised May 2006 capes the effects of the fumigant (Gubler 1992). Fungicides Copyright © 2006 by the American Society for Enology and Viticulture. examined for control of Armillaria root disease also do not All rights reserved. provide long-term control (Adaskaveg et al. 1999). These 408

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findings are consistent with the biology of the pathogen. made on 1.5% water agar (WA) with benomyl (4 μg a.i./ Armillaria mellea is a wood decay fungus that lives be- mL) and streptomycin sulfate (100 μg/mL) added after au- neath the bark of infected roots, where it decomposes root toclaving. Isolation plates were incubated at 24°C for 2 wood and creates breaks in the vascular system (Redfern wk. Subcultures were transferred to 1% malt extract agar and Filip 1991). Eradicating A. mellea from a living host (MEA) with an overlay of sterile cellophane. Identity was requires the difficult task of delivering a fungicide to root confirmed by PCR amplification of the nuclear rDNA wood that is decomposed and, thus, not connected to the intergenic spacer I (IGS-I) (Harrington and Wingfield vine’s vascular system. Indeed, fungicides injected into 1995). DNA was amplified directly from mycelium by gently infected almond trees were effective only to the extent scraping a pipette tip over the surface of the mycelium that they actually reached the A. mellea infections (Ad- and dipping it into the PCR vial. Digestions were per- askaveg et al. 1999). formed with AluI added directly to the PCR vial and incu- In contrast to soil fumigation or fungicide injections, a bation at 37°C for 2 hr. The isolate had a restriction pat- more sustainable approach to the control of Armillaria tern of two fragments, 322 and 155 bp, which is diagnostic root disease of grapevine may be the use of resistant of A. mellea. rootstocks. In a previous study, Vitis germplasm with re- Inoculum was prepared by inoculating segments (1.5 cm sistance to Armillaria root disease was identified through diam x 5 cm length) of dormant pear shoots collected from inoculation of potted plants in the greenhouse (Raabe our experimental orchard (Armstrong Experimental Station, 1979). However, the identity of the Armillaria isolates is Plant Pathology Department, UC Davis). Glass jars were unknown because the study was completed before the filled with 30 segments per jar and autoclaved for two 1.5- discovery that A. mellea consists of multiple biological hr periods, separated by 12 hr at 24°C. After cooling, 60 species (Anderson and Ullrich 1979), which have since agar plugs from A. mellea cultures were added per jar. been named as distinct Armillaria species (Guillaumin et Screw-top lids were tightened atop each jar and sealed al. 1991, Volk et al. 1996). Furthermore, the 30 examined with parafilm to prevent contamination. Jars were incu- selections (Raabe 1979) consisted primarily of Vitis spe- bated in the dark at 24°C for 6 mo (months). cies (such as V. girdiana and V. treleasei) that are neither Wounding and inoculation rate. The factors examined used as rootstocks nor definitively resistant to grape in the first experiment were artificial wounding (wounded phylloxera, Daktulosphaira vitifoliae (Fitch), a wide- or intact bark) and inoculation rate (one pear segment spread vineyard pest. Our goal was to identify A. mellea- colonized by A. mellea at 1x inoculum or three pear seg- resistant rootstocks that are both widely available to ments at 3x inoculum). There were four experimental treat- growers and are resistant to phylloxera. ments: intact bark–1x inoculum, intact bark–3x inoculum, Our first objective was to identify the best technique wounded bark–1x inoculum, and wounded bark–3x inocu- to hasten infection by conducting a series of experiments lum. Inoculum was prepared 6 mo before inoculating the on St. George, a rootstock identified as susceptible (Raabe plants. One-yr-old, field-grown dormant rootings of St. 1979). Raabe’s technique of inoculating vines by burying George rootstock were potted in 11.4-L pots in sterile inoculum beneath their root systems was successful at potting mix. Potting mix consisted of a 1:1:1 v/v ratio of causing infection, but it took many months to complete peat moss, perlite, and Supersoil (peat moss, ground fir the experiment. To address the problem of slow coloniza- bark, redwood compost, sand, 2-4-2 slow release fertilizer, tion for which greenhouse experiments with Armillaria oyster shell flour, and ground dolomite; Rod McClellan species are known (Singh 1980), modifications of Raabe’s Co., South San Francisco, CA) that was autoclaved for 1 inoculation technique were tested. Included in the testing hr. Plants were fertilized with Hoagland’s nutrient solution were the use of artificial wounding, inoculation of younger twice per month. plants, and placement of inoculum directly on the root Treatments were imposed on plants 2 mo after planting. collar, all factors that affect infection of other hosts by Wounds 30 mm x 5 mm were made with a sterile scalpel other Armillaria species (Garrett 1956, Sinclair et al. 1987, by removing the root bark and the attached vascular Wahlstrom and Johansson 1992, Solla et al. 2002). Our cambium to expose the underlying root wood. Wounds second objective was to use an improved inoculation were positioned on the root collar ~20 cm below the soil technique to evaluate the resistance of eight rootstocks surface and 2 cm above the base of the rooting, where to infection by A. mellea. the majority of roots originated. Plants inoculated with three pieces of inoculum (3x inoculum) were wounded in Materials and Methods the same relative position to the soil surface and the base of the rooting, equidistant around the circumference of The isolate of A. mellea used for all inoculations was the root collar. At the time of inoculation, slivers of wood collected from a symptomatic grapevine (Healdsburg, from each piece of inoculum were plated on WA to con- Sonoma County, CA) that had been part of a previous firm their viability. Inoculum was secured to the root col- study (Baumgartner and Rizzo 2001a). Tissues were axeni- lar with cotton string, directly over the wound site on cally removed in the laboratory from portions of mycelial wounded plants and in the same relative position on fans that were previously unexposed. Isolations were plants with intact bark. Sterile segments of pear wood

Am. J. Enol. Vitic. 57:4 (2006) 410 – Baumgartner and Rizzo served as inoculum for noninoculated plants. Plants were Fine root inoculation. In a third experiment, the fre- arranged in a completely randomized design (CRD) on one quency of infection of 2-mo and 12-mo-old roots of St. greenhouse bench, with 12 inoculated and 12 noninoc- George was compared to determine if roots, as opposed to ulated plants per treatment. the root collar, are susceptible to infection. Two-mo and At 1, 4, and 12 mo after inoculation, four inoculated and 12-mo-old roots were chosen because they represented four noninoculated plants per treatment were destructively nonlignified and lignified roots, respectively. Inoculum sampled and their root collar tissues were cultured for was prepared 6 mo before inoculating the plants. One-yr- confirmation of infection by A. mellea. Root collar wood old dormant rootings were potted in 11.4-L pots with ster- located directly beneath the inoculum and underneath bark ile potting mix. After two months, we inoculated three 2- surrounding the inoculation site, from the soil line to the mm diam nonlignified roots (2-mo-old) and three 5-mm base of the trunk, was examined for mycelial fans by re- diam lignified roots (12-mo-old). Inoculum (one pear seg- moving thin layers of bark with a sterile scalpel. Underly- ment) was secured to each of three roots per age per plant ing wood and pieces of mycelial fans were plated on WA. with narrow strips of cheesecloth. At the time of inocula- The viability of the residual inoculum was confirmed at tion, slivers of wood from each piece of inoculum were each sampling time by plating four small pieces per pear plated on WA to confirm their viability. Sterile segments segment on WA. Isolation plates were incubated at 24°C of pear wood served as inoculum for noninoculated and examined after 2 wk. Subcultures were transferred to plants. Plants were arranged in a CRD on one greenhouse MEA for identification. Frequency of infection for each bench, with 12 inoculated and 12 noninoculated plants. treatment x sampling time combination was calculated as Plants were fertilized with Hoagland’s nutrient solution the percentage of inoculated plants from which A. mellea twice per month. was cultured. The effects of wounding (intact or wounded At 1, 4, and 12 mo after inoculation, four inoculated and bark), inoculation rate (1x or 3x inoculum), and their inter- four noninoculated plants were destructively sampled and actions on frequency of infection were determined with inoculated roots of each age were cultured for confirma- analysis of variance (ANOVA) using the MIXED proce- tion of infection by A. mellea. The viability of the residual dure in SAS (SAS System, version 8.2; SAS Institute Inc., inoculum was confirmed at each sampling time. Frequency Cary, NC), with Kenward–Roger as the denominator de- of infection for each root age was calculated per plant as grees of freedom method. All effects were considered fixed the percentage of inoculated roots from which A. mellea effects. Tukey’s tests were used for mean comparisons. was cultured and was averaged across plants for each Host age. In a second experiment, the rate of infection sampling time. The FREQ procedure in SAS was used to of 1-yr and 3-yr-old St. George was compared. Inoculum perform a chi-square test of independence to determine was prepared 6 mo before inoculating both sets of plants. the relationship between root age and frequency of infec- The first set of plants consisted of 1-yr-old dormant tion for sampling times when frequency of infection varied rootings. These were potted in 11.4-L pots in sterile pot- between root age groups. ting mix. After two years, a second set of 1-yr-old dormant Relative resistance of eight rootstocks. In a fourth rootings was potted. After an additional 2 mo, inoculum experiment, dormant rootings of Riparia Gloire (V. riparia (one pear segment per plant) was secured to the root col- Michx.), St. George (V. rupestris Scheele), 3309C (V. lars of both sets of plants with cotton string, ~20 cm be- riparia x V. rupestris), 5C (V. berlandieri Planch. x V. low the soil surface and 2 cm above the base of the root- riparia), 110R (V. berlandieri x V. rupestris), Ramsey (V. ing. At the time of inoculation, slivers of wood from each champinii Planch.), O39-16 (V. vinifera x Muscadinia piece of inoculum were plated on WA to confirm their vi- rotundifolia Michx.), and Freedom (1613C seedling x Dog ability. Sterile segments of pear wood served as inoculum Ridge seedling) were potted in 11.4-L pots with sterile for noninoculated plants. Plants were arranged in a CRD potting mix. This experiment was conducted after the three on one greenhouse bench, with 12 inoculated and 12 experiments described above were completed and the fast- noninoculated plants per age group. Plants were fertilized est method of infecting plants had been identified. Inocu- with Hoagland’s nutrient solution twice per month. lum was prepared 6 mo before inoculating the plants. Af- At 1, 4, and 12 mo after inoculation, four inoculated and ter 2 mo, inoculum (one pear segment per plant) was four noninoculated plants per age group were destruc- secured to the root collars of 15 to 26 plants per rootstock tively sampled and their root collar tissues were cultured with cotton string. The inoculum was secured to the root for confirmation of infection by A. mellea. Viability of the collars without wounding the root bark, ~20 cm below the residual inoculum was confirmed at each sampling time. soil surface and 2 cm above the base of the rooting. Un- Frequency of infection for each plant age x sampling time even sample sizes among rootstocks were due to variable combination was calculated as the percentage of inocu- rooting success. At the time of inoculation, slivers of lated plants from which A. mellea was cultured. The FREQ wood from each piece of inoculum were plated on WA to procedure in SAS was used to perform a chi-square test confirm their viability. Sterile segments of pear wood of independence to determine the relationship between served as inoculum for noninoculated plants. Plants were plant age and frequency of infection for sampling times arranged in a randomized complete block design (RCBD) when frequency of infection varied between age groups. on two greenhouse benches that served as replicate

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blocks, with 7 to 13 inoculated plants and 6 noninoculated That was also true for inoculum secured to all plants that plants per rootstock per block. Plants were fertilized with were sampled at 1 mo and 4 mo after inoculation. After 12 Hoagland’s nutrient solution twice per month. mo, however, the inoculum was dark brown and friable, In contrast to the first three experiments on St. George, and none of it was viable. Foliage of infected St. George which were terminated 12 mo after inoculation, this fourth was not visibly distinguishable from that of the noninoc- experiment was observed for 24 mo after inoculation. This ulated St. George at any of the sampling times; foliar permitted a comparison with Raabe’s findings, who also symptoms of Armillaria root disease that typically occur observed his inoculated plants for 24 mo (Raabe 1979). It on infected grapevines in the field (stunted shoots, also increased the probability of mortality, another pos- dwarfed leaves, wilting, premature defoliation) did not ap- sible measure of resistance. At 24 mo after inoculation, all pear on our potted plants. Armillaria mellea was not cul- plants were destructively sampled and their tissues were tured from noninoculated St. George at any sampling time, examined for the presence of A. mellea infection. Plants demonstrating that the nursery stock was not a source of that died during the 24-mo period were immediately infection among the inoculated plants. sampled for the presence of A. mellea infection. Inoculum Host age. The younger St. George plants were infected viability was confirmed at the time of death for dead faster, with a significantly higher frequency of infection plants and 24 mo after inoculation for surviving plants. among 1-yr-old plants at 1 mo after inoculation (χ2 = 4.80, Frequency of infection for each rootstock was calculated p = 0.03), compared to no infection among 3-yr-old plants as the percentage of inoculated plants (surviving plants + at this time (Table 1). At 4 and 12 mo after inoculation, all dead plants) from which A. mellea was cultured. Fre- inoculated plants were infected, regardless of plant age quency of mortality for each rootstock was calculated as (Table 1). No plants died, even among those examined 12 the percentage of inoculated plants that died. mo after inoculation. Mycelial fans (Figure 1) were present Statistical analysis. ANOVA was used to determine on the root collars immediately beneath the inoculum on the main effects of rootstock (Riparia Gloire, St. George, all inoculated vines from which A. mellea was cultured. 3309C, 5C, 110R, Ramsey, O39-16, or Freedom) and block (A or B) on frequencies of infection and mortality, using the GLM procedure in SAS. Tukey’s test was used for Table 1 Effects of wounding, inoculation rate, plant age, and mean comparisons. A log transformation was applied to root age on frequency of Armillaria mellea infection of potted St. George rootstock. Plants were destructively sampled at frequencies of mortality to homogenize variances; reverse- 1, 4, and 12 mo after inoculation. transformed means are presented in the text. Tukey’s single degree of freedom test for non-additivity was used Frequency of infection after inoculation to test the linear portion of the block x rootstock interac- (% infected plants among total inoculated) tion on frequencies of infection and mortality. The CORR Treatment 1 mo 4 mo 12 mo procedure was applied to a data set that included fre- Wounding and quency of infection and frequency of mortality as depen- inoculation ratea dent variables, with rootstock as the independent variable. Intact bark Linear correlations were used to measure the intensity of 1x inoculum 75 ab 100 a 100 a association between frequency of infection and frequency 3x inoculum 50 a 100 a 100 a of mortality. Wounded bark 1x inoculum 50 a 100 a 100 a Results 3x inoculum 75 a 100 a 100 a Plant agec Wounding and inoculation rate. Wounding (p = 1.0), 1 yr 75 a 100 a 100 a inoculation rate (p = 1.0), and their interaction (p = 0.5) 3 yr 0 b 100 a 100 a had no significant effects on frequency of infection of in- oculated St. George. At 1 mo after inoculation, frequency Root aged of infection among inoculated plants ranged from 50 to 2 mo 0 a 0 a 0 a 75% (Table 1). At 4 and 12 mo after inoculation, frequency 12 mo 0 a 0 a 0 a of infection was 100% in all four treatments; every inocu- aThe root collars of 1-yr-old plants were inoculated with A. mellea. lated plant was infected, regardless of treatment (Table 1). Wounds were made to the depth of the root wood. 1x inoculum: None of the inoculated plants died, even among the 16 one piece of pear wood colonized by A. mellea; 3x inoculum: three pieces of pear wood colonized by A. mellea. plants examined 12 mo after inoculation. Mycelial fans bMeans (n = 4 plants) within a column, for each experiment, followed (Figure 1) were present on the root collars immediately by different letters are significantly different at p ≤ 0.05, Tukey’s beneath the inoculum on all inoculated vines from which test. A. mellea was cultured. cThe root collars of 1-yr and 3-yr-old plants were inoculated with Based on successful culture of A. mellea from slivers 1x inoculum, without wounding the root bark. dThe fine roots of 1-yr-old plants were inoculated with 1x inoculum of inoculum plated at the time of inoculation, all 32 pieces for each of three 2-mo-old roots and three 12-mo-old roots per plant, of inoculum were viable at the start of the experiment. without wounding the roots.

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As in the first experiment, all eight pieces of inoculum age of the roots to which it had been secured. Foliage of secured to the root collars of the 1-yr-old plants and the infected St. George was not visibly distinguishable from 3-yr-old plants were viable at the time of inoculation. That that of the noninoculated St. George at any of the sam- was also true for inoculum secured to all plants that were pling times. Armillaria mellea was not cultured from sampled at 1 mo after inoculation and at 4 mo after inocu- noninoculated St. George at any sampling time. lation. After 12 mo, the inoculum was dark brown and fri- Relative resistance of eight rootstocks. Based on the able, and none of it was viable. Foliage of infected St. results of the three experiments presented above, which George was not visibly distinguishable from that of the were completed before conducting this fourth experiment, noninoculated St. George at any of the sampling times. the best method for inoculation was determined. This Armillaria mellea was not cultured from noninoculated St. choice was based on the speed of infection when one George at any sampling time. piece of inoculum was secured directly to the root collars Fine root inoculation. None of the root inoculations of 1-yr-old plants without wounding their root bark. This resulted in A. mellea infection (Table 1). Mycelial fans fourth experiment was extended to 24 mo after inoculation, were absent from all plants and A. mellea was not isolated since 12 mo after inoculation proved to be insufficient to from discolored tissue sampled from roots where inoculum bring about mortality in our first three experiments, and in had been secured. Trichoderma sp., Fusarium oxysporum, order to compare our results to those of Raabe (1979). and Cylindrocarpon sp. were isolated from discolored Tukey’s single degree of freedom test for non-additivity root tissue. At the time of inoculation, all 24 pieces of in- detected no significant effects of the block x rootstock in- oculum secured to all roots were viable. At 1 mo after in- teraction on frequency of infection (p = 0.0883) or fre- oculation, only 33% and 50% of the inoculum attached to quency of mortality (p = 0.4186). There were no significant 2-mo and 12-mo-old roots, respectively, was still viable. differences between blocks for either frequency of infec- None of the inoculum was viable among any of the plants tion (p = 0.2240) or frequency of mortality (p = 0.7281). sampled at 4 and 12 mo after inoculation, regardless of the There were significant differences in frequency of infection among rootstocks (p = 0.0016). Freedom had the lowest frequency of infection and, thus, the highest resistance compared to all other rootstocks examined (Table 2). Rootstocks with the least resistance to infection—that is, the most susceptible rootstocks—were 3309C, Riparia

Table 2 Frequencies of infection and mortality of potted grape- vine rootstocks inoculated with Armillaria mellea (% of plants among total inoculated). Plants were destructively sampled 24 mo after inoculation. Plants that died during the 24-mo period were sampled immediately after death.

Frequency of Frequency of infection mortality Rootstocka (% infected plants) (% dead plants)

Freedom 7 ab 0a (n = 15) St. George 35 b 6 a (n = 17) Ramsey 38 b 5 a (n = 21) 110R 44 bc 6 a (n = 18) O39-16 63 cd 13 a (n = 24) 5C 73 cd 8 a (n = 26) Riparia Gloire 79 d 26 a (n = 19) 3309C 85 d 0 a (n = 26) Figure 1 Mycelial fans (thick, white sheets of fungal tissue) of Armillaria mellea underneath the root bark, on the root collar of potted St. George aRoot collars of 1-yr-old plants of each rootstock were inoculated rootstock. This plant, which was inoculated without wounding the bark with 1x inoculum, without wounding the root bark (n: number of in- and was sampled at 12 mo after inoculation, showed none of the foliar oculated plants per rootstock, summed over two replicate blocks). symptoms of Armillaria root disease (stunted shoots, dwarfed leaves, bMeans within a column with different letters are significantly different wilting, premature defoliation) typically found in the field. at p ≤ 0.05, Tukey’s test.

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Gloire, 5C, and O39-16. St. George, Ramsey, and 110R had hyphae to penetrate 10 times as many cell layers in the relatively intermediate resistance, with fewer than 50% of same amount of time, demonstrating that the outer root plants infected. bark of Picea sitchensis inhibits A. mellea infection of the Frequency of mortality was very low. The first plant underlying vascular cambium. It is possible that St. died 14 mo after inoculation and only 12 of 166 inoculated George, the rootstock on which we examined the effects of plants died 24 mo after inoculation. Frequency of mortal- wounding, is so tolerant of A. mellea that removing the ity did not vary significantly among rootstocks (p = bark does not significantly increase the rate at which the 0.3857; Table 2). There was no correlation between fre- pathogen colonizes wood beneath the bark. Inoculations quency of infection and frequency of mortality (r = with A. ostoyae showed that the hyphae of this 0.4237, p = 0.1020). At the time of inoculation, all 166 pathogen penetrate wounded and nonwounded tissue at a pieces of inoculum were viable. After 24 mo, none of the faster rate in a susceptible species, Pseudotsuga men- inoculum was viable. Armillaria mellea was not cultured ziesii, than in a resistant species, Larix occidentalis (Rob- from noninoculated rootstocks at any sampling time in ei- inson et al. 2004). Both conifer species formed necro- ther block. phylactic periderm in response to infection and wounding, but A. ostoyae more frequently breached the periderm of Discussion the susceptible species. St. George was found to be mod- Our finding that Freedom had the lowest frequency of erately resistant to infection in our comparisons of the infection by A. mellea indicates that this rootstock is more eight rootstocks. If we had used the most susceptible likely to resist infection than the other seven rootstocks rootstock, 3309C, instead of St. George in our first experi- that were tested. In contrast, the high frequency of infec- ment, we may have detected significant effects of wound- tion of 3309C, Riparia Gloire, 5C, and O39-16 indicates that ing on infection rate. It is likely that the moderate resis- vineyards established on infested sites with these root- tance of St. George to A. mellea infection is also the stocks are likely to suffer from more severe Armillaria root reason that tripling the amount of inoculum had no effect disease. Although none of the 3309C plants died, despite on infection rate. the fact that 85% were infected, it is important to note Controlled inoculations have not been used previously that none of the differences in mortality among rootstocks to demonstrate the effects of host age on infection by an were statistically significant. The low frequency of mortal- Armillaria species. Our finding that younger grapevines ity of all eight rootstocks is consistent with low frequency are infected faster than older grapevines is consistent with of mortality of peach rootstocks inoculated with A. ta- observations of more severe mortality in young forest bescens (Beckman and Pusey 2001) and various conifer plantations, compared to older stands of the same conifer species inoculated with A. ostoyae (Entry et al. 1992). species (Sinclair et al. 1987). Increased resistance of older In comparison with past inoculations of Vitis germ- plants may be related to the fact that they have more lay- plasm (Raabe 1979), we had different results for the only ers of outer root bark and, thus, a thicker barrier to slow two rootstocks, Riparia Gloire and St. George, that our ex- penetration by A. mellea. periments shared in common. We found that St. George Our inability to bring about infection of fine roots, had a significantly lower frequency of infection than both lignified and nonlignified, is not surprising. Although Riparia Gloire (35% versus 79%). In contrast, Raabe found A. mellea can grow on a variety of substrates in culture that St. George had a higher frequency of infection than and has a broad host range (Raabe 1962), the few annual Riparia Gloire (80% versus 10%). Differences in our find- plants that it has been shown to infect are mainly those ings are likely due to method of inoculation. Raabe buried that produce fleshy roots, such as potato, parsnip, and inoculum in the potting media instead of attaching it di- carrot (Thomas 1934). It is surprising that rhizomorphs did rectly to the root collars, as we did. Therefore, it is pos- not grow out from the inoculum that we attached to the sible that the susceptible, woody roots (roots ≥5 mm diam) fine roots toward woody roots or the root collar. Raabe of St. George inoculated by Raabe grew to encounter the (1979) inoculated grapevines by burying inoculum in pots inoculum before those of Riparia Gloire, based on the sub- beneath their root systems, much like how our fine root in- stantially higher vigor and deeper rooting habit of St. oculations were conducted. His technique was successful George (Pongracz 1983). Although our modifications to at infecting grapevines, although he did not state whether Raabe’s inoculation technique did not bring about signifi- the pathogen spread from the inoculum to the plants as cant mortality in the same amount of time (24 mo), they did rhizomorphs or whether the grapevines became infected increase the likelihood that all inoculated plants were chal- when their roots grew into direct contact with the inocu- lenged by the pathogen. lum. The reduced viability of our inoculum in the fine root Our finding that wounding did not increase suscepti- inoculations, relative to our other experiments, may have bility to infection is in contrast to past research on coni- been due to the strips of cheesecloth that held the inocu- fers (Wahlstrom and Johansson 1992, Solla et al. 2002). lum next to the roots, possibly by creating a moist envi- Solla et al. (2002) detected hyphae of A. mellea within in- ronment that accelerated decomposition of the inoculum. tact, outer root bark of Picea sitchensis as soon as eight We used one isolate of A. mellea for all four experi- days after inoculation. Removal of the outer bark allowed ments. Although this isolate was virulent on grapevines,

Am. J. Enol. Vitic. 57:4 (2006) 414 – Baumgartner and Rizzo because it was isolated from a symptomatic vine and Garrett, S. 1956. Rhizomorph behaviour in Armillaria mellea (Vahl) based on our consistent findings of infection among in- Quel., II. Logistics of infection. Annal. Bot. 20:193-209. oculated rootstocks, results may have been different with Gubler, W.D. 1992. Armillaria root rot. In Grape Pest Management. another isolate. Variation in virulence among Armillaria D.L. Flaherty et al. (Eds.), pp. 92-93. Publication 3343. University isolates has been assessed in the past (Raabe 1967, Wil- of California Division of Agriculture and Natural Resources, Oak- bur et al. 1972), but these studies were conducted before land. A. mellea was divided into multiple species. Therefore, Guillaumin, J.J., J.B. Anderson, and K. Korhonen. 1991. Life cycle, the isolates used by others (Raabe 1967, Wilbur et al. interfertility, and biological species. In Armillaria Root Disease. 1972) may have represented different Armillaria species, C.G. Shaw III and G.A. Kile (Eds.), pp. 10-47. U.S. Department which more recent studies have clearly demonstrated to of Agriculture, Forest Service, Agriculture Handbook 691, Wash- ington, DC. vary in virulence (Mallet and Hiratsuka 1988, Shaw and Loopstra 1988). Harrington, T.C., and B.D. Wingfield. 1995. A PCR-based identi- fication method for species of Armillaria. Mycologia 87:280-288. Mallet, K.I., and Y. Hiratsuka. 1988. Inoculation studies of lodgepole Conclusions pine with Alberta isolates of the Armillaria mellea complex. Can. J. For. Res. 18:292-296. Our finding of variation in susceptibility to infection by A. mellea among eight rootstocks of different genetic Munnecke, D.E., M.J. Kolbezen, W.D. Wilbur, and H.D. Ohr. 1981. Interactions involved in controlling Armillaria mellea. Plant Dis. backgrounds demonstrates that resistance exists in the 65:384-389. Vitis genome. However, the inoculation technique we used is too time-consuming for investigating additional root- Pongracz, D.P. 1983. Rootstocks for Grape-vines. David Philip Publisher, London. stocks or for use in a breeding program. It took us three years to complete our screening experiment and there are Raabe, R.D. 1962. Host list of the root rot fungus Armillaria mellea. still more rootstocks that need to be evaluated. A faster Hilgardia 33:25-88. screening technique is needed to identify additional resis- Raabe, R.D. 1967. Variation in pathogenicity and virulence in tant grapevine rootstocks, as well as resistant varieties Armillaria mellea. Phytopathology 57:73-75. and rootstocks of the many other hosts of A. mellea. Raabe, R.D. 1979. Testing grape rootstocks for resistance to the oak root fungus. California Plant Pathol. 46:3-4. Redfern, D.B., and G.M. Filip. 1991. Inoculum and infection. In Literature Cited Armillaria Root Disease. C.G. Shaw III and G.A. Kile (Eds.), pp. 48-61. U.S. Department of Agriculture, Forest Service, Agricul- Adaskaveg, J.E., H. Forster, L. Wade, D.F. Thompson, and J.H. ture Handbook 691, Washington, DC. Connell. 1999. Efficacy of sodium tetrathiocarbonate and pro- piconazole in managing Armillaria root rot of almond on peach Robinson, R.M., D. Morrison, and G.D. Jensen. 2004. Necro- rootstock. Plant Dis. 83:240-246. phylactic periderm formation in the roots of western and Douglas-fir trees infected with Armillaria ostoyae. II. The response Anderson, J.B., and R.C. Ullrich. 1979. Biological species of Ar- to the pathogen. For. Pathol. 34:119-129. millaria mellea in North America. Mycologia 71:402-414. Shaw, C.G., III, and E.M. Loopstra. 1988. Identification and patho- Baumgartner, K. 2004. Root collar excavation for postinfection con- genicity of some Alaskan isolates of Armillaria. Phytopathology trol of Armillaria root disease of grapevine. Plant Dis. 88:1235- 78:971-974. 1240. Sinclair, W.A., H.H. Lyon, and W.T. Johnson. 1987. Diseases of Baumgartner, K., and D.M. Rizzo. 2001a. Distribution of Armillaria Trees and Shrubs. Cornell University Press, Ithaca, NY. species in California. Mycologia 93:821-830. Singh, P. 1980. Armillaria root rot: Artificial inoculation and devel- Baumgartner, K., and D.M. Rizzo. 2001b. Ecology of Armillaria opment of the disease in greenhouse. Eur. J. For. Pathol. 10:420- species in mixed- forests of California. Plant Dis. 85: 431. 947-951. Solla, A., F. Tomlinson, and S. Woodward. 2002. Penetration of Picea Baumgartner, K., and D.M. Rizzo. 2002. Spread of Armillaria root sitchensis root bark by Armillaria mellea, Armillaria ostoyae and disease in a California vineyard. Am. J. Enol. Vitic. 53:197-203. Heterobasidion annosum. For. Pathol. 32:55-70. Baumgartner, K., and A.E. Warnock. 2006. A soil inoculant inhibits Thomas, H.E. 1934. Studies on Armillaria mellea (Vahl) Quelet: Armillaria mellea in vitro and improves productivity of grape- Infection, parasitism, and host resistance. J. Agric. Res. 48:187- vines with root disease. Plant Dis. 90:439-444. 218. Beckman, T.G., and P.L. Pusey. 2001. Field testing peach root- Volk, T.J., H.H. Burdsall Jr., and M.T. Banik. 1996. Armillaria stocks for resistance to Armillaria root rot. HortScience 36:101- nabsnona, a new species from western North America. Mycologia 103. 88:484-491. Bliss, D.E. 1951. The destruction of Armillaria mellea in citrus Wahlstrom, K.T., and M. Johansson. 1992. Structural responses soils. Phytopathology 41:665-683. in bark to mechanical wounding and Armillaria ostoyae infection Entry, J.A., N.E. Martin, R.G. Kelsey, and K.J. Cromack. 1992. in seedlings of Pinus sylvestris. Eur. J. For. Pathol. 22:65-76. Chemical constituents in root bark on five species of western co- Wilbur, W., D.E. Munnecke, and E.F. Darley. 1972. Seasonal de- nifer saplings and infection by Armillaria ostoyae. Phytopathol- velopment of Armillaria root rot of peach as influenced by fun- ogy 82:393-397. gal isolates. 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